Withaferin A was procured from Aptus laboratories (Hyderabad, India) and TGF-β1 was obtained from Bio-legend (United States); Bleomycin sulfate was procured from Cipla labs (India); Masson’s trichrome staining kit, Sirius red, Chloramine-T, Hydroxyproline, and Ehrlich reagent were procured from Sigma-Aldrich, Anti-ZO-1, anti-E-cadherin, anti-Smad 2/3, anti-p Smad 2/3, anti-vimentin, anti-NF-κB p65, anti-p VEGF, anti-p p38 MAPK, anti-p FAK, and anti-p PLCγ1 were procured from Cell Signaling Technology, while anti-Col 1A2, anti-Col 3A1, anti-smooth muscle actin, anti-CTGF, anti-fibronectin, and anti-TGF-β1 were obtained from Santa Cruz Biotechnology (United States). ELISA kits were purchased from eBioscience, United States. TGF-β bioplex kit was procured from Merck-Millipore. Rest of the chemicals and reagents were of analytical grade and obtained from commercially available sources.

HFL1 cells were procured from ATCC (ATCC^® CCL153^TM) and A549 cells were purchased from National Centre for Cell Science (NCCS, Pune, India). HFL1 and A549 cells were cultured in F-12K medium (ATCC) and RPMI medium (Sigma-Aldrich, United States) respectively; supplemented with 10% fetal bovine serum and 1% anti-biotic solution (Invitrogen, United States). TGF-β1 was selected to induce fibrotic events at a concentration of 10 ng/mL in both HFL1 and A549 cell lines. WFA was dissolved in DMSO and a stock concentration of 10 mM was prepared, stored at -20°C and diluted with respective media at required concentrations before use. Cells were cultured and treated with WFA at various concentrations (0.25, 0.5, and 1 μM) 2 h prior to induction of TGF-β1 and incubated for 24 h at 37°C maintained in 5% CO₂ incubator. All the experiments were performed in three internal replicates.

The effect of WFA on viability of HFL1 and A549 cells was evaluated using MTT assay (Maurya et al., 2011). Briefly, 10,000 cells per well of HFL1 and A549 were seeded in 96-well plates and after 24 h of incubation, WFA was treated at concentrations in serial dilution from 0.78 to 50 μM (7 serial dilutions). Post 24 h of incubation, MTT solution was added at a final concentration of 0.5 mg/mL, incubated for 4 h followed by addition of DMSO to dissolve the formazan crystals then incubated for 30 min and absorbance was measured at 570 nm with a multi-mode spectrophotometer. Experiments were performed thrice in triplicates and data was expressed as percentage cell viability versus concentration of the compound by taking control cells as 100% viable cells.

The ability of the cells to migrate was assessed using cell migration assay. Briefly, HFL1 cells were plated at a density of 1 × 10⁵ cells in 12 well culture plates and allowed to grow till 70–80% confluence was attained. The monolayer of the cells was disrupted to create a gap in a straight line using a sterile 200 μL micropipette tip, and the wells were washed with PBS to remove the detached cells (Choi and Kim, 2015). Cells were treated with TGF-β1 and WFA for 24 h. Images of the gap between cells were captured at time points of 0 and 24 h, the width of the gap was measured in all groups using microscopic scaling and quantified using image J software.

Cells were cultured on cover slips and after 24 h of plating, cells underwent starvation (media with no serum) for 6 h, followed by WFA treatment 2 h before TGF-β induction. After 24 h of incubation, cells were fixed with 4% paraformaldehyde for 15 min, washed with PBS and permeabilized with 0.1% Triton X-100 in PBS for about 30 min. Cells were washed twice with PBS and incubated for 1 h with blocking solution (3% BSA in PBS). The blocked cells were incubated with primary antibodies anti-vimentin and anti-α SMA at 1:100 dilutions in 3% BSA overnight at 4°C. On the following day, cells were washed thrice with PBS, incubated with secondary Fluorescein isothiocyanate (FITC) anti-rabbit and rhodamine anti-mouse antibodies for 2 h at room temperature. Cells were washed thrice in PBS (5 min per wash) and then cover slips were mounted with vectashield hardset antifade mounting medium^® with DAPI (Vector Labs, Burlingame, CA, United States) on a glass slide. Slides were visualized using a confocal microscope (Leica TCS SP8 Laser Scanning Spectral Confocal).

