SIRT1’s role in skin cell aging and energy problems caused by long-term cadmium exposure

Analytical-grade Cadmium chloride (Cd), nicotinamide adenine dinucleotide (NAD⁺), and other analytical-grade chemicals were supplied by Sigma-Aldrich (St. Louis, MO, USA). For protein quantification, primary antibodies against β-actin, acetylated P53 (Lys382), acetylated SOD2 (Lys122), and SIRT1 were sourced from the vendors specified in Table 1. Secondary horseradish peroxidase (HRP)-conjugated antibodies were supplied by Abcam (Cambridge, UK).

Adult female Sprague–Dawley rats, weighing 200–220 g (8 weeks of age), were maintained in a controlled environment with ad libitum access to a standard diet and water. Ethical clearance for all experimental protocols was granted by the Institutional Animal Care and Use Committee of Yangzhou University (Permit No. SYXK [Su] 2022–0044). The animals were allocated into either a control group or a Cd-treatment cohort through a randomized process (n = 6/group). To induce SIRT1 overexpression, rats in the SIRT1-OVE group received a total volume of 100 μL of AAV-r-SIRT1 (1 × 10¹² VG/mL) via multi-point intradermal injections into the abdominal skin at the beginning of the third month of the study. Successful overexpression of SIRT1 was validated 1 month post-injection (at the end of the fourth month) via Western blot analysis. Following this confirmation, the Cd-exposed group was allowed free access to drinking water containing Cd (50 mg/L) for 6 months, while the control group received normal drinking water. The AAV-ove-SIRT1 viruses were obtained from Hanheng Biotechnology Co., Ltd.

The primary rat skin fibroblasts were derived from Pregnant Sprague–Dawley (SD) rats at 18 days gestation. The dorsal and abdominal skin of the fetuses was dissected and minced into small pieces approximately 1 mm × 1 mm in size and transferred into a culture flask. The resulting cell suspension was collected after 2 days and cultured under conditions of 37 °C, 5% CO2, in Dulbecco’s Modified Eagle Medium (DMEM) supplemented with 10% fetal bovine serum (FBS). For the dose–response experiments, primary fibroblasts were seeded in 6-well plates and exposed to a gradient of Cd concentrations (0, 0.2, 0.5, and 1 μM) for 48 h.

Due to the inherent limitations of primary rat skin fibroblasts, such as their low transfection efficiency and limited stability during long-term genetic manipulation, the C3H/10 T1/2, Clone 8 (C3H) cell line was selected for the establishment of the SIRT1-overexpression model. C3H cells are a murine embryonic fibroblast cell line derived from C3H mouse embryos, widely utilized as a model for studying mesenchymal differentiation, toxicological responses, and molecular mechanisms of gene regulation due to their stable growth characteristics and multipotent differentiation potential (23, 24). These cells were cultured in DMEM supplemented with 10% FBS, 100 U/mL penicillin, and 100 μg/mL streptomycin at 37 °C in a humidified atmosphere containing 5% CO₂. For Cd exposure experiments, cells were treated with Cd for 48 h. To determine the optimal concentration for the rescue studies, C3H cells were initially treated with a gradient of Cd concentrations (0, 2, 4, and 6 μM) for 48 h. Based on the assessment of cell viability and senescence markers, 6 μM was identified as the optimal concentration for subsequent SIRT1-overexpression experiments.

The C3H/10T1/2, Clone 8 cell line utilized in this study was supplied by Procell Life Science & Technology Co., Ltd. The construction of lentivirus-mediated SIRT1 shRNA was carried out by Shanghai GeneChem. In various experiments, after transduction with lentivirus-mediated SIRT1 shRNA, cells were treated with 6 μM CdCl2 for 48 h, using scramble shRNA as the control.

