Patients with interstitial lung diseases, such as idiopathic pulmonary fibrosis (IPF) and bronchopulmonary dysplasia (BPD), suffer from lung fibrosis secondary to myofibroblast-mediated excessive ECM deposition and destruction of lung architecture. Transforming growth factor (TGF)-β1 induces epithelial-mesenchymal transition (EMT) of alveolar epithelial cells (AEC) to myofibroblasts both in vitro and in vivo. Inhaled nitric oxide (NO) attenuates ECM accumulation, enhances lung growth, and decreases alveolar myofibroblast number in experimental models. We therefore hypothesized that NO attenuates TGF-β1-induced EMT in cultured AEC. Studies of the capacity for endogenous NO production in AEC revealed that endothelial nitric oxide synthase (eNOS) and inducible nitric oxide synthase (iNOS) are expressed and active in AEC. Total NOS activity was 1.3 pmol·mg protein−1·min−1 with 67% derived from eNOS. TGF-β1 (50 pM) suppressed eNOS expression by more than 60% and activity by 83% but did not affect iNOS expression or activity. Inhibition of endogenous NOS with l-NAME led to spontaneous EMT, manifested by increased α-smooth muscle actin (α-SMA) expression and a fibroblast-like morphology. Provision of exogenous NO to TGF-β1-treated AEC decreased stress fiber-associated α-SMA expression and decreased collagen I expression by 80%. NO-treated AEC also retained an epithelial morphology and expressed increased lamellar protein, E-cadherin, and pro-surfactant protein B compared with those treated with TGF-β alone. These findings indicate that NO serves a critical role in preserving an epithelial phenotype and in attenuating EMT in AEC. NO-mediated regulation of AEC fate may have important implications in the pathophysiology and treatment of diseases such as IPF and BPD.
- alveolar epithelium
- lung injury
- nitric oxide synthases
- pulmonary fibrosis
- transforming growth factor-β
idiopathic pulmonary fibrosis (IPF), bronchopulmonary dysplasia (BPD), and post-acute respiratory distress syndrome fibrosing alveolitis are interstitial lung diseases that result in significant long-term morbidity and mortality for affected patients (5, 9, 11, 27). Clinical morbidity associated with interstitial lung disease arises from excessive and disordered extracellular matrix (ECM) production, proliferation and activation of pulmonary fibroblasts, destruction of lung architecture, and fibrosis. Once thought to be exclusively secondary to dysregulated inflammation in the lung, recent studies suggest that the ECM accumulation and fibrosis associated with these conditions result instead from chronic epithelial injury and activation independent of inflammation (34).
Whereas the normal response to lung injury is migration and proliferation of type II pneumocytes to reestablish an intact epithelial lining (3), the chronic and repetitive injury that culminates in fibrotic lung disease promotes hyperplasia and dysplasia of alveolar epithelial cells (AEC). Injured and activated AEC then participate in aberrant cytokine signaling that perpetuates the fibrotic response (3). Of particular recent interest is the possibility that AEC contribute directly to fibrosis through epithelial-mesenchymal transition (EMT) to a myofibroblast-like phenotype. Although myofibroblasts are integral to normal repair mechanisms, the persistence of the myofibroblast beyond a period of normal repair has been associated with ECM deposition, structural remodeling, and destruction of alveocapillary units (30). Recent studies by our group (43) and others (20) have demonstrated alveolar EMT both in vitro and in vivo and have shown that the majority of myofibroblast-like cells after experimental injury are the result of alveolar EMT (20). There is mounting evidence that alveolar EMT is primarily mediated by local production and activation of transforming growth factor (TGF)-β1 (20, 43). However, factors that modify the effects of TGF-β1 and the induction of alveolar EMT have not been identified.
Nitric oxide (NO), which is a potent mediator of alveolarization and lung growth (1, 24), is produced by three isoforms of the enzyme nitric oxide synthase (NOS) designated neuronal NOS (nNOS), inducible NOS (iNOS), and endothelial NOS (eNOS). Mice that overexpress eNOS in the lung are protected from ventilator-induced lung injury (38). In a baboon model of BPD, prematurely delivered animals that received inhaled NO had increased cell proliferation in terminal bronchioles, decreased α-smooth muscle actin (α-SMA) expression, and decreased elastin deposition in their lungs compared with controls subjected to ventilation without NO (26). Additionally, TGF-β1 downregulates NOS expression in a variety of nonpulmonary cell types (10, 29, 37), and NO production is dramatically increased in TGF-β1 null mice (40), indicating a reciprocal relationship between TGF-β1 and NO.