Stimulated and unstimulated cell tissue lysate were used to detect changes in phosphorylated Smad2 (Ser465/Ser467) and Smad3 (Ser423/Ser425) using the Luminex system. This assay was performed with customized highly sensitive MILLIPLEXMAP signaling 6-plex magnetic bead human TGF-β kit (Cat # 48-614MAG Millipore) based on protocol provided by manufacturer.

Animal studies were carried out in male Swiss albino mice, 8 weeks old, purchased from Teena labs (Hyderabad, India) and acclimatized for a week before study initiation. Animals were allowed free access to food and water ad libitium and maintained with 12/12 h light and dark cycle. All the animal experiments were approved by institutional animal ethics committee (IAEC) of NIPER-Hyderabad, in accordance with approved protocol. Animals were randomly divided into five groups (n = 12): (1) Control group, (2) BLM alone group (1 U/kg/day × 2), (3) BLM+WFA (2 mg/kg), (4) BLM+WFA (4 mg/kg), and (5) WFA alone (4 mg/kg). Mice were administered with 1 U/kg/day × 2 of BLM via oropharyngeal route while WFA was administered once daily from day 1 for 28 days by intraperitoneal (i.p.) route. To avoid pulmonary hemorrhage due to CO₂ method of euthanasia, animals were euthanized using 5% Isoflurane (SurgiVet isoflurane vaporizer-4215061) before bronchoalveolar lavage (BAL) fluid collection. Lungs were harvested after euthanizing animals with CO₂ asphyxiation on the day of termination for the pathological and molecular studies.

Bronchoalveolar lavage fluid was collected thrice from each animal by passing 1 mL of ice-cold PBS into the trachea, followed by gentle aspiration of the fluid by inserting a catheter in the trachea (Bale et al., 2016). The percentage recovery of lavage fluid was approximately 85% and did not differ among the animal groups significantly. BAL fluid collected from each animal was pooled and subjected to total and differential cell count using automatic hematology Advia 2120i system (Siemens). Subsequently, BAL fluid was centrifuged at 300 g, for 10 min at 4°C, and the supernatant was stored at -80°C or used immediately to perform total protein content of BAL using Bradford assay and lactate dehydrogenase (LDH) levels were estimated using commercially available kit (Accurex, India).

Animal body weights were recorded at the termination of the study. Lungs were collected after necropsy; cleaned and weighed. Lung weight index was calculated as ratio of lung weight to body weight.

Left lung tissues collected on the day of sacrifice were fixed in 10% non-buffered formalin solution and embedded in paraffin. Sections of 5 μm thickness were cut using microtome and subjected to H&E staining to observe pathologic morphological changes in the tissue. Further, to estimate the collagen deposition in the lung tissues, sections were stained with Masson’s trichrome and picrosirius red while the mast cell accumulation in lungs was estimated by toluidine blue staining as per the standard protocols (Veerappan et al., 2012; Godugu et al., 2013; Vogel et al., 2015).

Lung tissues were weighed, homogenized, and subjected to centrifugation maintaining at 4°C. Tissue supernatants were stored at -80°C in aliquots or used immediately to estimate cytokine expression. Pro-inflammatory cytokines including TNF-α and IL-1β were evaluated for their expression in lung tissues using commercially available ELISA kits (eBioscience, TNF-α Catalogue No-50-173-67; IL-1β Catalogue No-50-112-8807).

