4‐Octyl Itaconate Attenuates Age‐Induced Parotid Gland Dysfunction Through the Suppression of Fibrosis and Lipid Deposition by AMPKα/PPARα and the Inhibition of Oxidative Stress via the AMPKα/Nrf2 Signaling Pathway

ABSTRACT

Parotid gland aging is a key factor for reduced salivary production in older individuals. The role of 4‐Octyl itaconate (4‐OI) in age‐related parotid gland dysfunction remains unclear. Thirteen‐month‐old male mice were administered 4‐OI (25 mg/kg) or vehicle once daily for 4 weeks. The salivary flow rate was calculated to assess gland secretion function. Hematoxylin and eosin staining was used to analyze morphological alterations. Masson staining was used to quantify collagen deposition. Oil Red O staining was used to evaluate lipid droplet accumulation. Western blotting was used to detect protein expression levels. Metabolomics, transcriptome, and molecular docking analyses were performed to identify potential mechanisms of action. Functional assays revealed that 4‐OI increased the salivary flow rate. AMPKα has been identified as a direct target of 4‐OI by molecular docking. 4‐OI upregulates AMPKα phosphorylation, decreases lipid droplet deposition, and relieves fibrosis to restore the aging characteristics of the gland. 4‐OI regulates fatty acid oxidation (FAO) in the parotid gland through AMPKα/PPARα signaling pathways. The protective effects of 4‐OI in aged mice were abolished by Compound C and GW6471, which inhibit AMPKα and PPARα, respectively. In addition, we found that 4‐OI treatment ameliorated parotid gland aging‐induced oxidative reactions, enhanced mitochondrial biogenesis, and activated the Nrf2 pathway. Moreover, the Nrf2 inhibitor ML385 was used to validate this mechanism. Treatment with 4‐OI improved the aging of the parotid gland by increasing gland FAO, decreasing gland fibrosis, and preventing oxidative reactions. This was achieved by upregulating the AMPKα/Nrf2 and AMPKα/PPARα signaling pathways.

4‐Octyl itaconate (4‐OI) attenuates age‐induced parotid gland dysfunction through the suppression of fibrosis and lipid deposition by AMPKα/PPARα and the inhibition of oxidative stress via the AMPKα/Nrf2 signaling pathway. 4‐OI activates AMPKα, which in turn stimulates both the PPARα and Nrf2 pathways. Activation of PPARα enhances FAO and reduces lipid deposition and fibrosis. Concurrently, Nrf2 activation boosts antioxidant defenses and mitochondrial function, reducing oxidative stress and inflammation. These combined effects ultimately improve parotid gland function. The protective effects were reversed by the inhibitors Compound C (AMPKα), GW6471 (PPARα), and ML385 (Nrf2).

1. Introduction

Salivary hypofunction is among the most prevalent oral health issues in elderly individuals and is caused primarily by age‐related changes in acinar cells that lead to reduced salivary flow and altered saliva composition [1, 2]. This condition contributes to a range of clinical problems, including dental caries, oral candidiasis, oral dysbiosis, xerostomia, difficulties in speech and swallowing, impaired taste perception, and halitosis [3, 4]. Studies indicate that older adults experience significant reductions in both unstimulated and stimulated salivary flow by approximately 38.5% and 38.0%, respectively [5]. As a vital component of oral homeostasis, saliva plays key roles in lubrication, taste, mastication, deglutition, and initial immune defense [6]. By 2050, the percentage of the world's elderly population is predicted to increase to 22% [7]. Given the expanding aging population, developing treatments that improve saliva generation and dental health in elderly individuals is crucial.

Aging is a complicated biological process that involves many different systems and mechanisms whose ability to function is gradually decreased because of environmental, stochastic, genetic, and epigenetic events in various cell types and tissues [8, 9]. The hallmarks of cellular senescence are senescence‐associated β‐galactosidase (SA‐β‐gal) activation and persistent cell cycle arrest caused by P21 and P16^INK4A upregulation [10]. Systemic, low‐grade, chronic inflammation in aging tissues is a common characteristic of most age‐related disorders [11, 12]. Proinflammatory factors and other senescence‐associated secretory phenotype (SASP) components are secreted by these senescent cells, which compromise tissue integrity and promote aging [13]. Normal tissue function is impaired by cellular senescence, which decreases the quantity of functional cells within tissues [14], and their elimination can attenuate aging in tissue and improve health [15, 16, 17]. Among the three main salivary glands, the parotid gland is the largest and produces approximately 25% of the total saliva at rest and 53% upon stimulation [18]. Age is significantly associated with fat content in human parotid glands [19]. To date, no effective strategy for delaying the process of parotid gland aging has been reported.

D‐galactose‐induced aging rats, an aging‐acceleration model, exhibit increased production of reactive oxygen species (ROS) and acinar cell apoptosis in the salivary glands [20]. ROS are primarily generated by the mitochondrial respiratory chain. Previous studies have shown that mitochondrial dysfunction contributes to various aging‐related disorders [21, 22]. With the accumulation of advanced glycation end products in the salivary glands, D‐galactose‐induced aging mice also exhibit chronic inflammation and loss of acinar epithelial cells [23].

Immune response gene 1 (IRG1), which encodes the active enzyme aconitate decarboxylase (ACOD1), is primarily responsible for the synthesis of itaconate, a naturally occurring chemical molecule that is categorized as a dicarboxylic acid [24]. Itaconate decreases the production of inflammatory cytokines such as tumor necrosis factor‐α (TNF‐α) and interleukin‐1β (IL‐1β). Itaconate suppresses inflammatory signaling pathways, including nuclear factor‐κB (NF‐κB) and mitochondria‐related pathways, which reduce inflammation [25, 26]. In addition, itaconate has antioxidant effects that minimize oxidative stress damage through the scavenging of free radicals. Itaconate reduces oxidative stress by succinate dehydrogenase (SDH) to prevent the generation of ROS and activating nuclear factor erythroid 2‐related factor 2 (Nrf2) to induce the expression of antioxidant genes [27]. In addition, some studies have shown that itaconate derivatives can regulate oxidative stress and lipid metabolism via AMP‐dependent protein kinase (AMPK) phosphorylation and suppress inflammation [28, 29]. Supplementation with exogenous 4‐octyl itaconate (4‐OI), a derivative of itaconate, offers a promising strategy for aging‐related disorder treatment [30, 31, 32]. Although the anti‐inflammatory characteristics of itaconate indicate a potential benefit in treating parotid gland aging, further research is necessary to clarify its actual mechanisms and effectiveness in this context.

Therefore, the aim of this study was to investigate the antiaging effects of 4‐OI on the parotid gland in mice, with a focus on its ability to inhibit parotid gland fibrosis and lipid deposition via the AMPKα/PPARα and AMPKα/TGF‐β pathways and mitigate oxidative stress and inflammation via the AMPKα/Nrf2 signaling pathway.

2. Material and Methods

2.1. Animals

The 2‐month‐old and 13‐month‐old male C57BL/6 mice were purchased from Nanjing ZHISHU YANTU Biotechnology Co. Ltd. (Nanjing, China). All the mice were maintained under controlled environmental conditions with a 12‐h light/12‐h dark cycle at 20°C–26°C and 40%–70% humidity with free access to food and water. All procedures involving animal experiments were approved by the Ethics Committees of Second Xiangya Hospital of Central South University (Approval No. 20260981) and complies with the ARRIVE 2.0 guidelines. The study was divided into four sections. The first section examined the efficacy of 4‐OI in the exploration of its involvement in aging parotid glands, and the other three sections explored potential pathways (n = 10 per group).