Considering these findings, we hypothesized that NO attenuates EMT in AEC. The aims of our investigation were to: 1) determine whether NOS is expressed and enzymatically active in AEC; 2) determine the role of endogenous NO production in modulation of alveolar EMT; and 3) determine the effects of exogenous NO on alveolar EMT induced by TGF-β1.
MATERIALS AND METHODS
Cell culture and treatment.
RLE-6TN cells, which are a type II AEC line, were obtained from American Type Culture Collection (Manassas, VA) and maintained in DMEM, nutrient mixture Ham's F-12 supplemented with 10% fetal bovine serum, 20 mM/l HEPES, and 100 μg/ml Primocin (InvivoGen, San Diego, CA). In studies investigating the impact of endogenous NOS on EMT, RLE-6TN were cultured under control conditions in the presence of the NOS antagonist Nω-nitro-l-arginine methyl ester hydrochloride (l-NAME; 2 mM; Sigma, St. Louis, MO) or in the presence of 2 mM L-NAME and 100 μM diethylenetriamine NONOate [DETA-NONOate; half-life (t½) = 20 h; NO flux rate = 1.7 × 10−10 mol NO per min/plate; Alexis, San Diego, CA]. Cells were treated for 6 days based on prior studies demonstrating alveolar EMT within that time period (43). In studies investigating the impact of exogenous NO on TGF-β1-induced EMT, RLE-6TN were cultured in media alone, media supplemented with 50 pM (1.25 ng/ml) TGF-β1 (R&D Systems, Minneapolis, MN) alone, or with 50 pM TGF-β1 and 100 μM DETA-NONOate. Equivalent amounts of NO-depleted DETA-NONOate were used as controls in all experiments. Cultures were maintained in a humidified 5% CO2 incubator at 37°C, and all media were changed every 24 h.
In primary cell experiments, rat lung AEC were isolated using previously described methods (43). Briefly, type II AEC were isolated from adult male Sprague-Dawley rats by disaggregation with elastase (2.0–2.5 U/ml; Worthington Biochemical, Freehold, NJ), followed by differential adherence on IgG-coated bacteriological plates. Primary cell purities were 85–90% in all preparations. All animals were treated in accordance with the guidelines of and with the approval of the University of Texas Southwestern Medical Center Institutional Animal Care and Use Committee. Freshly isolated AEC were plated in a minimal defined serum-free medium on 1.1-cm2 tissue culture-treated polycarbonate filter cups (Transwell; Corning Costar, Cambridge, MA). For the first 24–48 h of culture, media were supplemented with 100 μg/ml cis-OH-proline (Sigma) to selectively eliminate fibroblasts from cultures (19). AEC were then cultured under control conditions with TGF-β1 or with TGF-β1 and DETA-NONOate added for 10 days as described above.
The following antibodies were used for immunofluorescence: mouse monoclonal anti-α-SMA (Sigma) and mouse monoclonal anti-p180 lamellar protein antibody (Covance, Berkeley, CA). Nonspecific mouse IgG at equivalent concentrations was used as a control in all experiments. The following antibodies were used for immunoblotting: mouse monoclonal anti-α-SMA (Sigma); mouse monoclonal anti-collagen I (Abcam, Cambridge, MA); mouse monoclonal anti-E-cadherin (BD Transduction, San Diego, CA); and mouse monoclonal anti-pro-surfactant protein B (Lab Vision, Fremont, CA).
NOS enzymatic activity assay.
