Progranulin deficiency induces premature vascular senescence and dysfunction

Abstract

Significance:

Cellular senescence—an age-associated state characterized by irreversible cell cycle arrest and pro-inflammatory signaling—contributes to vascular dysfunction and cardiovascular disease. However, the molecular mechanisms linking senescence to vascular impairment remain incompletely defined. Progranulin (PGRN) is a multifunctional protein involved in inflammation, lysosomal function, and cellular homeostasis, but its role in vascular aging is not well understood.

Methods:

We assessed PGRN expression in human and mouse arteries and in senescent vascular smooth muscle cells (VSMCs). Vascular function was examined in PGRN-deficient (PGRN^−/−) mice. Cellular senescence was pharmacologically targeted using the senolytic agents navitoclax (ABT-263) and fisetin, and vascular phenotypes were evaluated in adult (6-month-old) and aged (18-month-old) mice.

Results:

PGRN expression increased with age in human and mouse arteries and strongly correlated with p21 expression. In adult mice, PGRN deficiency induced endothelial dysfunction, enhanced vasoconstriction, and promoted vascular inflammation and remodeling. Transcriptomic profiling of PGRN^−/− VSMCs revealed a senescence-associated signature characterized by impaired oxidative phosphorylation, epigenetic dysregulation, and enrichment of collagen-related pathways. Senolytic treatment improved endothelial-dependent relaxation but increased vascular contractility in PGRN^−/− mice. In aged mice, PGRN deficiency exacerbated vascular dysfunction, remodeling, and renal injury without further increases in senescence markers, consistent with premature rather than progressive vascular senescence.

Conclusion:

PGRN deficiency promotes premature vascular dysfunction through coordinated mitochondrial, epigenetic, inflammatory, and structural mechanisms. These findings identify PGRN as a key modulator of vascular homeostasis and suggest that impaired PGRN signaling may predispose to early-onset vascular and cardiorenal dysfunction.

New & Noteworthy

This study identifies progranulin (PGRN) as a key regulator of premature vascular dysfunction. Using human and mouse arteries and integrated multi-omics analyses, we show that PGRN deficiency induces early vascular senescence, mitochondrial dysfunction, epigenetic remodeling, and fibrosis. Senolytic treatment uncovers divergent roles of endothelial versus smooth muscle cell senescence in vascular function, highlighting PGRN as an important modulator of vascular homeostasis during aging.

INTRODUCTION

Aging is a major risk factor for cardiovascular diseases (CVD), which significantly contribute to global mortality (1, 2). As individuals age, arteries experience pathological adaptation that leads to various vascular conditions, including hypertension, atherosclerosis, and coronary artery disease (3). Among the mechanisms driving these changes, cellular senescence—defined as a state of irreversible cell cycle arrest accompanied by pro-inflammatory signaling and metabolic dysfunction (4–6)—has emerged as a key contributor to vascular dysfunction (5, 7, 8). Despite significant progress, the molecular pathways linking senescence to vascular impairment remain incompletely defined.

Progranulin (PGRN) is a multifunctional glycoprotein involved in inflammation, lysosomal function, and cellular homeostasis (9, 10). Alterations in PGRN signaling have been implicated in neurodegenerative diseases, including frontotemporal dementia (FTD) (11, 12), as well as in metabolic and inflammatory disorders (9, 13). We and others have shown that PGRN plays an important role in vascular biology. Specifically, PGRN deficiency alters vascular tone in both resistance and conduit vessels, with increased vasoconstriction in mesenteric arteries (14) and markedly reduced contractility of aortic vascular smooth muscle cells (VSMCs) (15). These changes are associated with disrupted mitochondrial quality control and autophagy, processes closely linked to cellular senescence and vascular aging (16–18). Notably, PGRN expression has been reported to increase with age in cardiac tissue, and its deficiency exacerbates age-related cardiac remodeling and senescence (19). Together, these observations suggest that PGRN may act as a context-dependent modulator of vascular stress responses, although its role in vascular senescence and aging remains unclear.

Based on these observations, we hypothesized that PGRN regulates vascular dysfunction associated with aging by modulating cellular senescence and related molecular pathways. The objective of this study was to determine whether PGRN influences vascular senescence and function across aging and to define the associated mitochondrial, epigenetic, and inflammatory mechanisms.

METHODS

Mice

Male and female C57BL/6J wild-type (WT, PGRN^+/+) and global PGRN knockout (PGRN^−/−; B6(Cg)-^{Grntm1.1Aidi/J}) mice aged 6 and 18 months were used in this study. All animals were maintained on standard chow with ad libitum access to tap water. The 18-month-old WT mice were obtained from the Translational Research Branch, Division of Aging Biology at the National Institute on Aging (NIA), National Institutes of Health (NIH), through a special grant (BRUDER092724SE). Upon arrival at the University of South Alabama vivarium, these mice were allowed a 2–3-week acclimation period prior to experimentation.

