Modafinil Suppresses Hypertrophic Scar Formation by Inhibiting Adenosine Deaminase and Activating Adenosine Signaling

Modafinil (MF) is a widely used wake-promoting compound initially developed for the management of sleep-wake disorders such as narcolepsy [1]. Its clinical utility has since expanded to conditions including chronic fatigue syndrome, attention deficit hyperactivity disorder, cocaine dependence, and obesity [2,3,4,5,6]. MF is also recognized for enhancing attention, learning, and other cognitive domains, and exhibits notable neuroprotective properties [1,5,7,8].

Recent work has highlighted an additional biological profile of MF: robust anti-inflammatory and anti-fibrotic activity across diverse experimental systems [9,10,11,12]. MF has been shown to limit ischemia–reperfusion injury [13,14,15] and reduce inflammation in models of testicular torsion [16], inflammatory bowel disease [17], multiple sclerosis [18], and inflammatory liver disease [11]. In clinical settings, prolonged MF administration (three years or more) has been associated with slower disability progression in patients with multiple sclerosis [19]. Mechanistically, MF diminishes immune cell infiltration [11,18] and reduces circulating pro-inflammatory cytokines, including interleukin 1β, interleukin 6, and tissue necrosis factors [11,14,17,20]. Beyond immunomodulation, MF suppresses fibrogenic pathways such as hepatic stellate cell activation in liver fibrosis [11] and fibroblast activation in pulmonary fibrosis models [10]. Collectively, these findings indicate that MF regulates fibroblast and immune cell activation, proliferation, infiltration, and their production of inflammatory and fibrogenic mediators. Despite these broad actions, the primary molecular target(s) responsible for MF’s anti-inflammatory and anti-fibrotic effects remain incompletely defined.

Current evidence indicates that MF is associated with increased cAMP signaling involving the A_2A and A_2B adenosine receptor (A_2AAR and A_2BAR) [10,21,22]. The cAMP cascade serves as a key endogenous regulator of inflammation and fibrosis through its downstream effectors, exchange protein directly activated by cAMP (Epac) and protein kinase A (PKA) [23,24,25,26,27]. MF has been shown to inhibit K_Ca3.1 currents via PKA [22] and to reduce expression of K_Ca3.1 and K_Ca2.3 through Epac–dependent mechanisms [11]. Alterations in these channels closely align with fibrotic phenotypes, as decreased Epac signaling and subsequent increases in K_Ca3.1 expression have been documented in various human and experimental fibrotic conditions, including transforming growth factor-β (TGFβ)-stimulated human hepatic stellate cells [11,28]. Conversely, activation of Epac or inhibition of K_Ca3.1 attenuates inflammatory and fibrotic responses [11,28,29,30]. Although these data strongly implicate cAMP signaling in MF’s therapeutic actions, the upstream trigger responsible for MF-induced cAMP elevation and AR engagement remains unresolved. To address this gap, we examined the hypothesis that MF increases local adenosine availability by inhibiting adenosine deaminase (ADA), thereby initiating A_2AAR/A_2BAR–dependent cAMP signaling in fibroblasts and immune cells.

The S- and R-isomers of MF (MF-S and MF-R) were synthesized by Cellion Biomed Inc. (Daejeon, Republic of Korea) according to a previously described protocol [31]. The PKA inhibitor H-89, the K_Ca3.1 channel blocker TRAM-34, and TGFβ were purchased from Sigma-Aldrich (St. Louis, MO, USA). The K_Ca3.1 channel activator 1-ethyl-2-benzimidazolinone (1-EBIO) and the non-selective AR agonist 5′-N-Ethylcarboxamidoadenosine (NECA) were obtained from Tocris Bioscience (Minneapolis, MN, USA). The CD73 inhibitor AB680 was purchased from MedChemExpress (Monmouth Junction, NJ, USA). Contractubex^® Gel was obtained from Merz Therapeutics (Frankfurt, Germany). Unless otherwise specified, all other reagents were obtained from Sigma-Aldrich. MF-S, MF-R, TRAM-34, and 1-EBIO were initially dissolved in dimethyl sulfoxide (DMSO), and working solutions were prepared in distilled water with a final DMSO concentration of 0.5%.