Determinations of NOS enzymatic activity in the presence of excess substrate and cofactors provide a reliable quantitative assessment of enzyme abundance (28). RLE-6TN were cultured in media alone or in the presence of 50 pM TGF-β1 for 6 days. At the end of this period, cells were washed with ice-cold PBS, pelleted, and resuspended on ice in 50 mM Tris buffer (pH 7.5) containing 10 μg/ml pepstatin A, 10 μg/ml leupeptin, 10 μg/ml aprotinin, 10 μg/ml N-α-p-tosyl-l-lysine choromethyl ketone, 20 μM tetrahydrobiopterin, 3.0 mM dithiothreitol, 1.0 mM phenylmethylsulfonyl fluoride, and 10 mM 3-[(3-cholamidopropyl)dimethylammonio]-1-propanesulfonate. The cells were lysed by freeze-thawing in liquid nitrogen. NOS activity was determined by measuring the conversion of [14C]-l-arginine to [14C]-l-citrulline (28). Briefly, cell preparations (50 μl) were added to 50 μl of buffer, yielding final concentrations of reagents as follows: 2 mM β-NADPH, 2 μM tetrahydrobiopterin, 10 μM flavin adenine dinucleotide, 10 μM flavin mononucleotide, 0.5 mM CaCl2 in excess of EDTA, 15 nM calmodulin, 2 μM cold l-arginine, and 2.0 μCi/ml [14C]-l-arginine. After incubation at 37°C for 30 min, the assay was terminated by the addition of 400 μl of 40 mM HEPES buffer (pH 5.5), with 2 mM EDTA and 2 mM EGTA. The terminated reactions were applied to 1 ml of Dowex AG50WX-8 (Tris form) and eluted with 1 ml of the 40 mM HEPES buffer. [14C]-l-citrulline was collected in scintillation vials and quantified by liquid scintillation spectroscopy. NOS activity was fully inhibited by 2.0 mM Nω-nitro-l-arginine methyl ester hydro-chloride (l-NAME). The calcium dependence of NOS activity was evaluated by the addition of 2.5 mM EGTA to the incubation mixture.
NOS mRNA expression by RT-PCR.
RLE-6TN and primary rat AEC were cultured for 6 days in media or in the presence of TGF-β1, and RNA was harvested by chloroform extraction. Reverse transcription (RT) was carried out using 500 ng RNA/25 μl reaction. eNOS mRNA was specifically primed for RT using the rat eNOS reverse primer 5′-CACCGTG-CCCATGAGTGA-3′ (Genosys, St. Louis, MO). nNOS mRNA was specifically primed for RT using the rat nNOS reverse primer 5′-TTAGGAGCTGAAAACCTCATC-3′ (Genosys), and random priming was used for iNOS detection. RT reactions were performed using manufacturer-recommended concentrations of RT reagents (Taqman Reverse Transcription Reagents Kit; ABI, Foster City, CA) at 25°C for 10 min, at 48°C for 30 min, and at 95°C for 5 min, respectively, for eNOS, nNOS, and iNOS. Rat brain and lung were used as positive controls and for generating standard curves for nNOS, eNOS, and iNOS, respectively. RT negative controls were performed on initial experiments to ensure the specificity of the PCR reaction.
Real-time-PCR was performed using the Taqman method. Following RT, 3 μl of cDNA was used for each 20-μl PCR reaction. nNOS mRNA and iNOS mRNA were detected using Taqman Gene Expression Assays (nNOS: Rn00583793_m1, GenBank ref. NM_052799.1; iNOS: Rn00561646_m1, GenBank ref. NM_012611.2) (Applied Biosystems). The eNOS PCR reaction was carried out using a forward primer specific to rat eNOS 5′-CACCAGGAAGAAGACTTTTAGGA-3′ (Genosys), a Taqman eNOS probe 5′-CAACCAGCGTCCTGCAAACCGTG-3′ (Applied Biosystems), and the reverse primer 5′-CACCGTGCCCATGAGTGA-3′. PCR reactions were performed in a Prism 7700 Sequence Detection System, and data were analyzed using SDS 1.9.1 software (Applied Biosystems). A standard curve was generated for each gene product, and each sample was normalized to the relative expression of 18S RNA.
RLE-6TN and primary AEC monolayers treated as described were harvested on days 6 and 10, respectively. Filters or culture plates were rinsed, fixed with formaldehyde, and exposed to 0.2% Triton X-100 for 10 min. After being rinsed in Tris-buffered saline (TBS) and blocked with a casein-based blocking solution (CAS Block; Zymed, San Francisco, CA), filters or plates were incubated with the appropriate primary antibody overnight at 4°C. After being washed and blocked with CAS Block, filters or slides were incubated with secondary antibodies for 45 min: for α-SMA staining, anti-mouse IgG conjugated to Alexa 488 (Invitrogen, Carlsbad, CA) at a dilution of 1:250, and for lamellar protein, biotinylated anti-mouse IgG at a dilution of 1:250. For lamellar protein staining, cells were incubated with streptavidin conjugated to Alexa 594 at a dilution of 1:250 for 15 min. Monolayers were rinsed again and postfixed in 3.7% formalin. Coverslips were applied to all filters and culture dishes using Prolong Gold with DAPI anti-fade mounting medium (Invitrogen). Filters or plates were then viewed with an Axioskop 40 microscope equipped with epifluorescence optics (Zeiss, Thornwood, NY). Images were captured with an Axiocam HRC using Axiovision color digital software (Zeiss).
