Rationale: Recent findings suggesting transforming growth factor (TGF)-β1 activation by mechanical stimuli in vitro raised the question of whether this phenomenon was relevant in vivo in the context of pulmonary fibrosis.
Objectives: To explore the effect of mechanical stress on TGF-β1 activation and its signaling pathway in rat and human fibrotic lung tissue using a novel ex vivo model.
Methods: Rat lung fibrosis was induced using transient gene expression of active TGF-β1. Lungs were harvested at Day 14 or 21 and submitted to various stimuli in a tissue bath equipped with a force transducer and servo-controlled arm.
Measurements and Main Results: Fibrotic lung strips responded to tensile force by releasing active TGF-β1 from latent stores with subsequent increase in tissue phospho-Smad2/3. In contrast, measurable active TGF-β1 and phospho-Smad2/3 were not induced by mechanical stress in nonfibrotic lungs. Protease inhibition did not affect the release of active TGF-β1. A TGF-β1 receptor inhibitor, Rho-associated protein kinase inhibitor, and αv integrin inhibitor all attenuated mechanical stretch–induced phospho-Smad2/3 in fibrotic lung strips. Furthermore, the induction of phospho-Smad2/3 was enhanced in whole fibrotic rat lungs undergoing ventilation pressure challenge compared with control lungs. Last, tissue slices from human lung with usual interstitial pneumonia submitted to mechanical force showed an increase in TGF-β1 activation and induction of phospho-Smad2/3 in contrast with human nonfibrotic lungs.
Conclusions: Mechanical tissue stretch contributes to the development of pulmonary fibrosis via mechanotransduced activation of TGF-β1 in rodent and human pulmonary fibrosis.
Culturing on substrates and matrices with higher stiffness properties typically induces a more fibrogenic phenotype of fibroblast. Furthermore, recent in vitro work has shown that mechanical forces are able to activate transforming growth factor (TGF)-β1 and release one of the key profibrotic growth factors in vitro. This is mediated by physically opening the binding between the latent TGF-β1–binding protein, the latency-associated peptide, and the αv integrins. However, there is no direct evidence, to date, to confirm that mechanical stretch induces TGF-β1 activation in fibrotic lungs.
We demonstrated that fibrotic lung tissue activates more TGF-β1 in response to mechanical stretch than normal lung tissue via a Rho kinase and αv integrin–mediated pathway. This is the first study to show that this pathway is exaggerated in fibrotic lung disease. The translational impact of our work is highlighted by the fact that we reproduced the same results after stretching fresh human lung tissue from patients with idiopathic pulmonary fibrosis. In a ventilation study, high inspiratory pressures resulted in activation of the TGF-β1 system, presumably through mechanical stretch of the cell–matrix interface. This highlights the importance of developing drugs that reduce the stiffness in fibrotic lungs.
Idiopathic pulmonary fibrosis (IPF) is a devastating lung disease of unknown origin with a median survival of 3 to 4 years from time of diagnosis. In IPF, scar tissue accumulates and matures until normal lung tissue is slowly overtaken, eventually compromising blood-gas exchange and leading to death (1). The critical dependence of fibrogenesis on transforming growth factor (TGF)-β activity has been a central focus of research over the past two decades (2, 3). The TGF-β1 isoform is of primary importance in IPF (4). It is produced as a large latent complex (LLC) consisting of the latent TGF-β–binding protein (LTBP) attached to the extracellular matrix (ECM) and the covalently attached latency-associated peptide, which in turn encompasses the bioactive TGF-β homodimer in a noncovalent manner. The biology of TGF-β1 activation and release of the bioactive dimer from the resting LLC is a complex process initiated by various stimuli, including reactive oxygen species, serine protease, and matrix metalloproteinase (MMP), each of which dissociates the active TGF-β from the latency-associated peptide (5–8). In addition, recent studies suggest that mechanical stimuli induce latent TGF-β activation through αv-integrins in several cell types (9–16), followed by fibrogenesis (15–19). However, whether mechanical stress induces TGF-β1 activation in vivo with respect to fibrogenesis in the lung is still an unresolved question. Contraction-dependent TGF-β1 activation is an important mechanism, termed “mechanotransduction,” by which parenchymal cells can sense and respond to the rigidity of the ECM surrounding them. It is also likely that cells respond to changes in ECM rigidity during breathing, which could be a potential TGF-β1 activation stimulus promoting the persistence of fibrosis, particularly in already fibrotic and thereby stiffened lungs.