Primary human lung fibroblasts from a normal adult (NHLFs) were purchased from Lonza (Basel, Switzerland). HFL-1 human fibroblast cell line from human fetal lung tissue (CCL-153), NIH-3T3 murine fibroblasts (CRL-2795), and Jurkat T cells (clone E6-1) were obtained from the American Type Culture Collection (ATCC; Manassas, VA, USA). NHLFs and HFL-1 cells were maintained in FGM-2 (Lonza, CC-3131) and F-12K medium (ATCC 30-2004), respectively. NIH-3T3 cells were cultured in DMEM (Hyclone, Logan, UT, USA) supplemented with 10% fetal bovine serum (FBS), 100 U/mL penicillin, and 100 μg/mL streptomycin. Jurkat T cells were maintained in DMEM/F12 medium (Thermo Fisher Scientific, Seoul, Republic of Korea) with 10% FBS and 80 U/mL penicillin and 80 μg/mL streptomycin. All cell lines were incubated at 37 °C in a humidified 5% CO₂ atmosphere. Medium was replaced twice weekly, and cells were subcultured every 7 days.

MF-S dissolved in DMSO was incorporated into ointment formulations consisting of pure petroleum jelly and liquid paraffin at ratios of 1:7:2, 0.5:7.5:2, 0.1:7.9:2, and 0.03:7.97:2. The ointment base (petroleum jelly:liquid paraffin = 8:2) served as the placebo control.

New Zealand White rabbits (n = 10; 2.6–3.0 kg) were obtained from Koatech (Pyeongtaek-si, Republic of Korea) and maintained under controlled environmental conditions (16–22 °C, 40–60% humidity, <60 dB noise, positive pressure, 12-h light/dark cycle). Animals were anesthetized with intramuscular administration of alfaxan (6 mg/kg) and Rompun (5 mg/kg). Hypertrophic scars were induced by generating full-thickness excisional wounds (8-mm biopsy punch) extending through the epidermis, dermis, and perichondrium on the ventral surface of each ear (four wounds per ear). After postoperative disinfection, wounds were left to heal spontaneously for 4 weeks.

Scar maturation and re-epithelialization were confirmed by gross inspection. Scars were allocated to eight groups (six scars per group): one placebo, four MF-S (0.3%, 1%, 5%, 10%), one Contractubex (positive control), and one negative control. Treatments were applied topically once daily for 4 weeks (0.05 mL per scar). At the end of treatment, rabbits were euthanized, and scar tissues were harvested, fixed in 10% neutral-buffered formalin, and processed for histological evaluation.

Macroscopic imaging of each scar site was performed at baseline (initiation of treatment) and week 4 using a dermatoscope (Folliscope PS, Lead M, Seoul, Republic of Korea). Hypertrophic scar severity was graded according to the following criteria (Table 1).

Formalin-fixed tissues were dehydrated, embedded in paraffin, and sectioned at 3-μm slices. Sections were stained with hematoxylin and eosin (H&E) or Masson’s Trichrome (DAKO, Santa Clara, CA, USA). Measurements were performed using the Zeiss ZEN microscope software (version 2.1; Carl Zeiss AG, Oberkochen, Germany). Measurements included epidermis-to-cartilage distance at the thickest point of each scar, epidermal thickness, and the corresponding distance in adjacent normal skin. The scar elevation index was calculated as the ratio of the epidermis-to-cartilage distance at the thickest point of the scar to that of normal skin. Collagen deposition was quantified from 400× images using ImageJ (version 1.54k; National Institutes of Health, Bethesda, MD, USA) by measuring the proportion of blue-stained collagen.