RLE-6TN monolayers were lysed in 2% SDS sample buffer at 37°C for 15 min. Protein concentrations were determined using a standard protein assay (Bio-Rad, Hercules, CA) with BSA as the standard. Equal amounts of protein from control and treated samples were resolved on a 10% SDS-PAGE and electrophorectically transferred to Immobilon-P nylon membranes (Millipore, Billerica, MA). Membranes were blocked overnight at 4°C with 5% nonfat dry milk in TBS with 0.2% Tween at pH 7.5. They were then incubated with primary antibody at either room temperature for 2 h (α-SMA) or overnight at 4°C (collagen I, E-cadherin, and pro-surfactant protein B). Blots were incubated with horseradish peroxidase-linked anti-mouse IgG conjugates for 2 h at room temperature, and antibody complexes were visualized using SuperSignal West Pico Chemiluminescent Substrate (Pierce, Milwaukee, WI). Glyceraldehyde-3-phosphate dehydrogenase (GAPDH) was used as a loading control, densitometric analyses were performed, and protein abundance was normalized to GAPDH. Blots were scanned using MultiDoc-It Imaging Systems (UVP Bioimaging Systems, Upland, CA) and analyzed using LabWorks 4.0 software.
Cell number determination.
RLE-6TN were seeded onto 60-mm tissue culture plates at a density of 4.2 × 103/cm2. The number of adherent cells on day 6 was then determined after culture under control conditions with TGF-β1 (50 pM) or with TGF-β1 and DETA-NONOate (100 μM). Cells were washed with cold PBS and lysed, and nuclei were stained by incubating at 37°C overnight in a solution containing 0.1% Nonidet P-40, 0.1 M citric acid, and 0.1% crystal violet. Stained nuclei were counted using a hemacytometer (42).
TdT-mediated dUTP nick end labeling assay for apoptosis.
RLE-6TN cells were cultured under control conditions in the presence of TGF-β1 (50 pM) or in the presence of TGF-β1 and NO (100 μM) for 6 days, and the proportion of cells undergoing apoptosis was determined using a fluorometric TUNEL Apoptosis Detection Kit (Upstate, Temecula, CA). The TdT-mediated dUTP nick end labeling (TUNEL) assay identified apoptotic cells by using terminal deoxynucleotidyl transferase to transfer biotin-dUDP to the free 3′-OH of cleaved DNA. The biotin-labeled cleavage sites were then incubated with avidin-FITC, nuclei were counterstained with propidium iodide (Vector Labs, Burlingame, CA), and cells were visualized by fluorescent microscopy. The proportion of TUNEL-positive cells to total cells was determined and expressed as fold change from control.
Cellular proliferation determination.
Procedures described in Immunofluorescence were used. After fixation and permeabilization, RLE-6TN cells were incubated with Ki67 primary antibody (Santa Cruz Biotechnology, Santa Cruz, CA) overnight at a 1:500 dilution and then conjugated to goat anti-rabbit Alexa 594 1:500 dilution for visualization under fluorescent microscopy. Nuclei were counterstained with DAPI, and the percentage of Ki67-positive cells to total cells was determined and expressed as fold change from control.
Data are expressed as means ± SE. NOS activity assay and RT-PCR results were analyzed by unpaired Student's t-test. Global differences in densitometric analyses were analyzed using the Kruskal-Wallis test, and specific comparisons between treatment arms were assessed using the Mann-Whitney U-test. Cell number and TUNEL apoptosis data were analyzed using ANOVA followed by Dunn's comparison between groups. Significance was defined as P < 0.05.
Expression of NOS isoforms in AEC.