Mechanical ventilation of patients with IPF, particularly single-lung ventilation to facilitate lung biopsy or cancer surgery, may mediate further deterioration of lung function and promote progressive fibrogenesis, suggested by perioperative mortality rates of patients with IPF that can be as high as 20% (20–22). The exact mechanisms behind this observation remain unclear; however, TGF-β1 activation via mechanotransduction due to increased tissue stiffness represents a plausible explanation.
To further investigate the role of TGF-β1 activation via mechanotransduction in lung fibrogenesis, we used several in vitro and ex vivo models, including a novel approach of stretching rodent and human lung strips in a tissue bath and isolated rodent lung ventilation. These systems allow the study of the effect of mechanical stretch on the activation of the TGF-β1/Smad2/3 signal transduction pathway ex vivo.
A horizontal tissue bath was constructed with dimensions 3 × 1 × 1 cm. A force transducer and a servo-control arm were used in tandem with a digital controller interface (Models 400A, 322C, 604C; Aurora Scientific Inc., Aurora, Canada). Trimmed tissue strips were pasted to metal clips via ethyl 2-cyanoacrylate–based adhesive (Electron Microscopy Sciences, Inc., Hatfield, PA) and attached to a hook on the force transducer and servo-arm in the tissue bath (Figure 1D). Lungs were removed en bloc and cut transversely in 10 × 2 × 2-mm sections. Tissue strips were equilibrated in a solution (Krebs buffer) in the tissue bath for 16 minutes to bring them to 37°C, after which the solution was changed and tissue strips were left at rest in the bath without application of stretch. Samples from the tissue bath were taken at this point (thus, before the strips had undergone stretch) and termed “Before” samples. Next, 10 to 15 seconds of 5 mN was applied to obtain the Young’s modulus measurements. After these measurements, 5 to 15 minutes of 5- to 20-mN cyclical stretch was administered, and samples from the tissue bath solution samples were collected (termed “After”). Lung strips were incubated for 6 hours in Krebs buffer with the Complete EDTA-free Protease inhibitor cocktail (1X; Roche Diagnostics GmbH, Mannheim, Germany) without any mechanical stimuli and then frozen in liquid nitrogen before storage at −80°C for further analysis. Nonstretched lung tissues were termed “Wash,” and stretched lung tissue were called “Stretched.”
We aimed to model the tension of breathing in the tissue bath by setting the tension of cyclic stretch to 15 mN for 4 minutes oscillating at a frequency of 2 Hz, to a length of 1.1 times the original resting length of the tissue strips (lung strips measured 10 × 2 × 2 mm). The estimated pressure was 15 mN/4 mm2 = 0.37 mN/mm2; 0.37 mN/mm2 is equal to 37.73 cm/H2O, which is within the pressure range observed in the lungs of patients with IPF (23). TGF-β1 activity in the bath solution measured 4 minutes before stretching (Before) served as internal control to the activity of TGF-β1 measured after stretched bath solution (After). In the nonstretched Wash group, lung strips were incubated in the bath for 16 minutes, after which the tissue wash solution was taken after 4 minutes of resting and taken out of the bath. Both fibrotic lung (adenovirus encoding active TGF-β1; AdTGF-β1) and nonfibrotic lung (AdDelete [AdDL]) were used for this tissue bath experiment.
Lung tissues were collected with consent of the patients (approved by the Research Ethics Board of St. Joseph’s Healthcare Hamilton). Control lung tissue was collected from patients undergoing surgery for cancer. Lung fibrosis tissue was collected from patients undergoing biopsy for the diagnosis of unclear interstitial lung disease. The biopsies analyzed in this study revealed a usual interstitial pneumonia (UIP) pattern on histopathology. UIP is the distinct histological pattern that is observed in IPF lung. Nonfibrotic human lungs were used as nonfibrotic control. After biopsy, all tissue was stored in RPMI cell culture medium for 1.5 hours. In preparation for analysis in the tissue bath apparatus, the tissue was washed in cold Krebs plus bovine serum albumin and sectioned into 10 × 2 × 2-mm strips.
