Despite evidence that implicates transforming growth factor-α (TGF-α) in the pathogenesis of acute lung injury, the contribution of TGF-α to the fibroproliferative response is unknown. To determine whether the development of pulmonary fibrosis depends on TGF-α, we induced lung injury with bleomycin in TGF-α null-mutation transgenic mice and wild-type mice. Lung hydroxyproline content was 1.3, 1.2, and 1.6 times greater in wild-genotype mice than in TGF-α–deficient animals at Days 10, 21, and 28, respectively, after a single intratracheal injection of bleomycin. At Days 7 and 10 after bleomycin treatment, lung total RNA content was 1.5 times greater in wild-genotype mice than in TGF-α–deficient animals. There was no significant difference between mice of the two genotypes in lung total DNA content or nuclear labeling indices after bleomycin administration. Wild-genotype mice had significantly higher lung fibrosis scores at Days 7 and 14 after bleomycin treatment than did TGF-α–deficient animals. There was no significant difference between TGF-α–deficient mice and wild-genotype mice in lung inflammation scores after bleomycin administration. To determine whether expression of other members of the epidermal growth factor (EGF) family is increased after bleomycin-induced injury, we measured lung EGF and heparin-binding– epidermal growth factor (HB–EGF) mRNA levels. Steady-state HB–EGF mRNA levels were 321% and 478% of control values in bleomycin-treated lungs at Days 7 and 10, respectively, but were not significantly different in TGF-α–deficient and in wild-genotype mice. EGF mRNA was not detected in normal or bleomycin-treated lungs of mice of either genotype. These results show that TGF-α contributes significantly to the pathogenesis of pulmonary fibrosis after bleomycin-induced injury, and that compensatory increases in other EGF family members do not occur in TGF-α–deficient mice.
The fibroproliferative response to lung injury is characterized by mesenchymal cell proliferation and collagen accumulation within the alveolar and interstitial compartments of the lung. This response to injury results at least in part from the increased expression of growth factors and cytokines within the tissue microenvironment. Transforming growth factor-α (TGF-α), a member of the epidermal growth factor (EGF) family that includes TGF-α, EGF, and heparin-binding–EGF (HB–EGF), could play a prominent role in modulating the proliferative and fibrotic responses of the injured lung. Human TGF-α shares 42% and 30% homology with human EGF and HB–EGF (1), respectively, and binds to the EGF receptor (2). TGF-α stimulates the proliferation of cultured epithelial cells (3), fibroblasts (4), and endothelial cells (5). Activation of the EGF receptor stimulates collagen and glycosaminoglycan synthesis by mesenchymal cells (6, 7). In addition, TGF-α induces the expression of matrix metalloproteinase-1, -2, -3, and -9, and tissue inhibitor of metalloproteinase (TIMP)-1 and -3 by epithelial cells and fibroblasts in vitro (8-11).
Previous studies implicate TGF-α in the fibroproliferative response to acute lung injury. The expression of mRNA for TGF-α and of TGF-α protein is increased within areas of cellular proliferation and collagen accumulation in a rat model of bleomycin-induced acute lung injury (12). Lung fibroblasts isolated from hamsters injured by hyperoxia transcribe TGF-α mRNA and secrete TGF-α–immunoreactive protein (13). Epithelial lining fluid recovered from silica-exposed rats contains TGF-α–immunoreactive protein that accounts for 85% of the mitogenic activity found in this fluid (14). Transgenic mice expressing human TGF-α under control of regulatory regions of the human surfactant protein-C gene develop pulmonary fibrosis characterized by peribronchiolar and pleural fibrotic lesions and markedly enlarged alveolar spaces (15). TGF-α–immunoreactive protein is present in pulmonary edema fluid recovered from patients within the first 24 h after the onset of acute lung injury (16). Furthermore, TGF-α levels in bronchoalveolar lavage fluid (BALF) were significantly higher in the vast majority of a large cohort of patients with established acute respiratory distress syndrome and in patients with idiopathic pulmonary fibrosis (IPF) than in normal subjects (17).
Despite circumstantial data that implicate TGF-α in the pathogenesis of pulmonary fibrosis, the contribution of TGF-α to the fibrotic response is unknown. To determine whether the development of fibrosis depends on TGF-α, we induced lung injury with bleomycin in transgenic mice engineered to be completely deficient of TGF-α. In this study our goals were to test the following hypotheses: (1) that lung fibrosis is decreased after bleomycin injury in TGF-α–deficient animals; and (2) that the expression of other EGF family members is not modified by the absence of TGF-α in bleomycin-injured lungs. The study showed that lung collagen accumulation after bleomycin injury is significantly reduced in animals lacking TGF-α, and that compensatory increases in EGF or HB–EGF do not occur in bleomycin-injured lungs of TGF-α–deficient animals. These results provide direct evidence that TGF-α contributes significantly to the pathogenesis of lung fibrosis after acute lung injury.