Section I: After 1 week of environmental adaptation, the 13‐month‐old mice were randomly assigned to one of two groups: 13‐month‐old mice group (13 M) or 13 M + 4‐OI. The mice in the 13 M + 4‐OI group were administered 4‐OI (25 mg/kg/day, HY‐112675, MedChemExpress, NJ, USA) via intraperitoneal (i.p.) injection for 4 weeks, whereas the mice in the 13 M group and 2 M groups were given vehicle by i.p. injection. Section II: After 1 week of environmental adaptation, the mice were separated into four groups: 2‐month‐old mice group (2 M), 13 M, 13 M + 4‐OI, and 13 M + 4‐OI + CC (AMPKα inhibitor). Mice in the 13 M + 4‐OI + CC group received an i.p. injection of an AMPKα inhibitor (Compound C, 10 mg/kg; HY‐13418A; MedChemExpress, NJ, USA) daily for 4 weeks, whereas the mice in the 13 M group and 2 M group were administered vehicle by i.p. injection. Section III: After 1 week of environmental adaptation, the mice were separated into four groups: 2 M, 13 M, 13 M + 4‐OI, and 13 M + 4‐OI + ML385 (an Nrf2 inhibitor). Mice in the 13 M + 4‐OI + ML385 group received an i.p. injection of an Nrf2 inhibitor (ML385, 30 mg/kg; HY‐13418A; MedChemExpress, NJ, USA) daily for 4 weeks, whereas the mice in the 13 M group and 2 M group were given vehicle by i.p. injection. Section IV: After 1 week of environmental adaptation, the mice were separated into 4 groups: 2 M, 13 M, 13 M + 4‐OI, and 13 M + 4‐OI + GW6471 (PPARα antagonist). Mice in the 13 M + 4‐OI + GW6471 group received an i.p. injection of a PPARα antagonist (GW6471, 10 mg/kg; HY‐13418A; MedChemExpress, NJ, USA) daily for 4 weeks, whereas the mice in the 13 M group and 2 M group were given vehicle by i.p. injection. 4‐OI, compound C, ML385, and GW6471 were dissolved with 10% DMSO (HY‐Y0320, MedChemExpress) and 90% corn oil (HY‐Y1888, MedChemExpress). The mice in the 2 M and 13 M groups were only administered vehicle containing 10% DMSO and 90% corn oil by i.p. injection.

2.2. Measurement of Parotid Gland Function

To measure the stimulated salivary flow rate, mice received pilocarpine (5 mg/kg; #HY‐B1006; MedChemExpress, NJ, USA) by i.p. injection after anesthesia. Whole saliva was carefully collected from the bottom of the mouth using a micropipette for 10 min at room temperature. The salivary flow rate (μL/min) was determined by dividing the total volume (μL) of the collected saliva by the collection duration (min). To reduce swallowing and prevent suffocation, the mice's heads were turned downward to one side.

2.3. Histological Analysis

Fresh mouse parotid gland tissues were fixed in 4% paraformaldehyde solution for 24 h. The tissues were subsequently dehydrated, embedded in paraffin, and sliced into paraffin sections (4 μm thick). The paraffin sections were dewaxed in xylene for 30 min and stained using hematoxylin and eosin (H&E) staining solution, Masson staining solution, periodic acid–Schiff (PAS) staining solution and Sirius Red staining solution. The remaining tissues were embedded in Tissue‐Tek OCT compound (Sakura, Tokyo, Japan) and sliced into sections (8 μm thick) using a cryostat microtome (Leica CM1860) for Oil Red O staining, according to the manufacturer's instructions. Photomicrographs were captured with a microscope (Leica DM2000). In accordance with a previous study, we assessed the degree of histological damage to the parotid glands [33]. Briefly, scores of 0, 1, 2, 3, 4, and 5 indicated that > 90%, 70%–90%, 50%–70%, 30%–50%, 10%–30%, and < 10% of the acini remained, respectively. The mucin‐containing area (PAS) and fibrotic region (Masson and Sirius Red) were examined quantitatively with ImageJ software (Version 1.46; NIH, Bethesda, MD, USA).

2.4. SA‐β‐Gal Staining

The senescent area in frozen parotid gland sections was analyzed using an SA‐β‐gal staining kit (Beyotime, Shanghai, China) at 37°C for 20 h in the dark according to the manufacturer's instructions. Photomicrographs were captured with a microscope (Leica DM2000). SA‐β‐gal‐positive cells were stained green. Senescent cells were quantified by assessing the percentage of the SA‐β‐gal‐positive area in six different fields per sample.

2.5. BODIPY Staining

To assess the accumulation level of neutral lipids, frozen parotid gland sections were incubated with a BODIPY 493/503 staining kit (GlpBio, CA, USA) for 30 min in the dark according to the manufacturer's instructions, after which the nuclei were stained with DAPI. All fluorescence images of the sections were captured using a fluorescence microscope (Axio Imager M2 microscope; Carl Zeiss, Germany).

2.6. TUNEL Staining

The tissue sections were fixed in 4% paraformaldehyde solution for 24 h, washed twice with phosphate‐buffered saline (PBS), and then permeabilized with Proteinase K. After three washes with PBS, TUNEL working solution (#C1089; Beyotime, Shanghai, China) was added, and the samples were incubated for 2 h at 37°C, after which the nuclei were stained with DAPI. All fluorescence images of the sections were captured using a fluorescence microscope (Axio Imager M2 microscope; Carl Zeiss, Germany).

2.7. Western Blot Analysis

Parotid gland tissues were homogenized and lysed with RIPA lysis buffer (Beyotime, Shanghai, China) supplemented with a protease and phosphatase inhibitor cocktail (Beyotime, Shanghai, China) on ice. The protein concentrations of the samples were subsequently quantified with a BCA protein assay kit (Beyotime, China). Next, samples with equal protein concentrations were separated by 10%–12% sodium dodecyl sulfate‐polyacrylamide gel electrophoresis (SDS‐PAGE) and transferred to polyvinylidene fluoride (PVDF) membranes. After blocking with 5% BSA for 2 h at room temperature, the PVDF membranes were subsequently incubated with primary antibodies at 4°C overnight. After being washed with TBST (0.1% Tween in Tris), the PVDF membranes were incubated with secondary peroxidase‐conjugated antibodies, after which the protein bands were detected by chemiluminescence using ECL luminescence solution (Tanon, China). The primary antibodies used for the immunohistochemistry or immunofluorescence experiments are listed in Table S1.

2.8. Immunohistochemistry (IHC) and Immunofluorescence Staining

The paraffin‐embedded tissue sections were removed using xylene and rehydrated in graded alcohol. For immunofluorescence staining, antigen retrieval was performed by boiling the samples in Tris‐EDTA buffer solution (pH 9.0, 98°C for 45 min). After blocking with 5% BSA and washing with PBS, the sections were incubated with primary antibodies overnight at 4°C. After being washed with PBS, the sections were incubated with HRP‐conjugated secondary antibodies, and a DAB substrate kit (#34002; Thermo Fisher Scientific) was used for IHC. The sections were counterstained with hematoxylin and observed using a microscope (Leica DM2000). For immunofluorescence, the sections were incubated with fluorescence‐labeled secondary antibodies. Nuclei were stained with DAPI. All fluorescence images of the sections were captured using a fluorescence microscope (Axio Imager M2 microscope; Carl Zeiss, Germany). The primary antibodies used for the IHC or immunofluorescence experiments are listed in Table S1.

For colocalization analysis, a white line with arrows was drawn across representative regions of the merged images using ImageJ software. The length of the white line (in μm) was recorded and corresponded directly to the x‐axis of the fluorescence intensity profile. Fluorescence intensity profiles of the two or three channels along this line were generated, where the x‐axis represents the distance (μm) along the white line and the y‐axis represents fluorescence intensity (a.u.). The degree of colocalization was assessed by comparing the spatial overlap of the peak intensities of the two or three channels along the line and a shorter distance between the peak intensities of the two or three channels indicated a higher degree of colocalization.

2.9. Flow Cytometry Analysis

Cut the parotid gland tissues into 1–5 mm minced pieces, add tissue dissociation buffer to digest into a single‐cell suspension for 30 min. Filter through a 70 μm sterile filter, then centrifuge the cell suspension at 1200 rpm for 5 min. Resuspend cells once with PBS. Add 10 μL Propidium Iodide and 5 μL Annexin V‐FITC. Incubate at room temperature in the dark for 10 min. Samples were analysis with a flow cytometer (Beckman Coulter), and data were processed with FlowJo V10. software.

2.10. Mitochondrial Membrane Potential (MMP) Detection

The mitochondrial membrane potential was measured with a JC‐1 detection kit (C2003S; Beyotime, Shanghai, China) following the manufacturer's instructions. After the medium was added, a fluorescence microscope was used to detect the red and green fluorescence.

2.11. ROS Detection

The frozen parotid gland sections were incubated with 10 μM dihydroethidium (DHE) solution (#Beyotime, Shanghai, China) for 30 min at 37°C in the dark. The sections were subsequently washed three times using PBS and stained with DAPI for 5 min. Fluorescence images of the sections were captured using a fluorescence microscope (Axio Imager M2 microscope; Carl Zeiss, Germany).

2.12. Transmission Electron Microscopy (TEM)

Parotid gland tissue was fixed with 2.5% glutaraldehyde and treated with 1% osmium at 4°C. The parotid gland tissue was dehydrated in graded alcohol and then treated in graded acetone. The samples were cut into ultrathin slices (60–80 nm) for TEM analysis.

