Effect of pirfenidone on pulmonary fibrosis in acute lung injury via the regulation of the miR-34a-5p/TGF-β1/SMAD pathway

Introduction

Acute lung injury (ALI) is a common lung disease characterized by severe acute inflammation, acute respiratory insufficiency with tachypnea, and diffuse alveolar infiltrates on chest X-ray, with a serious heterogenous pulmonary disorder and high mortality (1-3). Moreover, ALI is a clinical disorder and a global health problem (4). Pulmonary fibrosis (PF), commonly seen in ALI, involves inflammation, fibroblast activation, and excessive accumulation of extracellular matrix (ECM), which induces irreversible scar formation and remodeling of lung tissue (5,6). Smoking, infection, obesity, and environmental pollution are potential contributing factors of PF (7,8). Hence, further exploration of the pathogenesis and treatment targets of PF in ALI is necessary.

The time and degree of macrophage polarization play a role in the severity of PF (9,10). Various phenotypes of macrophages exert different effects on PF, which are affected by a number of factors, including cytokines, tissue-specific triggers, and microbial signals (11). Under the stimulation of inflammatory factors, macrophages can differentiate into the M2 type and thus contribute to the pathogenesis of PF (12), with PF in ALI being associated with this process. Th2 cytokines typically induce M2 macrophage activation, leading to the substantial production of anti-inflammatory cytokines and the expression of surface markers like Arg-1 and CD206 in the later stages of silicosis (13). IL4, a key Th2 cytokine, is a well-established inducer of M2 polarization, simulating the Th2-dominated immune microenvironment observed in PF (14,15). Studies have shown that IL4 promotes the differentiation of macrophages into the M2 subtype by activating the STAT6 signaling pathway, which is associated with fibrotic processes, including collagen deposition and inflammation resolution (14,15). Elevated IL4 levels are clinically associated with enhanced M2 macrophage infiltration and fibrotic progression in ALI patients (16,17). Pirfenidone (PFD) has been applied in a number of studies on PF. For instance, PFD was found to alleviate lipopolysaccharide-induced lung injury via the strengthening of BAP31 modulation of endoplasmic reticulum stress and mitochondrial injury (18). PFD also ameliorates liposaccharide-stimulated PF and inflammation via the inactivation of NLRP3 inflammasome (19). However, whether PFD can regulate macrophage M2 polarization to participate in PF of acute kidney injury (AKI) has not been clarified.

miR-34a-5p figures prominently in injury-like illness. For example, in intestinal ischemia-reperfusion injury, the knockdown of miR-34a-5p alleviates the accumulation of reactive oxygen species and apoptosis upon promotion via SIRT1 signaling (20). miR-34a-5p also regulates Krüppel-like factor 4 level in hyperoxia-stimulated senescence in lung epithelial cells (21). Moreover, hepatic expression of miR-34a-5p is elevated in metabolic-associated fatty-liver disease/nonalcoholic steatohepatitis, which can be reduced after PFD treatment (22). miR-34a-5p can inactivate Notch signaling to cause imbalance in macrophage M1–M2 polarization and affect inflammation in liver injury due to the coexposure to dibutyl phthalate (DBP) and benzo(a)pyrene (BaP) (23). However, the role of miR-34a-5p and its effect on M2 macrophage polarization in ALI under PFD treatment remain unclear.

In the study, we conducted in vitro and in vivo experiments aiming to determine how PFD exerts an effect on PF in ALI via the regulation of the miR-34a-5p/TGF-β1/SMAD pathway. We present this article in accordance with the ARRIVE and MDAR reporting checklists (available at https://jtd.amegroups.com/article/view/10.21037/jtd-2025-829/rc).

Methods

Cell treatment and transfection

SV40-transformed mouse alveolar macrophage (MH-S) cells (American Type Culture Collection, Manassas, VI, USA) were cultivated in RPMI-1640 medium (Gibco, Thermo Fisher Scientific, Waltham, MA, USA) together with 1% penicillin/streptomycin and 10% fetal bovine serum (FBS; Gibco) at 37 ℃ in an atmosphere containing 5% CO₂. For macrophage M2 polarization, MH-S cells were treated with 10 ng/mL of IL4 (R&D Systems, Minneapolis, MN, USA) for 1 day, which was followed by stimulation with 100 µg/mL of PFD (Shionogi & Co., Ltd., Osaka, Japan) for 2 days. The PFD concentration (100 µg/mL) was selected based on previous in vitro studies (24,25).