TGF-α null-mutation and wild-genotype mice were bred from C57BL/6 mice heterozygous for a targeted disruption of exon 3 of the TGF-α gene (18). The genotypes of wild mice and TGF-α–deficient mice were confirmed by polymerase chain reaction (PCR) analysis performed on DNA prepared from the tails of 3-wk-old animals. PCR was done as previously described (18). To identify the disrupted TGF-α allele, we used the following primer pair: Primer 1, complimentary to genomic DNA that was upstream of exon 3, with the base sequence 5′-dGACTAGCCTGGGCTACACAGTG-3′; and Primer 2, complimentary to sequences at the 3′ terminus of the neo gene inserted into exon 3, with the base 5′-dCCGCTTCCTCGTGCTTTACGGT-3′. To identify the wild-type allele, Primer 1 was used in conjunction with Primer 3, which was complimentary to the TGF-α sequences on the side 3′ of the disruption site, and had the sequence 5′-dACATGCTGGCTTCTCTTCCTGC-3′.
The null-mutation phenotype was confirmed at both the level of gene transcription and protein production. TGF-α mRNA expression in the lung tissue of homozygous TGF-α null-mutation and wild-genotype mice was analyzed with reverse transcription (RT)–PCR, using the Titan RT–PCR system (Boehringer-Mannheim, Indianapolis, IN). One microgram of lung total RNA was added to RT–PCR buffer containing 10 U of ribonuclease inhibitor (RNasin), 5 mM dithiothreitol, 0.2 mM deoxynucleotide triphosphates (dNTPs), 2.5 mM MgCl2, 0.3 mM oligonucleotide primers, and 1 μl of Avian myeloblastosis virus (AMV)/Expand High Fidelity PCR enzyme mix (Boehringer Mannheim Inc., Indianapolis, IN) in 50 μl of reaction volume. The oligonucleotide primers were derived from exon 2 and exon 5 of the TGF-α gene, and had the sequences 5′-GTCAGGCTCTGGAGAACAGC-3′ and 5′-CGGCACCACTCACAGTGCTTG-3′, respectively. Reverse transcription was done at 50°C for 30 min, and amplification was done through 50 cycles at 94°C for 1 min, 65°C for 1 min, and 72°C for 3 min. The reaction products were resolved in 1% agarose gels. The DNA was transferred onto nylon membranes (Nytran, 0.45 μm pore size; Schleicher & Schuell, Keene, NH) and was hybridized with 32P-labeled oligonucleotide in 6× saline sodium citrate (SSC), 0.01 M NaH2PO4, 0.5% sodium dodecyl sulfate (SDS), 100 μg/ml single-stranded DNA, 0.1% nonfat dried milk, and 10 mM ethylenediaminetetraacetic acid (EDTA) at 37°C for 18 h. The filters were then washed in 6× SSC for 10 min at room temperature before autoradiography. The oligonucleotide probe used in Southern blot analysis was complementary to a portion of exon 3 of TGF-α, and had the sequence 5′-dTCCTGCACCAAAAACCTGCAGGT-3′.
The concentration of TGF-α–immunoreactive protein in the lungs of homozygous TGF-α null-mutation and wild-genotype mice was determined with the acid-ethanol extraction method and radioimmunoassay (RIA) as previously described (12). Protein concentrations of the lung extracts were determined according to the method of Lowry as modified by Ohnishi and Barr (19). The protein extracts were fractionated on Waters tC18 Sep-Pak cartridges (Millipore, Milford, MA). After loading, the cartridges were washed with 2 ml of 10% acetonitrile and 0.005% trifluoroacetic acid before elution of the TGF-α-containing fraction with 2 ml of 40% acetonitrile and 0.05% trifluoroacetic acid. The fractions were evaporated in a Speed-Vac concentrator (Savant Instruments, Inc., Hicksville, NY) and resolubilized in RIA buffer. The TGF-α concentration of the fractionated lung extracts was measured in duplicate, using a commercially available TGF-α RIA (Peninsula Laboratories, Belmont, CA), and the results were expressed as picograms of TGF-α per lung.
The specific pathogen-free, female and male, 8-wk old homozygous TGF-α null-mutation and wild-genotype mice, subjected to bleomycin-induced lung injury had initial body weights of 21.4 ± 0.62 g and 22.8 ± 0.65 g (mean ± SE, P = 0.52), respectively, at the time of bleomycin instillation. Intratracheal administration of 0.075 U of bleomycin sulfate (Blenoxane; Bristol Laboratories, Syracuse, NY) in 50 μl of sterile saline was done via a tracheostomy under intraperitoneal avertin anesthesia (20). Control mice received saline alone. The bleomycin dose used was shown to consistently produce pulmonary fibrosis with a mortality rate of < 10% in preliminary experiments with mice of similar genetic background.
