Senotherapeutic Potential of Araliadiol in Senescent Human Dermal Fibroblasts: An In Vitro Study Using Three Senescence Models

Etoposide, used as a senescence inducer, was obtained from MedChemExpress (#HY-13629; Monmouth Junction, NJ, USA), and H₂O₂ was acquired from Daejung Chemicals (#4104-4400; Siheung-si, Republic of Korea). Araliadiol was isolated from C. asiatica extracts as previously described [47,49]. The purified compound (>98% purity) was kindly supplied by ASK Company Co., Ltd. (Daegu, Republic of Korea) and dissolved in dimethyl sulfoxide (DMSO; #D2650; Sigma-Aldrich, St. Louis, MO, USA) for use in cell-based experiments. A 10 mg/mL stock solution was prepared in DMSO, aliquoted, and kept at -20 °C to ensure stability. Working solutions were freshly diluted in culture medium immediately before each experiment. ABT-737, used as a positive control, was purchased from Selleckchem (#S1002; Houston, TX, USA).

Human dermal fibroblasts (HDFs) were sourced from Promocell (#C-12302; Heidelberg, Germany) and grown in Dulbecco's modified Eagle's medium (DMEM; #LB001-05; Welgene, Gyeongsan-si, Republic of Korea) supplemented with 10% (v/v) fetal bovine serum (FBS; #35-015-CV; Corning, NY, USA). Following previously established methods [50,51], only cells between passages 10 and 15 were used for experiments. Cells were subcultured at approximately 80% confluence and maintained at 37 °C in a humidified incubator with 5% CO₂.

To evaluate the senotherapeutic potential of araliadiol, three stressors—etoposide, H₂O₂, and UVA—were used to induce senescence in HDFs. Non-senescent HDFs (≤passage 15) were dispensed into 6-well plates at 1 × 10⁴ cells/well and maintained for 24 h. Cells were then treated with etoposide (0, 0.2, 2, and 20 μM), H₂O₂ (0, 125, 250, and 500 μM), or irradiated with UVA (0, 5, 7, and 9 J/cm²) using Vilber BIO-LINK BLX Crosslinker (Vilber Lourmat, Collégien, France). Cell numbers were assessed on days 0, 1, 3, and 7 using 0.4% trypan blue (#15250061; Gibco, Grand Island, NY, USA), as previously described [52].

Cellular senescence is characterized by permanent cell cycle arrest in the absence of cell death [53]. Therefore, we determined the concentrations or irradiation doses at which cell proliferation was inhibited without inducing apoptosis. Based on trypan blue assays, senescence was reliably induced by treatment with 2 μM etoposide, 250 μM H₂O₂, or 7 J/cm² UVA, followed by culture for 7 days. These conditions were used in all subsequent experiments.

Senescence was evaluated using SA-β-gal staining, a widely recognized biomarker, following a modified protocol from a previous study [54]. Non-senescent and senescent HDFs were dispensed into 12-well plates at 5 × 10⁴ cells/well and maintained for 24 h. After removing the medium, the cells were washed once with Dulbecco's phosphate-buffered saline (DPBS; #LB001-02; Welgene), fixed with 2% formaldehyde (#F8775; Sigma-Aldrich) and 0.2% glutaraldehyde (#G5882; Sigma-Aldrich) for 5 min at room temperature, and washed twice with DPBS. Cells were then stained using Senescence β-Galactosidase Staining Kit (#9860; Cell Signaling Technology, Danvers, MA, USA) at 37 °C for 16 h. Images were captured under a bright-field microscope at 200× magnification, and the proportion of SA-β-gal–positive cells was determined based on the total number of cells.

Intracellular ROS levels were assessed using 2′,7′-dichlorofluorescein diacetate (H₂DCFDA; #D6883; Sigma-Aldrich; Merck KGaA), as described by Ng (2021) [55]. Briefly, non-senescent and senescent HDFs were dispensed into 96-well plates at 2.5 × 10³ cells/well and maintained for 24 h. After incubation, the cells were washed once with DPBS and incubated with 10 µM H₂DCFDA for 30 min at 37 °C. The intracellular ROS levels were quantified by detecting 2′,7′-dichlorofluorescein fluorescence with a microplate reader set to 485 nm excitation and 520 nm emission.

