Starfish-Derived Extracts Enhance Mitophagy and Suppress Senescence-Associated Markers in Human Dermal Fibroblasts

To evaluate the anti-aging effects of extracts derived from Asterias pectinifera (Ap) and Asterias amurensis (Aa) in human dermal fibroblasts (HDFs), we first established both non-senescent (nsHDF) and senescent (sHDF) cell models. Replicative senescence was induced through serial passaging, with senescence confirmed by SA-β-gal staining and mitochondrial-specific oxidative stress assessed via MitoSOX staining. Proliferative capacity was measured across passages 2–7 for nsHDF and beyond passage 20 for sHDF. As expected, sHDF exhibited a marked increase in SA-β-gal activity and significantly elevated mitochondrial reactive oxygen species (mtROS) levels compared to nsHDF (Figure 1A,B), validating the successful induction of cellular senescence.

Prior to assessing the anti-senescence effects of Ap and Aa, we evaluated their cytotoxicity in HDFs. Cells were treated with varying concentrations of Ap and Aa for 72 h, and viability was measured using the CCK-8 assay. While high concentrations (>100 μg/mL) reduced cell viability, doses below 50 μg/mL were non-cytotoxic in both nsHDF and sHDF (Figure 1C).

We then investigated whether Ap and Aa could regulate the senescence phenotypes in sHDF. Cells were treated with vehicle (Veh), Ap, Aa, or resveratrol—a polyphenolic antioxidant known for its anti-senescence properties and used here as a positive control [21,22]. Treatment with Ap, Aa or resveratrol resulted in a significant reduction in the proportion of SA-β-gal-positive cells by more than 20%, along with a noticeable decrease in staining intensity compared to vehicle-treated controls (Figure 1D). Consistent with this, the mRNA expression level of p21, a well-known senescence marker, was also reduced following treatment with Ap or Aa (Figure 1E). In addition, the expression levels of Bcl-2-associated X protein (Bax) and B-cell lymphoma 2 (Bcl-2), which have opposing roles in apoptosis regulation, were both significantly increased following Ap and Aa treatment (Figure 1F). Taken together, these findings demonstrate that Ap and Aa-derived extracts effectively attenuate senescence-associated phenotypes in HDFs.

To examine whether Ap and Aa exert anti-inflammatory effects through the modulation of SASP factors in sHDF, we first measured the mRNA expression levels of IL-6, IL-8 and MMP-1 (Figure 2A). Next, we comprehensively analyzed cytokine secretion profiles from sHDF following Ap- or Aa-derived extracts treatment, using the multiplex cytokine analysis Luminex and ELISA assays (Figure 2B,C). The results showed that Ap and Aa treatment significantly reduced the transcriptional and secretory levels of IL-6, IL-8, and MMP-1. Next, we aimed to investigate the effects of Ap and Aa treatment on extracellular signal-regulated kinase (ERK) signaling in HDFs. Following Ap and Aa treatment, the level of ERK phosphorylation was assessed and quantified using immunoblot analysis (Figure 2D). A decrease in ERK phosphorylation was observed in both nsHDF and sHDF treated with Ap or Aa. To investigate whether the observed anti-inflammatory effects were mediated through NF-κB–dependent inflammatory signaling, cells were transfected with an NF-κB luciferase reporter plasmid and subsequently treated with lipopolysaccharide (LPS), a known activator of NF-κB [23]. After LPS stimulation, cells were treated with Ap or Aa to assess whether the starfish extracts could modulate LPS-induced NF-κB transcriptional activity. Figure 2E demonstrates that LPS-induced increase in NF-κB transcriptional activity was markedly attenuated by Ap or Aa treatment. These findings suggest that Ap and Aa mitigate SASP-associated inflammation, thereby contributing to their overall anti-senescence activity.