For more details, see the online supplement.
The administration of adenovirus encoding active TGF-β1 (AdTGF-β1) causes transient synthesis of high levels of transgenic active TGF-β1 in the lungs, peaking after 4 to 7 days and completely disappearing by Day 10 (2). After 14 days, the lungs show marked and persistent fibrosis over a period of 56 days, suggesting that the fibrosis in this model depends on endogenous TGF-β1 activation (2, 3, 24). We demonstrate significant fibrosis accompanied by an up-regulation of collagen synthesis in AdTGF-β1–treated rat lungs at Day 14 as compared with nonfibrotic animals that received an empty vector (AdDL) (Figures 1A and 1B). We also showed that LTBP expression was increased in AdTGF-β1–treated rat lungs at Day 7 and this increase lasted until Day 21 (Figure 1C).
Figure 1D shows the model setup in which we placed a sensitive force transducer and a servo-controlled arm oriented horizontally in a tissue bath to apply mechanical stretch to lung strips. We investigated whether duration and intensity of stretch impacted the activation of TGF-β1 in rat fibrotic tissue. First, the tension of breathing in the tissue bath was set such that the length change of the lung strip was 10% of the resting tissue length, and cyclic frequency in 2Hz, corresponding to tissue stretch estimated in vivo during breathing. A constant force of 5 mN mechanical stimulation was applied to the fibrotic lung strips for 5 to 15 minutes. The activity of TGF-β1 was measured before and after stimulation of the lung strips. Interestingly, both active and total TGF-β1 were released into the bath solution after stretch. As the duration of stretch got longer, the released active TGF-β1 increased with a significant time-dependent effect on the active/total TGF-β1 ratio (Figure 1E). To evaluate the impact of stretch intensity on TGF-β1 activation, increasing forces from 5 to 20 mN for 4 minutes were applied during stretch to fibrotic lung strips. As the applied tension increased, the active/total TGF-β1 ratio and active TGF-β1 significantly increased in response to the stimulus (Figure 1F). These data indicate that mechanotransduction of active TGF-β1 is a direct response to varying tissue tension in respect to both duration and amplitude of stimulus. Based on these results, we modeled the tension of breathing in the tissue bath for the following experiments by setting the resting tissue tension to 15 mN for 4 minutes and cyclically stretching (2 Hz) the lung strip with a length change of 10% of the resting tissue length.
We investigated whether TGF-β1 activity and downstream TGF-β1 signaling were activated by cyclic stretch in the fibrotic rat lung tissue. First, the Young’s modulus, reflecting stiffness, of lung slices from fibrotic (AdTGF-β1) and nonfibrotic (AdDL) lungs was measured, showing a significant increase in stiffness in fibrotic lungs compared with nonfibrotic lungs (Figure 2A). The activation of TGF-β1 in bath solution was measured before and after mechanical cyclic stimulation. Although stretch did not enhance active TGF-β1 release in nonfibrotic lungs, an increase of active TGF-β1 release was observed in fibrotic lungs after mechanical stimulation (Figure 2B). In contrast, the total TGF-β1 level was not significantly increased by mechanical stimulation (Figure 2B). Moreover, we demonstrated that there was a moderate positive correlation between the stiffness of each lung strip and active TGF-β1 level released in the bath solution (Figure 2C; r = 0.5549, P = 0.004). Furthermore, protein extracts from lung strips were collected 6 hours after mechanical stimulation to assess Smad2/3 phosphorylation. Unlike nonfibrotic lungs, there was a significant increase in Smad2/3 phosphorylation, reflecting TGF-β1 signaling activation, in AdTGF-β1–treated lung tissue after stretch (Stimuli) compared with unstretched lung (Control) (Figure 2D). These data clearly suggest that the active TGF-β1 released by mechanical stretch is able to activate downstream signaling pathways. Next, we addressed the effect of serine protease and MMPs as activators of TGF-β1 in our system. We demonstrate that an equivalent amount of active TGF-β1 was released from tissue with or without inhibitors after two serial challenges (Figure 2E). Moreover, pan-protease inhibitor had no effect on Smad2 phosphorylation induced by mechanical stretch in fibrotic lung (Figure 2F). This result indicates that the activation of TGF-β1 is independent of serine proteases and MMP activity in our model.