Cell viability was assessed using an ATP content assay, as previously described [49]. Briefly, non-senescent and senescent HDFs were dispensed into 96-well plates at 2.5 × 10³ cells/well and maintained for 24 h. Cells were then treated with araliadiol (0–10 μM) or ABT-737 (0–5 μM) and incubated for up to 48 h. After treatment, the medium was removed and the cells were washed once with DPBS. Wells were replenished with 100 μL DPBS, followed by 100 μL CellTiter-Glo^® 2.0 reagent (#G9242; Promega, Madison, WI, USA). Plates were agitated on an orbital shaker at 25 °C for 5 min to facilitate cell lysis, incubated for 10 min in the dark to stabilize luminescence, and read using Synergy HTX Multi-Mode Microplate Reader (Biotek, Winooski, VT, USA). Relative luminescence units (RLU) were recorded and used to calculate cell viability.

qRT-PCR was performed to determine the effects of araliadiol on gene expression in senescent HDFs. Non-senescent and senescent HDFs were dispensed into 100 mm dishes at 3 × 10⁵ cells/dish and maintained for 24 h, then treated with araliadiol (0–5 μM) for up to 48 h. Total RNA was isolated using RiboEx reagent (#301-001; Geneall Biotechnology, Seoul, Republic of Korea). Complementary DNA (cDNA) was then generated from 1 μg of RNA using oligo dT primers, 0.1 M DTT, 2.5 mM dNTPs, 5× First-Strand Buffer, and M-MLV reverse transcriptase (#28025021; Thermo Fisher Scientific, Waltham, MA, USA).

Quantitative PCR amplification of target genes (GAPDH, IL1β, IL6, IL8, CXCL1, CCL2, MMP-1, and MMP-3) was conducted using HOT FIREPol^® EvaGreen^® qPCR Mix Plus (#08-24-0000; SOLIS BIODYNE, Tartu, Estonia), gene-specific primers, cDNA templates, and nuclease-free water on a Real-Time PCR System (Thermo Fisher Scientific). Relative gene expression was determined by the 2^−ΔΔCT method, with GAPDH serving as the reference gene. Primer sequences are provided in Table 1.

To evaluate the senotherapeutic potential of araliadiol in senescent HDFs, protein expression was assessed by Western blot analysis. Non-senescent and senescent HDFs were dispensed into 100 mm dishes at a density of 3 × 10⁵ cells/dish and maintained for 24 h, followed by treatment with araliadiol (0–5 μM) for up to 48 h. Following treatment, cells were incubated in RIPA buffer with protease (#04693116001; Roche, Darmstadt, Germany) and phosphatase inhibitor cocktails (#4906845001; Roche) for 30 min. The lysates were centrifuged at 12,000 rpm for 15 min, after which the supernatants were collected. Protein quantification was performed using the Pierce BCA Protein Assay Kit (#23225; Thermo Fisher Scientific). Protein samples (≤20 μg) were separated on 10% Tris-glycine sodium dodecyl sulfate-polyacrylamide gel electrophoresis (SDS-PAGE) gels and then transferred to nitrocellulose membranes (#88018; Thermo Fisher Scientific). Blocking was performed for 1 h using TBS-T buffer (1.0 mM Tris base, pH 8.0, 1.5 mM NaCl, 0.1% Tween-20) containing 5% skim milk (#SKI400-500; BIOPURE, Seoul, Republic of Korea). Membranes were then exposed to primary antibodies overnight at 4 °C, washed three times with TBS-T (5 min each), and incubated with horseradish peroxidase–conjugated secondary antibodies for 2 h at room temperature. Protein bands were detected using the Pierce ECL Western Blotting Substrate (#32106; Thermo Fisher Scientific). Detailed information on the primary antibodies used is presented in Table 2. All primary antibodies were obtained from Abcam (Cambridge, UK), Cell Signaling Technology (CST; Danvers, MA, USA), Santa Cruz Biotechnology (Dallas, TX, USA), R&D systems (Minneapolis, MN, USA), or Invitrogen (Carlsbad, CA, USA).