2.13. Metabolomic Analysis of Parotid Glands

The parotid tissue was homogenized for 6 min using a mixed solvent of methanol and water in a ratio of 4:1 (v/v), followed by low‐temperature ultrasonic extraction for 30 min. After centrifugation, the supernatant was collected and reserved for metabolomics analysis. Additionally, transfer 20 μL supernatants from each sample, mix, and use as quality control sample (QC). The instrument platform for this LC–MS/MS analysis was UHPLC‐Q Exactive HF‐X system (Thermo Fisher Scientific). Chromatographic conditions: Column was ACQUITY UPLC HSS T3 (100 mm × 2.1 mm i.d., 1.8 μm; Waters, Milford, USA); mobile phase A was 95% water +5% acetonitrile (containing 0.1% formic acid), mobile phase B was 47.5% acetonitrile +47.5% isopropanol +5% water (containing 0.1% formic acid), injection volume was 3 μL, and column temperature was 40°C. The orthogonal partial least squares discriminant analysis (OPLS‐DA) and bioinformatics analyses were conducted on the Majorbio Cloud Platform (https://cloud.majorbio.com/).

2.14. Transcriptome Analysis of Parotid Glands

Total RNA was extracted using TRIzol Reagent, and then cDNA was synthesized. NovaSeq Reagent Kit was used for sequencing, while StringTie and HISAT2 were used to process the data. DEGs (differential expression genes) with |log2FC| ≧ 1 and p‐adjust ≤ 0.05 were considered to be significantly different expressed genes. We conducted functional‐enrichment analysis using Kyoto Encyclopedia of Genes and Genomes (KEGG) to identify DEGs that were highly enriched in pathways relative to the whole‐transcriptome background.

2.15. Molecular Docking and Molecular Dynamics Simulations

The AMPKα protein information was obtained from the UniProt website (https://www.uniprot.org/uniprotkb) and the AMPKαprotein structure was obtained from the PDB database (https://www.rcsb.org/). Import the obtained structure into Discovery Studio for protein structure optimization. Retrieve four structures from the PubChem website and perform energy minimization using Discovery Studio 2019. Prepare the protein and small molecule files using Autodock 4.0, and perform docking using Autodock VINA 1.2.6. Finally, perform visualization analysis in PyMOL (v3.1) and Discovery Studio 2019.

A 100 ns molecular dynamics simulation of the protein‐ligand complex was performed using GROMACS 2025, following the procedure below: The protein was modeled using the AMBER99SB‐ILDN force field parameters, and the ligand topology was constructed using the GAFF2 force field parameters. The protein and ligand topology files were merged to ensure no conflicts in atom types. Periodic boundary conditions (PBC) were applied, and the complex was placed inside a cubic box with a minimum distance of 1.2 nm between the complex surface and the box walls. The solvent was filled with the TIP3P water model, and Na⁺/Cl⁻ counterions were added to neutralize the net charge of the system (ion concentration was maintained at 0.15 M to simulate physiological conditions). First, an energy minimization was performed, followed by 10 000 000 steps of isothermal‐isovolumetric (ITC) and isothermal‐isobaric (IIB) ensemble equilibration, with a coupling constant of 0.1 ps and a simulation duration of 2 ns. The temperature/pressure coupling parameters were the same as those in the equilibration phase, and the trajectory was saved every 1000 steps for subsequent analysis. Finally, a free molecular dynamics simulation was run, consisting of 50 000 000 steps with a step size of 2 fs, for a total duration of 100 ns. Finally, a molecular dynamics simulation was performed using GROMACS 2025 at constant temperature (310 K) and constant pressure (1 bar), with a total duration of 100 ns. The binding free energy was calculated using MMPBSA after the system reached equilibrium.

The following metrics were calculated using GROMACS 2025: root mean square deviation (RMSD, to assess overall structural stability), root mean square fluctuation (RMSF, to analyze residue flexibility), radius of gyration (Rg, to characterize molecular compactness), solvent accessible surface area (SASA, to reflect the degree of solvent exposure), and protein‐ligand (to assess interaction stability), Two‐ and three‐dimensional free energy landscape (FEL), and mmPBSA (mean binding free energy, residue decomposition energy).

2.16. Statistical Analysis

Statistical analysis was performed using GraphPad Prism (version 9.3.0; San Diego, CA, USA). The data are expressed as the mean ± standard deviation (SD). Two‐tailed Student's t‐test was used to evaluate differences between two groups. One‐way analysis of variance (ANOVA) followed by Tukey's post hoc multi‐comparison test was used to evaluate differences among multiple groups. In all the statistical analyses, significance was defined as a p value < 0.05.

3. Results

3.1. 4‐OI Alleviated Parotid Gland Secretion Dysfunction and the Functional Phenotype in Aged Mice

To explore the effect of 4‐OI on the parotid glands of 13 M mice, we subsequently injected 13 M mice with 25 mg/kg body weight 4‐OI. After 4 weeks of treatment, the salivary flow rate in mice was significantly greater in the 13 M + 4‐OI group than in the 13 M group (Figure 1A). To explore whether exogenous 4‐OI affects the histology of the glands, we used H&E and PAS staining to analyze the parotid glands of mice treated with 4‐OI for 4 weeks. H&E staining was used to evaluate the structure of the parotid glands, and PAS staining was used to assess mucin secretion. H&E staining revealed significant gland tissue damage, including tissue disarray and a reduction in cell density, in the 13 M group. On the other hand, the 13 M + 4‐OI group, like the 2 M group, presented a greater cell density and a more ordered glandular structure. The 13 M groups had deficient acini, as determined by PAS staining. Mucin‐producing acini were evaluated by measuring the regions stained purplish‐red. Compared with those in the 13 M group, the mucin‐containing regions in the 4‐OI treatment group were notably reduced (Figure 1B,C).

FIGURE 1.

4‐OI alleviates function and structure impairments in the parotid gland of aged mice. (A) Salivary flow rates in different groups of mice (n = 6). (B) Representative images and quantitative analysis of hematoxylin and eosin (H&E) staining in the parotid gland of different groups of mice (n = 6). Scale bars: 50 μm (top) and 20 μm (bottom). (C) Representative images and quantitative analysis of Periodic acid–Schiff (PAS) staining in the parotid gland of different groups of mice (n = 6). Scale bars: 50 μm (top) and 20 μm (bottom). (D) Representative images and quantitative analysis of senescence‐associated β‐galactosidase (SA‐β‐gal) staining in the parotid gland of different groups of mice (n = 6). Scale bars: 50 μm. (E, F) Representative Western blot and quantitative analysis the protein levels of P16^INK4A, P21, and P53 in the parotid gland of different groups of mice (n = 3). (G) Representative images of immunofluorescent staining for F4/80 (red) and DAPI (blue) and quantitative analysis of F4/80 positive cells in the parotid gland of different groups of mice (n = 6). Scale bars: 50 μm. (H) Representative images of immunofluorescent staining for CD45 (red) and DAPI (blue) and quantitative analysis of CD45 positive cells in the parotid gland of different groups of mice (n = 6). Scale bars: 50 μm. (I, J) Representative images of immunofluorescent staining for KRT7 (red, a marker for luminal ductal cells), KRT5 (green, a marker for basal ductal cells), and DAPI (blue) and quantitative analysis of KRT7 positive cells and KRT5 positive cells in the parotid gland of different groups of mice (n = 6). Scale bars: 50 μm (top) and 20 μm (bottom). (K, L) Representative images of immunofluorescent staining for KRT14 (red, a marker for myoepithelial cells), BHLHA15 (green, a marker for mature acinar cells), and DAPI (blue) and quantitative analysis of KRT14 positive cells and BHLHA15 positive cells in the parotid gland of different groups of mice (n = 6). Scale bars: 50 μm (top) and 20 μm (bottom). (M, N) Representative Western blot and quantitative analysis the protein levels of IRG1 in the parotid gland of different groups of mice (n = 3). (O) Representative images of immunofluorescent staining for IRG1 (red), BHLHA15 (green), and DAPI (blue) in the parotid gland of different groups of mice (n = 6). Scale bars: 50 μm (top) and 20 μm (bottom). The data are presented as the means ± SD. *p < 0.05, **p < 0.01, ***p < 0.001. ns, not significant.

A common method used to assess cellular senescence is SA‐β‐gal staining. Notably, compared with those of 2 M mice, the parotid glands of 13 M mice showed a considerable increase in SA‐β‐gal activity. On the other hand, the 13 M + 4‐OI group showed an obvious reduction in β‐gal activity (Figure 1D). Additionally, we evaluated several indicators of cellular senescence, including P16^INK4A, P21, and P53. Western blot analysis was conducted to investigate P16^INK4A, P21, and P53 expression. When 4‐OI was applied to 13 M parotid glands, the levels of P16^INK4A, P21, and P53 decreased compared with those in the 13 M group (Figure 1E,F). Immunofluorescence analysis was performed using antibodies against CD45 and F4/80 to investigate inflammatory cell and macrophage infiltration into the parotid glands (Figure 1G,H). Very few CD45‐positive and F4/80‐positive cells were identified in the parotid glands of either genotype in the 2 M group. 13 M mice showed significant accumulation of CD45‐positive and F4/80‐positive cells, whereas the 13 M + 4‐OI group showed considerable suppression of this accumulation.