Cells were treated with MF-S for 10 min prior to analysis. Intracellular cAMP levels were measured using a cAMP assay kit (R&D Systems, Minneapolis, MN, USA). Extracellular and intracellular adenosine concentrations were determined using a human adenosine ELISA kit (MyBioSource, San Diego, CA, USA). Purified ADA from calf intestine or cultured cells was treated with MF-S, and ADA enzymatic activity was subsequently measured using an ADA activity assay kit (Abcam, Cambridge, UK). ADA activity was determined using an ADA activity assay kit (Abcam, Cambridge, UK). Purified ADA from calf intestine was incubated with MF-S at the indicated concentrations prior to enzymatic analysis. For cellular measurements, cells were treated with MF-S for 10 min, lysed, and ADA activity was measured in the cell lysates.

Cells were rinsed with ice-cold PBS and lysed in protein extraction buffer containing protease inhibitors. Protein concentrations were determined using the Bradford assay. Lysates (30 μg protein) were separated by SDS-PAGE and transferred to nitrocellulose membranes. After blocking with 5% bovine serum albumin in TBST for 1 h, membranes were incubated overnight at 4 °C with primary antibodies and for 1 h with HRP-conjugated secondary antibodies. Primary antibodies included Epac1 (sc-28366), Epac2 (sc-28326), and GAPDH (sc-25778) from Santa Cruz Biotechnology (Santa Cruz, CA, USA); collagen 1α (Col1, #66948) from Cell Signaling Technology (Danvers, MA, USA); CD39 (PA5-86969) and CD73 (PA5-29750) from Thermo Fisher Scientific; and Smad4 (ab40759) from Abcam. Signals were detected by chemiluminescence, and band intensities were analyzed using LAS-4000 mini and IMAGE GAUGE software (version 1.2; Fuji Film, Minato-ku, Tokyo, Japan).

Whole-cell patch-clamp recordings were performed on NIH-3T3 cells using an EPC-9 amplifier (HEKA Elektronik, Lambrecht, Germany). Cells were voltage-clamped at 0 mV, and 650-ms voltage ramps from −100 to +100 mV were applied every 10 s. The extracellular solution contained (in mM): 150 NaCl, 6 KCl, 1.5 CaCl₂, 1 MgCl₂, 10 HEPES, and 10 glucose, with the pH adjusted to 7.4 using NaOH. The pipette solution contained (in mM): 40 KCl, 100 K-aspartate, 2 MgCl₂, 0.1 EGTA, 4 Na₂ATP, and 10 HEPES, with the pH adjusted to 7.2 using KOH. K_Ca3.1 currents were activated by loading 1 μM Ca²⁺ through the patch pipette and adding 1-EBIO (100 μM) to the bath solution.

Competitive radioligand binding assays were used to evaluate MF binding to A₁, A_2A, A_2B, and A₃ ARs, using membrane preparations and subtype-specific [³H] radioligands. MF was serially diluted (8-point, 4-fold series) starting at 2 mM. Reference ligands were similarly prepared. After incubation (1 h for A₁/A₃; 2 h for A_2A/A_2B), samples were filtered through pre-treated GF/C plates, washed, dried, and analyzed using a MicroBeta2 counter. Percent inhibition was calculated relative to background-corrected high-control values. The half-maximal inhibitory concentration (IC₅₀) values were determined by nonlinear regression and subsequently converted to K_i according to the Cheng–Prusoff equation [32].

Data are presented as mean ± standard error of the mean (SEM). Statistical differences were evaluated using one-way ANOVA followed by Bonferroni’s post hoc test or a two-tailed Student’s t-test. A p-value < 0.05 was considered statistically significant.