Treated cells were transfected with negative control (NC) inhibitor, miR-34a-5p inhibitor, NC mimic, and miR-34a-5p mimic (GenePharma, Shanghai, China) to silence or upregulate miR-34a-5p expression. These plasmids were transfected with Lipofectamine 2000 (Invitrogen, Thermo Fisher Scientific) as per the manufacturer’s instructions 1 day before any treatment.

Quantitative polymerase chain reaction (qPCR)

Total RNA was extracted from cells and tissues using a mirPremier microRNA (miRNA) Isolation Kit (Sigma-Aldrich, St. Louis, MO, USA) as per the manufacturer’s instructions. RNA was reverse transcribed to complement DNA (cDNA) via a PrimeScript miRNA cDNA Synthesis Kit (Takara Bio, Kusatsu, Japan). The levels of RNA transcripts were analyzed with a CFX96 real-time PCR system (Bio-Rad Laboratories, Hercules, CA, USA). miR-34a-5p expression was normalized to U6. The primers sequences were as follows: miR-34a-5p forward, 5'-ACACTCCAGCTGGGTGGCAGTGTCTTAGC-3'; miR-34a-5p reverse, 5'-CTCAACTGGTGTCGTGGA-3'; U6 forward, 5'-CTCGCTTCGGCAGCACA-3'; and U6 reverse, 5'-AACGCTTCACGAATTTGCGT-3'. Up to 20 µL, the reaction conditions were set as follows: predenaturation at 95 ℃ for 10 min, denaturation at 95 ℃ for 2 s, annealing at 60 ℃ for 20 s, and extension at 70 ℃ for 10 s for 40 cycles. The pooled analysis of each gene expression with triple repetition was conducted via the 2^−ΔΔCt method.

Cell Counting Kit-8 (CCK8) assay

Cells were seeded into 96-well culture dishes with 100 µL of medium including 10% serum. Subsequently, 10 µL of CCK8 solution (Dojindo Laboratories, Kumamoto, Japan) was added at 24, 48, and 72 h at 37 ℃ for approximately 2 h. Thereafter, the absorbance at 450 nm was estimated with a microplate reader (BioTek Instruments, Winooski, VT, USA).

Western blotting

Tissues and cells were lysed with radioimmunoprecipitation assay buffer with proteinase inhibitors. After fractionation via sodium dodecyl sulfate-polyacrylamide gel electrophoresis (SDS-PAGE), proteins were transferred onto polyvinylidene fluoride (PVDF) membranes, and 5% nonfat milk was applied for blocking. The primary antibodies were CD68 (ab283654; 1:1,000; Abcam, Cambridge, UK), CD206 (ab300621; 1:1,000; Abcam), Arg-1 (ab203490; 1:1,000; Abcam), YM-1 (ab93034; 1:1,000; Abcam), TGF-β1 (ab179695; 1:1,000; Abcam), p-SMAD2 (GTX03203; 1:1,000; GeneTex, Irvine, CA, USA), SMAD2 (ab33875; 1:1,000; Abcam), p-SMAD3 (GTX00969; 1:1,000; GeneTex), SMAD3 (ab84177; 1:1,000; Abcam), and GAPDH (ab125247; 1:1,000; Abcam). After being washed three times, membranes were grown with horseradish peroxidase (HRP)-conjugated secondary antibody for another 1 h. An enhanced chemiluminescence detection system (GE HealthCare, Chicago, IL, USA) was used to visualize the immunoreactive bands.