To investigate whether 4‐OI contributes to parotid gland epithelial cell survival, we used immunofluorescence staining with antibodies against KRT5, which is a marker of basal cells; KRT7, a marker of luminal cells; KRT14, a marker of myoepithelial cells; and BHLHA15, a marker of mature acinar cells. Four weeks after 4‐OI treatment, parotid gland tissues from 13 M + 4‐OI groups exhibited increased KRT5 and KRT7 expression compared to those in the 13 M group (Figure 1I,J). Similarly, KRT14 and BHLHA15 expression were restored in the 13 M + 4‐OI group (Figure 1K,L). The above results demonstrate that aging‐induced damage to the parotid glands is ameliorated by 4‐OI treatment.

To determine whether endogenous itaconate synthesis occurs in the parotid glands during aging, we examined IRG1 expression in the parotid glands. According to the western blot results, increased IRG1 expression was noted in the 13 M group compared with the 2 M group, but levels gradually decreased in the 13 M + 4‐OI group (Figure 1M,N). Next, we performed immunofluorescence staining to observe IRG1 localization in the parotid gland tissues to identify the primary site of IRG1 expression in the gland (Figure 1O). The results demonstrated that IGR1 expression in the 13 M group was greater than that in the 2 M group. However, IRG1 expression dramatically decreased following 4‐OI treatment, mostly in the cytoplasm of the acinar cells.

3.2. 4‐OI Treatment Prevents Apoptosis and Inflammation in Aged Parotid Glands

Low‐grade chronic inflammation is among the common hallmarks of aging. We hypothesized that 4‐OI treatment reduces parotid gland inflammation, which has antiaging effects. As expected, 4‐OI treatment significantly reduced the age‐related increases in the expression levels of proinflammatory cytokines and SASP factors (IL‐1β and IL‐6) in 13 M parotid glands (Figure 2A,B). Next, we performed TUNEL staining and flow cytometry analysis to determine the degree of apoptosis in the parotid glands. 13 M mice showed an increase in the number of aging‐induced TUNEL‐positive cells and increased PI/Annexin V double‐positive populations in the parotid glands (Figure 2C,D). We also measured the expression of the apoptosis‐related proteins Bcl‐2, Bax, and cleaved‐Caspase3 using Western blotting. Compared with the 2 M group, the 13 M group exhibited dramatically increased apoptosis levels. Bcl‐2 protein levels decreased, whereas Bax and cleaved‐Caspase3 protein levels increased (Figure 2E,F). These effects were markedly ameliorated following 4‐OI treatment. Western blot analysis revealed significant upregulation of NLRP3, ASC, and Caspase 1/p20 protein levels in the parotid gland tissues of 13 M mice compared with those in 2 M mice (Figure 2G,H). Immunofluorescence analysis revealed a significant change in NLRP3 expression in acinar cells. However, 4‐OI treatment significantly downregulated the expression of the abovementioned components of NLRP3 (Figure 2I,J).

FIGURE 2.

4‐OI decreased age‐induced apoptosis, inflammation, and NLRP3 activation in the parotid gland of aged mice. (A, B) Representative Western blot and quantitative analysis the protein levels of IL‐1β and IL‐6 in the parotid gland of different groups of mice (n = 3). (C) Representative images of immunofluorescent staining for TUNEL (red) and DAPI (blue) and quantitative analysis of TUNEL positive cells in the parotid gland of different groups of mice (n = 6). Scale bars: 50 μm. (D) Flow cytometry to detect cell apoptosis. (E, F) Representative Western blot and quantitative analysis the protein levels of Bcl‐2, Bax, Cleaved‐Caspase3 in the parotid gland of different groups of mice (n = 3). (G, H) Representative Western blot and quantitative analysis the protein levels of NLRP3, ASC, and Caspase 1/p20 in the parotid gland of different groups of mice (n = 3). (I) Representative images of immunofluorescent staining for NLRP3 (red), BHLHA15 (green), and DAPI (blue) in the parotid gland of different groups of mice. Scale bars: 50 μm (top) and 20 μm (bottom). The white line with arrows indicates the region selected for colocalization analysis. (J) Fluorescence intensity profiles of NLRP3 (red) and BHLHA15 (green) along the white line shown in (I). The x‐axis represents the distance (μm) along the white line with arrows, and the y‐axis represents fluorescence intensity (a.u.). A shorter distance between the peak intensities of the two channels indicates a higher degree of colocalization. The data are presented as the means ± SD. *p < 0.05, **p < 0.01, ***p < 0.001. ns, not significant.

3.3. 4‐OI Treatment Reduces Parotid Gland Fibrosis, Oxidative Stress, and Improves Mitochondrial Function in Aged Mice

Fibrosis results from excessive extracellular matrix (ECM) deposition because it disrupts normal tissue architecture and contributes to gland dysfunction. Next, we investigated whether 4‐OI affects parotid gland fibrosis. Compared with the 2 M group, 13 M mice presented increased amounts of collagen accumulation in the gland, but 4‐OI treatment decreased the amount of collagen deposition, as determined by Masson staining and Sirius Red staining (Figure 3A,B). 4‐OI‐treated mice consistently exhibited significantly decreased expression of fibrotic proteins, including FN and COL1 (Figure 3C,D). Consistent with these findings, immunofluorescence staining revealed that the mean fluorescence intensity of COL1 in the parotid glands was reduced in 4‐O‐treated mice (Figure 3E). Moreover, Western blot analysis of αSMA also showed that 4‐OI could relieve fibrosis in the parotid gland fibrosis of aged mice (Figure 3F).

FIGURE 3.

4‐OI inhibits fibrosis, excessive production of ROS, and reversed aged‐induced mitochondrial membrane potential decreased in the parotid gland of aged mice. (A) Representative images and quantitative analysis of Masson staining in the parotid gland of different groups of mice (n = 6). Scale bars: 50 μm (top) and 20 μm (bottom). (B) Representative images and quantitative analysis of Sirius Red staining in the parotid gland of different groups of mice (n = 6). Scale bars: 50 μm (top) and 20 μm (bottom). (C, D) Representative Western blot and quantitative analysis the protein levels of FN1 and COL1 in the parotid gland of different groups of mice (n = 3). (E) Representative images of immunofluorescent staining for COL1 (red) and DAPI (blue) and quantitative analysis of relative fluorescence intensity of COL1 in the parotid gland of different groups of mice (n = 6). Scale bars: 50 μm. (F) Representative Western blot and quantitative analysis the protein levels of αSMA in the parotid gland of different groups of mice (n = 3). (G) Representative ROS‐induced fluorescence staining images labeled with DHE dye (red) and DAPI (blue), accompanied by quantitative analysis of relative DHE fluorescence intensity in the parotid glands of mice from different groups (n = 6). (H) Representative images and quantitative analysis of JC‐1 staining in the parotid glands of mice from different groups (n = 6). Scale bar: 50 μm (top) and 20 μm (bottom). (I, J) Representative Western blot and quantitative analysis the protein levels of PGC‐1α, TFAM, and cytochrome c in the parotid gland of different groups of mice (n = 3). The data are presented as the means ± SD. **p < 0.01, ***p < 0.001. DHE, dihydroethidium; ns, not significant; ROS, reactive oxygen species.

The aging microenvironment is recognized for its persistent oxidative stress, which causes severe inflammation and cellular injury [34]. Thus, the antioxidant properties of 4‐OI were assessed. In the 13 M group, the mean fluorescence intensity of DHE was greater than that in the 2 M group. However, 4‐OI treatment decreased the intracellular red fluorescence in these groups and decreased the mean fluorescence intensity (Figure 3G). Because mitochondrial impairment is recognized as one of the primary causes of cellular senescence, we explored whether mitochondrial dysfunction contributes to cellular senescence in the glands. The MMP is an indicator of the function of mitochondria and can be measured with JC‐1, a fluorescent dye that detects changes in the MMP. The 13 M groups accumulated and produced more green fluorescence, indicating a low MMP. Treatment with 4‐OI efficiently restored mitochondrial function by preserving the MMP (Figure 3H).

To investigate the underlying mechanism, we used western blotting to measure the expression of key mitochondrial biogenesis proteins. Aging downregulated the expression of peroxisome proliferator‐activated receptor γ coactivator‐1α (PGC‐1α) and mitochondrial transcription factor A (TFAM) compared with that in the 2 M group, whereas 4‐OI significantly upregulated PGC‐1α and TFAM expression compared with that in the 13 M groups. Cytochrome c is a key mediator of mitochondria‐related apoptosis, so we explored cytochrome c expression levels. The results revealed that in the 13 M group, the level of cytochrome c in the parotid glands increased. After 4‐OI treatment, cytochrome c expression decreased (Figure 3I,J). Overall, our results suggest that 4‐OI rescues mitochondrial biogenesis from aging‐related damage to the parotid gland.