At 2, 4, 7, 10, 14, 21, and 28 d after injection, the mice were killed by exsanguination under deep anesthesia. The lungs were exposed by a midthoracotomy incision and the pulmonary arteries were perfused with ribonuclease (RNase)- free phosphate-buffered saline (PBS). The right lung was isolated with a ligature at the right hilum and was resected, rinsed in RNase-free PBS, and finely minced. The minced lung was divided into three aliquots, one each for tissue hydroxyproline, RNA, and DNA measurement. Each aliquot of the minced right lung was weighed, frozen in liquid nitrogen, and stored at −70°C for further analysis. The left lung was inflated with 4% neutral buffered paraformaldehyde instilled at 30 cm H2O pressure through the trachea for 120 min. The trachea was then tied and the lung immersed in the 4% buffered paraformaldehyde for 24 h before embedding in paraffin.
Total lung collagen content was measured by assaying lung hydroxyproline content after hydrolysis with 6 N HCl as previously described (21). The minced lung aliquot was added to 800 μl of 6 N HCl and hydrolyzed overnight at 110°C. To 200 μl of hydrolysate were added 100 μl of 0.02% methyl red and 20 μl of 0.04% bromthymol blue, and the sample volume was adjusted to 2 ml of 0.5× assay buffer (0.024 M C6H8O7 · H2O, 0.02 M CH3CO2H, 0.088 M C2H3O2Na · H2O, 0.085 M NaOH), and the pH was adjusted to 6.5 to 7.0. The colorimetric assay was performed by adding 1 ml of Chloramine-T solution to the sample, incubating at room temperature for 20 min, and adding 1 ml of dimethylbenzaldehyde solution followed by incubation at 60°C for 15 min. The absorbance at 550 nm was measured for each lung sample. To account for any loss of hydroxyproline during the hydrolysis procedure, each minced lung aliquot was spiked with a known quantity of tritiated hydroxyproline, and the residual radioactivity of the colorimeteric assay samples was quantified by scintillation spectroscopy. Results of the colorimeteric assay were corrected for the tritiated hydroxyproline recovered. Whole-lung hydroxyproline values were determined by normalizing the hydroxyproline values obtained with the colorimetric assay of the minced lung aliquots to whole-lung wet weight, and were expressed as μg/lung.
Total cellular RNA was isolated from the frozen tissue through a modification of the method of Chirgwin and colleagues, with cesium chloride density-gradient centrifugation (22, 23). Total cellular RNA was quantified in triplicate by optical density measurement at 260 nm. Whole-lung total cellular RNA content was determined by normalizing the values obtained for the aliquots of minced lung to whole-lung wet weight, and was expressed as μg/lung.
Lung DNA content was determined with fluorimetry, using a modification of the method of Aguayo and colleagues (24). Lung tissue was digested with proteinase K to a final concentration of 1 mg/ml in 400 μl of proteinase K digestion buffer (10 mM Tris-HCl, 0.1 M EDTA, 0.5% SDS, pH 8.0) at 55°C for 3 h, followed by freeze–thawing and sonication for 1 min. DNA was quantified in triplicate by adding 4 μl of lung homogenate to a cuvette containing 1,996 μl of TNE buffer (10 mM Tris, 10 mM EDTA, 2 M NaCl, pH 7.4) plus 1 ml of a 0.1 mg/ml Hoechst 33258 dye solution, and the fluorescence was measured with a spectrofluorimeter (Model TKO-100; Hoeffer Scientific, San Francisco, CA). The DNA content of each minced lung aliquot was calculated from a standard curve developed by measuring the fluorescence emission of known concentrations of calf thymus DNA. Whole-lung DNA values were determined by normalizing the DNA values obtained for the minced lung aliquots to whole-lung wet weight, and were expressed as μg/lung.
In situ cell proliferation was detected by bromodeoxyuridine (BrdU) immunohistochemistry (Boehringer-Mannheim) according to the manufacturer's instructions. At Day 9 after bleomycin or saline instillation, animals received an intraperitoneal injection of BrdU labeling reagent (1 ml/100 g body weight). Twenty-four hours later the lungs were harvested, fixed with 4% paraformaldehyde, and paraffin embedded, as described earlier. Lung sections were digested in proteinase K (10 μg/ml) at 37°C for 15 min, denatured in 1 M HCl for 8 min at 60°C, and then blocked for 1 h in PBS containing 5% fetal calf serum, 2 mM levamisole, and 0.05% Triton-X 100. The sections were stained with mouse monoclonal anti-BrdU antibody F(ab′)2 fragments conjugated with alkaline phosphatase (1:5 dilution), and BrdU-labeled nuclei were detected with Fast Red substrate. To assess total cellularity, nuclei in adjacent serial lung sections were stained with 4′,6-diamidine-2′-phenylindole dihydrochloride (DAPI) (2.5 μg/ml) in PBS.