Immunofluorescence staining was conducted as previously reported with slight changes [49]. Non-senescent and senescent HDFs were dispensed into 12-well plates at 2.5 × 10⁴ cells/well and maintained for 24 h, followed by treatment with araliadiol (0–5 μM) for up to 48 h. Cells were rinsed once with DPBS, fixed in 4% formaldehyde for 15 min, and then washed three additional times with DPBS (5 min each). Cells were treated with 0.5% Triton X-100 (#9002-03-1; BIOPURE) to permeabilize membranes and subsequently incubated in a blocking buffer composed of 10% normal goat serum (#005-000-121; Jackson ImmunoResearch, West Grove, PA, USA), 2% Tween-20 (#9005-64-5; BIOPURE), and 1% bovine serum albumin (#BSA025; Bovogen, Melbourne, VIC, Australia). After overnight incubation at 4 °C with primary antibodies (p-p65, MMP-3, and p-c-Jun), samples were washed and subsequently incubated for 1 h at 37 °C with Alexa Fluor^® 488 Goat anti-Rabbit IgG (#A-11008; Invitrogen; Thermo Fisher Scientific) or Alexa Fluor^® 568 Goat anti-Rabbit IgG (#A-11011; Invitrogen; Thermo Fisher Scientific). Fluorescent Images were obtained using a Zeiss Axiovert 200 M fluorescence microscope (Carl Zeiss AG, Oberkochen, Baden-Württemberg, Germany) with FITC, Rhodamine, or DAPI filters, corresponding to each fluorophore.

The extracellular procollagen type I was quantified using Procollagen Type I C-Peptide EIA Kit (TaKaRa, Kusatsu, Shiga, Japan). Briefly, non-senescent and senescent HDFs were dispensed into 12-well plates at 2.5 × 10⁴ cells/well and incubated for 24 h, followed by treatment with araliadiol (0–5 μM) for 48 h. Following treatment, supernatants were harvested by centrifugation at 12,000 rpm for 2 min, and procollagen type I levels were measured in accordance with the manufacturer's instructions.

Statistical analyses were based on a minimum of three independent experiments. Results are reported as the mean ± standard deviation (SD). Analyses were carried out using Prism version 8.0.1 (GraphPad Software, San Diego, CA, USA). Comparisons among groups were made using one-way analysis of variance (ANOVA) with Tukey's post hoc test. Significance was defined as p < 0.05 (* p < 0.05, ** p < 0.01, and *** p < 0.001).

Unlike aging in other tissues, skin aging is externally visible and clinically characterized by wrinkles and sagging [56]. Because these age-related skin changes profoundly impair the QoL [18,19], senescent dermal fibroblasts are frequently employed as primary cellular models for screening senotherapeutic agents [4,57]. In this study, we investigated the senotherapeutic potential of araliadiol by establishing three distinct senescence models of HDFs using etoposide, H₂O₂, and UVA as inducers.

As shown in Figure 1, HDFs underwent cell cycle arrest without any evidence of cell death under optimized exposure conditions. Cell numbers, assessed using trypan blue-based cell counting, remained stable for 0–7 days following treatment with 2 μM etoposide or 250 μM H₂O₂, whereas higher concentrations (20 μM etoposide, 500 μM H₂O₂) resulted in significant cell loss (Figure 1a,b). Similarly, UVA irradiation at 7 J/cm² maintained stable cell counts for 7 days, whereas higher exposure (9 J/cm²) induced cytotoxicity (Figure 1c). These results align with the observations of Childs et al. (2014), who demonstrated that moderate stress drives senescence, whereas excessive stress promotes apoptosis [58]. Based on these results, we established that 2 μM etoposide, 250 μM H₂O₂, and 7 J/cm² UVA effectively induce senescence in HDFs without cytotoxicity. For subsequent experiments, non-senescent fibroblasts were designated NSenDFs, whereas fibroblasts senescent by etoposide, H₂O₂, and UVA were designated DNA damage–induced senescent dermal fibroblasts (DISenDF), ROS-induced senescent dermal fibroblasts (RISenDF), and UVA-induced senescent dermal fibroblasts (UISenDF), respectively (Figure 1d).

Next, we examined whether the fibroblasts exhibited senescent phenotypes. SA-β-gal staining (Figure 2a) revealed that only 1.73% of NSenDF were positive, compared with 90.63%, 81.88%, and 90.91% of DISenDF, RISenDF, and UISenDF, respectively. As SA-β-gal is widely recognized as a reliable marker of cellular senescence [54], these results validate the successful establishment of senescent fibroblasts across all three models. Western blotting (Figure 2b) further showed strong upregulation of the cell cycle arrest markers p16 and p21 in all senescent fibroblasts, whereas NSenDFs displayed only weak expression. Intracellular ROS levels were markedly elevated in the senescent groups: 182.25% in DISenDF, 161.85% in RISenDF, and 218.95% in UISenDF, compared to NSenDF (Figure 2c).