Abdominal skin tissues (~0.2 g) were collected, rinsed with distilled water to remove surface contaminants and residual blood, blotted dry, and weighed to obtain wet weight. Samples were digested in a mixed acid solution containing 5 mL nitric acid (HNO₃) and 1 mL perchloric acid (HClO₄). All digestion procedures were performed in a chemical fume hood under strict safety protocols. Predigestion was carried out on a heating plate at 120 °C until the solution became nearly clear, followed by gradual heating to 180 °C until complete digestion was achieved and a colorless, transparent solution was obtained. After cooling to room temperature, the digested samples were diluted to a final volume of 25 mL with deionized water.

Blank solutions were prepared using the same procedure. Cadmium standard solutions were prepared from a certified reference standard (GBW(E)080193) to generate calibration curves. After filtration, Cd concentrations in the samples were measured using a flame atomic absorption spectrophotometer (PinAAcle 900F, PerkinElmer, USA). Calibration curves demonstrated excellent linearity (R² > 0.99). Method accuracy was verified using spike recovery tests and standard reference materials. Each sample was analyzed in technical triplicate, and the average value was used for calculation. Cd content was expressed as μg/g wet weight.

For the ultrastructural examination of fibroblasts and skin tissues after CdCl₂ treatment, both were fixed using a consistent method. The images of these samples were then acquired with a HITACHI HT7800 transmission electron microscope.

Assessment of SA-β-gal activity was performed utilizing a specific senescence detection kit (Cell Signaling Technology), strictly adhering to the instructions provided by the manufacturer. In brief, cellular specimens were immobilized using 0.5% glutaraldehyde for a 15-min duration at ambient temperature. Following a PBS rinse, the cells were subjected to overnight incubation with the chromogenic substrate at 37 °C. The presence of senescent cells, identified by their distinct blue pigmentation, was quantified across five stochastically selected areas per sample utilizing bright-field microscopy.

Intracellular ROS levels were measured using the fluorescent probe 2′,7′-dichlorofluorescin diacetate (DCFH-DA; Sigma-Aldrich). Cells were exposed to 10 μM DCFH-DA (Sigma-Aldrich) for 30 min at 37 °C in a light-shielded environment to label endogenous ROS. After removing excess probe with PBS washes, the fluorescent emission was monitored using an Olympus microscope. Quantitative analysis of the fluorescence intensity was subsequently performed through ImageJ software to assess redox status.

The Total Antioxidant Capacity (T-AOC) was assessed by homogenizing cells, obtaining supernatant through centrifugation, and mixing it with a T-AOC reagent (Beyotime, Shanghai, China). Specimens were maintained at approximately 25 °C for 6 min to facilitate the reaction. Absorbance was measured at 405 nm by microplate reader (BioTek Synergy HTX). T-AOC levels were then determined by comparison with a standard curve (T-AOC content / Protein mass).

Cells and skin tissue were homogenated in PBS (PH7.4), then centrifugation at 5000 g for 5 min at 4 °C. Supernatant was used to measure 8-Hydroxy-desoxyguanosine (8-OHdG) by ELISA kits (Mibo China). The optical density of each well was measured at 450 nm.

According to the protocol provided by Beyotime (#S0175), the quantification of total NAD involved an initial sample preparation where 1 × 10⁶ cells were spun down at 1000 g (4 °C) for 5 min. The resulting pellet was resuspended in 200 μL of ice-cold NAD+/NADH extraction solution after discarding the culture medium. Concurrently, 20 mg of skin tissue fragments were prepared in 400 μL of extraction buffer. Both cell and tissue lysates were then vortexed and cleared by centrifugation at 12,000 g for 10 min at 4 °C. The cleared supernatant was subsequently subjected to colorimetric analysis at 450 nm using a 96-well plate reader.

Mitochondrial membrane potential was determined via the JC-1 Mitochondrial Membrane Potential Assay Kit (Beyotime, China). Upon completing the 37 °C incubation (20 min) with the staining solution, specimens were cleansed and subjected to fluorescent imaging. The ΔΨm status was subsequently derived from the fluorescence intensity ratio of red J-aggregates to green J-monomers.