Animal experiments

Male C57BL/6 mice were purchased from the Shanghai Laboratory Animal Center, Chinese Academy of Sciences (Shanghai, China). For adaptive feeding, mice at 7–8 weeks old (18–22 g) were all kept in a specific pathogen-free environment for 1 week prior to assays, with access to food and water. To guarantee biological duplication, 15 mice were randomly divided into five groups: a control (Ctrl) group (intratracheal administration of 50-µL 0.9% saline; n=3), an IL4 group (intratracheal injection of 3 mg/kg bleomycin dissolved in saline; n=3), a PFD group (administration of 200 mg/kg of PFD 14 days after bleomycin treatment for 4 weeks; n=3), a PFD + NC mimic group (intraperitoneal injection of NC mimic into PFD mice; n=3), and a PFD + miR-34a-5p mimic group (intraperitoneal injection of miR-34a-5p mimic into PFD mice; n=3). No mice died, so these 15 mice were euthanized via cervical dislocation without anesthetics. Lung tissues were excised and rapidly frozen through liquid nitrogen within 10 min for histological analysis, qPCR, Western blotting, and immunohistochemistry (IHC). The animal experimental protocols were approved by the Experimental Animal Ethics Committee of Anhui Medical University (No. LLSC20220061), in compliance with national guidelines for the care and use of animals. A protocol was prepared before the study without registration.

Masson staining and hematoxylin-eosin (HE) staining

Tissue slides were stained with HE. After being dewaxed, rehydrated, and stained with hematoxylin (Sigma-Aldrich) for nearly 5 min, slices were rinsed for 2 min and treated with 1% eosin (Sinopharm Group, Shanghai, China) for about 5 min. Subsequently, sections were subjected to hydration with graded ethanol and vitrification with dimethylbenzene.

In Masson staining, sections were deparaffinized to water, dyed with mordant, and rinsed with water. Subsequently, azure blue staining for 3 min and hematoxylin staining for 3 min were implemented. To stain tissues red, 1% hydrochloric acid alcohol was employed for several seconds for differentiation. Following staining with ponceau-fuchsin dye for another 5–10 min, 1% phosphomolybdic acid aqueous solution was employed for differentiation. Thereafter, 5-min staining with aniline blue solution was implemented.

IHC

Paraffin-embedded slices (4 µm in size) were dried for about 1 h. Slices were deparaffinized with xylene for nearly 20 min, soaked in ethyl alcohol citrate buffer for 5 min for antigen retrieval, and rinsed with phosphate-buffered saline (PBS) and blocked with 5% bovine serum albumin for about 1 h. After incubation with the primary antibodies anti-CD206 (ab300621; 1:2,000; Abcam) and YM-1 (ab230610; 1:1,000; Abcam) overnight at 4 ℃, 50 µL of secondary antibody was added for 1-h treatment. Color development was achieved with 3,3'-diaminobenzidine tetrahydrochloride. Slices were counterstained with hematoxylin for another 10 min, followed by dehydration upon application of ethanol and xylene. Microscopy with a BX51 fluorescence microscope (Olympus, Tokyo, Japan) was applied for observation.

Statistical analysis

GraphPad Prism 9.0 (Dotmatics, Boston, MA, USA) was used to analyze the data obtained from three independently conducted assays. Data were presented as the mean ± standard deviation (SD). Comparisons of two or three groups were conducted via the t-test or one-way analysis of variance (ANOVA). P<0.05 indicated statistical significance.

Results

Cell viability and M2 macrophage polarization in IL4-treated MH-S cells were regulated by miR-34a-5p

To determine whether miR-34a-5p is involved in the ALI process, the miR-34a-5p level in MH-S cells in response to IL4 treatment was tested via qPCR. The expression of miR-34a-5p was significantly upregulated after IL4 treatment (Figure 1A). Subsequently, miR-34a-5p inhibitor was transfected to silence miR-34a-5p expression, which was verified through qPCR (Figure 1B). The CCK8 assay demonstrated that miR-34a-5p inhibition could improve the cell viability suppressed by IL4 (Figure 1C). Furthermore, Western blotting was conducted to assess the protein level of M2 macrophage polarization-related markers CD68 and CD206 and the inflammation factors Arg-1 and YM-1. It was observed that when induced by IL4, the protein levels of CD68, CD206, Arg-1, and YM-1 were significantly diminished, and the expression of miR-34a-5p was decreased. This finding suggests that miR-34a-5p inhibition could suppress M2 macrophage polarization and inflammation (Figure 1D). These data indicated that cell viability and M2 macrophage polarization are mediated by miR-34a-5p in IL4-treated MH-S cells.

