Effects of Chlorogenic Acid on Cellular Senescence in an In Vitro Model of 3T3-L1 Murine Adipocytes

Adipose tissue has long been considered an energy storage and endocrine organ. However, adipose tissue aging, associated with the accumulation of senescent cells within adipose tissue, leads to a progressive decline in tissue functionality, with significant implications for overall health and longevity.

In our experimental model, to investigate the induction of cellular senescence and to assess the potential protective effects of CGA, we evaluated two widely validated adipocyte markers of senescence: SA-β-Gal activity and Lamin B1 expression [27,28,29]. Our results revealed that, following exposure to H₂O₂, 3T3-L1 adipocytes displayed a marked increase in SA-β-Gal staining. Interestingly, CGA treatment significantly reduced SA-β-Gal staining in a dose-dependent way. In particular, at the highest tested concentration (20 µM), β-galactosidase activity was comparable to that observed in untreated controls, thereby supporting the protective effects of CGA against senescence in murine adipocytes (Figure 1A).

At the same time, we assessed the expression of Lamin B1, a key component of the nuclear envelope, whose downregulation reflects chromatin remodeling and transcriptional changes. Lamin B1 loss is recognized as a marker of senescence in response to all canonical signals of senescence [30]. In our experimental conditions, the data showed the marked downregulation of Lamin B1 levels following exposure to H₂O₂ compared to untreated controls, further confirming the establishment of a senescent state. Notably, CGA treatment dose-dependently restored its expression starting from 5 µM, suggesting a protective role in maintaining nuclear integrity (Figure 1B). For both parameters examined, no significant changes were found in 3T3-L1 cells exposed to CGA alone compared to control cells (see Figure S1 of Supplementary Materials).

It is well established that one of the features of senescent cells is the upregulation of cyclin-dependent kinase inhibitors (CDKIs), which determine irreversible cell cycle arrest [31]. In particular, the p53/p21 axis plays a pivotal role since, upon DNA damage, the tumor suppressor protein p53 acts as a transcription factor for several stress-responsive genes, including Cdkn1a, which encodes for the CDK inhibitor p21. Elevated p21 expression levels lead, in fact, to G1/S arrest, considered a hallmark for the onset and maintenance of the senescent phenotype [32,33]. Since p53 activation is regulated by post-translational modifications regulating its stability and transcriptional activity, the phosphorylated form (Phospho-p53) was analyzed as a specific marker of its functional activation under oxidative stress conditions. In contrast, p21 expression mainly reflects transcriptional induction downstream of activated p53.

Based on these considerations, to further explore the mechanisms underlying CGA's protective effects, we investigated its impacts on key regulators of the cell cycle checkpoint pathway associated with senescence. Our data demonstrated that H₂O₂ exposure led to a marked increase in the levels of Phospho-p53 (Figure 2A), along with its downstream effector p21 (Figure 2B), confirming the onset of senescence-induced growth arrest.

Interestingly, CGA treatment attenuated the activation of these senescence-related pathways in a dose-dependent way. Even at the lowest tested concentration, CGA was, in fact, able to significantly reduce Phospho-p53 and p21 expression.

Additionally, mitogen-activated protein kinase (MAPK) family members, particularly p38 MAPK and ERK1/2 (pERK, phosphorylated extracellular signal-regulated kinases 1/2), further contribute to senescence establishment by inhibiting cell proliferation and differentiation and by promoting the activation of SASP [34]. In fact, the growth arrest of senescent cells modulated by CDKIs is under the direct control of MAPKs, particularly ERK1/2 and p38 [34]. In our experimental conditions, elevated levels of phosphorylated p38 (Phospho-p38; Figure 2A) and pERK1/2 (Figure 2A) were detected in H₂O₂-exposed adipocytes, underscoring their involvement in cell cycle arrest. Moreover, in this case, CGA treatment was able to reduce Ph-p38 and pERK1/2 levels in a dose-dependent way, starting from 5 µM. CGA treatment alone did not alter the analyzed parameters with respect to control cells (see Figure S2 of Supplementary Materials). These findings therefore suggest that CGA may alleviate senescence by modulating upstream sensors and downstream effectors involved in checkpoint regulation.