Cellular and tissue proteins were isolated utilizing RIPA lysis buffer supplemented with a cocktail of phosphatase and protease inhibitors (Beyotime). After quantifying protein concentrations via a BCA assay (Thermo Fisher Scientific), 30 μg of each protein sample was resolved using SDS-PAGE and subsequently electroblotted onto PVDF membranes (Millipore). The membranes were then subjected to a 1-h blocking phase in 5% skim milk at ambient temperature, followed by an overnight primary antibody incubation at 4 °C. Post-washing, HRP-linked secondary antibodies were applied for 1 h. Finally, immunoreactive bands were developed using an ECL chemiluminescence system (Bio-Rad) and the signal intensities were analyzed through ImageJ.

Primary rat dermal fibroblasts were inoculated into E-Plates at a seeding density of 5 × 10³ cells/well. The assembly was then integrated into the Real-Time Cellular Analysis (RTCA) platform (Roche, Mannheim, Germany) within a CO₂ incubator (37 °C, 5%). The system automatically logged the Cell Index (CI)— a composite parameter of cell adhesion, morphology, and population—at 15-min frequencies. Upon reaching the logarithmic growth stage, the cultures were challenged with varying Cd concentrations. For comparative accuracy, the CI was normalized to a baseline of 1.0 at the onset of treatment, with subsequent proliferation kinetics tracked continuously for 48 h following Cd treatment.

The primary rat skin fibroblasts were immobilized in 70% ethanol for a 12-h duration. Subsequent to fixation, the cells were subjected to a 15-min staining process with propidium iodide (PI)/RNase buffer (Beyotime, C1052) under light-shielded conditions. The fluorescent profiles were then acquired utilizing a flow cytometer (LSRFortessa, BD Biosciences, USA).

The primary rat skin fibroblasts were immobilized in 4% paraformaldehyde, followed by permeabilization utilizing 0.1% Triton X-100. The specimens were then subjected to a 1-h incubation with the TUNEL reaction cocktail (Beyotime, China) at 37 °C in a light-shielded environment. Finally, apoptotic signals were captured using a Nikon fluorescence microscope.

Quantitative results are expressed as the mean ± standard error of the mean (SEM), derived from a minimum of three separate replicates. GraphPad Prism 8.0 (GraphPad Software, San Diego, CA, USA) was employed to conduct all statistical evaluations. To determine differences between two specific groups, the unpaired Student’s t-test was utilized. For assessments involving more than two cohorts, one-way analysis of variance (ANOVA) followed by Tukey’s multiple comparison test was applied. Statistical significance was predefined as a p-value less than 0.05.

To investigate the cellular effects of Cd on skin aging, we established an in vitro model using primary rat dermal fibroblasts. Cells were exposed to a gradient of Cd concentrations (0–1 μM) for 48 h. Cd treatment resulted in a dose-dependent decline in cell proliferation, with a significant reduction observed at 1 μM (Figure 1A), indicating substantial cytotoxic stress at this threshold. Morphologically, Cd-exposed fibroblasts exhibited decreased adhesion and spreading ability, along with an overall reduction in cell viability at higher Cd concentrations, reflecting impaired cellular attachment and survival under Cd-induced stress.

We next examined cellular senescence by analyzing the expression of classical senescence-associated markers. Western blotting revealed that exposure to Cd markedly upregulated the protein levels of P53, P21CIP1, and P16INK4A, key regulators of cell cycle arrest and senescence (Figure 1B) (25, 26). Consistent with the aforementioned observations, SA-β-gal histochemical staining revealed a marked elevation in the density of blue-pigmented senescent cells within the Cd-exposed cohorts relative to the control group. It is worth noting that while elevated Cd concentrations led to a reduction in total cell density and viability, the percentage of SA-β-gal-positive individuals among the surviving cells exhibited a substantial increase (Figure 1C) (27). Supporting these findings, flow cytometric profiling indicated that Cd exposure triggered a prominent G1/G0 phase arrest (Figure 1D), a hallmark characteristic of the senescence-associated growth cessation phenotype.