Stained tissue sections were imaged using a DeltaVision microscope system (Applied Precision, Inc., Issaquah, WA) on a Zeiss Axiovert 100 microscope (Zeiss, Thornwood, NY). Images were collected with a Zeiss Achromat ×10 lens. Fast Red was excited at 555 nm and imaged at 617 nm, and DAPI was excited at 360 nm and imaged at 457 nm. Entire sections of whole-lung lobes were imaged sequentially, using the automated stage of the DeltaVision system. These images were transferred to a Macintosh 9500/ 132 computer (Apple Computer Inc., Cupertino, CA) and analyzed with the NIH Image v1.62b7 system (National Institutes of Health, Bethesda, MA). Images were automatically thresholded, using a statistical method based on the intensity histogram. The total area of BrdU-positive nuclei was taken to be the area of thresholded BrdU image, and the total area of all nuclei was taken to be the area of the thresholded DAPI image. The ratio of the BrdU area to the DAPI area was defined as the replication fraction, and was found to agree well with visual counts of positive nuclei (data not shown).
Lung sections from bleomycin-injured and control mice were stained with hematoxylin and eosin (H&E) or Masson trichrome stains, coded, and scored blindly for inflammation and fibrosis. Lung inflammation was scored in tissue sections stained with H&E on a scale of 0 to 3 (Grade 0 = no inflammatory involvement; Grade 1 = mononuclear inflammatory cell infiltration of 3% to 29% of the parenchyma; Grade 2 = mononuclear inflammatory cell infiltration of 30% to 59% of the parenchyma; and Grade 3 = mononuclear inflammatory cell infiltration of 60% to 100% of the parenchyma). Lung fibrosis was scored in trichrome-stained sections on a scale of 0 to 4 (Grade 0 = no increase in connective tissue; Grade 1 = fine connective-tissue fibrils in less than 50% of the area occupied by inflammatory cells, without coarse collagen; Grade 2 = fine fibrils in 50% to 100% of the same area, without coarse collagen; Grade 3 = fine fibrils in 100% of the area, with coarse collagen bundles in 10% to 49% of the area; Grade 4 = fine fibrils in 100% of the area, with coarse collagen in 50% to 100% of the area).
A rat HB–EGF complementary RNA (cRNA) probe was prepared by subcloning the EcoRI–DdeI fragment of rat HB–EGF complementary DNA (cDNA) (provided by J. Abraham, Scios Nova, Inc., Mountain View, CA) into a plasmid vector (pBluescript SK; Stratagene, La Jolla, CA) downstream to the T3 promoter. The 32P-labeled, single-stranded HB–EGF cRNA, 332 bases in length, was transcribed from template DNA linearized with EcoRI through the use of an RNA in vitro transcription kit (Boehringer-Mannheim) and [α-32P]uridine triphosphate ([α-32P]UTP) (NEN, Wilmington, DE) (25).
A mouse EGF cRNA was prepared by subcloning the XhoI–SmaI fragment of mouse EGF cDNA (provided by G. Bell, University of Chicago, Chicago, IL) into a pBluescript SK (Stratagene) plasmid vector downstream of the T3 promoter. The 32P-labeled, single-stranded mEGF cRNA, 576 bases in length, was transcribed from template DNA linearized with EcoRI, using an RNA in vitro transcription kit as described earlier.
An 18S ribosomal RNA (rRNA) probe was used as an internal control to normalize for the amount of RNA applied to Northern blots (12). This 24-base oligonucleotide, with the sequence 5′-dACGGTATCTGATCGTCTTCGAACC-3′, and complementary to bases 1,043 to 1,066 of human 18S rRNA, was synthesized, purified, and 5′-end labeled as previously described (12).
Northern blot analysis for HB–EGF and EGF mRNA was performed as previously described (23). Briefly, total cytoplasmic RNA (20 μg/lane) was electrophoresed through 1% agarose/formaldehyde gels (26) and transferred to nylon membranes (Nytran). The membranes were hybridized with either HB–EGF or EGF cRNA probe, at 2 × 106 cpm of probe per ml of hybridization solution (0.125 M Na2HPO4, 0.25 M NaCl, 7% [wt/vol] SDS, 1 mM EDTA, 50% [vol/vol] formamide, 10% [wt/vol] polyethylene glycol [8,000], 0.1 mg/ml sonicated denatured salmon sperm DNA, 0.1 mg/ml yeast transfer RNA [tRNA]) at 55°C for 18 h. The membranes were washed in 2× SSC–0.1% (wt/ vol) SDS at room temperature for 15 min. The membranes hybridized with HB–EGF were then washed with 0.1× SSC–0.1% SDS (wt/vol) at 60°C for 30 min, and the membranes hybridized with EGF were washed in an identical solution at 65°C for 30 min. The membranes were cut just above the 18S rRNA bands. The bottom portions were saved for subsequent hybridization with the 18S oligonucleotide probe. Nonhybridized cRNA probe was digested from the top portions of the membranes by treatment with ribonuclease A (Sigma, St. Louis, MO), 1 μg/ml in 2× SSC, for 10 min at room temperature, and the membranes were washed again with 0.1× SSC–0.1% (wt/vol) SDS at 60°C and 65°C for HB–EGF and EGF, respectively.