Cellular senescence is characterized by cell proliferation arrest and resistance to apoptosis [35]. The inability of senescent cells to re-enter the cell cycle is indeed a key feature of this state, resulting in reduced tissue regeneration and homeostasis and in the formation of dysfunctional cells in aged or stressed tissues [36]. Our data confirmed that, in our experimental model, H₂O₂ significantly reduced the number of mature 3T3-L1 adipocytes (Figure 3A) and altered the expression of key apoptotic regulators. More specifically, H₂O₂ exposure resulted in elevated levels of the anti-apoptotic protein Bcl-2 (Figure 3B), accompanied by the marked downregulation of the pro-apoptotic protein BAX (Figure 3B). This shift toward an enhanced anti-apoptotic profile supports the survival of senescent cells and reflects a cellular environment favoring resistance to apoptosis. Notably, CGA treatment was able to reverse these alterations in a dose-dependent way. In fact, CGA treatment restored the proliferative capacity, as indicated by the increased cell number, and modulated the Bcl-2/BAX ratio to values comparable to those observed in controls. Meanwhile, CGA alone did not induce any detectable changes compared to control cells (see Figure S3 of Supplementary Materials). These findings underscore the ability of CGA to counteract impaired cellular proliferation and resistance to apoptosis, thereby reinforcing its therapeutic potential as a senomorphic agent in preserving adipose tissue homeostasis.

An important feature of SASP is the accumulation of ROS [37], which not only trigger oxidative damage but also act as activators of the inflammatory pathways defining the SASP profile. In our experimental model, H₂O₂ exposure significantly increased the intracellular ROS levels, indicating the onset of an oxidative stress state. Interestingly, CGA treatment was able to reduce them in a dose-dependent way, with values, for the highest tested concentration, that were even lower than those observed in control cells (Figure 4A).

Given the central role of ROS in promoting inflammatory signaling, we next assessed the expression of NF-κB, the master regulator of inflammation activated in response to oxidative stress. In fact, NF-κB acts as a key modulator able to induce and sustain the expression of multiple SASP factors, including pro-inflammatory cytokines and matrix remodeling enzymes [36]. The results obtained showed that H₂O₂ treatment markedly increased NF-κB expression, whereas CGA significantly reduced its levels in a dose-dependent way (Figure 4B).

Since NF-κB is a key modulator of the inflammatory SASP response, we then examined its principal downstream effectors.

It is widely known that SASP activation leads to an increase in the release of pro-inflammatory cytokines and matrix remodeling enzymes, which contribute to tissue dysfunction and chronic inflammation [38]. In this context, in our study, to further investigate the potential activity of CGA to modulate the senescence phenotype, we assessed the expression levels of the pro-inflammatory cytokines interleukin-6 (IL-6), interleukin-8 (IL-8), and tumor necrosis factor-α (TNF-α), which are well-established SASP components and important contributors to adipose tissue dysfunction. These cytokines play a central role in the onset of the low-grade chronic inflammatory state typical of senescent adipose tissue, where they promote metabolic dysregulation, immune cell infiltration, and extracellular matrix (ECM) remodeling [36]. IL-6 and TNF-α, in particular, are closely linked to impaired insulin signaling and adipocyte dysfunction. Meanwhile, IL-8 has an important role as a chemoattractant for immune cells and as a promoter of senescence propagation [39,40]. As expected, H₂O₂ exposure induced the strong upregulation of all three cytokines, confirming the establishment of a pro-inflammatory SASP phenotype. Notably, CGA treatment reduced IL-6, IL-8, and TNF-α expression in a dose-dependent way, significantly counteracting the inflammatory response triggered by oxidative stress (Figure 4C). To further characterize the SASP profile, we also examined the expression of matrix metalloproteinase 3 (MMP3), an important enzyme modulated by NF-κB, involved in the degradation of ECM components and considered not only a structural modulator but also a key SASP effector that amplifies inflammatory signaling and contributes to tissue remodeling in aging- and senescence-related pathologies [41,42].