3.4. 4‐OI Ameliorated Parotid Gland Injury in Aged Mice by Regulating the AMPKα and Nrf2 Pathways

We performed non‐targeted metabolomics and transcriptome analysis of parotid glands to explore the potential mechanism of 4‐OI in parotid glands of aged mice. The OPLS‐DA score plot of metabolites exhibited distinct separation between the 13 M and 2 M groups, as well as between the 13 M + 4‐OI and 13 M groups (Figure 4A,B). The Venn diagram and histogram showing the number of metabolites shared and unique to different groups (Figure 4C,D). In order to clarify the changes in metabolite regulation, a volcano plot was constructed (Figure 4E–G). Differential metabolite analysis revealed 155 up‐regulated and 58 downregulated metabolites between the 13 M and 2 M groups. In contrast, 85 up‐regulated and 36 downregulated metabolites were identified between the 13 M + 4‐OI and 13 M groups. To investigate the impact of 4‐OI therapy on specific pathways, KEGG pathway classification was applied to the determined differential metabolites between comparison groups. Compared to the 2 M group, the 13 M group had aberrant metabolic pathways including histidine and glutathione metabolism. 4‐OI profoundly regulated metabolic pathways such as alanine, aspartate and glutamate metabolism (Figure 4H–K). To elucidate the mechanisms behind the protective benefits of 4‐OI. Parotid glands transcriptional profiling was carried out in the 2 M, 13 M, and 13 M + 4‐OI groups. Between the 13 M and 2 M and 13 M + 4‐OI and 13 M groups, 163 and 267 DEGs were found, respectively (Figure 4L,N). KEGG pathway analysis revealed that DEGs between 13 M + 4‐OI and 13 M groups enriched in signaling pathways including PPAR and AMPKα (Figure 4O,P). PPAR and AMPKα are associated with lipid metabolism. Furthermore, the intracellular pathway that mediated the attenuation of aged parotid gland by 4‐OI was explored. We found that 4‐OI restored the reduced expression of AMPKα phosphorylation and PPARα in the parotid glands of 13 M mice to the levels observed in the 13 M group (Figure 4Q,R). Furthermore, 4‐OI restored Nrf2 expression in the parotid glands of 13 M mice while decreasing the expression of Kelch‐like ECH‐associated protein 1 (Keap1). In addition, the anti‐inflammatory factors heme oxygenase 1 (HO‐1) and NAD(P)H:quinone oxidoreductase 1 (NQO1), which are downstream molecules of the Nrf2 pathway, were also upregulated in 13 M mice after 4‐OI was administered (Figure 4S,T). Overall, these findings indicate that exogenous 4‐OI activates AMPKα/PPARα/Nrf2/NQO‐1/HO‐1 signaling pathways in 13 M mice.

FIGURE 4.

Effect of 4‐OI on the metabolomics and transcriptome profile of aged mice parotid glands and enhancement of AMPKα and Nrf2 pathway in the parotid gland of aged mice by 4‐OI. OPLS‐DA score plot of metabolites detected in (A) 13 M versus 2 M and (B) 13 M + 4‐OI versus 13 M. (C) The Venn diagram illustrates the differentially expressed metabolites that are particular or common within different comparison groups. (D) KEGG pathway classifications on differential metabolites. The vertical axis represents KEGG compound categories, while the horizontal axis shows the number of compounds annotated to each category. Bar colors indicate the primary classification level of the compounds. Volcano plots plot of differential metabolites detected in (E) 13 M versus 2 M and (F) 13 M + 4‐OI versus 13 M. (G) Histogram displaying the numbers of differential metabolites in 13 M versus 2 M and 13 M + 4‐OI versus 13 M. Heatmap showing the variable importance (VIP) scores of the top 30 differential metabolites in (H) 13 M versus 2 M and (I) 13 M + 4‐OI versus 13 M. Bubble plots illustrate the primary metabolic pathways responsible for the enrichment of differential metabolites in (J) 13 M versus 2 M and (K) 13 M + 4‐OI versus 13 M. Volcano plots plot of DEGs detected in (L) 13 M versus 2 M and (M) 13 M + 4‐OI versus 13 M. (N) Histogram of the numbers of up‐regulated and downregulated DEGs in 13 M versus 2 M and 13 M + 4‐OI versus 13 M. Bubble plots illustrate the primary KEGG pathways responsible for the enrichment of DEGs in (O) 13 M versus 2 M and (P) 13 M + 4‐OI versus 13 M. (Q, R) Representative Western blot and quantitative analysis the protein levels of AMPKα, p‐AMPKα, and PPARα in the parotid gland of different groups of mice (n = 3). (S, T) Representative Western blot and quantitative analysis the protein levels of Nrf2, Keap1, NQO‐1, and HO‐1 in the parotid gland of different groups of mice (n = 3). The data are presented as the means ± SD. *p < 0.05, **p < 0.01, ***p < 0.001. ns, not significant.

3.5. Molecular Docking and Molecular Dynamics Simulations Interaction of 4‐OI and AMPKα

4‐OI binds to AMPKα with a −5.6 kcal/mol affinity, establishing hydrogen bonds with SER‐459 and ARG‐475 residues (Figure 5A,B). The two‐dimensional structure revealed binding mode show that 4‐OI's interaction with AMPKαis predominantly based on hydrogen bonding connections established directly or indirectly between water molecules and surrounding amino acids (Figure 5C), supporting the high binding affinity between 4‐OI and AMPKα. The 4‐OI and AMPKα reach a stable state after 10 ns. The ligand binds stably to the site without significant movement (Figure 5D,E). Energy decomposition analysis indicates that the binding free energy during the 0–100 ns period is −21.37 kcal/mol, suggesting that the binding between the two is highly stable (Figure 5F). The residues contributing most significantly include TYR:442, SER:459, and ARG:475 (Figure 5G). The protein adopts a more compact conformation upon ligand binding, increasing its stability (Figure 5H,I). Hydrogen bond interactions contribute to the stability of the binding (Figure 5J). The FEL analysis indicated a dominating single low‐energy basin, indicating a stable conformational state with no large‐scale structural changes (Figure 5K,L). These data collectively indicate that the binding between the 4‐OI and AMPKα is stable in molecular dynamics simulations and may possess biological activity.

FIGURE 5.

The binding mode of 4‐OI with AMPKα through molecular docking and molecular dynamics simulations. (A) A three‐dimensional model of the complex. (B) the specific binding mode between 4‐OI and AMPKα. (C) The two‐dimensional structure of the binding mode during the kinetic simulation of 4‐OI and AMPKα. (D) The root mean square deviation (RMSD) of 4‐OI (blue) with AMPKα (black). (E) The root mean square fluctuation (RMSF) of 4‐OI with AMPKα. (F) Binding free energy components are detailed, including ΔVDWAALS, ΔEEL, ΔEPB, ΔENPOLAR, ΔGGAS, ΔGSOLV, ΔTOTAL. (G) The per‐residue decomposition energy plot of critical amino acid residues. (H–J) The radius of gyration (Rg), solvent accessible surface area (SASA), and hydrogen bond number of 4‐OI with AMPKα. (K, L) Two‐ and three‐dimensional free energy landscape (FEL) plots quantitatively depicting conformational energy states. Energy troughs (dark blue regions) correspond to high‐probability conformational clusters and represent the system's stable states. Energy plateaus or saddle points (yellow/red regions) indicate transition states or energy barriers that separate different conformational clusters, reflecting the kinetic bottlenecks of conformational transitions.