Previously hybridized nylon membranes were stripped of probe and hybridized with the 18S rRNA oligonucleotide probe as previously described (12). Briefly, the membranes were stripped by washing with 0.1× SSC–0.1% SDS at 97°C for 20 min. The membranes were prehybridized at 37°C for 2 h with 6× SSC, 0.1 mg/ml denatured salmon sperm DNA, 0.5% SDS, and 0.1% nonfat dried milk, They were then hybridized overnight at 37°C with the same solution as the prehybridization buffer. The membranes were then washed with 6× SSC at room temperature for 10 min and with 6× SSC at 55°C for 30 min.
Quantification of mRNA was done as previously described (12). Autoradiographs of the hybridized membranes were made by exposing PhosphorImager storage plates (Molecular Dynamics, Sunnyvale, CA) at room temperature for 18 to 72 h, as needed, to provide adequate intensity of bands. The plates were scanned with a PhosphorImager. Relative quantities of hybridized probe in each band were determined with Image Quant software (Molecular Dynamics). The signal intensity for HB–EGF or EGF mRNA bands was divided by the signal intensity of 18S rRNA bands in the same lanes to control for variations in the quantity of RNA loaded in each lane. Conventional autoradiographs of the hybridized membranes were made by exposing XAR-2 film (Kodak, Rochester, NY) at −70°C with Cronex intensifying screens (Dupont, Wilmington, DE) for 14 d (HB–EGF and EGF riboprobes) and 7 d (18S oligonucleotide probe).
and A and B are the group means, S is the group standard deviation, and N is the number of observations (27).
The absence of functional TGF-α in the lungs of null- mutation mice was demonstrated by RT–PCR analysis of lung RNA and quantitation of lung TGF-α protein. RNA isolated from the lungs of TGF-α null-mutation-homozygous mice directed the synthesis of an RT–PCR product that did not hybridize with an oligonucleotide probe for TGF-α exon 3, indicating disruption of the TGF-α gene (Figure 1). In addition, acid–ethanol lung extracts from TGF-α null-mutation mice contained no detectable TGF-α–immunoreactive protein (n = 4), whereas lung extracts from wild-genotype mice contained 9.9 ± 5.4 pg/100 μg protein (n = 4). These results confirm TGF-α gene inactivation in the lung of the TGF-α null-mutation mouse via disruption of exon 3, and are consistent with previous reports (18, 28).
Lung collagen, as measured by hydroxyproline content, was significantly increased in bleomycin-treated wild- genotype mice as compared with saline controls. At Day 10 after bleomycin injury, mean lung hydroxyproline content of the wild-genotype mice was 1.3 times that of the saline-instilled wild-type animals (153.1 ± 9.8 μg/lung, compared with 113.7 ± 11.3 μg/lung, P < 0.05) (Figure 2). At Days 21 and 28 after bleomycin administration, the mean lung hydroxyproline content of wild-genotype mice was 1.3 and 1.4 times, respectively, that of the saline controls (Day 21: 187.5 ± 10.5 μg/lung, compared with 139.9 ± 8.2 μg/lung, P < 0.05; Day 28: 225 ± 19.7 μg/lung, compared with 155.9 ± 4.8 μg/lung, P < 0.05). The lung hydroxyproline content of bleomycin-treated TGF-α null-mutation mice was significantly higher than that of saline-instilled mice only at Day 21 (158.5 ± 12.5 μg/lung, compared with 112.1 ± 3.6 μg/lung, P < 0.05).
The lung hydroxyproline content of bleomycin-treated TGF-α–deficient mice was significantly lower than that of bleomycin-treated wild-genotype animals. At Day 10 after bleomycin administration, mean lung hydroxyproline content of the wild-genotype mice was 1.3 times that of bleomycin-treated TGF-α–deficient mice (153.1 ± 19.9 μg/ lung, compared with 115.2 ± 8.9 μg/lung, P < 0.05). At Days 21 and 28 after bleomycin instillation, the mean lung hydroxyproline content of the wild-genotype mice was 1.2 and 1.6 times, respectively, that of the bleomycin-treated TGF-α null-mutation mice (Day 21: 187.5 ± 10.5 μg/lung, compared with 158.5 ± 12.5 μg/lung, P < 0.05; Day 28: 225 ± 19.7 μg/lung, compared with 139.1 ± 15.7 μg/lung, P < 0.05). No significant differences in lung hydroxyproline content of TGF-α–deficient and wild-genotype mice were observed when only saline was administered.