Our data demonstrated that H₂O₂ exposure resulted in the marked upregulation of MMP-3 (Figure 4D) gene expression, reinforcing the onset of a SASP profile in senescent 3T3-L1 adipocytes. Interestingly, in this case, CGA treatment was also able to reduce its expression starting from 5 µM.

Furthermore, we analyzed the expression of cyclooxygenase-2 (COX-2), an inducible enzyme involved in the synthesis of pro-inflammatory prostaglandins, commonly induced under oxidative and inflammatory conditions and contributing to the chronic inflammatory state associated with cellular senescence [43]. As shown in Figure 4E, we observed the marked upregulation of COX-2 following treatment with H₂O₂. Notably, CGA treatment downregulated COX-2 expression in a dose-dependent manner.

Moreover, in this case, CGA exposure alone did not induce any detectable changes in ROS, NF-κB, and COX-2 levels or in IL-6, TNF-α, IL-8, and MMP-3 gene expression compared to control cells (see). Figure S4 of Supplementary Materials

Altogether, these findings support CGA's ability to modulate oxidative stress and reduce the inflammatory and matrix remodeling components of SASP.

In adipose tissue, cellular senescence is not only associated with chronic inflammation and structural remodeling but also with the onset of several types of metabolic dysfunction [44]. In particular, one of the main consequences of senescence-related dysfunction in adipose tissue is the impairment of insulin signaling, which reduces glucose homeostasis, contributing to the development of an insulin resistance state [11]. In senescent adipocytes, oxidative stress interferes with insulin signaling by suppressing the phosphatidylinositol 3-kinase (PI3K)–AKT pathway, an important regulator of insulin-mediated metabolic responses [45]. This downregulation leads to the reduced expression and translocation of glucose transporter 4 (GLUT4), compromising glucose uptake and promoting systemic metabolic dysregulation [46].

Our results showed that exposure to H₂O₂ significantly inhibited PI3K activation (Figure 5A) with consequent reduced AKT phosphorylation (Figure 5B) and GLUT4 expression (Figure 5C), confirming the onset of an insulin-resistant state in senescent 3T3-L1 adipocytes. At the same time, glucose uptake (Figure 5D) was also markedly reduced, providing further evidence of impaired insulin signaling and highlighting the compromised metabolic functionality associated with cellular senescence. Interestingly, CGA treatment was able to counteract these alterations in a dose-dependent way. CGA, in fact, restored PI3K-AKT pathway activation and enhanced both GLUT4 expression levels and glucose uptake, suggesting an improvement in insulin responsiveness in senescent adipocytes. No changes were observed in cells treated with CGA alone compared to controls (see Figure S5 of Supplementary Materials). These results therefore suggest an important role of CGA in preventing insulin resistance associated with cellular senescence and aging.

One of the main consequences of cellular senescence is the disruption of adipogenic potential due to the alteration of functional markers involved in adipocyte differentiation [47]. In physiological conditions, in fact, the adipogenesis process is a well-orchestrated program that—through the sequential activation of early and late transcription factors, including peroxisome proliferator-activated receptor gamma (PPARγ)—plays a key role in the full differentiation of the mature adipocyte. Furthermore, the expression of PPARγ-regulated specific genes including fatty acid synthase (FASN) and fatty acid binding protein 4 (FABP4) further contributes to adipose tissue development by promoting lipid biosynthesis and accumulation [48,49].

In the presence of cellular senescence, an impaired differentiation capacity and dysfunctional lipid metabolism occur, which determine several alterations to the adipogenic program, thus contributing to metabolic decline in aging adipose tissue [33,50]. In our experimental model, H₂O₂ significantly impaired adipogenesis, as shown by Oil Red O staining, which revealed a marked reduction in intracellular lipid droplets compared to untreated control cells (0.63 ± 0.11 fold vs. CTR, p < 0.05) (Figure 6A,B). This effect was associated with the strong downregulation of PPARγ levels (Figure 6C) following H₂O₂ exposure, indicating an altered differentiation program. The transcriptional regulation of PPARγ was further supported by FASN gene expression (Figure 6D), which was significantly reduced by H₂O₂, thereby confirming the lipogenic function impairment in senescent cells. Remarkably, CGA treatment effectively counteracted these effects in a dose-dependent way. CGA treatment, in fact, enhanced lipid accumulation, as demonstrated by an increase in both the size and number of lipid droplets compared to control cells (0.78 ± 0.12, 0.93 ± 0.10, and 1.03 ± 0.08 for CGA at 5, 10, and 20 μM, respectively). At the same time, CGA restored both PPARγ protein expression (Figure 6C) and FASN mRNA levels (Figure 6D), suggesting that CGA not only preserves the differentiation capacity but also supports functional lipid metabolism in senescent adipocytes. Notably, also in this case, no significant changes were found in 3T3-L1 cells exposed to CGA alone compared to control cells (see Figure S6 of Supplementary Materials).