Recent studies suggest that the accumulation of damaged mitochondria amplifies skin aging phenotypes and the mitophagy, a selective removal process, is crucial for maintaining skin cell homeostasis [24,25]. To investigate the role of starfish-derived extracts on mitophagy, we first quantified whether autophagosomes are formed upon starfish-derived extract treatment in sHDF. To directly visualize autophagosomes, transmission electron microscopy (TEM) was employed. As expected, bafilomycin A1 (BafA) treatment markedly increased the number of autophagosomes, and Figure 3A demonstrates a further accumulation of autophagosomes containing fragmented mitochondria in starfish-treated sHDF, suggesting enhanced autophagic activity. To further assess mitophagy activation, we employed g MitoTracker and LysoTracker co-staining to visualize mitochondrial-lysosomal structures indicative of mitophagic flux. Notably, Ap-, Aa-, and resveratrol-treated cells exhibited enhanced colocalization of lysosomal (red) puncta with mitochondria, compared to vehicle-treated controls, confirming the induction of mitophagy by starfish-derived extracts (Figure 3B). To further quantify mitophagy, we generated HDF cells stably expressing mitochondria-targeted Keima (mtKeima; HDF-mtKeima) to measure the level of mitophagy induction following Ap or Aa treatment. As a positive control for mitophagy activation, cells were treated with 25 μM carbonyl cyanide m-chlorophenyl hydrazone (CCCP) for 2 h, a well-established inducer of mitochondrial depolarization and PINK1/Parkin-mediated mitophagy. We confirmed an approximately two-fold increase in the level of mitophagy activation with Ap-, Aa-, or resveratrol–treated cells (Figure 3C). Similar results were obtained in Hela-mtKeima cells, indicating enhanced mitophagy (Figure 3D). Next, we analyzed mitophagy level in HDFs by measuring the degradation of cytochrome C oxidase subunit II (COX II), a mtDNA-encoded inner membrane protein, following replicative senescence. Figure 3E shows that Ap or Aa treatment led to COX II degradation in sHDF. These findings suggest that Ap and Aa can activate mitophagy in various cell types, highlighting their potential for broader biological applications.

Since PINK1 is a key upstream regulator of canonical mitophagy initiation, we next sought to determine whether the mitophagy-activating and anti-senescence effect of starfish-derived extracts were dependent on PINK1 signaling. We first aimed to quantify the activation of mitophagy induced by Ap and Aa in the PINK1 knockdown condition using the HDF-mtKeima cell system. As a result, compared to the siCtl group, PINK1 knockdown significantly reduced the mitophagy-activating effects of the starfish extracts. These findings indicate that PINK1 is a key factor in Ap- and Aa-induced mitophagy activation, suggesting that Ap and Aa promote mitophagy in a PINK1-dependent manner (Figure 4A). Therefore, it was necessary to investigate the impact of PINK1 on the anti-senescence effects of Ap and Aa. To this end, we analyzed the expression of senescence-related genes in PINK1-knockdown HDFs following Ap or Aa treatment using qRT-PCR. First, to verify the efficiency of PINK1 siRNA-mediated knockdown, we assessed PINK1 mRNA expression and confirmed that its expression was effectively suppressed. Next, we examined the mRNA expression of the skin aging markers p21 and MMP-1 and found that their levels were elevated by at least 2-fold and up to 5-fold in the siPINK1 group compared to the siCtl group (Figure 4B). We next investigated whether the absence of PINK1 also affects SASP profiles. Under the same experimental conditions, we performed Luminex assays and ELISA using the supernatants collected from these samples. Through the results of the Luminex and ELISA assays, we found that PINK1 knockdown abolished Ap- or Aa-mediated regulation of pro-inflammatory cytokines, leading instead to an increase in their secretion. Moreover, MMP-1 secretion was significantly elevated under PINK1-deficient conditions. Taken together, these findings demonstrate that anti-inflammatory effects of the starfish extracts are mediated through the mitophagy key regulator PINK1 (Figure 4C,D). In summary, we demonstrate that PINK1, a key regulator of mitophagy, plays a critical role in the anti-senescence effects of the starfish extracts.