Lung total RNA content was significantly increased in wild-genotype mice during the first 2 wk after bleomycin administration as compared with that of saline controls. At Day 7 after instillation, the mean lung RNA content of the wild-genotype mice that received bleomycin was 2.0 times that of the saline-instilled animals (210.2 ± 21.4 μg/lung, compared with 104.9 ± 10.6 μg/lung, P < 0.05) (Figure 3). At Days 10 and 14 after instillation, the mean lung RNA content of wild-genotype mice that received bleomycin was 2.8 times that of the saline controls (238.1 ± 23.4 μg/ lung, compared with 84.4 ± 10.9 μg/lung, P < 0.05, and 236.1 ± 42.5 μg/lung, compared with 84.9 ± 11.5 μg/lung, P < 0.05, respectively). The lung RNA content of bleomycin-treated, TGF-α–deficient mice was 1.7 times higher than that of saline controls at Day 7 (141.7 ± 9.1 μg/lung, compared with 84.9 ± 10.6 μg/lung, P < 0.05) and 1.9 times higher at Day 10 (156.9 ± 9.2 μg/lung, compared with 82.5 ± 12.0 μg/lung, P < 0.05).
At Days 7 and 10 after bleomycin administration, the mean lung RNA content of wild-genotype mice was 1.5 times higher than that of TGF-α–deficient animals (210.2 ± 21.4 μg/lung, compared with 141.7 ± 9.1 μg/lung, P < 0.05, and 238.1 ± 23.4 μg/lung, compared with 156.9 ± 9.2 μg/ lung, P < 0.05, respectively). No significant difference in the lung RNA content of wild-genotype and TGF-α–deficient mice was observed in the absence of bleomycin treatment (P = 0.065).
There was no significant difference in the lung DNA content of TGF-α–deficient and wild-genotype mice either with (P = 0.9) or without (P = 0.06) bleomycin administration (Figure 4). Likewise, lung DNA content in wild-genotype mice was not different after bleomycin administration as opposed to saline instillation (P = 0.19). Only at Day 28 after bleomycin administration was the mean lung DNA content of TGF-α–deficient mice 1.4 times greater than that of the saline controls (995.0 ± 63.7 μg/lung, compared with 706.8 ± 66.3 μg/lung, P < 0.05).
Quantitative analysis of cell proliferation in bleomycin-treated and control lungs was done with BrdU as a specific marker of DNA synthesis (29). There was a trend toward an increased mean labeling index in bleomycin-injured lungs of wild-genotype mice at Day 10 as compared with saline controls; however, the difference was not significant (1.77 ± 0.7%, compared with 0.35 ± 0.02%, P = 0.18). The mean labeling indices were not significantly different for TGF-α–deficient and wild-genotype mice at Day 10 after bleomycin injury (1.75 ± 0.79%, compared with 1.77 ± 0.7%, P = 0.99). We observed the highest number of BrdU-labeled nuclei within inflammatory foci of bleomycin-injured lungs. BrdU-labeled nuclei were observed in bronchiolar epithelium, alveolar mononuclear cells, and interstitial cells with equal frequency in injured wild-genotype and TGF-α–deficient lungs.
To provide a visual correlate to the quantitative hydroxyproline data, we examined Masson trichrome-stained sections of lung tissue by light microscopy. The lungs of saline-treated animals appeared normal, regardless of their TGF-α genotype. The lungs of animals receiving intratracheal saline showed only thin bands of collagen immediately adjacent to large vessels and airways (data not shown). After bleomycin instillation, the lungs of wild-genotype mice contained dense bands of collagen replacing large areas of lung parenchyma (Figures 5A and 5B). In contrast, the areas of collagen accumulation in the lungs of bleomycin-treated, TGF-α–deficient mice were fewer in number and considerably less dense (Figures 5C and 5D). In both the wild-genotype and TGF-α–deficient mice, areas of lung tissue with increased collagen also contained increased numbers of inflammatory cells.
To quantitate the histologic changes observed after bleomycin administration, lung sections were stained, coded, and then blindly scored for inflammation and fibrosis. At Day 14 after bleomycin instillation, the mean inflammation scores were significantly higher in TGF-α–deficient and wild-genotype mice than in saline controls (Table 1). Likewise, TGF-α–deficient and wild-genotype mice had significantly higher mean fibrosis scores at Days 7, 10, 14, and 28 after bleomycin administration than did saline-treated animals. There was a trend toward higher lung inflammation scores at Days 7, 10, 14, and 28 after bleomycin instillation in the wild-genotype mice than in TGF-α–deficient animals, but this difference was not statistically significant. In contrast, the mean lung fibrosis scores were significantly higher for wild-genotype mice than for TGF-α–deficient animals at Days 7 and 14 after bleomycin instillation. In addition, there was a trend toward higher lung fibrosis scores at Days 10 and 28 in the wild-genotype mice, but this difference did not achieve statistical significance. No difference was observed in the mean fibrosis scores of TGF-α–deficient and wild-genotype mice in the absence of bleomycin administration.