Lung tissue slides were deparaffinised in xylene and rehydrated in series of gradient alcohol. Antigen retrieval was performed by heating sections for 10 min in citrate buffer followed by eliminating endogenous peroxidase using 3% H₂O₂. The sections were incubated with 3% BSA to avoid non-specific background prior to incubation with primary antibodies including mouse anti-α-SMA antibody (1:100), rabbit anti-vimentin antibody (1:100), and rabbit anti-NF-κB p65 for 30 min at room temperature. HRP-linked polymer detection system was used to develop color reaction, sections were then counterstained with hematoxylin, mounted with resinous mounting solution and visualized for immunoreactivity and percentage of positive cells were calculated (Godugu et al., 2017).

Procedure for western blot analysis was adopted from (Patel et al., 2015) with slight modifications. Briefly, cells were lysed using radio immunoprecipitation assay lysis buffer (RIPA) (Sigma-Aldrich) and lung tissues were homogenized using tissue protein extraction reagent (TEPER) lysis buffer, mixed with protease and phosphatase inhibitors (Sigma-Aldrich) maintained at 4°C, to obtain supernatants. Protein concentration was determined using Bicinchonic acid commercial assay kit (Sigma-Aldrich). Equal amounts of proteins were separated on SDS polyacrylamide gels and transferred onto polyvinyldifluoride (PVDF) membrane (Bio-Rad). After transfer, protein of interest was blocked for non-specific binding (3% BSA) and incubated with primary antibody overnight. Blots were washed thrice with TBST, each time for 10 min and incubated with HRP-conjugated secondary antibody for 1 h at room temperature. Membranes were visualized using enhanced chemiluminescence reagents by chemdoc (Vilber fusion Fx) and the protein bands were subjected to densitometry analysis using Image J software. The membranes were probed with β-actin antibody as an internal control. Each blot shown represents at least three independent experiments.

All results (in vitro and in vivo) were expressed as mean ± SEM of three independent experiments. Statistical analysis was performed by ANOVA followed by Tukey’s post hoc test. All statistical analyses were performed using GraphPad Prism^® Version 5 software. Probability values less than 0.05 levels were considered as statistically significant.

In vitro effects of WFA on cell viability were tested on A549 (human alveolar adenocarcinomic basal epithelial cells) and HFL1 (human fetal lung fibroblasts) cells; IC₅₀ values were found to be 3.46 ± 0.2 and 4.45 ± 0.12 μM respectively at 24 h studied using MTT [3-(4, 5-dimethylythiazol-2-yl)-2, 5-diphenyl tetrazolium bromide] assay (Figure 1A). Based on the results of cell viability assay performed at various concentrations, maximum safe concentrations (<1 μM) were selected so that viability of the cells was not affected upon WFA treatment and concentration dependent studies were performed with WFA at 0.25, 0.5, and 1 μM in cultured cells. TGF-β1 stimulation promotes the rate of cellular migration (Barrett et al., 2017). Therefore, to delineate the effect of WFA on cell migration, wound scratch assay was performed with WFA at concentrations of 0.25, 0.5, and 1 μM on TGF-β1 (10 ng/mL) treatment for 24 h in HFL1 cells. WFA ameliorated TGF-β1 induced rate of cell migration (Figures 1B,C) in a concentration dependent manner demonstrating the decrease in the inducible cell proliferation rate by WFA. It was also evident that there was a significant decrease in migration rate of HFL1 cells by WFA at 0.5 and 1 μM, compared to control fibroblast cells suggesting the inhibitory effect of WFA on constitutive cell proliferation where in WFA attenuated the basal cell migration rate of fibroblasts.

EMT, the transformation of fibroblasts to myofibroblasts is crucial in pathogenesis of fibrotic disorders of different organs (Li et al., 2016). In in vitro studies, TGF-β1 is a key cytokine used to induce EMT events in various fibroblasts, specifically in A549 cells where EMT plays a prominent role. Therefore expression of some of the EMT related proteins were investigated upon WFA treatment on both A549 and HFL1 cell lines (Figure 2). TGF-β1 stimulated cells resulted in significant damping of both epithelial markers E-cadherin, Zonula occludens (ZO-1) and enhanced expression of vimentin. WFA treatment increased the expression of E-cadherin as well as ZO-1 and decreased the expression of mesenchymal vimentin in A549 cells concentration dependently.