Previous studies have shown that racemic modafinil (MF) increases intracellular cAMP levels in fibroblasts, vascular smooth muscle cells, and osteoblasts [21,22]. Based on these findings, we investigated whether the individual enantiomers, MF-S and MF-R, similarly elevate cAMP levels in fibroblasts, and we compared the effect of MF-S with that of NECA, a potent, non-selective AR agonist. Treatment of primary human lung fibroblasts (NHLFs) with MF-S for 10 min resulted in a concentration–dependent increase in intracellular cAMP levels (Figure 1A, left panel). NECA (Figure 1A, middle panel) and MF-R (Figure 1A, right panel) induced comparable increases in cAMP. At a concentration of 100 nM, MF-S did not differ significantly from either NECA or MF-R in its capacity to elevate intracellular cAMP levels. Because cAMP signaling inhibits K_Ca3.1 activity via PKA activation, we next examined whether MF-S and MF-R suppress K_Ca3.1 currents in NIH-3T3 fibroblasts (Figure 1B–D). Outward currents were elicited by intracellular application of 1 μM Ca²⁺ via the patch pipette and subsequent stimulation with 100 μM 1-EBIO in the bath solution. These currents were markedly reduced by the selective K_Ca3.1 blocker TRAM-34 (10 μM) (Figure 1D), confirming their identity as K_Ca3.1–mediated currents. Both MF-S and MF-R inhibited K_Ca3.1 currents in a concentration–dependent manner (Figure 1B,C). The IC₅₀ values were 5.5 ± 0.6 nM for MF-S and 11.1 ± 1.5 nM for MF-R. Importantly, pretreatment with the PKA inhibitor H-89 abolished the inhibitory effect of MF-S on K_Ca3.1 currents (Figure 1C), indicating that MF-S–induced K_Ca3.1 inhibition is mediated through a PKA–dependent mechanism.

Previous reports have suggested that MF and MF-S activate cAMP signaling via A_2AAR and A_2BAR [10,21]. To determine whether MF-S and MF-R directly engage ARs, we performed competitive radioligand binding assays using membrane preparations from HEK293 or CHO-K1 cells stably expressing individual human AR subtypes (HEK293/A₁AR/G_α15, CHO/A_2AAR/G_α15, HEK293/A_2BAR/G_α15, CHO/A₃AR/G_α15). These assays revealed that both MF-S and MF-R exhibit very low or undetectable binding affinity for all tested AR subtypes (Table 2). These findings indicate that the cAMP-elevating effects of MF-S and MF-R are unlikely to be attributable to direct agonistic interactions with ARs.

Adenosine increases intracellular cAMP through activation of A_2AAR and A_2BAR. Extracellular adenosine is generated from ATP via the sequential enzymatic actions of the ectonucleotidases CD39 and CD73 [33], whose expression and activity are known to be positively regulated by cAMP [34,35]. On this basis, we investigated whether MF-S modulates CD39 and CD73 expression. Exposure of human lung fibroblasts (NHLFs and HLF-1) to MF-S (100 nM) for 24 h resulted in a marked increase in the protein expression of CD39 (Figure 2A) and CD73 (Figure 2B).

Because extracellular adenosine is subsequently transported into cells via nucleoside transporters [36], we examined intracellular adenosine levels following MF-S treatment. Short-term exposure (10 min) to MF-S significantly increased intracellular adenosine concentrations in both NHLFs and Jurkat T cells (Figure 3). Kinetic analysis using Hill’s equation revealed that the concentration required to elicit the half-maximal effective concentration (EC₅₀) for MF-S–induced adenosine elevation was 18.37 ± 15.17 nM in NHLFs and 12.9 ± 5.9 nM in Jurkat cells (Figure 3, right panels). MF-S also increased adenosine levels in Jurkat cell supernatant (data not shown).

To determine whether adenosine generation is required for MF-S–induced cAMP signaling, we examined the effect of the selective CD73 blocker AB680 on MF-S–mediated cAMP signaling. AB680 (1 nM) alone did not alter intracellular cAMP levels or Epac1 and Epac2 expression, whereas MF-S (100 nM) treatment significantly increased all three parameters. Notably, co-treatment with AB680 effectively abolished MF-S–induced increases in cAMP, Epac1, and Epac2 (Figure 4A,B). In addition, MF-S reversed TGFβ–induced Col1 upregulation in NHLFs (Figure 4C), an effect that was also blocked by AB680. These findings indicate that MF-S activates cAMP signaling predominantly through enhanced adenosine production.