|Day||Condition||Inflammation Score||Fibrosis Score|
|7||Bleomycin TGF-α+/+||1.6 ± 0.4||2.4 ± 0.4|
|Bleomycin TGF-α−/−||1.2 ± 0.4||1.2 ± 0.4*|
|Control TGF-α+/+||0.4 ± 0.2*||0*|
|Control TGF-α−/−||0.8 ± 0.6||0†|
|14||Bleomycin TGF-α+/+||2.0 ± 0||3.0 ± 0.3|
|Bleomycin TGF-α−/−||1.8 ± 0.4||1.8 ± 0.4*|
|Control TGF-α+/+||0.4 ± 0.2*||0*|
|Control TGF-α−/−||0.8 ± 0.2†||0†|
|28||Bleomycin TGF-α+/+||1.6 ± 0.4||2.0 ± 0.6|
|Bleomycin TGF-α−/−||1.4 ± 0.2||1.6 ± 0.5|
|Control TGF-α+/+||0.6 ± 0.5||0*|
|Control TGF-α−/−||1.0 ± 0||0†|
Total cellular RNA extracted from the lungs of control and bleomycin-injured mice was examined with Northern blot analysis for the presence of EGF and HB–EGF. HB– EGF was detected at low levels as an ∼ 2.5-kb transcript in the lungs of saline-treated animals (Figure 6). At Days 7 and 10 after bleomycin administration, lung steady-state HB–EGF mRNA levels of wild-genotype mice increased to 321% (n = 5, P < 0.05) and 478% (n = 5, P = 0.05) of control values, respectively (Figures 6 and 7). A similar increase was observed in wild-genotype animals at Day 14 after bleomycin injury, but did not achieve statistical significance. At Days 2, 4, and 28 after injury, HB–EGF mRNA levels were comparable to those of control animals. HB–EGF steady-state mRNA levels increased to the same extent and with the same temporal profile in the lungs of TGF-α–deficient and wild-genotype mice after bleomycin-induced lung injury. In contrast to HB–EGF mRNA, EGF mRNA was undetectable in the lungs of TGF-α–deficient and wild-genotype mice at all time points examined after saline or bleomycin administration (data not shown). As a positive control, EGF was detected as an ∼ 4.5 kb transcript in mouse kidney.
The major goal of this study was to investigate the role of TGF-α in the pathogenesis of lung fibrosis. Our strategy was to determine whether collagen accumulation in the acutely injured lung was decreased in mice genetically engineered to lack TGF-α. We found that pulmonary collagen accumulation and lung fibrosis were significantly lower after bleomycin injury in TGF-α–deficient mice than in wild-genotype animals. In contrast, there was no difference in lung inflammation between the two genotypes after bleomycin treatment. We also demonstrated that HB–EGF expression was increased in the lung after bleomycin-induced injury, and that the expression of HB– EGF in the injured lung was not modified in the absence of TGF-α. Furthermore, we observed that EGF gene expression was not induced in the lung after bleomycin injury.
Our study provides direct evidence that TGF-α contributes significantly to the pathogenesis of lung fibrosis after acute lung injury. The reduction in lung hydroxyproline content and lung fibrosis scores of TGF-α–deficient animals supports the hypothesis that TGF-α amplifies the fibrotic response of the injured lung. Our results are consistent with our previous finding that TGF-α is localized in areas of collagen accumulation in bleomycin-injured rat lungs (12), and with the observation that overexpression of TGF-α by respiratory epithelial cells induces lung fibrosis in transgenic mice (15). The elimination of TGF-α alone, however, is not sufficient to completely inhibit the fibrotic response, as indicated by the increase in lung hydroxyproline content at Days 14 and 21 after bleomycin treatment of TGF-α–deficient mice. This is not unexpected, since other inflammatory cytokines, such as interleukin-1 (30), macrophage inflammatory protein-1α (31), and macrophage chemotactic protein-1 (32), and growth factors such as platelet-derived growth factor (33) and TGF-β (34), have been reported to be modulated in the injured lung. Despite these potentially redundant pathways leading to fibrosis, TGF-α deficiency has a significant effect on the remodeling of the extracellular matrix that occurs in response to lung injury.