Figure 1 miR-34a-5p regulated M2 macrophage polarization in IL4-treated MH-S cells. (A) miR-34a-5p expression after IL4 treatment in MH-S cells according to qPCR. (B) The transfection efficacy of NC inhibitor and miR-34a-5p inhibitor was assessed via qPCR. Cells were divided into Ctrl, IL4, IL4 + NC inhibitor, and IL4 + miR-34a-5p inhibitor groups. (C) CCK8 assay was used to assess cell viability. (D) The protein expression of CD68, CD206, Arg-1, and YM-1 was determined via Western blotting. **, P<0.01 vs. Ctrl; ^##, P<0.01 vs. IL4 + NC inhibitor. CCK8, Cell Counting Kit-8; Ctrl, control; NC, negative control; qPCR, quantitative polymerase chain reaction.

PFD modulated cell viability and M2 macrophage polarization through miR-34a-5p

To determine whether PFD regulates M2 macrophage polarization via miR-34a-5p, we treated MH-S cells with PFD. After PFD treatment, miR-34a-5p expression was silenced, as indicated by qPCR (Figure 2A). To confirm the activity of the PFD-miR-34a-5p axis in IL4-treated MH-S cells, miR-34a-5p mimic was employed to promote miR-34a-5p expression, the efficacy of which was confirmed via qPCR (Figure 2B). In the CCK8 experiment, the cell viability of IL4-treated MH-S cells was increased after PFD treatment, and this effect was reversed when miR-34a-5p expression was elevated (Figure 2C). Additionally, Western blotting demonstrated that CD68, CD206, Arg-1, and YM-1 protein expression was significantly reduced with PFD treatment, and this effect was significantly abrogated via the upregulation of miR-34a-5p (Figure 2D). Taken together, our findings suggest that PFD regulates miR-34a-5p to increase cell viability and M2 macrophage polarization.

Figure 2 PFD mediated M2 macrophage polarization through miR-34a-5p. (A) PFD was added to MH-S cells, and the expression change of miR-34a-5p was observed. (B) Transfection of miR-34a-5p mimic was assessed via qPCR. Cells were divided into the Ctrl, IL4, PFD, PFD + NC mimic, and PFD + miR-34a-5p mimic groups. (C) Viability of the five groups of cells was evaluated through CCK8 assay. (D) The protein expression of CD68, CD206, Arg-1, and YM-1 in the five groups was examined via Western blotting. **, P<0.01 vs. Ctrl; ^#, P<0.05, ^##, P<0.01 vs. IL4; ^&&, P<0.01 vs. PFD + NC mimic. CCK8, Cell Counting Kit-8; Ctrl, control; NC, negative control; PFD, pirfenidone; qPCR, quantitative polymerase chain reaction.

miR-34a-5p mediated the TGF-β1/SMAD pathway under PFD treatment

Considering that miR-34a-5p can regulate the TGF-β1/SMAD pathway, a critical functional pathway in ALI, we assessed the effect of miR-34a-5p on TGF-β1/SMAD pathway-related proteins via Western blotting. The results indicated that the expression of TGF-β1, p-SMAD2, and p-SMAD3 proteins was significantly increased with IL4 treatment and decreased with the addition of PF, with these changes being reversed after miR-34a-5p was overexpressed (Figure 3). Thus, PFD downregulated miR-34a-5p to mediate the TGF-β1/SMAD pathway in IL4-treated MH-S cells.

Figure 3 miR-34a-5p modulated the TGF-β1/SMAD pathway. TGF-β1/SMAD pathway-related proteins including TGF-β1, p-SMAD2,SMAD2, p-SMAD3, SMAD3 in the Ctrl, IL4, PFD, PFD + NC mimic, and PFD + miR-34a-5p mimic groups were analyzed via Western blotting. **, P<0.01 vs. Ctrl; ^#, P<0.05, ^##, P<0.01 vs. IL4; ^&, P<0.05, ^&&, P<0.01 vs. PFD + NC mimic. Ctrl, control; NC, negative control; PFD, pirfenidone.