Because Smad4 is a central mediator of TGFβ–driven fibrotic signaling [37], we next examined its expression. TGFβ treatment for 24 h increased Smad4 protein levels, whereas co-treatment with MF-S markedly attenuated this effect (Supplementary Figure S1). These results suggest that MF-S suppresses profibrotic signaling downstream of TGFβ. We therefore evaluated the anti-fibrotic effects of MF-S in an in vivo model.

Given our previous findings that MF or MF-S suppresses fibrosis in murine models of liver and lung fibrosis, we assessed the therapeutic efficacy of MF-S in a rabbit ear model of hypertrophic scarring. Unlike murine fibrosis models, this model allows pharmacological intervention after the inflammatory phase has largely subsided. Topical administration of MF-S significantly reduced scar formation. Macroscopic scar scores, scar elevation indices, epidermal thickness, and collagen density were all markedly increased in untreated scars but were significantly attenuated by MF-S treatment (Figure 5). The anti-scar effects of MF-S were comparable to those observed with Contractubex, a clinically used positive control.

Prior studies have demonstrated that phenyl ring moieties contribute to the binding of ADA [38,39]. Because MF-S contains phenyl rings, we hypothesized that MF-S increases adenosine levels by inhibiting ADA activity. ADA catalyzes the conversion of adenosine to inosine; therefore, inosine production was used as an indirect measure of ADA activity. Treatment of purified calf intestine ADA with MF-S resulted in a concentration–dependent reduction in enzymatic activity (Figure 6A, left panel). Nonlinear regression analysis using the Hill equation yielded an IC₅₀ value of 0.32 ± 0.21 nM for MF-S (Figure 6A, right panel). We next examined whether MF-S inhibits ADA activity in a cellular context. Treatment of NIH-3T3, HFL-1, and Jurkat T cells with MF-S led to a concentration–dependent decrease in ADA activity in all cell types tested (Figure 6B). These collective findings strongly indicate that MF-S directly inhibits ADA activity. Finally, the inhibitory potential of MF-S was compared with that of pentostatin, a potent and irreversible ADA inhibitor. No statistically significant difference in ADA inhibition was observed between MF-S (100 nM) and pentostatin (1 nM), indicating that MF-S exhibits a comparable inhibitory effect on ADA activity under the tested conditions.

In the present study, MF-S inhibited ADA activity in a concentration–dependent manner, resulting in increased adenosine and cAMP levels in both fibroblasts and immune cells. Importantly, MF-S–induced activation of adenosine–cAMP signaling was abolished by pharmacological inhibition of adenosine production using the selective CD73 inhibitor AB680. These findings indicate that MF-S enhances adenosine availability primarily through ADA inhibition, thereby activating downstream cAMP signaling pathways. In addition, MF-S increased the expression of CD39 and CD73, which may further amplify adenosine signaling by promoting extracellular adenosine generation. Because adenosine exerts immunosuppressive and anti-inflammatory effects [24,40], and cAMP signaling suppresses fibrotic responses through inhibition of Smad signaling and activation of Epac- and PKA–dependent pathways [25], activation of the adenosine–cAMP axis provides a mechanistic framework for the anti-inflammatory and anti-fibrotic effects of MF and its isomers observed in vitro and in vivo. Collectively, these findings suggest that MF and its isomers may represent promising therapeutic candidates for inflammatory, autoimmune, and fibrotic disorders.