Consistent with the effect of EMT inhibition of WFA on A549 cells, WFA also decreased the expression of key mesenchymal markers including α-smooth muscle actin (α-SMA) and fibronectin in TGF-β1 induced fibroblast cells in concentration dependent manner. To further confirm the effects of WFA on EMT markers, immunofluoresence assay was carried out on fibroblast cells (Figure 3). Results of immunocytochemistry assay clearly revealed the enhanced expression of mesenchymal markers α-SMA and vimentin in TGF-β1 treated cells compared to control cells. The distribution and expression of both α-SMA and vimentin were decreased upon WFA treatment (Figure 3) which was in consistency with the key findings of EMT events obtained by western blot.

As WFA ameliorated EMT in vitro (A549 and HFL1), we further examined the molecular mechanism through which WFA suppresses TGF-β1 induced EMT. TGF-β1 is a key fibrotic cytokine that drives EMT by excess ECM synthesis (Moustakas and Heldin, 2016). TGF-β1 stimulation phosphorylates Smad 2/3 and forms heterocomplex with Smad 4 which modulates the target gene expression (Nakao et al., 1997). We assessed the effects of WFA on TGF-β1/Smad signaling pathway and as expected, there was a marked increase of p-Smad 2/3 protein expression in TGF-β1 treated cells compared to control cells and WFA significantly attenuated the phosphorylation of Smad 2/3 (Figure 4A,I).

Connective tissue growth factor (CTGF) is recognized as a major matricellular protein induced by TGF β1, functions as a central mediator in fibrosis was significantly increased in TGF-β1 treated cells and WFA decreased CTGF expression in a concentration dependent manner (Figures 4A,H). WFA effects on collagen 1A1 and collagen 3A2 were evaluated for their expression on TGF-β1 stimulation and found significant reduction in the expression of these proteins compared to TGF-β1 treated cells (Figures 4F,G). To confirm the anti-fibrotic effects of WFA mediated by Smad 2/3 signaling, in vitro fibroblasts cell lysates were investigated for their expression using TGF-β signaling 6-plex magnetic bead based kit (Figures 4D,E). Bioplex assay results implicated the amelioration of p-Smad 2 and p-Smad 3 proteins by WFA intervention on TGF-β1 stimulation in a concentration dependent manner which correlated with the results of p-Smad 2/3 expression obtained from western blot. These results demonstrated that WFA may attenuate PF through inhibiting TGF β1-Smad 2/3 signaling pathway.

WFA is reported to exert anti-angiogenic effects in some cancers and to effectuate the role of WFA as anti-angiogenic agent in PF, some of the significant target proteins of angiogenesis like p-p38 MAPK (mitogen activated protein kinase), p-VEGF (vascular endothelial growth factor), p-FAK (focal adhesion kinase), and p-PLCγ1 (phospholipase C) were evaluated (Figure 4B). Notable increase in expression of the aforementioned proteins was observed in TGF-β1 treated HFL1 cells. WFA treatment resulted in significant decreased expression of p-p38 MAPK and its key effector molecule p-VEGF concentration dependently. Further, direct intracellular signaling molecules of VEGF including p-FAK and p-PLC γ1 were significantly attenuated by WFA, suggesting the possible role of WFA as anti-angiogenic agent in ameliorating PF. Consistent with this, p-VEGF expression in BLM treated lung sections was significantly ameliorated by WFA at both doses (Figures 4J–M). As inflammation is one of the critical phases in fibrotic progression, effect of WFA on key inflammatory mediator, NF-κB p65 was evaluated. WFA significantly decreased the enhanced expression of NF-κB p65 in a concentration dependent manner (Figure 4C,N).