All mice were housed in an AAALAC-accredited facility within the College of Medicine at the University of South Alabama. Euthanasia was performed using CO₂ overdose (20–30% chamber volume per minute). All procedures were approved by the Institutional Animal Care and Use Committee (IACUC) at the University of South Alabama (protocol #2219557) and conducted in accordance with the Guide for the Care and Use of Laboratory Animals.

Senolytic Drug Treatments

Navitoclax:

Six-month-old PGRN^−/− mice received intraperitoneal injections of navitoclax (ABT263) at a dose of 50 mg/kg/day for 28 days (2 cycles of 5 days on and 14 days off) (7, 20). Navitoclax, obtained from Adooq Bioscience-CA-USA, was initially dissolved in dimethyl sulfoxide (DMSO) and then resuspended in corn oil. Six-month-old PGRN^−/− mice receiving corn oil were used as a control group. At the time of euthanasia, blood samples were collected, and tissues were harvested and prepared for subsequent analyses.

Fisetin:

Six-month-old PGRN^−/− mice received intraperitoneal injections of Fisetin at a dose of 100 mg/kg/day for 28 days (2 cycles of 5 days on and 14 days off) adapted from Mahoney et al. (21). Fisetin, obtained from Adooq Bioscience-CA-USA, was initially dissolved in DMSO and then resuspended in corn oil. Six-month-old PGRN^−/− mice receiving corn oil were used as a control group. At the time of euthanasia, blood samples were collected, and tissues were harvested and prepared for subsequent analyses.

Human Coronary Arteries Collection

De-identified human tissues were obtained from cadaveric donor tissues via the Center for Organ Recovery and Education (CORE) under the approval by the University of Pittsburgh Committee for Oversight of Research and Clinical Training Involving Decedents (CORID). A detailed protocol describing tissue collection and handling was published previously(22). Briefly, coronary arteries were recovered from cadaveric organs and stored in cold Belzer UW Cold Storage Transplant Solution (Bridge to Life) at 4°C for transporting. Coronary arteries were excised and washed with a sterile rinsing solution (sterile PBS supplemented with 2.5 μg/mL of fungicide (Gibco, 15290026), 0.05 mg/mL of gentamicin162 (Gibco, 15710064), and 5 μg/mL of plasmocin (InvivoGen, ant-mpt-1) and maintained at −80°C for immunoblotting. Demographic and clinical information for human arterial samples, including age, sex, and cause of death, is provided in Supplementary Table 1.

Vascular Function

Rings from second-order mesenteric resistance arteries were mounted in a wire myograph (Danish Myo Technology) for isometric tension recordings using PowerLab software (AD Instruments) as described before(14, 23). Mesenteric resistance arteries are resistance vessels play a central role in the regulation of peripheral vascular resistance and are particularly susceptible to age-associated vascular dysfunction and remodeling. Briefly, rings (2 mm) were placed in tissue baths containing warmed (37 °C), aerated (95% O₂, 5% CO₂) Krebs Henseleit Solution (in mmol/L: 130 NaCl, 4.7 KCl, 1.17 MgSO₄, 0.03 EDTA, 1.6 CaCl₂, 14.9 NaHCO₃, 1.18 KH₂PO₄, and 5.5 glucose). After 30 minutes of stabilization, curves of tension were performed to adjust the ideal tension for each segment, followed by incubation with potassium chloride (KCl, 120mM). Contractility of mesenteric arteries was tested by cumulative applications of the phenylephrine (PE, 10⁻¹⁰–10⁻⁴ M) and thromboxane A₂ receptor agonist, U46619 (10⁻¹¹–10⁻⁶ M). Vasodilation of mesenteric arteries was tested by cumulative applications of the acetylcholine (ACh, 10⁻¹⁰–10⁻⁴ M) and sodium nitroprusside (SNP, 10⁻¹⁰–10⁻⁴ M). A statistical approach was employed to create a curve that depicts the relationship between the concentrations of agonists and the resulting contraction and dilation. This allows for the evaluation of the maximal effect (Emax) and pD2 values for the agonists. The “Emax” refers to the maximum impact achievable by the agonists, and “pD2” represents the negative logarithm of the molar concentration of the agonists required to achieve 50% of the maximum effect (EC50).