To evaluate the responsiveness of our models to senolytic intervention, we tested ABT-737, a BCL-2/BCL-xL inhibitor previously reported by Kim et al. (2022), to selectively eliminate senescent cells [23]. Treatment with 2.5 μM ABT-737 did not affect NSenDF viability (97.42%) but significantly reduced viability in DISenDF, RISenDF, and UISenDF to 42.11%, 53.18%, and 44.93%, respectively. At 5 μM, ABT-737 caused only mild toxicity in NSenDF (89.64%), while dramatically reducing viability in senescent fibroblasts to 16.91%, 23.29%, and 18.37%, respectively (Figure 2d). Taken together, the results shown in Figure 1 and Figure 2 confirm that the three senescence models established in this study reliably induced fibroblast senescence without triggering apoptosis and responded predictably to a known senolytic agent. Thus, these models provide a robust and valid platform for evaluating the senotherapeutic potential of candidate compounds.

Therapies for aging are typically categorized as senolytic, which selectively killing senescent cells, or senomorphic, which suppress the deleterious phenotypes of senescent cells to reduce chronic inflammation and limit the spread of senescence [21,22]. The accumulation of senescent cells accelerates tissue aging by enhancing SASP secretion, thereby stimulating paracrine senescence and inflammation in adjacent cells [8] and impairing the immune-mediated clearance of senescent cells [4,7]. Thus, removing senescent cells or neutralizing their harmful secretory output is a central strategy for mitigating skin aging.

After establishing three senescent dermal fibroblast models (Figure 1 and Figure 2), we examined whether araliadiol, a polyacetylene compound, has senotherapeutic potential. First, we assessed its senolytic activity (Figure S1). As expected, the positive control ABT-737 (2.5 μM) did not affect the viability of NSenDF (109.02%) but selectively induced cell death in all three types of senescent fibroblasts. By contrast, araliadiol, even at the highest tested concentration (10 μM), failed to induce selective senescent cell death (Figure S1).

Notably, Thomas et al. (2023) reported that isofalcarintriol, a polyacetylene compound, prolongs the lifespan of Caenorhabditis elegans and alleviates frailty indices in aged mice [59]. Furthermore, recent studies have demonstrated the antioxidant and anti-inflammatory activities of araliadiol, a polyacetylene compound, suggesting its potential to suppress SASP factors [46,49]. Based on these observations, we investigated whether araliadiol exerts senomorphic effects.

As depicted in Figure 3a–c, the expression profile of representative SASP factors (IL1β, IL6, IL8, CXCL1, and CCL2) was markedly elevated in DISenDF, RISenDF, and UISenDF compared with NSenDF. Treatment with araliadiol (5 μM) significantly reduced SASP expression across all three senescent models, with IL1β and CXCL1 showing the most pronounced decreases. Specifically, IL1β expression increased 81.17-, 156.24-, and 162.85-fold in DISenDF, RISenDF, and UISenDF, respectively, but decreased to 55.39-, 109.79-, and 93.64-fold after araliadiol (5 μM) treatment. Similarly, CXCL1 expression increased 114.71-, 116.73-, and 139.45-fold in DISenDF, RISenDF, and UISenDF, respectively, but decreased to 64.77-, 79.42-, and 81.01-fold, respectively, after treatment.

Because NF-κB p65 is a major transcription factor regulating SASP expression, we next evaluated its protein levels. Western blotting (Figure 3d–f) revealed that total p65 levels were unchanged in senescent cells compared to NSenDF, whereas the phosphorylation of p65 at Ser536 (p-p65) was strongly increased. Treatment with araliadiol (5 μM) markedly reduced p-p65 levels without altering total p65. Immunofluorescence staining (Figure 3g–i) further confirmed this finding, showing stronger cytoplasmic and nuclear p-p65 signals in senescent fibroblasts than in NSenDF, which were substantially diminished upon araliadiol (5 μM) treatment. Collectively, these findings indicate that araliadiol does not exert senolytic activity in dermal fibroblasts, but instead exhibits potent senomorphic effects, suppressing SASP gene expression through inhibition of p65 phosphorylation.

MMPs are a family of proteases predominantly produced by dermal fibroblasts and critically involved in the breakdown of extracellular matrix (ECM) constituents, including collagen and elastin [60]. Among them, MMP-1 is a collagenase that primarily degrades type I and III fibrillar collagens, whereas MMP-3, classified as stromelysin, degrades nonfibrillar ECM components and activates MMP-1 [13,60]. Consequently, MMPs are considered the key drivers of skin aging because of their roles in disrupting structural proteins and impairing skin elasticity. Therefore, considerable research has focused on identifying anti-aging agents that inhibit MMP activity [61,62].