To address the underlying mechanisms of mechanical stretch–induced TGF-β1 activation, inhibitors related to different pathways of TGF-β1 activation and expression were used in our system. First, we used the TGF-β1 kinase (ALK5) inhibitor SD208, which attenuated Smad2/3 phosphorylation induced by mechanical stretch in fibrotic lung (Figure 3A), suggesting that activated TGF-β1 acts through the ALK5 TGF-β receptor. To investigate whether the enhanced active TGF-β1 observed in the bath wash After stretch was due to enhanced secretion and release of de novo synthesized TGF-β1 from cells, a protein secretion inhibitor (PTI) was used. PTI did not have any effect on stretch-induced pSmad2/3 expression in the lung (Figure 3B), indicating that mechanical stretch did not influence the release of TGF-β1 from cells. Rho kinase has been shown to be central in mechanotransduction-induced activation of TGF-β1 (15, 25–27); therefore, we used a Rho-associated protein kinase (ROCK) inhibitor, Y-27632, to assess the effect of this pathway in our model. Y-27632 significantly attenuated Smad2/3 phosphorylation induced by mechanical stretch in fibrotic lung (Figure 3C). This result suggests that the activation of TGF-β1 induced by mechanical stretch occurs, at least partly, through cellular mechanotransduction involving Rho kinase. Next, the effect of an αv integrin small molecular inhibitor (CWHM-000012-8) was assessed. This inhibitor attenuated stretch-induced Smad2/3 phosphorylation in fibrotic lung (Figure 3D) at both concentrations of αv inhibitor used. Overall, these data suggest that mechanical stretch enhances TGF-β signaling via a mechanotransduced integrin–mediated TGF-β activation pathway rather than via secretion and increased release of TGF-β.
To investigate whether mechanical stretch can activate TGF-β1 in the whole lung, we used an ex vivo lung ventilation system. For each lung, the left lung received control ventilation and the right lung was given challenge ventilation. During control ventilation, the inspiratory pressure for the lungs was 5 cm H2O, and during the expiratory cycle lungs were allowed to expire to an imposed positive end-expiratory pressure of 2 cm H2O. The lungs were ventilated by performing three total lung capacity maneuvers each minute, which involved a 6-second inflation to a pressure of 30 cm H2O, followed by relaxation against a positive end-expiratory pressure of 2 cm H2O; 30 cm H2O is a pressure that would be reached by many patients with IPF at lung volumes well below total lung capacity. The significant increase in elastance of AdTGF-β1–treated rat lungs in comparison to AdDL nonfibrotic lungs confirmed increased lung stiffness in fibrotic lungs before ventilation (Figure 4A). Six hours after ventilation there was a significant increase in Smad2/3 phosphorylation in fibrotic lung tissue receiving challenge ventilation compared with control ventilation (Figure 4B). Nonfibrotic lungs did not show any differences in phosphorylated Smad2/3 between control and challenge ventilation. Immunohistochemistry was performed on rodent lungs to understand the regional effects of ventilation of fibrotic lungs. AdDL-treated lungs did not show any differences in pSmad2 staining between control and challenge ventilation. In AdTGF-β1–treated rodents, whether subjected to control or challenge ventilation, there was strong pSmad2 staining in the fibrotic areas. Interestingly, the AdTGF-β1–treated lungs subjected to challenge ventilation showed more pSmad2-stained cells in the nonfibrotic areas than in AdTGF-β1–treated lungs subjected to control ventilation (Figure 4C). We also assessed the effect of ventilation on lung tissue by hematoxylin and eosin staining, which shows that ventilation did not induce any significant damage to the lung structure in both nonfibrotic and fibrotic lung (Figure 4D).