Previous studies have demonstrated that MF and MF-S elevate intracellular cAMP levels in diverse cell types, including fibroblasts and smooth muscle cells [21,22]. Notably, MF– or MF-S–induced cAMP signaling and anti-fibrotic effects were suppressed by pharmacological antagonism of A_2AAR or A_2BAR [10,21]. However, the present radioligand binding assays revealed no evidence of direct interaction between MF-S and ARs, indicating that cAMP activation occurs through an indirect mechanism. Given that MF-S inhibits ADA and increases adenosine levels, it is likely that endogenously generated adenosine subsequently activates A_2AAR and A_2BAR, leading to cAMP elevation. Structural studies have suggested that the presence of a phenyl ring is important for ADA inhibition [38,39]. Thus, the phenyl moiety of MF and its isomers may contribute to their ADA-inhibitory activity. Consistent with this mechanism, blockade of CD73–mediated adenosine production using AB680 effectively suppressed MF-S–induced cAMP signaling, further supporting the conclusion that MF-S activates cAMP signaling via enhanced adenosine generation.

Accumulating evidence indicates that cAMP-elevating agents upregulate CD39 and CD73 expression and activity. NECA, forskolin, and prostaglandin E₂ increase CD73 expression and function in human microvascular endothelial cells [35], while membrane-permeable cAMP analogs like 8-bromo-cAMP and cAMP-elevating agents such as forskolin and prostaglandin E₂ enhance CD39 expression and enzymatic activity via PKA–dependent pathways in macrophages [34]. These observations suggest that cAMP positively regulates the ectonucleotidase system. Accordingly, MF and its isomers, by elevating intracellular cAMP, may promote CD39 and CD73 expression through cAMP–dependent mechanisms, thereby establishing a feed-forward loop that enhances adenosine generation and signaling.

Cyclic AMP plays a central role in modulating inflammatory and fibrotic responses through coordinated regulation of Smad, Epac, and PKA signaling pathways. Smad proteins are key mediators of tissue fibrogenesis [41], and TGFβ1 induces fibrosis primarily via activation of Smad signaling [37]. In the present study, MF-S reversed TGFβ-induced upregulation of Smad4, suggesting that MF-S suppresses profibrotic signaling downstream of TGFβ (Supplementary Figure S1). Smad4 serves as an essential and potentially limiting co-mediator for Smad3-dependent transcription. Thus, suppression of TGFβ–induced Smad4 upregulation by MF-S may effectively dampen canonical Smad signaling, consistent with previous reports demonstrating that cAMP signaling functionally inhibits TGFβ/Smad3/4 transcriptional responses [42]. Furthermore, cAMP exerts anti-fibrotic effects by inhibiting epithelial–mesenchymal transition and extracellular matrix production through Epac–mediated pathways. Notably, profibrotic stimuli, including TGFβ and angiotensin II, downregulate Epac expression, and this reduction contributes to tissue fibrosis [25]. Additionally, cAMP inhibits TGFβ–induced fibrosis by suppressing the expression and production of connective tissue growth factor [43,44]. Parallel to its anti-fibrotic role, cAMP signaling suppresses immune cell activation and proliferation, thereby attenuating inflammatory responses [24,45]. Therefore, cAMP-elevating agents such as MF-S may counteract fibrogenesis by restoring cAMP signaling and suppressing TGFβ–driven profibrotic pathways.

Upregulation of the Ca²⁺-activated K⁺ channels K_Ca3.1 and K_Ca2.3 upregulation has been reported in immune cells and fibrotic tissues in both human patients and animal models of inflammatory, autoimmune, and fibrotic diseases. Increased expression of these channels enhances Ca²⁺ influx, thereby promoting immune activation, inflammation, and fibrogenesis. Profibrotic and pro-inflammatory mediators, including TGFβ and cytokines, induce K_Ca3.1 and K_Ca2.3 expression [46,47,48], whereas pharmacological or genetic inhibition of these channels attenuates fibrotic and inflammatory responses [28,46]. Consistent with these findings, MF and its isomers inhibit K_Ca3.1 currents via PKA–dependent pathways and downregulate K_Ca3.1 and K_Ca2.3 expression via Epac–mediated mechanism [11,22]. Thus, suppression of K_Ca channel activity represents an additional mechanism by which MF and MF-S mitigate inflammatory and fibrotic responses.