NOS enzymatic activity was detectable under basal conditions in RLE-6TN cells (Fig. 1A). Total NOS activity was 1.3 pmol·mg protein−1·min−1, which is equivalent to that found in bronchiolar epithelial cells (35). Sixty-seven percent of total activity was calcium dependent, signifying the expression of either eNOS or nNOS, and 33% of NOS activity was calcium independent and indicative of iNOS activity (Fig. 1A). To specifically identify the NOS isoforms that are expressed in AEC, RT-PCR was performed for eNOS, nNOS, and iNOS. eNOS and iNOS mRNA were detected in both RLE-6TN and primary AEC under basal conditions (Fig. 1, E–H). In contrast, nNOS mRNA was not detected.
The effects of TGF-β1 on RLE-6TN NOS were also determined. Treatment with TGF-β1 decreased total NOS activity by 78% (Fig. 1B). The decline in total activity was primarily due to an 83% drop in calcium-dependent activity (Fig. 1C), whereas calcium-independent NOS activity was unaffected (Fig. 1D). Similarly, RLE-6TN and primary rat AEC cultured in the presence of TGF-β1 had 87% (Fig. 1E) and 64% less eNOS mRNA (Fig. 1G) than control cells, respectively. The relative abundance of iNOS mRNA tended to increase with TGF-β1 treatment, but the observed difference was not statistically significant (Fig. 1, F and H). Similar to control AEC, TGF-β1-treated AEC expressed no nNOS mRNA. These data suggest that TGF-β1 treatment suppresses eNOS expression and activity but does not impact iNOS in AEC.
Modulation of EMT by endogenous NOS.
To examine the role of endogenous NOS in the regulation of alveolar EMT, we suppressed endogenous NOS activity with the NOS antagonist l-NAME and performed immunofluorescence microscopy for the myofibroblast marker α-SMA. Compared with control cells, l-NAME-treated cells expressed dramatically more α-SMA protein (Fig. 2, A and B, respectively). In addition, l-NAME-treated epithelial cells displayed an altered morphology and resembled elongated fibroblasts, similar to the phenotype of TGF-β1-treated AEC (Fig. 2, F and H, respectively). In contrast, RLE-6TN treated with l-NAME and DETA-NONOate retained an epithelial morphology (Fig. 2G) and did not express α-SMA (Fig. 2C). These findings demonstrate that endogenous NOS activity is an important regulator of EMT in AEC.
Inhibition of TGF-β1-induced EMT by exogenous NO.
We next examined the effect of exogenous NO on TGF-β1-induced alveolar EMT. Control AEC (RLE-6TN) on day 6 demonstrated a rounded, cobblestone-like appearance (Fig. 3A). In contrast, TGF-β1-treated cells displayed a stellate, fibroblast-like morphology (Fig. 3E). Importantly, AEC cultured with both TGF-β1 and the NO donor DETA-NONOate retained an epithelial morphology and were virtually indistinguishable from control cells (Fig. 3I). Immunofluorescence for α-SMA was also performed on RLE-6TN cultured under the above conditions. Unlike control cells that expressed very little α-SMA (Fig. 3B), TGF-β1-cultured cells displayed a fibroblast-like morphology and expressed large amounts of stress fiber-associated α-SMA (Fig. 3F). Cells cultured with both TGF-β1 and NO retained an epithelial morphology (Fig. 3J), and α-SMA was detected diffusely throughout the cytoplasm rather than being confined to stress fibers. NO-depleted DETA-NONOate had no effect on TGF-β1-induced EMT (data not shown).
To ensure that these findings were not unique to a cell line, we performed parallel experiments using primary rat AEC cultured for 10 days. Control cells expressed very little α-SMA protein (Fig. 3, C and D), AEC treated with TGF-β1 expressed α-SMA organized into stress fibers and exhibited a fibroblast-like morphology (Fig. 3, G and H), and AEC treated with both TGF-β1 and DETA-NONOate retained an epithelial morphology and generally expressed α-SMA diffusely throughout the cytoplasm (Fig. 3, K and L).
To evaluate changes in the expression of myofibroblast markers, immunoblots for type I collagen and α-SMA were performed. Minimal collagen I or α-SMA was detected in control cells, and a dramatic increase in both collagen I and α-SMA expression occurred with TGF-β1 treatment (Fig. 4, A and B). The addition of NO dramatically reduced the expression of collagen I in TGF-β1-treated cells to control levels, but it had no effect on α-SMA expression.