3.6. 4‐OI Attenuates Lipid Accumulation in the Parotid Gland via a PPARα‐Dependent Mechanism

We evaluated whether GW6471, a PPARα antagonist, could effectively reverse the protective effects of 4‐OI in 13 M mice. The PPARα antagonist resulted in a substantial increase in parotid gland damage scores, eliminating the protective effects of 4‐OI on 13 M mice (Figure 6A). Oil Red O staining and BODIPY staining of gland sections confirmed that the number of lipid droplets was significantly lower in the 13 M + 4‐OI group than in the 2 M group, and GW6471 inhibited the protective effects of 4‐OI (Figure 6B). We evaluated the relationship between fatty acid oxidation (FAO) and lipid deposition by analyzing the expression of FAO pathway‐related regulators. CPT1A and ACOX1 are key enzymes that regulate long‐chain fatty acids in mitochondria and β‐oxidation in peroxisomes [35, 36]. IHC revealed decreased ACOX1 and CPT1A expression in the 13 M parotid glands compared with that in the 2 M group, and this reduction was effectively corrected by 4‐OI (Figure 6C). Hence, GW6471 reversed the protective effects of 4‐OI on the glands of 13 M mice. According to the western blot results, GW6471 decreased the expression of FAO pathway‐related regulators, including ACOX1 and CPT1A, in the parotid gland tissues of 13 M mice (Figure 6D,E). Immunofluorescence staining revealed decreased PPARα, ACOX1, and CPT1A expression after GW6471 treatment; however, compared with that in the 13 M + 4‐OI group, COL1 expression in the 13 M + 4‐OI group increased (Figure 6F–H). Furthermore, GW6471 decreased the expression of genes related to the PPARα signaling pathway and its downstream targets, including p‐ACC [37, 38], in the parotid gland tissues of 13 M mice (Figure 6I,J). Thus, 4‐OI appears to alleviate lipid overaccumulation in the parotid gland via the PPARα‐mediated FAO signaling pathway.

FIGURE 6.

PPARα antagonist verified that 4‐OI alleviates parotid gland lipids accumulation via the PPARα‐mediated FAO signaling pathway in aged mice. (A) Representative images and quantitative analysis of hematoxylin and eosin (H&E) staining in the parotid gland of different groups of mice (n = 6). Scale bars: 50 μm (top) and 20 μm (bottom). (B) Representative images and quantitative analysis of Oil Red O staining and BODIPY staining in the parotid gland of different groups of mice (n = 6). Scale bars: 50 μm. (C) Representative IHC staining images and quantitative analysis of the ACOX1 and CPT1A in the parotid gland of different groups of mice. Scale bar: 50 μm. (D, E) Representative Western blot and quantitative analysis the protein levels of ACOX1 and CPT1A in the parotid gland of different groups of mice (n = 3). (F) Representative images of immunofluorescent staining for CPT1A (red), ACOX1 (green), PPARα (pink), and DAPI (blue) in the parotid gland of different groups of mice. Scale bars: 50 μm (top) and 20 μm (bottom). The white line with arrows indicates the region selected for colocalization analysis. (G) Fluorescence intensity profiles of CPT1A (red), ACOX1 (green), and PPARα (pink) along the white line shown in (F). The x‐axis represents the distance (μm) along the white line with arrows, and the y‐axis represents fluorescence intensity (a.u.). A shorter distance between the peak intensities of the three channels indicates a higher degree of colocalization. (H) Representative images of immunofluorescent staining for COL1 (red) and DAPI (blue) and quantitative analysis of relative fluorescence intensity of COL1 in the parotid gland of different groups of mice (n = 6). Scale bars: 50 μm. (I, J) Representative Western blot and quantitative analysis the protein levels of ACC, p‐ACC, and PPARα in the parotid gland of different groups of mice (n = 3). The data are presented as the means ± SD.*p < 0.05 **p < 0.01, ***p < 0.001. ns, not significant.

3.7. Nrf2 Inhibition Effectively Reversed the Protective Effects of 4‐OI

Previous research has identified Nrf2 as a key target that controls the bioactivity of itaconate [39]. As an inhibitor of Nrf2, ML385 binds to Nrf2 and inhibits the downstream expression of its target genes. The 13 M + 4‐OI + ML385 group exhibited significantly increased histological damage and gland damage scores and decreased salivary flow rates, reversing the protective effects of 4‐OI on parotid gland function in 13 M mice (Figure 7A,B). Additionally, ML385 treatment decreased Nrf2, NQO‐1, and HO‐1 expression in parotid gland tissues (Figure 7C,D). The immunofluorescence results were consistent with the western blot results, which revealed that ML385 inhibited Nrf2 expression in tissues (Figure 7E). These results demonstrate that 4‐OI exerts its antiaging effects in a Nrf2‐dependent manner.

FIGURE 7.

Nrf2 inhibitor reverses 4‐OI's restoration of function and structure impairments in parotid gland in aged mice. (A) Representative images and quantitative analysis of hematoxylin and eosin (H&E) staining in the parotid gland of different groups of mice (n = 6). Scale bars: 50 μm (top) and 20 μm (bottom). (B) Salivary flow rates in different groups of mice (n = 6). (C, D) Representative Western blot and quantitative analysis the protein levels of Nrf2, Keap1, NQO‐1, and HO‐1 in the parotid gland of different groups of mice (n = 3). (E) Representative images of immunofluorescent staining for Nrf2 (red) and DAPI (blue) and quantitative analysis of relative fluorescence intensity of Nrf2 in the parotid gland of different groups of mice (n = 6). Scale bars: 50 μm. The data are presented as the means ± SD. *p < 0.05, **p < 0.01, ***p < 0.001. ns, not significant.

3.8. The Alleviation of Parotid Gland Histological Impairment and Gland Fibrosis in Aged Mice by 4‐OI Through the AMPKα/TGF‐β Pathway Was Confirmed Using the AMPKα Inhibitor Compound C

To determine whether AMPKα plays a role in enhancing aged parotid glands, mice were treated with 4‐OI and intraperitoneally injected with CC, an AMPKα inhibitor, during the 4 weeks of 4‐OI treatment. Histological investigations using H&E staining revealed that 4‐OI therapy decreased gland damage scores and parotid gland destruction compared to the aged group. However, CC treatment reversed these effects (Figure 8A). Immunofluorescence staining revealed that KRT5, KRT7, KRT14, and BHLHA15 expression levels were significantly elevated by 4‐OI, whereas their expression was downregulated after AMPKα inhibition (Figure 8B–E).

FIGURE 8.

AMPKα inhibitor reverses 4‐OI's reduction of fibrosis in parotid gland in aged mice. (A) Representative images and quantitative analysis of hematoxylin and eosin (H&E) staining in the parotid gland of different groups of mice (n = 6). Scale bars: 50 μm (top) and 20 μm (bottom). (B) Representative images of immunofluorescent staining for KRT7 (red, a marker for luminal ductal cells), KRT5 (green, a marker for basal ductal cells), and DAPI (blue) and (D) quantitative analysis of KRT7 positive cells and KRT5 positive cells in the parotid gland of different groups of mice (n = 6). Scale bars: 50 μm (top) and 20 μm (bottom). (C) Representative images of immunofluorescent staining for KRT14 (red, a marker for myoepithelial cells), BHLHA15 (green, a marker for mature acinar cells), and DAPI (blue) and (E) quantitative analysis of KRT14 positive cells and BHLHA15 positive cells in the parotid gland of different groups of mice (n = 6). Scale bars: 50 μm (top) and 20 μm (bottom). (F) Representative images and (I) quantitative analysis of Masson staining in the parotid gland of different groups of mice (n = 6). Scale bars: 50 μm (top) and 20 μm (bottom). (G) Representative images and (J) quantitative analysis of Sirius Red staining in the parotid gland of different groups of mice (n = 6). Scale bars: 50 μm (top) and 20 μm (bottom). (H) Representative images of immunofluorescent staining for COL1 (red) and DAPI (blue) and (K) quantitative analysis of relative fluorescence intensity of COL1 in the parotid gland of different groups of mice (n = 6). Scale bars: 50 μm. (L–M) Representative Western blot and quantitative analysis the protein levels of FN1, COL1, and TGF‐β in the parotid gland of different groups of mice (n = 3). (N) Representative Western blot and quantitative analysis the protein levels of αSMA in the parotid gland of different groups of mice (n = 3). (O) Representative images of immunofluorescent staining for αSMA (red), VIMENTIN (green), and DAPI (blue) in the parotid gland of different groups of mice. Scale bars: 50 μm (top) and 20 μm (bottom). The white line with arrows indicates the region selected for colocalization analysis. (P) Fluorescence intensity profiles of αSMA (red) and VIMENTIN (green) along the white line shown in (O). The x‐axis represents the distance (μm) along the white line with arrows, and the y‐axis represents fluorescence intensity (a.u.). A shorter distance between the peak intensities of the two channels indicates a higher degree of colocalization. The data are presented as the means ± SD. *p < 0.05, **p < 0.01, ***p < 0.001. ns, not significant.

Assessment of interstitial fibrosis in the gland using Masson and Sirius Red staining revealed that 4‐OI significantly decreased fibrosis in tissue and that these beneficial effects were blocked by CC (Figure 8F,G,I,J). As expected, the decrease in the average fluorescence intensity of COL1 in the parotid glands of mice treated with 4‐OI was reversed by CC treatment (Figure 8H,K). Consistent with these results, CC reversed the protective effects of 4‐OI on FN1 and COL1 protein levels (Figure 8L,M). Western blotting revealed that CC increased the expression of the downstream target of AMPKα signaling, TGF‐β (Figure 8L,M). Consistent with these findings, the expression of fibrotic proteins such as αSMA and VIMENTIN was significantly inhibited following treatment with 4‐OI, whereas the expression of these proteins increased after AMPKα inhibition (Figure 8N–P). Therefore, these results demonstrate that 4‐OI inhibited gland interstitial fibrosis through AMPKα/TGF‐β signaling pathways.