Withaferin A significantly decreased the total number of infiltrated cells in BALF which were elevated in BALF collected from BLM induced murine lungs (Figure 5A). Likewise, WFA significantly decreased the infiltrations of lymphocytes and neutrophils in a dose-dependent manner in BLM treated mice (Figures 5B,C). In parallel, WFA decreased the levels of BALF total protein content in BLM challenged mice, suggesting that WFA may attenuate lung microvascular leakage (Figure 5D). Furthermore, WFA also inhibited the lung damage induced by BLM illustrated by the decrease in LDH activity in BALF (Figure 5E). In addition to the recruitment of lymphocytes and neutrophils, the inflammatory phase of BLM is also featured by significant lung edema due to severe lung injury and ECM deposition, as measured by lung to body weight index which was substantially decreased by WFA (Figure 6A). Excessive collagen deposition in lung tissues of BLM treated group was reflected by increase in soluble collagen levels and hydroxyproline content respectively (Figures 6D,E). WFA reduced this BLM induced hydroxyproline and soluble collagen levels in a dose-dependent manner demonstrating the anti-fibrotic effect of WFA. Further, we examined the effect of WFA on surrogate parameters of oxidative and nitrative stress including glutathione levels and nitric oxide content respectively (Figures 6B,C). WFA augmented the expression of glutathione and decreased the nitric oxide levels in lung tissues.

To validate whether WFA exerts anti-fibrotic effects by modulating histological changes, we performed Hematoxylin and Eosin (H&E) staining to observe key morphological alveolar changes while Masson’s trichrome and picrosirius red stains were performed to evaluate collagen deposition and toluidine blue staining to detect extent of mast cell infiltration (Figure 7). A standard quantitative analysis based on improved Ashcroft scoring system was done in H&E stained sections to examine the histological changes (Figure 6F). H&E stained BLM treated lung tissues showed markedly increased alveolar thickness and obliteration of pulmonary musculature with fibrotic masses, while control lung tissues exhibited no fibrotic burden with normal lung architecture. WFA treatment at 2 mg/kg reduced the fibrous obliteration; excluding entailed single fibrotic masses while WFA at 4 mg/kg resulted in lung architecture devoid of fibrotic masses but with slightly enlarged alveoli and was comparable to normal lung architecture. In accordance with the observed restoration of histological changes, the anti-fibrotic effect of WFA was further supported by Masson’s trichrome and picrosirius red stains, in which WFA reduced the increased collagen deposition induced by BLM. Toluidine blue staining of lung sections illustrated decreased density of accumulated mast cell infiltration in WFA treated groups compared to BLM challenged mice.

To investigate whether WFA modulates the production of inflammatory cytokines including IL-1β and TNF-α, we evaluated their expression by ELISA in lung tissue supernatants, which was ameliorated upon treatment with WFA (Figures 5F,G). Further, immunoblot assay revealed that WFA treatment decreased the expression of decisive inflammatory cytokine, NF-κB p65 in lung tissue. In addition, WFA dramatically ameliorated the EMT signaling matrix proteins and fibrotic events following BLM induction. WFA substantially increased the expression of epithelial marker E-cadherin, ZO-1 and decreased mesenchymal markers including α-SMA. Moreover, WFA attenuated the protein expression of pivotal fibrotic markers including TGF-β1, vimentin, collagen 3A1 and p-Smad 2/3 in dose-dependent manner, thus illustrating that the anti-fibrotic potential of WFA was mediated via TGF-β/Smad signaling (Figure 8). Next, we performed immunohistochemical staining for proteins α-SMA, vimentin, NF-κB p65, and p-VEGF in lung sections to measure the extent of myofibroblast transformation, inflammation, and angiogenesis, respectively (Figure 9). Immunostaining in various regions of the lung revealed significant decrease in the distribution and immunoreactivity of α-SMA, vimentin, and NF-κB p65 proteins by WFA dose-dependently. These results indicate that WFA bears the potential to inhibit inflammation, EMT, ECM, fibrotic responses, and angiogenesis in BLM challenged mice.