Dulbecco's modified eagle medium (DMEM) and Dulbecco's phosphate-buffered saline (DPBS) were purchased from Thermo Fisher Scientific (Monza, Italy). Hydrogen peroxide solution (3%—10 Vols) was acquired from Liofilchem (Teramo, Italy). Chlorogenic acid (CGA) and all other chemicals and solvents used in this study, unless otherwise indicated, were obtained from Merck Life Science (Milan, Italy).

Mouse 3T3-L1 preadipocytes (ATCC, Manassas, VA, USA) were cultured and differentiated into mature adipocytes according to the method described in our previous work [88]. Briefly, the cells were seeded at 1.3 × 10⁴ cells/cm² in multiwell plates and, upon reaching confluence, were maintained in culture for 10 days, until complete differentiation into mature adipocytes. In detail, during the first 4 days (induction phase), cells were incubated in DMEM supplemented with 10% fetal bovine serum (FBS), 4 mM L-glutamine, 100 U/mL penicillin/streptomycin, and 25 mM HEPES buffer (DMEM-full), containing a mixture of 3 isobutyl-1-methylxanthine (0.5 mM), dexamethasone (Dex) 1 μM, and insulin (1 μg/mL) (MDI). Afterwards, this medium was replaced, and the cells were maintained, from day 4 to day 7, in DMEM-full containing only 1 μg/mL insulin. Finally, from day 7 to day 10, cells were cultured in insulin-free DMEM-full to obtain completely differentiated adipocytes.

To induce senescence, on days 5, 6, and 7 of the cell differentiation process, cells were exposed intermittently to a sub-lethal concentration of H₂O₂ (100 µM) for 3 h, following the method established by Zoico et al. [25]. In particular, on days 5 and 6, cells were treated for 3 h with H₂O₂; after this exposure period, cells were gently washed with DPBS, and insulin DMEM-full supplemented with different CGA concentrations was added until the next H₂O₂ exposure. On day 7, after the final H₂O₂ exposure, cells were gently washed with DPBS, and CGA was added in insulin-free DMEM-full and maintained until the end of the differentiation period (day 10). The CGA concentrations (5, 10, and 20 µM) were selected according to oral bioavailability studies [89] and consistent with physiologically relevant levels achievable through pharmacological or dietary intake [90].

Cells treated with the CGA vehicle alone (DMSO 0.1% v/v) and exposed only to the differentiation medium were used as controls. At the end of the treatment period (day 10), cells were processed immediately or stored at –80 °C until further analysis. A schematic representation of the experimental design is shown in Figure 7.

The senescence-associated β-galactosidase (SA-β-Gal) staining assay is a method that is widely used to identify senescent cells. In fact, increased activity of the β-galactosidase enzyme at pH 6 is a typical feature of senescent but not proliferating cells [91]. This test is based on the enzymatic hydrolysis of the chromogenic substrate 5-bromo-4-chloro-3-indolyl β-D-galactopyranoside (X-Gal), which produces an insoluble blue precipitate that accumulates in senescent cells and can be seen under an optical microscope. In particular, in this study, the SA-β-Gal staining procedure was performed following the protocol described by Itahana et al. [27], with minor modifications. In short, after the removal of the culture medium, cells were washed with DPBS and fixed with 4% formaldehyde for 5 min at room temperature. Subsequently, cells were washed again with DPBS and incubated with a freshly prepared SA-β-Gal staining solution at 37 °C for 12–16 h. At the end of the incubation period, the staining solution was removed, and the wells were rinsed with DPBS in order to eliminate the unbound dye. Blue-stained SA-β-Gal-positive cells were observed using an inverted microscope, and representative images were captured for further analyses.