Next, we wanted to evaluate the effect of starfish-derived extracts on the mitochondrial metabolism. Dynamin-related protein 1 (Drp1) is a central player in mitochondrial quality control, particularly through its role in mitochondrial fission. Drp1 is recruited from the cytosol to the outer mitochondrial membrane, where it assembles into spiral structures that constrict and divide mitochondria [26]. There was an increase in phosphorylation of Drp1, which is a key regulatory event that promotes mitochondrial fission, upon the treatment of starfish-derived extracts in HDFs (Figure 5A). Next we evaluated the intracellular antioxidant activity of starfish-treated HDFs in response to hydrogen peroxide (H₂O₂). Starfish-derived extracts treatment of cells alleviated the production of reactive oxygen species (ROS) and mitochondrial ROS production (Figure 5B,C). To determine whether Ap- or Aa-derived extracts lead to improved mitochondrial function, we measured their effects using Seahorse assay, particularly through the oxygen consumption rate (OCR), a key indicator of mitochondrial respiration. Starfish extracts increased basal respiration, as well as maximal respiratory capacity in sHDF (Figure 5D). Starfish-derived extracts' ability to modulate mitochondrial respiration, as shown by oxygen consumption rate, supports its potential as a therapeutic agent in oxidative stress-related diseases.

Asterias pectinifera (Ap) and Asterias amurensis (Aa) were collected from the coastal waters of Dangjin, Republic of Korea. Ap and Aa were washed thoroughly, frozen, and stored at −20 °C until further use. Each species was immersed in a 7% aqueous potassium hydroxide (KOH) solution at a 1:1 ratio (w/v) for 24 h at room temperature to remove non-collagen substances. The resulting materials were rinsed with distilled water until the pH reached 8.0. Ap and Aa ossicle extract was mixed with distilled water at a ratio of 1:10 (w/v). DL-tartaric acid and ascorbic acid were added to the mixture at final concentrations of 0.65% (w/v) each. The sample was subjected to ultrasonic treatment at 20% Ampl for 30 min using an ultrasonic processor (HD4400, Bandelin, Berlin, Germany). The solution was stirred at 4 °C, and its pH was adjusted to 7.0 during stirring. Subsequently, complex protease ZF101, (Angel Yeast Co., Ltd., Yichang, Hubei, China) was added at a concentration of 0.01% (w/v), and enzymatic hydrolysis was carried out by continuous stirring at 4 °C for 24 h. After removing the precipitation by centrifugation, the mixture was heated at 80 °C for 5 min to inactivate the enzyme. The resulting starfish-derived extracts were lyophilized using a freeze dryer (TFD8501, Ilsinbiobase, Dongducheon, Korea) for 72 h. The lyophilized powders were reconstituted in distilled water filtered through a 0.22 µm pore-size filter unit (Corning Inc., Corning, NY, USA, Cat. No. 431097) and used as samples in subsequent experiments.

To conduct a qualitative analysis of starfish extracts, samples were analyzed using a Vanquish UHPLC system (Thermo Fisher Scientific, Waltham, MA, USA) coupled with a heated electrospray ionization (H-ESI) source. Briefly, starfish extracts were centrifuged at 12,000× g for 10 min at 4 °C and filtered through a 0.22 µm PTFE syringe filter prior to analysis. Chromatographic separation was achieved on a Hypersil Gold C18 column (100 × 2.1 mm, 1.9 µm) maintained at 40 °C. The mobile phases consisted of solvent A (0.1% formic acid in water) and solvent B (0.1% formic acid in methanol), delivered at a flow rate of 0.3 mL/min under the following gradient: 0–3 min, 90% A; 3–13 min, 10–100% B; 13–17 min, 100% B; 17–17.5 min, 100–10% B; 17.5–20 min, 10% B. The injection volume was 5 µL. Mass spectrometric detection was performed with an Orbitrap analyzer at a resolution of 120,000 over an m/z range of 70–1000. Ionization was carried out in both positive (3.5 kV) and negative (2.5 kV) modes. Data-dependent MS/MS fragmentation was conducted using a normalized collision energy (NCE) of 30. Compound identification was performed by comparing accurate mass and MS/MS spectra with reference databases using Thermo Scientific Compound Discoverer 3.3 software.