Our tissue bath model was used on human pulmonary fibrosis biopsy specimens and nonfibrotic control lungs. The histology of all fibrosis samples included in this study showed a typical UIP pattern with fibroblastic foci (Figure 5A). The human fibrotic lung slices had significantly increased stiffness compared with nonfibrotic control lung (Figure 5B), with tissue Young’s modulus similar to the fibrotic rat lungs shown in Figure 2A. After being exposed to the same stretch protocol as rat lung slices, human fibrotic lung strips showed an increased active/total TGF-β1 ratio in the bath solution to a similar degree as observed in fibrotic rat lungs, whereas nonfibrotic control lungs showed no TGF-β1 activation after mechanical stimulation (Figure 5C). Similarly, unlike nonfibrotic controls, we found that phosphorylated Smad2/3 expressions were increased in UIP lung after cyclic mechanical stimulation (Figure 5D). These data suggest that the active TGF-β1 released by mechanical stretch is able to activate Smad2/3 phosphorylation in UIP lungs.
In this paper, we demonstrate that cyclic mechanical stress and mechanical ventilation are physiologically relevant stimuli of the TGF-β1 signal transduction system during lung fibrogenesis. TGF-β1 is an immunomodulatory and fibrogenic factor that is assembled as a latent complex in the endoplasmic reticulum and processed in the Golgi before it is secreted and stored as a latent complex in the tissue (28). Extracellular activation of TGF-β1 is required before it acts on its receptors and can be mediated by various stimuli. The biological activation is a complex, tightly regulated process, and studies in vivo have yet to define the biological significance of simple proteolysis of latent TGF-β1 (29). TGF-β1 induces the differentiation of myofibroblasts (30), key cells in fibrogenesis. Myofibroblasts can contribute to release active TGF-β1 from a matrix-bound latent complex via contraction of their actin filaments and tethering to integrins and thus can perpetuate fibroblast activation and matrix deposition (9, 31, 32). When contraction occurs in rigid ECM, myofibroblasts are able to physically pull apart the LLC to release the TGF-β1 active homodimer (9). Although this has been elegantly demonstrated for skin tissue, there is no evidence, to date, to show whether mechanical stretch could induce TGF-β1 activation in fibrotic lungs. We here show that mechanical forces that occur in the lung tissue during the breathing cycle are indeed able to promote fibrogenic signals and thus contribute to disease progression in pulmonary fibrosis.
In our experiments, we found that active TGF-β1 is released from latent complexes in fibrotic rat lung and IPF tissue by mechanical force and that the release is dependent on duration and intensity of stimulation. These results suggest that the mechanical stretch may worsen fibrosis in vivo through enhanced TGF-β1 activation. In contrast to fibrotic lungs, we did not detect TGF-β1 activation in our experiments in nonfibrotic rat or human tissues, but we cannot exclude the possibility that physiological activation of TGF-β1 may also play a role in tissue homeostasis in normal lungs. The most plausible reason for the difference of response to mechanical stress between fibrotic and nonfibrotic lung may be lung stiffness. Tissue stiffness as measured by Young’s modulus was significantly increased in fibrotic compared with control lungs. Furthermore, lung stiffness was positively correlated with the amount of active TGF-β1 released after mechanical stretch. Previous studies have shown that the stiffness of bleomycin-induced fibrotic lung is significantly increased compared with normal tissue when measured by atomic force microscopy (32). In patients, increased lung stiffness is indirectly measured by spirometry, which typically shows a loss of FVC (33). In IPF, lung stiffness is likely a major contributor to the persistence and progression, perhaps even initiation of fibrosis. Similar phenomena have been reported to exist in liver fibrosis, where it has been suggested that increased tissue stiffness may precede the fibrotic response (34). Fibroblasts isolated from fibrotic human lung maintain responsiveness to variations in stiffness of the matrix on which they are cultured, and studies showed that tissue stiffness cues and mechanical transduction may offer unique and potent targets for inactivating IPF fibroblasts (35). The biomechanical properties of tissue are complex and, among other factors, regulated by crosslinking enzymes, such as transglutaminases (TG) and lysyl oxidases (LOX/LOXL). These enzymes are attractive therapeutic targets in fibrosis, considering that they are up-regulated in IPF lung, and mice deficient in TG2 or LOXL2 are resistant to experimental lung fibrosis (36, 37).