To determine if the preservation of an epithelial morphology by exogenous NO is accompanied by the conservation of other epithelial characteristics, immunofluorescence and immunoblotting were performed for three characteristic epithelial markers. While control RLE-6TN expressed lamellar protein in distinct intracytoplasmic granules (Fig. 5A), TGF-β1-treated cells lost nearly all lamellar protein expression (Fig. 5B). In contrast, cells treated with both TGF-β1 and NO donor had abundant intracytoplasmic lamellar protein, similar to control cells (Fig. 5C). AEC treated with TGF-β1 and NO-depleted DETA-NONOate expressed lamellar protein at levels similar to TGF-β1-treated cells (data not shown).
E-cadherin is a critical tight junction protein that is characteristic of epithelial cells and may play a role in the initiation of EMT (16). Immunoblotting revealed that TGF-β1-treated AEC express half as much E-cadherin as untreated AEC (Fig. 6A). However, TGF-β1-treated AEC that were cotreated with NO donor expressed E-cadherin at control levels. Finally, changes in surfactant protein were determined between the treatment groups. Surfactant production is a critical indicator of AEC function (25). Immunoblots for pro-SP-B revealed that TGF-β1 decreased pro-SP-B expression by more than 50%, whereas the addition of NO maintained pro-SP-B expression at control levels (Fig. 6B).
Inhibition of TGF-β1-induced apoptosis by exogenous NO.
Based on experimental observations that TGF-β1 negatively affected cell number, we quantified adherent cell number on day 6 after seeding equal numbers of RLE-6TN under control conditions with TGF-β1 or with TGF-β1 and NO. Although TGF-β1 halved adherent cell number, TGF-β1 combined with a NO donor restored the number of AEC back to control levels (Fig. 7A). To further explore whether changes in cell number could be explained by the effects of NO on TGF-β1-mediated apoptosis, a TUNEL assay was performed on cells cultured under similar conditions. Compared with control cells, apoptosis was increased 8.6-fold in TGF-β1-exposed cells (Fig. 7, B and C). In contrast, the presence of exogenous NO more than halved the number of AEC undergoing apoptosis (Fig. 7, B and C). Cellular proliferation, as assessed by Ki67 expression, was equal in control cells, TGF-β1-exposed cells, and TGF-β1 and NO-exposed cells (data not shown).
The beneficial impact of NO on lung injury has been suggested by multiple animal and human studies. Recent studies suggest that inhaled NO may decrease the incidence of death and BPD in extremely premature neonates (2). Ventilated primates given low-dose inhaled NO have fewer myofibroblasts and less elastin deposition in alveoli than control animals (26). eNOS transgenic animals are less susceptible to ventilator-induced lung injury and have less fibrosis following injury with bleomycin (38, 44). In the present study, we have demonstrated that both endogenous and exogenous NO attenuates EMT in AEC. Our data suggest a potential mechanism by which NO impacts alveolar fate and protects against fibrosis.
Although NO performs many critical functions in the lung (31), the role of NO in the alveolar epithelium has not been well characterized. We demonstrate that eNOS and iNOS are expressed and enzymatically active in AEC. The level of calcium-dependent activity in AEC, derived from eNOS, was similar to that previously demonstrated in H441 human bronchiolar epithelial cells (35). Our finding that the suppression of endogenous NOS activity in AEC with l-NAME leads to a transition to a myofibroblast phenotype suggests that endogenous NO prevents EMT and the generation of myofibroblasts. Studies from eNOS knockout mice suggest a crucial role for eNOS in lung growth (23), in the normal response to injury, and in the prevention of fibrosis (6). Our work raises the novel possibility that alveolar eNOS directly regulates AEC fate.
We further found that TGF-β1 caused a >80% decline in eNOS activity and expression in RLE-6TN and primary rat AEC, whereas iNOS activity and mRNA were relatively unaffected. TGF-β1-mediated counterregulation of NOS activity and expression has been demonstrated in multiple nonpulmonary cell types (12, 13, 32). Elevated TGF-β1 in tracheal aspirates has been correlated with poorer prognosis in patients with IPF and with the need for home oxygen in patients with BPD (17, 21). Our finding that TGF-β1 decreases eNOS activity and expression in AEC provides a mechanism by which high TGF-β1 levels, such as those found in the tracheal aspirates of patients with IPF or BPD, may promote myofibroblast formation and initiate interstitial fibrosis.