Western Blot

Mesenteric arteries from mice, as well as in coronary artery samples from adult and aged humans were extracted using radioimmunoprecipitation assay buffer (RIPA) buffer (30 mM HEPES, pH 7.4,150 mM NaCl, 1% Nonidet P-40, 0.5% sodium deoxycholate, 0.1% sodium dodecyl sulfate, 5 mM EDTA, 1 mM NaV04, 50 mM NaF, 1 mM PMSF, 10% pepstatin A, 10 μg/mL leupeptin, and 10 μg/mL aprotinin). Total protein extracts were centrifuged at 15,000 rpm/10 min and the pellet was discarded. Proteins from homogenates of mesenteric arteries and kidney (20 μg) were used. Proteins were separated by electrophoresis on a polyacrylamide gradient gel (BioRad Hercules) and transferred to Immobilon-P poly (vinylidene fluoride) membranes. Non-specific binding sites were blocked with 5% skim milk or 1% bovine serum albumin (BSA) in tris-buffered saline solution with tween for 1h at 24 °C. Membranes were then incubated with specific antibodies overnight at 4 °C (Supplementary table 2). After incubation with secondary antibodies, the enhanced chemiluminescence luminol reagent (SuperSignal^™ West Femto Maximum Sensitivity Substrate, Thermo-Scientific #34095, Massachusetts, USA) was used for antibody detection.

Real-Time Polymerase Chain Reaction (qPCR)

mRNA from mesenteric arteries, aorta and kidney were extracted using RNeasy Mini Kit (Quiagen #74106, North Rhine-Westphalia, GER). Complementary DNA (cDNA) was generated by reverse transcription polymerase chain reaction (qPCR) with MultiScribe Reverse Transcriptase (Thermo-Scientific #4319983, Massachusetts, USA). Reverse transcription was performed at 58 °C for 50 min; the enzyme was heat inactivated at 85 °C for 5 min, and real-time quantitative RT-PCR was performed with the PowerTrack^™ SYBR Green Master Mix (Thermo-Scientific #A46109, Massachusetts, USA). Sequences of genes as listed in Supplementary Table 3. Experiments were performed in a CFX Opus 384 Real-Time PCR System (Bio-Rad Laboratories, Inc.). Data were quantified by 2ΔΔ Ct and are presented by fold changes versus the control group. Fold change is an indicative of either upregulation or downregulation.

Histology

The aorta and kidney from PGRN^+/+ and PGRN^−/− mice were collected and placed in a 4% paraformaldehyde (PFA) solution for histologic analysis. After 12 hours in PFA, tissues were placed in 70% ethanol until the day of preparing the samples for histology. Aorta and kidney were embedded in paraffin, then samples were sectioned (10μm) and stained with Masson’s trichrome to analyze the remodeling and structure. Arterial wall cross-sectional area was quantified using the Echo microscope software. Images were acquired in a Keyence microscope (BZ-X series). Aortic fibrosis was evaluated in a blinded manner by five independent observers using a semi-quantitative scoring system (0–4), where 0 indicates no fibrosis and 4 indicates severe fibrosis. The final fibrosis score for each sample was calculated as the average of the five independent evaluations.

Vascular Smooth Muscle Cells Isolation

We conducted the isolation of VSMC from the aortas of male PGRN^+/+ and PGRN^−/− mice(15). The isolation procedure followed a well-established enzymatic dissociation protocol. Following the isolation, the VSMC were cultured in DMEM from Invitrogen Life Technologies. To maintain cell health and preserve their physiological characteristics, the culture medium was supplemented with 10% fetal bovine serum (FBS) obtained from HyClone, along with 100 U/ml penicillin, 100 μg/ml streptomycin, and 10 mmol/L Hepes (pH 7.4) from Sigma-Aldrich. To ensure the viability and functionality of the arterial smooth muscle cells during experimentation, we utilized cells within passages 4 to 8.

Reactive Oxygen Species Quantification Via Lucigenin-Derived Chemiluminescence Assay

As described before(24), VSMC were washed with PBS 1x and harvested in 70-μL lysis buffer (2×10–2 M KH2PO4; 10⁻³ M EGTA, and protease inhibitors: 1μg/mL of aprotinin, 1 μg/mL of leupeptin, and 1 μg/mL of pep-statin). About 50 μL of the sample was added to 175-μL assay buffer (50 mM KH2PO4, 1 mM EGTA, and 150 mM sucrose, pH 7.4 and 5×10⁻⁶ M lucigenin). Then, the first reading was performed and considered as basal reading. Nicotinamide adenine dinucleotide phosphate [NADPH (10⁻⁴ M)] was added to each sample, and the luminescence signal was measured, for 30 cycles of 30 s each, in a FlexSation 3 microplate reader (Molecular Devices, San Jose). Basal buffer readings were subtracted from the respective sample readings.

Cell Cycle Analyzes

Cell cycle arrest in PGRN+/+ and PGRN^−/− VSMCs was assessed using the Cell Cycle Phase Determination Kit according to the manufacturer’s instructions (Cayman, MI, USA).