Cell proliferation was carried out using the Erythrosin B exclusion test. Erythrosin B solution (0.1% w/v) was prepared in DPBS, following the manufacturer's instructions (Sigma-Aldrich, Milan, Italy). After trypsinization, cell counts were performed using a hemocytometer, and results were expressed as a percentage relative to untreated control cells.

To evaluate intracellular lipid accumulation, adipogenic cultures were stained with Oil Red O (ORO), following the method described by Molonia et al. [92]. Subsequently, after fixation and staining, cells were visualized under an inverted microscope and photographed for further analysis. Quantification of lipid content was performed using the ImageJ software (v1.54g) [93]. Lastly, the Oil Red O stain retained in the cells was eluted with 100% isopropanol, and lipid accumulation was evaluated spectrophotometrically at 490 nm using a microplate reader (GloMax^® Discover System—TM397, Promega Corporation, Madison, Wisconsin, USA). Results were reported as fold changes relative to control cells.

Following the treatments, 3T3-L1 cells were washed with DPBS and harvested using a cell scraper. Total protein content was determined by incubating the samples for one hour at a low temperature in a lysis buffer (1 mM EDTANa₂, 10 mM Tris–HCl, 150 mM NaCl, 0.1% SDS, 5% glycerol, and 1% Triton). Nuclear and cytoplasmatic extracts were prepared as previously described [94]. In more detail, the cells were firstly treated with a hypotonic buffer (10 mM HEPES, 10 mM KCl, 1.5 mM MgCl₂, and 5% glycerol) to extract cytoplasmatic proteins and then incubated with a hypertonic buffer (20 mM HEPES, 1 mM MgCl₂, 400 mM NaCl, 1 mM EGTA, 0.1 mM EDTA, and 10% glycerol) for the nuclear fractions. All lysis buffers contained protease inhibitors (1 μg/mL leupeptine, 2 μg/mL aprotinine, 1 mM benzamidine, and 5 mM NaF) and 1 mM dithiothreitol (DTT). Protein fractions were stored at −20 °C until further analysis, and protein concentrations were determined using the Bradford assay [95].

For immunoblot analyses, 30 µg of lysate was subjected to SDS-PAGE and then electrotransferred onto a PVDF membrane. Membranes were then blocked using 5% non-fat dry milk solution and incubated overnight at 4 °C with the following specific primary antibodies: rabbit anti-Lamin B1 monoclonal antibody (Cell Signaling Technology, Danvers, MA, USA) (1:1000), mouse anti-PPAR-γ monoclonal antibody (Santa Cruz Biotechnology, Dallas, TX, USA) (1:1500), rabbit anti-PI3K p85 monoclonal antibody (Cell Signaling Technology) (1:1000), mouse anti-p21 (Waf1/Cip1) monoclonal antibody (Santa Cruz Biotechnology) (1:500), mouse anti-phospho-p53 (Ser15) monoclonal antibody (Cell Signaling Technology) (1:1000), rabbit anti-phospho-p38 (Thr180/Tyr182) monoclonal antibody (Cell Signaling Technology) (1:1000), rabbit anti-phospho-ERK1/2 (Thr202/Tyr204) monoclonal antibody (Cell Signaling Technology) (1:1000), rabbit anti-Bax monoclonal antibody (Cell Signaling Technology) (1:1000), rabbit anti-Bcl-2 monoclonal antibody (Cell Signaling Technology) (1:1000), rabbit anti-Phospho-Akt (Ser473) monoclonal antibody (Cell Signaling Technology) (1:1000), rabbit anti-GLUT-4 monoclonal antibody (Cell Signaling Technology) (1:1000), rabbit anti-NF-κB p65 polyclonal antibody (Invitrogen, Milan, Italy) (1:1000), mouse anti-COX-2 monoclonal antibody (Santa Cruz Biotechnology) (1:500), and rabbit anti-β-actin monoclonal antibody (Cell Signaling Technology) (1:6000).