PFD modulated miR-34a-5p to affect ALI in vivo

HE and Masson staining results indicated that IL4-induced tissue injury could be ameliorated by PFD treatment, while the injury was aggravated with the overexpression of miR-34a-5p (Figure 4A,4B). qPCR assay was used to assess the change in miR-34a-5p expression in tissues. The IL4-induced increase in miR-34a-5p expression was reduced after the addition PFD, but this inhibitory effect was significantly neutralized via the transfection of the miR-34a-5p mimic (Figure 4C). Western blotting indicated that after IL4 treatment, PFD inactivated the TGF-β1/SMAD pathway and that this effect was abolished via the increase in miR-34a-5p expression (Figure 4D). Additionally, IHC analysis revealed that in the PFD group, the protein expression of CD206 and YM-1 declined markedly when compared with the IL4 group; while in the PFD + miR-34a-5p mimic group, the protein expression of CD206 and YM-1 increased compared with the PFD + NC mimic group. These results suggested that miR-34a-5p can promote M2 macrophage polarization and inflammation (Figure 4E). In summary, PFD can regulate miR-34a-5p to improve ALI in vivo.

Figure 4 PFD regulated miR-34a-5p to affect ALI in vivo. Mice were randomly divided into Ctrl, IL4, PFD, PFD + NC mimic, and PFD + miR-34a-5p mimic groups. (A,B) Histomorphological analysis of tissues was performed via HE and Masson staining. Scale bar: 50 µm. (C) The expression of miR-34a-5p in tissues was determined via qPCR. (D) Western blot analysis of proteins in the TGF-β1/SMAD pathway. (E) IHC analysis of CD206 and YM-1 expression. Scale bar: 50 µm. **, P<0.01 vs. Ctrl; ^#, P<0.05, ^##, P<0.01 vs. IL4; ^&, P<0.05, ^&&, P<0.01 vs. PFD + NC mimic. ALI, acute lung injury; Ctrl, control; HE, hematoxylin-eosin; IHC, immunohistochemistry; NC, negative control; PFD, pirfenidone; qPCR, quantitative polymerase chain reaction.

Discussion

PFD is beneficial for the treatment of PF, as it can ameliorate PF through the inhibition of macrophage M2 polarization and JAK2/STAT3 signaling (26). For rat lung fibroblasts, PFD suppresses polarization to the M2 macrophage type and fibrogenic activity (27). Previous studies found that PFD ameliorates PF via downstream molecules, primarily miRNAs (28,29). According to a study by Escutia-Gutiérrez et al., PFD treatment can inhibit the overexpression of miR-34a-5p in metabolic-associated fatty liver disease or nonalcoholic steatohepatitis (22). Moreover, miR-34a-5p inactivates Notch signaling to produce an imbalance in M1–M2 macrophage polarization and increase the inflammation in liver injury due to coexposure of DBP and BaP (23). Thus, it is reasonable to assume that miR-34a-5p may function in the PFD treatment of ALI through inhibiting M1–M2 macrophage polarization. In our study, we confirmed that miR-34a-5p promoted M2 macrophage polarization in IL4-treated MH-S cells, and this phenomenon was improved by PFD.

We further sought to clarify the molecular mechanisms underlying the relationship between the PFD-miR-34a-5p axis and ALI. It is widely known that miRNAs exert a degree of suppressive or promotive function via downstream proteins. Studies have shown that the upregulation of miR-34a-5p may promote M2 macrophage polarization through multiple mechanistic pathways (23,30). For instance, miR-34a-5p is known to target negative regulators of M2 polarization, including SIRT1 (30). Besides, miR-34a-5p has been shown to induce M2 polarization in liver injury via Notch signaling activation, a pathway potentially conserved in PF (23). Our demonstration that miR-34a-5p enhances TGF-β1/SMAD signaling provides another pathway, as TGF-β1 is a potent inducer of alternative macrophage activation. TGF-β1/SMAD pathway, a key pathway in ALI (31), has been confirmed to be mediated by miR-34a-5p. Long noncoding RNA (lncRNA) CCAT2 was found to reduce miR-34a-5p expression to regulate TGF-β1/SMAD4 signaling and activate hepatic stellate cell proliferation (32). Meanwhile, miR-34a-5p was reported to regulate epithelial-mesenchymal transition partially by targeting the TGF-β1/SMAD4 pathway in silica-induced PF (33). Additionally, miR-34a-5p can reduce liver fibrosis by regulating the TGF-β1/SMAD3 pathway (34). Importantly, the TGF-β1/SMAD pathway has been shown to contribute to the polarization of macrophages to the M2 type, but PFD can suppress the TGF-β1/SMAD3 pathway to inhibit this polarization and radiation-generated lung fibrosis (25). Eucalyptol can also abrogate M2 macrophage polarization and inhibit macrophage-secreted profibrotic factor TGF-β1 (35). We thus speculated that miR-34a-5p is involved in the PFD regulation of M2 macrophage polarization via targeting of the TGF-β1/SMAD pathway. Western blot experiments confirmed that miR-34a-5p elevated the levels of STGF-β1/SMAD pathway-related proteins. Animal experiments further demonstrated that PFD downregulated miR-34a-5p to improve ALI via the TGF-β1/SMAD pathway.