HDF cells were cultured in RPMI-1640 (Corning Inc., Corning, NY, USA) supplemented with 10% fetal bovine serum (FBS) (GenDEPOT, Barker, TX, USA), 25 mM HEPES (GenDEPOT, Barker, TX, USA), 100 U/mL penicillin, and 100 µg/mL streptomycin (Gibco, Grand Island, NY, USA). Cells were maintained at 37 °C in a 5% CO₂ incubator and passaged every 3–4 days. A replicative senescence model has been introduced previously. HDF cells were cultured beyond passage 20 to induce replicative senescence, following a method similar to that previously described [38]. Senescence was quantified by SA-β-galactosidase staining kit (Cell Signaling Technology, Danvers, MA, USA; 9860) following the manufacturer's instructions.

HDF cells were seeded in 96-well plates. After 24 h, the medium was replaced, with or without Ap or Aa in various concentrations. To measure cell viability, at 72 h after treatment, the old medium was removed and cells were incubated with CCK8 (CK04-11, Dojindo, Kumamoto, Japan) solution for 1 h. The absorbance at 450 nm was measured using a Varioskan™ LUX multimode microplate reader (Thermo Scientific, Waltham, MA, USA).

To quantify the induction level of mitophagy, HDF and Hela stably expressing mt-Keima were generated (a kind gift of Prof. Jinho Yun, DongA University School of Medicine, Korea) and analyzed using an LSR Fortessa X-20 flow cytometer (BD Biosciences, San Jose, CA, USA) equipped with a 405 nm/561 nm laser and BV605/PE-CF594 detector, as described previously [34,39]. The percentage of cells undergoing mitophagy was determined by gating cells exhibiting a high ratio of emission at 561 nm/405 nm excitation.

To measure intracellular ROS, cells were seeded in a 96-well plate. After Veh, Ap, Aa, or resveratrol treatment with or without H₂O₂, cells were stained with 10 μM 2′,7′-dichlorofluorescein diacetate (DCF-DA; Sigma-Aldrich, St. Louis, MO, USA) in serum-free (SF) media. For mitochondrial-mediated ROS measurement, 1 μM MitoSOX™ Green (Invitrogen, Waltham, MA, USA) in SF media was incubated with cells for staining. After 30 min, the medium was removed, and the cells were washed with phosphate-buffered saline (PBS). Fluorescence was measured using a Varioskan™ LUX multimode microplate reader (Thermo Scientific) at 490 nm excitation/520 nm emission for DCF-DA, and 488 nm excitation/510 nm emission for MitoSOX Green. For additional analysis of mitochondrial ROS, MitoSOX Green-stained cells were also analyzed by flow cytometry using an LSR Fortessa X-20 equipped with a 488 nm laser and FITC detector.

HDF cells were transfected with pNF-κB-Luc plasmid using Lipofectamine 2000 (Thermo Fisher Scientific) according to the manufacturer's protocol. After 24 h, cells were treated with vehicle, Ap (10 μg/mL), Aa (10 μg/mL), or resveratrol (10 μg/mL), either in the presence or absence of lipopolysaccharide (LPS; 100 ng/mL) co-treatment to stimulate NF-κB activity. NF-κB transcriptional activity was measured 24 h after treatment using a Dual-Glo luciferase assay kit (Promega, Madison, WI, USA).

Total RNA was isolated using a Direct-zol RNA Miniprep Kit (Zymo Research, Irvine, CA, USA; R2050) following the manufacturer's instructions. RNA quality was assessed using a NanoDrop ND-2000c (Thermo Fisher Scientific). cDNA synthesis was performed using the High-Capacity cDNA Reverse Transcription Kit (Applied Biosystems, Waltham, MA, USA; 4368814), and relative mRNA levels were analyzed on the QuantStudio 1 Real-Time PCR System (Applied Biosystems) with the Power SYBR Green PCR Master Mix (Applied Biosystems; 4368706). The GADPH gene served as housekeeping control. Primer sequences were previously reported [40].