TGF-α could promote the profibrotic response through several mechanisms. Although TGF-α stimulates the proliferation of fibroblasts in vitro (4), our results showed no significant difference in cellular proliferation in the lungs of TGF-α–deficient and wild-genotype animals at Day 10 after bleomycin injury. Since EGF receptor activation has been shown to stimulate macrophage chemotaxis in vitro (35), TGF-α could be recruiting macrophages that subsequently secrete profibrotic factors. However, lung inflammation scores were not significantly different for bleomycin-injured, TGF-α–deficient mice and bleomycin-treated, wild-genotype mice in our study, suggesting that the profibrotic response is not likely to be the consequence of differences in macrophage recruitment. Alternatively, TGF-α could promote extracellular-matrix deposition directly, through its ability to stimulate collagen synthesis (36), or indirectly, through its ability to induce TGF-β secretion (37). TGF-α expression in the injured lung could also inhibit collagen degradation through induction of TIMPs (9, 11), and thereby promote collagen accumulation during injury-induced remodeling of connective tissue.
Strong evidence that TGF-α is involved in connective tissue remodeling in the developing lung has been provided by SP-C/TGF-α–transgenic mice. Overexpression of TGF-α in the lungs of these mice was shown to disrupt postnatal alveolarization, leading to enlarged air spaces and fibrosis in the interstitium and pleura (38). Elastic fiber formation in bronchiolar regions and alveolar septae of these lungs was found to be dysmorphic or absent (15). Others have observed that TGF-α deficiency did not appear to modify wound repair. Wound healing after tail amputation was comparable for TGF-α–deficient and wild-genotype mice (18). Likewise, full thickness skin wounds closed at comparable rates and displayed similar histologic features in TGF-α–deficient and wild-genotype animals (28). Corneal wound healing, as measured by fluoroscein staining, also was comparable in both genotypes (28). In these studies, however, detailed analyses of connective tissue remodeling were not performed, and subtle changes may therefore have been missed.
The EGF family comprises several growth factors, including TGF-α, HB–EGF, and EGF. HB–EGF is expressed in normal and hyperoxia-injured lung (39, 40). We detected HB–EGF mRNA in control mouse lung, and observed that HB–EGF expression was induced in a temporally defined manner after bleomycin injury. However, we found no evidence that HB–EGF mRNA was further increased in compensation for TGF-α deficiency in this model. Therefore, even though potentially redundant pathways exist for growth factor-mediated lung remodeling, TGF-α plays a unique role within the EGF family in the fibrotic response to lung injury.
We did not detect EGF mRNA in lung homogenates of control or bleomycin-treated animals at any time, suggesting that EGF expression was not induced after bleomycin injury to the lung. However, the absence of detectable EGF transcript in whole-lung RNA does not preclude the possibility that EGF expression might be induced in certain cell types of the injured lung. EGF-immunoreactive protein has been detected in normal and injured developing human lung (41) and in normal rat lung (42-44). In addition, cultured rat type II pneumocytes transcribe and secrete EGF (45). The difference between our results and those previously reported may represent developmental or species-specific differences in EGF expression.
Pulmonary fibrosis is a frequent consequence of acute and chronic inflammatory lung diseases. Autopsy series of patients with acute respiratory distress syndrome (ARDS) identify pulmonary fibrosis as a common feature (46, 47). Pulmonary collagen content is increased in those patients dying later than 10 d after the onset of ARDS (47). Analysis of lung tissue obtained by open lung (48) and transbronchial biopsies (49) suggests an association between mortality and pulmonary fibrosis in established ARDS. Similarly, the degree of interstitial fibrosis is an important variable in predicting response to therapy and prognosis in patients with IPF (50-53).
A number of growth factors and inflammatory cytokines have been detected during the evolution of fibrotic lung lesions. We have shown that TGF-α levels in bronchoalveolar lavage fluid are increased in the vast majority of a large cohort of patients with established ARDS and in patients with IPF as compared with normal subjects (17). The presence of TGF-α in the lavage fluid of patients with ARDS supports the hypothesis that TGF-α may modulate the fibroproliferative response following acute lung injury in humans. TGF-α in the alveolar microenvironment could contribute to the pulmonary fibrosis found in patients with delayed resolution of ARDS (47), and could in part account for the restrictive pulmonary impairment observed in many survivors of ARDS (54). The presence of TGF-α in the alveolar lining fluid recovered from patients with IPF provides additional support for a role of TGF-α in the pathogenesis of fibrotic lung disease (17). The present study provides in vivo evidence that TGF-α participates in the fibrotic response to lung injury. Moreover, the results of the study suggest that therapeutic interventions designed to inhibit TGF-α expression or TGF-α–mediated signal transduction may limit the development of pulmonary fibrosis in the injured lung.
This study was supported by an American Lung Association of Washington Research Award (D.K.M.), and by grants HL 49401 (D.K.M.), HL30542 (J.G.C.), and CA18029 (R.C.H.), from the National Institutes of Health. The authors thank Dr. Judith Abraham of Scios Nova, Inc., for providing the rat HB–EGF cDNA probe, and Dr. Graeme I. Bell of the University of Chicago for providing the mouse EGF cDNA probe. They also thank Margaret K. Greer and Linda O'Neal for excellent technical assistance, and Ted Gooley for assistance in the statistical analysis.
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