Although this study did not directly assess pulmonary fibroblast activation, the regulation of TGF-β1/SMAD signaling by miR-34a-5p suggests a potential mechanism linking M2 macrophage polarization to fibroblast activation and collagen deposition. Previous studies have established that M2 macrophage-derived TGF-β1 directly activates fibroblasts and promotes collagen synthesis (36,37). Thus, the observed inhibition of TGF-β1/SMAD pathway by PFD and miR-34a-5p suppression likely attenuates fibroblast-mediated fibrosis, a hypothesis warranting direct validation in future studies.

Notably, this study has several limitations. First, although we demonstrated that miR-34a-5p regulated TGF-β1/SMAD pathway, the direct target gene of miR-34a-5p was not validated. Future work will prioritize experimental validation of this interaction to solidify the molecular mechanism. Second, while the MH-S cell line offers advantages in terms of reproducibility and genetic manipulation, it cannot fully recapitulate the heterogeneity of primary alveolar macrophages or their dynamic interactions with other lung cell types. Recent single-cell RNA sequencing studies have revealed remarkable diversity among pulmonary macrophage subsets, suggesting our findings may need validation across different macrophage populations. Future studies should incorporate primary human alveolar macrophages from PF patients, or air-liquid interface cultures of primary human bronchial epithelial cells, to preserve native tissue architecture while allowing mechanistic interrogation. Third, while we focused on macrophage polarization, we did not directly examine the effects of conditioned media from polarized macrophages on fibroblast activation or collagen production. Future studies should investigate this important aspect of macrophage-fibroblast interaction to fully understand the implications for PF treatment. Finally, we acknowledge that translating in vitro drug concentrations to clinical doses requires careful consideration of pharmacokinetic differences. Future studies could establish dose-response relationships across different experimental systems.

Conclusions

M2 macrophage polarization was mediated by miR-34a-5p in IL4-treated MH-S cells under PFD treatment. Moreover, miR-34a-5p mediated the TGF-β1/SMAD pathway in IL4-treated MH-S cells under PFD treatment. Animal assays verified that PFD modulates miR-34a-5p to influence ALI in vivo. These findings may suggest novel molecular options for the treatment of ALI.

Acknowledgments

None.

Footnote

Reporting Checklist: The authors have completed the ARRIVE and MDAR reporting checklist. Available at https://jtd.amegroups.com/article/view/10.21037/jtd-2025-829/rc

Data Sharing Statement: Available at https://jtd.amegroups.com/article/view/10.21037/jtd-2025-829/dss

Peer Review File: Available at https://jtd.amegroups.com/article/view/10.21037/jtd-2025-829/prf

Funding: This study was supported by grants from the Natural Science Research Project of Colleges and Universities in Anhui Province, China (to L.F.) (No. KJ2021A0302).

Conflicts of Interest: All authors have completed the ICMJE uniform disclosure form (available at https://jtd.amegroups.com/article/view/10.21037/jtd-2025-829/coif). The authors have no conflicts of interest to declare.

Ethical Statement: The authors are accountable for all aspects of the work in ensuring that questions related to the accuracy or integrity of any part of the work are appropriately investigated and resolved. The study was approved by the Experimental Animal Ethics Committee of Anhui Medical University (No. LLSC20220061), in compliance with national guidelines for the care and use of animals.

Open Access Statement: This is an Open Access article distributed in accordance with the Creative Commons Attribution-NonCommercial-NoDerivs 4.0 International License (CC BY-NC-ND 4.0), which permits the non-commercial replication and distribution of the article with the strict proviso that no changes or edits are made and the original work is properly cited (including links to both the formal publication through the relevant DOI and the license). See: https://creativecommons.org/licenses/by-nc-nd/4.0/.

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(English Language Editor: J. Gray)