Subsequently, the membranes were incubated for 2 h at room temperature with peroxidase-conjugated secondary antibody HRP-labeled goat anti-rabbit Ig (Cell Signaling Technology) (1:6000) or goat anti-mouse IgM secondary antibody HRP conjugate (Cell Signaling Technology) (1:6000), and they were visualized using an ECL plus detection system (Amersham; GE Healthcare Life Sciences, Marlborough, MA, USA). Blots were detected using a ChemiDoc Imaging System (Bio-Rad, Hercules, CA, USA). The intensities of the bands were quantified using the Image Lab program, and the equivalent loading of proteins in each well was confirmed by Ponceau staining or β-actin.

Intracellular reactive oxygen species (ROS) levels were determined using the fluorescent 2′,7′-dichlorodihydrofluorescein diacetate (DCFH-DA), following the protocol described by Bashllari et al. [96]. After treatment, cells were rinsed twice with DPBS and then incubated with 50 μM DCFH-DA at 37 °C for 30 min. The fluorescence intensity was measured using a microplate reader (GloMax^® Discover System—TM397) (excitation 485 nm and emission 530 nm). The assay was performed in triplicate. ROS levels were normalized to the total protein content and expressed as the DCFH-DA relative fluorescence intensity per mg of protein against controls.

Total RNA was extracted using the E.Z.N.A.^® Total RNA kit, following the manufacturer's instructions (OMEGA bio-tek, Norcross, GA, USA); quantified by a Quant-iT TM RNA assay kit (Invitrogen, Milan, Italy); and reverse-transcribed with M-MLV Reverse Transcriptase. A quantitative real-time polymerase chain reaction (PCR; Applied Bio systems 7300 Real-Time PCR System, Thermo Fisher, Milan, Italy) coupled with SYBR green chemistry (SYBR green JumpStart Taq Ready Mix, Sigma-Aldrich, Milan, Italy) was carried out for the identification of the mRNA levels of FASN (FW 5′-GGAGGTGGTGATAGCCGGTAT-3′, RV 5′-TGGGTAATCCATAGAGCCCAG-3′ [97]), IL-8 (FW 5′-GCACTTGGGAAGTTAACGCA-3′, RV 5′-GCACAGTGTCCCTATAGCCC-3′ [98]), MMP-3 (FW 5′-TGATGAACGATGGACAGAGG-3′, RV 5′-GAGAGATGGAAACGGGACAA-3′ [99,100]), and p21 (FW 5′-CTAGGGGAATTGGAGTCAGGC-3′, RV 5′-GAACAGGTCGGACATCACCA-3′ [100]), and 18S rRNA (FW 5′-GTAACCCGTTGAACCCCATT-3′, RV 5′-CCATCCAATCGGTAGTAGCG-3′ [101]) was used as a reference gene.

Data were elaborated by the SDS 1.3.1 software (Applied Biosystems, Foster City, CA, USA) and expressed as the threshold cycle (Ct). The fold increase in mRNA expression compared with the control cells not pre-treated and not exposed to H₂O₂ and CGA was determined using the 2^−ΔΔCt method.

Glucose uptake was determined using the fluorescent D-glucose analog 2-[N-(7-nitrobenz-2-oxa-1,3-diazol-4-yl) amino]-2-deoxy-D-glucose (2-NBDG), followed by the detection of cell-produced fluorescence according to the protocol described by Molonia et al. [102], with slight modifications. The fluorescence intensity was measured using a fluorescence microplate reader (GloMax^® Discover System—TM397) (excitation 465 nm, emission 540 nm). The measured signal, proportional to glucose uptake, was normalized to the total protein content and expressed as the relative fluorescence intensity per mg of protein, with values reported in comparison to untreated controls.

Experiments were performed in triplicate and replicated three times. Data are expressed as the mean ± S.D. from three independent experiments. Statistical analysis was carried out by a one-way ANOVA test, followed by Tukey's HSD, using the software ezANOVA (https://people.cas.sc.edu/rorden/ezanova/index.html↗ accessed on 6 November 2025). Statistical significance was set at p < 0.05 and p < 0.01. Exact p-values are available upon request.