Collectively, these results establish that Cd exposure induces a robust senescent phenotype in dermal fibroblasts, characterized by impaired proliferation, upregulation of senescence markers, and cell cycle arrest. This model effectively recapitulates key molecular features of heavy metal–induced dermal aging and provides a foundation for further mechanistic investigation.

To elucidate the mechanisms underlying Cd-induced cellular senescence, we examined markers of oxidative stress and genotoxic damage. Fluorescent imaging using DCFH-DA staining demonstrated a significant elevation of intracellular reactive oxygen species (ROS) levels following Cd exposure, indicating heightened oxidative stress in treated fibroblasts (Figure 2A). This oxidative derangement was further substantiated by a prominent decline in total antioxidant capacity (AOC), alongside a simultaneous elevation of 8-hydroxy-2′-deoxyguanosine (8-OHdG), which serves as a classic indicator of oxidative-mediated DNA lesions (Figures 2B,C). Western blot analysis revealed upregulation of key DNA damage response proteins, including ATM and γH2AX, in Cd-treated cells (Figure 2D). Direct evidence of apoptosis and DNA damage was provided by TUNEL analysis, where Cd-treated fibroblasts exhibited pronounced nuclear fragmentation and apoptotic morphology (Figure 2E).

Taken together, these results demonstrate that Cd induces profound oxidative stress and DNA damage in dermal fibroblasts, contributing to cellular dysfunction and potentially promoting senescence.

To further investigate the intracellular consequences of Cd exposure, we examined mitochondrial ultrastructure and function in dermal fibroblasts. Ultrastructural analysis via transmission electron microscopy (TEM) pronounced mitochondrial abnormalities, including swelling, cristae disruption, and loss of matrix density, indicative of structural mitochondrial damage (Figure 3A). Functional impairment was corroborated by JC-10 staining, which demonstrated a significant reduction in ΔΨm in Cd-treated cells compared to controls (Figure 3B), suggesting mitochondrial depolarization and dysfunction. In parallel, ELISA quantification of cellular NAD⁺ levels revealed a marked depletion following Cd exposure (Figure 3C), implicating energetic stress and potential inhibition of NAD⁺-dependent enzymes.

The enzymatic status of SIRT1 and the integrity of its downstream signaling pathways were investigated, predicated on its biological identity as an NAD⁺-reliant protein deacetylase. Western blot analysis showed a significant decrease in SIRT1 protein expression, along with hyperacetylation of its known substrates P53 (at Lys382) and SOD2 (at Lys122), consistent with impaired SIRT1 enzymatic function (Figures 3D,E). Collectively, these findings suggest that Cd exposure disrupts mitochondrial structure and function, depletes cellular NAD⁺ levels, and suppresses SIRT1-mediated protective signaling, thereby amplifying oxidative and senescence-related cellular damage.

Notably, our initial experiments revealed that the C3H immortalized cell line exhibited a significantly higher tolerance to Cd toxicity compared to primary rat fibroblasts. To account for this difference in cellular sensitivity and to ensure a reproducible senescence model, we expanded the testing range for C3H cells. Specifically, treatment with increasing concentrations of Cd (0, 2, 4, 6 μM) progressively elevated the expression of senescence-associated proteins and increased SA-β-gal positivity, with 6 μM showing the most pronounced effects (Supplementary Figures S1A,B). Based on our preliminary dose–response experiments, 6 μM Cd was selected as an optimal concentration for subsequent mechanistic studies, as it induced a robust senescent phenotype in C3H cells while maintaining sufficient cell viability for downstream analyses.