In experiments designed to investigate the effect of exogenous NO replacement on AEC fate, we observed a conservation of epithelial morphology, decreased collagen I expression, and retention of the epithelial markers E-cadherin and pro-SP-B in the face of TGF-β1 exposure. A marked accumulation of alveolar collagen, of which collagen I is a major component (8), is found in autopsy specimens of the lungs of patients with fibrosing alveolitis and BPD (18, 41). Our finding that NO decreased collagen I expression in TGF-β1-exposed AEC suggests a pathway by which aberrant ECM deposition and subsequent fibrosis can be prevented.
Our data demonstrating that E-cadherin expression is preserved in TGF-β1-exposed AEC may have important implications in the prevention of EMT. With suppression of E-cadherin levels during EMT, epithelial cells relinquish their cell-cell adhesive properties, acquire the migratory properties of mesenchymal cells (22), and release β-catenin from tight junctions, allowing for the activation of genes involved in EMT (16). A bidirectionality to these processes is suggested by the observation that E-cadherin overexpression reverts mesenchymal cells back to epithelium (16). Therefore, inhibition of the TGF-β1-induced downregulation of E-cadherin by NO may be a critical first step in the prevention of EMT.
We also observed that pro-SP-B expression as well as lamellar protein expression is retained in AEC treated with TGF-β1 and NO. Pro-SP-B, once proteolytically processed in type II AEC to mature SP-B, is important not only to the arrangement of surfactant phospholipids in lamellar bodies and to lung function in the early neonatal period (15) but also to the prevention of respiratory distress in adult animals (7). In agreement with our findings, studies in intact lambs have shown that ventilation with inhaled NO increased SP-B mRNA in the lung (36). Together, these data suggest that NO has multiple important functions in the preservation of normal AEC phenotype.
Additionally, we demonstrated that exogenous NO more than halves the amount of TGF-β1-induced apoptosis of AEC. Animal models and human studies in patients with IPF demonstrate that AEC apoptosis contributes to fibrosis in the lung (14, 39). The precise relationship between apoptosis and EMT remains to be determined, but it may be the case that AEC, when exposed to TGF-β1 during lung injury, will undergo apoptosis or EMT. The factors regulating this “fate determination” are unknown, and one study suggests that they are likely influenced by the underlying ECM (20). Our findings that NO not only prevents TGF-β1-induced apoptosis but also attenuates EMT suggest that NO contributes to the survival of alveolar epithelial cells and to the maintenance of a healthy phenotype.
Although AEC treated with TGF-β1 and NO retained epithelial characteristics, they continued to express α-SMA protein in quantities similar to cells treated with TGF-β1 alone. Immunofluorescence demonstrated that the α-SMA in TGF-β1-treated AEC is localized to stress fibers, whereas the α-SMA in TGF-β1 and NO-treated AEC is predominantly distributed diffusely throughout the cytoplasm. The small GTPase, RhoA, which is activated by TGF-β1, may be pertinent to these observations. Once activated directly by the TGF-β1 receptor complex without the involvement of the canonical Smad-dependent TGF-β1 signaling pathway, RhoA is critical to the cytoskeletal rearrangements and stress fiber formation necessary for mesenchymal movement (4). NO directly inhibits RhoA activation through protein kinase G activity (33) in other cell types. We propose that NO may inhibit TGF-β1-induced cytoskeletal rearrangements and stress fiber formation without affecting Smad-dependent α-SMA expression via the inhibition of Rho activation. Detailed studies of Rho, NO, and EMT are now warranted.
Based on our overall findings, we postulate that under normal conditions, NO generated by endogenous NOS in the alveolar epithelium functions to maintain alveolar epithelial cell phenotype, thereby contributing to optimal alveolar development and repair following injury. In contrast, with repetitive injury or genetic predispositions that lead to chronic elevations in profibrotic cytokines, including TGF-β1 AEC, NOS is downregulated, AEC transitions to a myofibroblast phenotype, and pulmonary fibrosis ensues. A greater understanding of AEC NOS and alveolar EMT will potentially enable us to better harness the potent actions of NO and its effectors and thereby prevent or treat fibrotic lung diseases such as BPD and IPF.
This work was supported by Children's Medical Center of Dallas Clinical Research Advisory Committee Award (to S. Vyas-Read) and National Institutes of Health Grants K12-HD-047349 (Pediatric Critical Care Scientist Development Program to B. C. Willis) and U01-HL-63399 (to P. W. Shaul).
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