As araliadiol suppressed SASP expression in senescent fibroblasts (Figure 3), we examined its effects on MMP-1 and MMP-3, which are also recognized as SASP factors. As shown in Figure 4a–c, both MMP-1 and MMP-3 mRNA levels were markedly elevated in DISenDF, RISenDF, and UISenDF compared with NSenDF. Treatment with araliadiol (5 μM) significantly reduced expression across all three senescent models. For example, in DISenDF, MMP-1 expression decreased from 4.94- to 2.59-fold, and MMP-3 expression decreased from 53.48- to 29.86-fold. Similarly, in RISenDF, MMP-1 decreased from 5.03- to 3.43-fold and MMP-3 from 55.48- to 40.53-fold, whereas in UISenDF, MMP-1 decreased from 5.90- to 3.70-fold and MMP-3 from 70.38- to 39.85-fold.

The inhibitory effects of araliadiol were validated at the protein level. Western blotting (Figure 4d–f) confirmed that MMP-1 and MMP-3 protein levels, elevated in senescent fibroblasts, were significantly reduced following treatment with araliadiol (5 μM). Immunofluorescence staining of MMP-3 (Figure 4g–i) provided additional support, showing weak fluorescence in NSenDF, strong signals in all senescent fibroblast models, and markedly reduced fluorescence after araliadiol (5 μM) treatment.

The expression of MMP genes, including MMP-1 and MMP-3, is largely regulated by the transcriptional activity of AP-1 [63]. Given that araliadiol downregulated MMP expression at both the transcript and protein levels (Figure 4a–i), we assessed its effect on AP-1 activation. While araliadiol did not alter total c-Fos or c-Jun levels, it significantly reduced phosphorylated c-Fos (Ser374) and c-Jun (Ser63), both of which were markedly elevated in DISenDF, RISenDF, and UISenDF compared to NSenDF (Figure 5a–c). Immunofluorescence staining corroborated these results, showing enhanced nuclear localization of p-c-Jun in senescent fibroblasts, which was substantially diminished by araliadiol treatment (Figure 5d–f). Collectively, these findings indicate that araliadiol attenuates AP-1 activation and suppresses MMP-1 and MMP-3 expression in senescent dermal fibroblasts, thereby potentially protecting against ECM degradation and skin aging.

Collagen makes up roughly 70–80% of the skin's dry weight, with type I and type III fibrillar collagens representing the predominant forms [64]. Relative to young skin (18–29 years), skin in older adults (≥80 years) exhibits more than a 30% decline in total collagen content, contributing to visible aging features such as wrinkles and sagging [11]. Therefore, whether araliadiol can restore collagen levels in senescent fibroblasts is an important indicator of its senotherapeutic potential.

We first assessed intracellular collagen protein expression using Western blotting (Figure 6a–c). As expected, DISenDF, RISenDF, and UISenDF displayed markedly reduced levels of collagen types I and III compared to NSenDF. However, araliadiol treatment did not restore intracellular collagen expression. Considering our earlier findings (Figure 4) that araliadiol suppressed MMP-1 and MMP-3 expression, we next investigated whether it could enhance extracellular collagen levels without affecting intracellular collagen expression.

ELISA (Figure 6d–f) revealed that extracellular pro-collagen type I levels were significantly reduced in all three senescent models compared to NSenDFs. Importantly, treatment with araliadiol (0–5 μM) increased extracellular collagen content in a dose-dependent manner. In DISenDF, extracellular collagen levels increased from 52.53% in untreated cells to 70.88% following treatment with 5 μM araliadiol (Figure 6d). Similarly, in RISenDF and UISenDF, extracellular collagen levels, which were reduced to 61.82% and 55.08% in untreated cells, were restored to 72.14% and 70.22%, respectively, after treatment with 5 μM araliadiol. These results indicate that araliadiol does not influence intracellular collagen expression but significantly enhances extracellular collagen levels, most likely through inhibition of MMP activity.

Taken together, our findings have shown that in three distinct senescent dermal fibroblast models, araliadiol suppressed SASP expression, downregulated MMP activity, and increased extracellular collagen content. These combined effects highlight the senotherapeutic potential of araliadiol for mitigating skin aging.