To determine whether SIRT1 plays a protective role against Cd-induced cellular stress, we overexpressed SIRT1 in C3H cells via lentiviral vector and subsequently exposed the cells to 6 μM Cd. Compared with negative control, SIRT1-overexpressing fibroblasts exhibited markedly reduced susceptibility to Cd-induced senescence and apoptosis. Specifically, Western blot analysis revealed decreased expression of senescence-associated proteins P53, P21CIP1, and P16INK4A in the SIRT1-overexpressing group (Figure 4A). Consistent with these molecular findings, SA-β-gal staining showed a significant reduction in senescent cell burden (Figure 4B), and flow cytometric analysis demonstrated a lower proportion of cells arrested in the G1/G0 phase (Figure 4C), indicating partial restoration of cell cycle progression. Furthermore, overexpression of SIRT1 significantly rescued intracellular NAD⁺ levels (Figure 4D), which are otherwise depleted by Cd treatment, supporting the functional activity of the NAD⁺-dependent deacetylase. TUNEL assays also confirmed a notable decrease in apoptotic nuclear fragmentation among SIRT1-overexpressing cells compared to Cd-treated controls (Figure 4E).

Taken together, these observations confirm that SIRT1 exerts a multifaceted protective effect against Cd-induced injury by quenching reactive species, upholding cell cycle homeostasis, and diminishing the apoptotic burden in primary fibroblasts.

To delve into the protective molecular network of SIRT1, we evaluated its impact on oxidative stress, mitochondrial function, and DNA integrity in Cd-exposed fibroblasts. Overexpression of SIRT1 significantly suppressed the accumulation of ROS, as demonstrated by reduced DCFH-DA fluorescence intensity (Figure 5A). This attenuation of oxidative stress was accompanied by normalization of T-AOC, suggesting restored redox homeostasis (Figure 5B). In addition, JC-10 staining revealed a substantial recovery of ΔΨm in SIRT1-overexpressing cells, indicating preserved mitochondrial function under Cd stress (Figure 5C).

At the genomic level, SIRT1 overexpression led to marked reductions in the expression of DNA damage response proteins, including γH2AX and ATM (Figure 5D). Levels of 8-OHdG, a sensitive marker of oxidative DNA damage, were also significantly decreased (Figure 5E). Western blot analysis further demonstrated that SIRT1 overexpression reversed Cd-induced hyperacetylation of SOD2 and P53 (Figure 5F), indicating restored SIRT1 enzymatic activity and downstream regulation. Collectively, these findings confirm that SIRT1 confers substantial protection against Cd-induced mitochondrial dysfunction and DNA damage by modulating redox balance, maintaining mitochondrial integrity, and promoting genomic stability.

To validate the protective effects of SIRT1 in vivo, we employed a chronic Cd exposure rat model combined with skin-targeted SIRT1 overexpression. Immunohistochemical staining confirmed successful upregulation of SIRT1 protein in the skin tissues of AAV-SIRT1–treated rats (Figure 6A). Atomic absorption spectroscopy revealed significantly lower Cd accumulation in the skin of the SIRT1-overexpressing group versus the group treated with Cd alone (Figure 6B), suggesting a potential enhancement of detoxification or metal efflux mechanisms. TEM further showed preserved mitochondrial morphology in SIRT1-overexpressing skin, with intact cristae and reduced swelling relative to Cd-treated controls (Figure 6C). Biochemical assays demonstrated that SIRT1 overexpression restored NAD⁺ content (Figure 6D) and significantly reduced 8-OHdG levels in skin tissue, indicating diminished oxidative DNA damage (Figure 6E). Consistent with a reduction in cellular senescence, Western blot results indicated that the protein abundance of P53, P21CIP1, and P16INK4A was lower (Figures 6F,G), confirming the mitigation of senescence-associated pathways.

These in vivo results reinforce the protective role of SIRT1 in mitigating Cd-induced dermal aging, highlighting its ability to preserve mitochondrial integrity, reduce oxidative stress, and suppress senescence pathways.