3.9. Compound C Eliminated the Protective Effect of 4‐OI on Parotid Gland Mitochondrial Function and Inflammation in Aged Mice

AMPKα is a key regulator of energy metabolism. The administration of 4‐OI alleviated ROS production in aged parotid glands. However, CC reversed the protective effect of 4‐OI (Figure 9A,C). An AMPKα inhibitor was used to confirm that 4‐OI treatment improves mitochondrial function by increasing AMPKα activity in the parotid gland. The inhibition of AMPKα expression reduced the MMP in 4‐OI‐treated aged mice, resulting in decreased red fluorescence of JC‐1 (Figure 9B,D). Subsequently, TEM was used to investigate the microstructural changes of mitochondria in parotid gland tissues. In the 13 M group, parotid gland tissues showed severe mitochondrial abnormalities, including irregular size distribution, reduced cristae density, and disturbed cristae structure. Conversely, in the 13 M + 4‐OI group, the microstructural changes of mitochondria were alleviated, with CC reversing the protective effects of 4‐OI (Figure 9E). In addition, the western blot results revealed that 4‐OI treatment increased the expression of mitochondrial biogenesis‐related proteins (PGC‐1α and TFAM) and decreased cytochrome c protein expression. The inhibitory effect of CC on AMPKα expression reversed these phenomena (Figure 9F,G). Thus, our results suggested that 4‐OI appears to rescue mitochondrial function via the AMPKα/PGC‐1α signaling pathway. SIRT1, an upstream enzyme in the AMPKα pathway, regulates cellular metabolism, promotes longevity, and prevents the manifestation of age‐related disorders [40, 41]. To confirm this hypothesis, the effects of 4‐OI on AMPKα/SIRT1 and its downstream pathways, including the NF‐κB pathway and Nrf2 pathway, were examined in aged mice using western blot analysis. In the parotid glands of aged mice, SIRT1 expression levels clearly decreased, but 4‐OI treatment increased p‐AMPKα, SIRT1, and Nrf2 expression levels and decreased Keap1, TLR4, p‐NF‐κB p65, and p‐IκB‐α expression levels. In contrast, cotreatment with 4‐OI and CC reversed the therapeutic effects of 4‐OI on NF‐κB‐mediated inflammation in aged parotid glands by upregulating p‐NF‐κB p65 and p‐IκB‐α expression (Figure 9H,I,K,L). 4‐OI treatment increased Nrf2 and HO‐1 expression in the parotid glands of aged mice, whereas these alterations were inhibited by CC (Figure 9J). To further explore the upstream regulator of Nrf2 upregulation activated by 4‐OI via AMPKα‐mediated GSK3β phosphorylation, Western blotting was performed. According to these findings, GSK3β is significantly phosphorylated in gland tissues after 4‐OI treatment. Furthermore, Nrf2 protein levels and the 4‐OI‐mediated phosphorylation of AMPKα and GSK3β were inhibited when 13 M + 4‐OI mice were cotreated with CC (Figure 9H,I). Additionally, CC prevented 4‐OI from reversing the expression levels of NLRP3 and the SASP cytokine TNF‐α in the 13 M group (Figure 9M,N).

FIGURE 9.

AMPKα inhibitor reverses 4‐OI's alleviation of apoptosis, inflammation, and mitochondrial dysfunction in parotid gland in aged mice. (A) Representative ROS‐induced fluorescence staining images labeled with DHE dye (red) and DAPI (blue), accompanied by (C) quantitative analysis of relative DHE fluorescence intensity in the parotid glands of mice from different groups (n = 6). (B) Representative images and (D) quantitative analysis of JC‐1 staining in the parotid glands of mice from different groups (n = 6). Scale bar: 50 μm (top) and 20 μm (bottom). (E) Representative transmission electron microscopy image of mitochondria in the parotid glands of 2 M, 13 M, 13 M + 4‐OI, and 13 M + 4‐OI + CC groups. Scale bars: 1 μm (top) and 500 nm (bottom). (F, G) Representative Western blot and quantitative analysis the protein levels of PGC‐1α, TFAM, and cytochrome c in the parotid gland of different groups of mice (n = 3). (H, I) Representative Western blot and quantitative analysis the protein levels of AMPKα, p‐AMPKα, SIRT1, GSK3β, p‐GSK3β, Nrf2, and Keap1 in the parotid gland of different groups of mice (n = 3). (J) Representative images of immunofluorescent staining for Nrf2 (red) and DAPI (blue) and HO‐1 (red) and DAPI (blue) in the parotid gland of different groups of mice. Scale bars: 50 μm. (K, L) Representative Western blot and quantitative analysis the protein levels of TLR4, NF‐κB p65, p‐NF‐κB p65, IκB‐α, and p‐IκB‐α in the parotid gland of different groups of mice (n = 3). (M, N) Representative Western blot and quantitative analysis the protein levels of NLRP3, TNF‐α, P16^INK4A, P21, and P53 in the parotid gland of different groups of mice (n = 3). (O) Representative images and quantitative analysis of senescence‐associated β‐galactosidase (SA‐β‐gal) staining in the parotid gland of different groups of mice (n = 6). Scale bars: 50 μm. (P) Representative images of immunofluorescent staining for 8‐OHdG (red) and DAPI (blue) in the parotid gland of different groups of mice. Scale bars: 50 μm. (Q) Representative images of immunofluorescent staining for TUNEL (red) and DAPI (blue) and quantitative analysis of TUNEL positive cells in the parotid gland of different groups of mice (n = 6). Scale bars: 50 μm. The data are presented as the means ± SD. *p < 0.05, **p < 0.01, ***p < 0.001. DHE, dihydroethidium; ns, not significant; ROS, reactive oxygen species.

Additionally, during cellular senescence, SIRT1 modulates cell cycle arrest by suppressing P53 and P21 expression [42, 43]. CC prevented 4‐OI from reducing P16^INK4A, P21, and P53 protein expression in the activated aged group (Figure 9M,N). Similarly, CC prevented 4‐OI from reversing the decrease in the percentage of SA‐β‐gal‐positive cells in aged parotid glands, as determined by SA‐β‐gal staining (Figure 9O). Stress, including inflammation, can induce persistent DNA damage [44], leading to cellular senescence, and we initially examined the expression of DNA damage indicators. The expression level of 8‐hydroxy‐2′‐deoxyguanosine (8‐OHdG) was significantly reduced after 4‐OI treatment, whereas 8‐OHdG expression increased after the inhibition of AMPKα (Figure 9P). The TUNEL staining results demonstrated that the antiapoptotic effect of 4‐OI was significantly inhibited by CC (Figure 9Q). The above results suggested that 4‐OI activated AMPKα/SIRT1 and exhibited an antiaging effect in aged mice.

4. Discussion

In summary, 4‐OI enhanced parotid gland secretory function, reduced gland fibrosis, decreased parotid gland lipid deposition, increased FAO, and ultimately restored parotid gland function in aging mice through the AMPKα/SIRT1/Nrf2 and PPARα signaling pathways. Additionally, the AMPKα, Nrf2, and PPARα inhibitors or antagonists, Compound C, ML385, and GW6471, respectively, reversed the protective effects of 4‐OI, highlighting its considerable therapeutic potential for aged parotid glands.

One of the main pathological mechanisms underlying aging is inflammation. The inflammatory pathway is activated by mitochondrial dysfunction, which can cause the release of proinflammatory substances into the circulation [45]. The accumulation of debris observed during aging is caused by inflammatory substances released by aging cells, which disrupt the function of adjacent normal cells and the microenvironment [46]. As people age, their immune systems undergo changes, such as a decline in adaptive immunity, which can result in elevated inflammation [47]. Itaconate, particularly a cell‐permeable variant such as 4‐OI, is an immunoregulatory metabolite [48]. According to Yang et al. [49] and Li et al. [50], 4‐OI reduces the expression of pyroptosis‐induced gasdermin E N‐terminal (GSDME‐NT) activity and relieves colitis by inhibiting pyroptosis. Although Irg1‐mediated itaconate production is classically associated with macrophages, systemically administered 4‐OI may exert beneficial effects on other epithelial tissues. Irg1 can be upregulated in hepatocytes under stress conditions [51]. Notably, we observed Irg1 upregulation predominantly in parotid gland aged mice. However, despite this endogenous upregulation, it appears insufficient to counteract the sustained inflammatory and oxidative burden associated with aging. Thus, exogenous 4‐OI supplementation likely provides a supraphysiological concentration of itaconate that surpasses the endogenous capacity, achieving effective therapeutic protection. This interpretation is supported by studies showing that 4‐OI can attenuate inflammation independently of tissue itaconate concentrations, acting through direct alkylation of KEAP1 to activate Nrf2 and inhibit NF‐κB signaling [25, 32]. Itaconate secreted from myeloid cells can act in a paracrine manner on adjacent epithelial cells [52, 53]. Furthermore, cell‐permeable 4‐OI has been shown to exert direct anti‐inflammatory and cytoprotective effects in various epithelial tissues, including retinal pigment epithelium and intestinal epithelium [50, 54]. Taken together, the protective effects of 4‐OI observed in our study are likely mediated by both direct actions on parotid epithelial cells and indirect modulation of the local immune microenvironment.