Using a set of pharmacological inhibitor studies, we show that mechanical stretch induces the activation of the TGF-β1 signaling pathway via Rho kinase and αv integrins. In contrast to Rho kinase and integrin inhibitors, a potent protein secretion inhibitor (PTI) did not affect TGF-β1 signaling in our experiments. These data suggest that mechanical forces acting on fibrotic tissue induce the activation of TGF-β1 through the ROCK and αv integrin without involving newly synthesized or secreted latent TGF-β1. ROCK activity is known to increase with matrix stiffness (38), and ROCK inhibitor has been shown to prevent myofibroblast differentiation and bleomycin-induced pulmonary fibrosis in mice (39, 40). Furthermore, recent studies have identified a role for mechanical matrix properties in regulating baseline and TGF-β1–stimulated contraction of lung fibroblasts and suggested that stiff fibrotic lung tissue may promote myofibroblast activation through contractility-driven events (41). Serine/threonine proteases and MMPs are enzymes that can directly cleave the TGF-β1 latent complex to activate TGF-β1. Protease and MMP inhibition did not change the activation of TGF-β1 from fibrotic lung strips, indicating that TGF-β1 activation during mechanical stimuli was not related to serine/threonine proteases and MMPs in this model. Other major TGF-β1 activators depend on integrin cytoskeleton interactions with the LLC in the ECM (9, 11, 15–19). It is quite likely that integrins may turn out to be the most prominent mediator of TGF-β1 activation during mechanotransduction. Integrin subtypes containing αv and β 1, 3, or 5 isoforms physically link the LLC with the cell cytoskeleton and are directly involved in TGF-β1 activation and fibrosis in vivo (9, 11, 15–19, 42–44). In addition to TGF-β1 activation, integrin signaling also activates TGF-β1 downstream pathways and regulates actin remodeling in fibroblasts through focal adhesion kinase (45). Not surprisingly, the αv integrin inhibitor used in our study significantly attenuated stretch-induced TGF-β1 activation and Smad2/3 phosphorylation. These data highlight that αv integrins play a critical role in mechanical stress–induced TGF-β1 activation in fibrotic lungs and support previously published in vitro and in vivo studies (9, 11, 15–19, 25–27, 42).
This published evidence together with our work indicates the presence of a mechanotransduction feedback loop leading to continued pathologic tissue stimulation in the fibrotic and stiffened microenvironment with ongoing TGF-β1 activation from latent stores. It is well known that the ECM is a large reservoir for cytokines and growth factors including TGF-β1 (30), which are released immediately on injury to start repair processes without delay. In the lungs of patients with IPF, the TGF-β1 storage protein, LTBP-1, is significantly increased, especially in fibrotic foci, demonstrating that abnormal ECM stores even more of this potent growth factor than normal tissue (46). TGF-β1 can signal via many downstream pathways, but the canonical Smad pathway is critically important to the maintenance of fibrogenic stimuli (47–49). The Smad pathway is induced in our model of mechanical stretch, and it can be blocked by several pharmacological inhibitors, all of them having therapeutic potential. Thus, our ex vivo work not only shows that TGF-β1 mechanotransduction activates a positive feedback loop in IPF tissue that leads to a vicious cycle and expansion of a profibrotic microenvironment but also describes a novel model that may become a useful tool to assess the efficacy of novel compounds that interact with the mechanotransduction process.