Several limitations of this study should be acknowledged. First, although MF-S was shown to inhibit ADA activity and elevate adenosine levels, direct structural or crystallographic evidence for MF-S binding to ADA was not obtained. While the presence of phenyl rings in modafinil suggests a potential for interaction with ADA, the precise molecular determinants and inhibitory mode of action remain unclear and warrant further structural and biochemical investigation. Second, while pharmacological inhibitors were used to dissect signaling pathways, genetic approaches (e.g., ADA or CD73 knockdown) were not employed to definitively establish causality. Third, the therapeutic efficacy of MF-S has been confirmed across multiple in vivo models, including murine models of liver and lung fibrosis [10,11] as well as a rabbit ear model of hypertrophic scarring, supporting the robustness of its anti-fibrotic effects; however, differences in tissue context and disease mechanisms among these models should be considered when interpreting translational relevance. Finally, although modafinil and its R-isomer have been used clinically for the treatment of narcolepsy for decades with a well-established safety profile in humans, long-term safety, pharmacokinetics, and potential off-target effects of MF and its isomers in the context of chronic anti-fibrotic therapy have not been systematically evaluated. Elucidation of this newly identified mechanism may therefore enable rational drug repurposing of modafinil or its isomers for novel fibrotic indications, while underscoring the need for indication-specific safety and pharmacological assessments.

MF-S inhibits ADA activity, thereby increasing adenosine availability and promoting the subsequent activation of cAMP signaling in both fibroblasts and immune cells. In addition, MF-S upregulates the ectonucleotidases CD39 and CD73, which may further enhance adenosine generation and signaling. Augmented adenosine signaling contributes to the suppression of immune and inflammatory responses. Moreover, activation of the adenosine–cAMP axis exerts anti-fibrotic effects by attenuating Smad–dependent signaling while engaging Epac- and PKA–mediated pathways. Collectively, these mechanisms provide a plausible explanation for the anti-inflammatory and anti-fibrotic effects of MF-S observed in both in vitro and in vivo models [11]. Taken together, our findings support MF and MF-S as promising therapeutic candidates for the treatment of inflammatory, autoimmune, and fibrotic disorders.

The following supporting information can be found at: https://www.sciepublish.com/article/pii/842, Figure S1: MF-S inhibited Smad4 upregulation by TGFβ in human lung fibroblasts. Smad4 protein levels were measured by Western blot. Blots shown are representative of 9 experiments performed with 9 different cultures. Data are presented as the mean ± SEM values. ** p < 0.01.

During the preparation of this manuscript, the authors used ChatGPT (version 4.0) solely for grammar checking. After using this tool, the authors reviewed and edited the content as needed and assume full responsibility for the content of the published article.

Conceptualization, S.C., K.-C.K. and S.-H.S.; methodology, S.C., J.-A.K. and S.-H.S.; formal analysis, S.C., J.-A.K. and S.-H.S.; writing—original draft preparation, S.C. and K.-C.K.; writing—review and editing, S.C. and S.-H.S.; supervision, S.C. and S.-H.S.; funding acquisition, S.C., K.-C.K. and S.-H.S. All authors have read and agreed to the published version of the manuscript.

The study was conducted according to the guidelines of the Declaration of Helsinki, and approved by the Institutional Review Board (or Ethics Committee) of Seoul National University Bundang Hospital, Gyeonggi-do, Republic of Korea (BA-2108-326-008-02, issued at 27 August 2021).

Not applicable.

The original contributions presented in this study are included in the article. Further inquiries can be directed to the corresponding author.

This work was supported by the National Research Foundation of Korea (NRF) grant funded by the Korea government (MSIT) (NRF-2022R1A2C1092484 and NRF-2022R1A2C1007823), Republic of Korea and intramural research promotion grants from Ewha Womans University, School of Medicine.

The authors declare that they have no known competing financial interests or personal relationships that could have appeared to influence the work reported in this paper.