Oxidative stress, which can be detrimental to organisms, is caused by an imbalance in the formation and removal of ROS [55]. Our study revealed that by enhancing Nrf2 expression and decreasing cellular ROS levels, 4‐OI protects the parotid gland from oxidative stress. Consequently, 4‐OI has the potential to protect against oxidative damage. Keap1, the master regulator of Nrf2, is one of the primary targets responsible for the protective effects of itaconate [25, 56]. Itaconate‐modified Keap1 inhibited Nrf2 degradation via the ubiquitin–proteasome system, activating the Nrf2 antioxidant pathway [25]. Previous research has shown that oxidative stress is critical for apoptosis in D‐Gal‐induced aged mice [57]. Almost every type of cell contains NF‐κB, a transcription factor that controls oxidative stress, inflammation, and impairment [58]. Furthermore, age‐related diseases such as cancer, dementia, and heart disease exhibit persistent NF‐κB activation [59]. Our research revealed that 4‐OI reduces inflammation by reducing NF‐κB signaling. Notably, NF‐κB activity and aging are not directly linked; however, this highlights the need for more research into this correlation.

AMPKα, a key kinase essential for energy regulation, is activated to improve mitochondrial function, accelerate fatty acid and cholesterol synthesis, reduce inflammatory responses, and modulate cellular aging [60]. The AMPKα signaling pathway regulates aging by interacting with SIRT1, P53, and NF‐κB, among other molecules. Thus, the AMPKα signaling pathway offers new options for the design of antiaging treatments. SIRT1 is involved in the regulation of inflammation, antioxidative stress, DNA repair, and autophagy. All of these processes are directly related to longevity and metabolic processes, which link SIRT1 to aging‐related disorders [61]. SIRT is a nicotinamide adenine dinucleotide (NAD+)‐dependent enzyme, and the reduction in NAD+ function with age is suggested to play a major role in the senescence‐related decrease in SIRT1 activity [62]. NAD+ is the substrate for SIRT1 synthesis. Research on NAD+ has demonstrated its critical involvement in regulating numerous cellular activities associated with aging [63]. Decreased levels of NAD+ combined with compromised SIRT1 disrupt mitochondrial biogenesis in aging mice [64]. According to Lin [65], PGC‐1α is a coregulated factor that controls mitochondrial metabolism and biogenesis. Our experiments demonstrated that 4‐OI has antiaging effects through the activation of the AMPKα/SIRT1 pathway. 4‐OI increased AMPKα levels, which increased NAD+ levels and thus increased SIRT1 expression. Increased SIRT1 levels led to decreased P53 content and decreased NF‐κB p65 expression, resulting in the downregulation of P16^INK4A and P21 expression and cell cycle arrest. Additionally, 4‐OI increases SIRT1 induction, which activates PGC‐1α to improve mitochondrial biogenesis.

Our research demonstrated that 4‐OI reprogrammed lipid metabolism by increasing β‐oxidation and enhancing mitochondrial function, thereby reducing aging‐stimulated lipid accumulation in the parotid glands of mice. Additional evidence that the aged group of mice accumulated more parotid gland lipid droplets than the 2 M group of mice was obtained by examining gland tissue slides stained with Oil Red O. On the other hand, 4‐OI treatment substantially reduced the accumulation of lipid droplets inside the gland. The disturbances in phospholipids, eicosanoids, and fatty acid metabolism observed in aged parotid glands reflected abnormal membrane lipid remodeling and inflammatory mediator metabolism. Previous studies found that itaconic acid could restore lipid balance in various disorders [66, 67, 68, 69]. Western blot analysis was used to evaluate the expression levels of proteins associated with the FAO pathway in the parotid gland to obtain insight into the exact processes through which 4‐OI prevents aging‐related parotid gland dysfunction. An important metabolic pathway for processing FFAs into acetyl‐CoA for energy production is mitochondrial FAO. This pathway is controlled mainly by PPARα, which stimulates the transcription of genes related to peroxisomal and mitochondrial β‐oxidation [70, 71]. FFA transport into mitochondria is facilitated by CPT1, the rate‐limiting enzyme in FAO, which is transcriptionally controlled by PPARs [72]. By phosphorylating and activating ACC, AMPKα, a key energy sensor, reduces malonyl‐CoA, an allosteric regulator of CPT1, which in turn increases FAO and inhibits de novo lipogenesis, hence modulating lipid metabolism [73]. Notably, total ACC protein levels remained unchanged between 2 M and 13 M parotid glands, suggesting that de novo lipogenesis was not substantially altered during aging. Therefore, the lipid‐lowering effect of 4‐OI is most likely attributable to restoration of FAO, rather than suppression of lipogenesis. Moreover, FAO‐related protein expression was markedly increased by 4‐OI therapy, suggesting that the reduced FAO activity observed in the aged parotid gland was successfully restored. In particular, 4‐OI efficiently stimulates the phosphorylation of AMPKα and its downstream target ACC [38], indicating activation of lipid metabolism. The expression of important regulators of mitochondrial fatty acid β‐oxidation, CPT1A and PPARα, markedly increased after 4‐OI treatment. All of these results point to the dual regulatory actions of 4‐OI, which improve parotid gland lipid homeostasis in lipid accumulation in aged parotid gland by promoting FAO.

As a carboxylic acid, itaconate is highly polar and cannot easily pass through cell membranes [25], indicating that it is not appropriate for use in both in vitro and in vivo models. 4‐OI, a cell‐permeable itaconate derivative, was created as an appropriate itaconate substitute to overcome these limitations. However, after itaconate supplementation, we did not assess the amount of itaconate present in the gland tissues. This information might have offered a more thorough understanding of the connection between endogenous itaconate production and exogenous itaconate supplementation, and further experiments are needed to investigate potential correlations. Although our molecular docking and molecular dynamics simulations suggested a stable physical interaction between 4‐OI and AMPKα, these in silico findings needed biological experiment to verify.

This study demonstrated that 4‐OI supplementation ameliorates parotid gland secretion dysfunction, ROS accumulation, and fibrosis in 13 M mice via the AMPKα/SIRT1/Nrf2 and AMPKα/TGF‐β pathways. Furthermore, 4‐OI may alleviate lipid deposition in 13 M mice by increasing parotid gland FAO, promoting the expression of proteins associated with FAO, and activating the AMPKα/PPARα/CPT1A signaling pathway. When translated to the clinical setting, 4‐OI supplementation may represent a potentially effective medication option for managing aging‐induced salivary hypofunction.

Author Contributions

Yun Chen: writing – original draft, methodology, investigation, formal analysis, validation, funding acquisition, conceptualization. Tianyuan Gong: investigation, formal analysis, validation. Zhongyan Shan: investigation, validation, formal analysis. Jing Lai: investigation, validation, formal analysis. Yiming Xu: investigation, validation, formal analysis. Han Zhao: methodology, investigation, formal analysis, project administration, funding acquisition, conceptualization. Lilei Zhu: writing – review and editing, supervision, funding acquisition, supervision, resources, project administration, conceptualization.

Funding

This work was supported by the National Natural Science Foundation of China (82401225), the health commission of Changsha (KJ‐A2023013), and Natural Science Foundation Hunan Provincial (2024JJ9538 and 2026JJ81700).

Consent

The authors have nothing to report.

Conflicts of Interest

The authors declare no conflicts of interest.

Supporting information

Appendix S1: fsb272335‐sup‐0001‐AppendixS1.zip.

Data Availability Statement

The molecular docking and molecular dynamics simulations in this study are available at Mendeley Data. https://data.mendeley.com/preview/st3895yvxb?a=3563086a‐2ec7‐4d06‐ba76‐9689adcc672d. The raw sequence data reported in this study are available at the National Genomics Data Center (https://ngdc.cncb.ac.cn/omix/releaseList) (PRJCA063582). The raw data files are under embargo and will be made publicly available upon publication.