We next designed an ex vivo experiment to model a situation that has been repeatedly described for patients with IPF (i.e., the progression or exacerbation of fibrosis after mechanical ventilation) (20–22). We used a mechanical rodent ventilator to ventilate whole rat lungs ex vivo at low and high inspiratory pressures. Ventilation of fibrotic (and stiffened) lungs showed a significant phosphorylation of Smad2 and Smad3 6 hours after ventilation challenge similar to that seen with fibrotic lung strips in the tissue bath model. Challenge ventilation with high inspiratory pressure also induced pSmad2 expression in nonfibrotic areas of fibrotic lungs without causing visible morphological damage to the lung structure. Other groups have previously shown, using an ARDS model of acid instillation, that rodent lungs produce more TGF-β1 3 days after ventilation challenge (50, 51). They also showed that ventilator-induced lung injury was capable of inducing epithelial–mesenchymal transition (51). In our study, mechanical ventilation did not change phosphorylation of Smad2/3 in nonfibrotic lungs. Similar observations were found in a previous study that investigated the effect of mechanical ventilation on the lungs during experimental ARDS; in this study, mechanical ventilation did not enhance TGF-β1 expression in the lungs of normal mice, but it induced the expression of TGF-β1 expression in mice with ARDS, followed by epithelial–mesenchymal transition (50). Both this study and our experiments do not allow us to exclude that longer or more disruptive ventilation strategies could activate TGF-β in normal lungs. However, our data clearly support the possibility that ventilation of fibrotic lungs may promote further fibrogenesis and may help explain the high mortality risk for patients with IPF undergoing lung surgery (20–22).
The main limitation of our report is that the experimental approach does not fully reproduce the physiological pressures present in the lung during breathing. Indeed, the pattern of stretch that we applied in the tissue strip model does not mimic the pressure/tension fluctuations that are seen with normal inspiratory and expiratory cycles. The tension used for the mechanistic and intervention studies corresponds to a pressure of 37 cm H2O. Although this may be above the physiologic range of pressures experienced by healthy individuals, it has been proposed that patients with lung fibrosis could achieve these pressures while breathing close to the inspiratory capacity during activity (23). It is reassuring that we found comparable activation of the TGF-β1 signaling when we modeled tension fluctuations more physiologically using the whole lung ventilation system, where the lungs were repeatedly exposed to inspiratory and expiratory cycles with pressures up to 30 cm H2O (similar to the 37 cm H2O that was applied to the tissue strips).
In summary, we have shown that the key fibrogenic factor TGF-β1 is activated in stiff fibrotic lung tissue after mechanical stretch applied to lung strips by a force transducer or to whole lungs by a rodent ventilator. We demonstrated that this activation of TGF-β1 is independent of protease activity but due to the effect of cellular mechanotransduction through Rho kinase and αv integrins. Moreover, TGF-β1 mechanotransduction is already enhanced in fibrotic tissue, suggesting that tissue stiffness is of major importance in driving fibrogenesis. Based on these data, it is reasonable to propose that invasive ventilation may induce exacerbation, or progression, of fibrosis in patients with established IPF through growth factor release from fibrotic tissue characterized by abnormal biomechanical properties. It is tempting to speculate that IPF progression might, at least in part, be driven by tissue stretch imposed by breathing against increasing tissue rigidity in patients with IPF, once they reach a certain threshold of lung stiffness. Therefore, therapeutic strategies that reduce lung stiffness are of significant interest, as they may dampen the biomechanical signals that drive disease progression via blocking the vicious cycle of TGF-β1 activation from latent stores in fibrotic lungs.
The authors thank Jiaji Xia for help with the Western blots and reporter cell assays. They also thank Katherine Radford for help with patient consent forms; Drs. Miller, Findlay, Schieman, and Cox for helping collect human tissue samples for this study; Mara Ludwig from McGill University in Montreal for help with tissue bath apparatus designs; and Mark Inman and Jennifer Wattie for help with the rodent ventilator experiments.
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*These authors contributed equally to the presented work.
Supported by the Pulmonary Fibrosis Foundation I. M. Rosenzweig Junior Investigator Award (C.S.); and the Fonds de Dotation “Recherche en Santé Respiratoire,” the Fondation du Souffle, and the Canadian Pulmonary Fibrosis Foundation (P.-S.B.).
Author Contributions: Conception and design: A.R.F., C.S., P.-S.B., J.G., and M.K.; analysis and interpretation: all authors; drafting the manuscript for important intellectual content: A.R.F., C.S., P.-S.B., K.A., M.I., G.J., J.G., and M.K.
This article has an online supplement, which is accessible from this issue’s table of contents at www.atsjournals.org
Originally Published in Press as DOI: 10.1164/rccm.201508-1638OC on January 15, 2016