IL-6, IL-8 and MMP-1 were measured from cell supernatants using R&D ELISA kits (R&D Systems, Minneapolis, MN, USA). ELISA was performed using the manufacturer's instructions, and absorbance at 450 nm was measured using a microplate spectrophotometer. For multiple screening, a magnetic Luminex screening assay with a Human Premixed Multi-Analyte Kit (R&D Systems) was used.

HDF cells were transfected with control siRNA or PINK1-specific siRNA (Bioneer, Daejeon, Korea) using Lipofectamine RNAiMAX (13778100, Invitrogen) according to the manufacturer's instructions.

Protein samples were separated by 8–15% SDS-PAGE and transferred to polyvinylidene difluoride membranes (PVDF; IPVH00010, Millipore, Burlington, MA, USA). Membranes were blocked for 1 h at room temperature in 5% skim milk prepared in Tris-buffered saline containing 0.1% Tween-20 (TBS/Tw; T2008, Biosesang, Youngin, Korea) and then incubated RT 1 h or overnight at 4 °C with primary antibodies. The following antibodies were used: phospho-p44/42 MAPK (Erk1/2) (Thr202/Tyr204) (9101, Cell Signaling Technology), p44/42 MAPK (Erk1/2) (137F5) (4695, Cell Signaling Technology), phospho-DRP1 (Ser616) (3455, Cell Signaling Technology), DRP1 (D8H5) (5391, Cell Signaling Technology), COX II (ab198286, Abcam, Cambridge, UK), and β-actin (AM1021B, Abcepta, San Diego, CA, USA). After washing, membranes were incubated with HRP-conjugated secondary antibodies (anti-mouse or anti-rabbit) and visualized using the Fusion Solo Imaging System (Vilber Lourmat STE, Collegien, France). Band intensities were quantified using ImageJ software (version 1.53e) (National Institutes of Health, Bethesda, MD, USA).

HDF cells were treated with vehicle, Ap, Aa, or resveratrol for 24 h, then incubated in serum-free medium containing MitoTracker Green (Thermo Fisher Scientific) and LysoTracker Deep Red for 30 min. Mitochondrial and lysosomal signals were subsequently visualized using a confocal microscope (LSM900, Carl Zeiss, Oberkochen, Germany).

HDF cells were treated with vehicle, Ap, Aa, or resveratrol. Cells were then fixed in 2.5% glutaraldehyde for 12 h, followed by post-fixation in reduced osmium tetroxide for 1.5 h at 4 °C. After three washes, samples were dehydrated through a graded ethanol series and embedded in Epon resin. Ultrathin sections were prepared, stained with uranyl acetate and lead nitrate, and examined using a Hitachi H-7000 transmission electron microscope(Hitachi High-Tech Corporation, Tokyo, Japan) to visualize autophagosome formation and assess mitochondrial ultrastructure.

Cells were plated in mini-plates (Agilent, Santa Clara, CA, USA, Cat. No. 103022-100) at 10,000 cells/well and placed in a non-CO₂ incubator for 1 h prior to oxygen consumption rate (OCR) measurement. The Seahorse XFp Cell Mito Stress Test Kit (Agilent, Cat. No. 103010-100) was employed following the manufacturer's protocol. In brief, basal OCR was recorded first, followed by sequential injections of 1 μM oligomycin, 2 μM fluoro-carbonyl cyanide phenylhydrazone (FCCP), and a mixture of 1 μM rotenone with 1 μM antimycin A at the designated assay intervals.

All data are expressed as the mean ± standard deviation (SD) from a minimum of three independent experiments. Statistical significance between treatment groups was assessed using either the Mann–Whitney U test, two-tailed unpaired Student's t-test, or two-way ANOVA, as appropriate. Significance levels are indicated as follows: * p < 0.05; ** p < 0.01; *** p < 0.001; ns, not significant. Graph generation and statistical analyses were conducted using GraphPad Prism 9 software (GraphPad Software, San Diego, CA, USA).