Senescence-associated β-galactosidase (SAβG) staining

SAβG staining in PGRN^+/+ and PGRN^−/− VSMC was performed using a Senescence Detection Kit (MilliporeSigma, KAA002)(25). VSMC cultured in 12-well plates were washed twice with PBS and fixed for 10 minutes using the diluted fixation solution provided in the kit, following the manufacturer’s instructions. After fixation, cells were washed three times with PBS, and 1 mL of freshly prepared SAβG staining solution was added to each well. Plates were then wrapped in aluminum foil to protect from light and incubated at 37°C for 24 hours. Following incubation, cells were washed three times with PBS, and images were captured using an Echo microscope.

Senescence Induction In VSMC

To evaluate PGRN gene expression in aged VSMCs, cells were exposed to 10 Gy of gamma irradiation(26) and analyzed five days post-irradiation. Expression of p21 gene expression was assessed to confirm the senescent/aged VSMC phenotype.

Sequencing and Gene Count Generation

Total RNA was isolated from PGRN+/+ and PGRN^−/− VSMCs using the RNeasy Mini Kit (Qiagen #74106, Germany). RNA quality and concentration were assessed by NanoDrop spectrophotometry. Samples were shipped on dry ice to Novogene (Sacramento, CA, USA) for sequencing.

Triplicate samples per group were processed. mRNA was enriched using poly-T oligo magnetic beads, fragmented (150–350 bp), and reverse-transcribed to generate cDNA libraries. Libraries were quantified (Qubit) and size-validated (Bioanalyzer), pooled, and sequenced on an Illumina platform to generate paired-end reads. Adapter sequences were removed, and clean reads were aligned to the mouse reference genome using HISAT2 (v2.0.5). Gene counts were generated using feature Counts (v1.5.0).

Transcriptomic Data Processing and Differential Expression Analysis

Raw counts were analyzed in R (v4.3.1) using DESeq2 (v1.42.0), comparing wild type (WT; n = 3) and knockout (KO; n = 3). Based on principal component analysis, one KO replicate was excluded. The remaining samples showed consistent clustering and concordant differential expression patterns, supporting dataset robustness.

Normalized counts were obtained using size-factor normalization and scaled for visualization. For heatmaps, representative WT samples were matched to KO samples for clarity, while all WT samples were included in differential expression testing.

Senescence-associated genes were identified using Reactome pathways (msigdbr v7.5) containing the term “senescence,” mapped to mouse orthologs. These gene sets were collapsed into a non-redundant list and intersected with significantly differentially expressed genes (FDR < 0.05). Identified genes were categorized into functional groups including cell cycle arrest, chromatin remodeling, DNA damage/stress signaling, SASP regulation, and proteostasis. Expression patterns were visualized using ComplexHeatmap (v2.26.1).

Analysis of Differential DNA Methylation in Mesenteric Arteries

DNA was extracted from mesenteric arteries of 6-month-old PGRN+/+ (n = 4) and PGRN^−/− (n = 3) mice using the DNeasy Mini Kit (Qiagen). DNA was fragmented (g-TUBEs, Covaris), and 2 μg was used for library preparation with the Oxford Nanopore Ligation Sequencing Kit V14. Libraries were sequenced on MinION flow cells, with reloading to optimize output.

Reads were basecalled using dorado (v1.3.0) with a modified-base detection model (5mC/5hmC) and aligned to the mm10 genome. Replicates were merged (samtools v1.22.1) to ensure comparable coverage. Methylation frequencies were calculated using modkit (v0.5.0), and differential methylation was assessed at CpG sites, promoters (−5 kb to +500 bp), and CpG islands (±4 kb).

Sites were filtered for coverage (≥10 reads per condition) and effect size (Cohen’s h ≥ 0.8). Annotated genomic regions included promoters, gene bodies, CpG islands, and regulatory elements from the SCREEN database. Methylation signals were computed in 50 bp bins and converted to bigWig format (bedtools v2.28.0) for visualization using deepTools (v3.5.6).

Integrative Transcriptomic and DNA Methylation Analysis

To evaluate the relationship between DNA methylation and transcriptional changes following PGRN loss, base-resolution methylation data were integrated with RNA-seq results at gene and pathway levels.

At the gene level, CpG sites were annotated to gene-associated regions using annotatr (v1.36.0). A representative methylation score per gene was defined as the CpG site with the largest absolute methylation difference (Δmethylation = KO − WT). These values were compared to RNA-seq log₂ fold changes. Gene-level coverage was assessed by comparing RNA expression with the number of CpG sites meeting coverage thresholds (≥10x).

To assess robustness, genes were stratified by methylation effect size (Cohen’s h ≥ 0.8, 0.5, 0.3). Overlap between methylation-associated genes and differentially expressed genes was quantified using the Jaccard index and Fisher’s exact test. Associations between methylation and gene expression were evaluated using Spearman correlation.

To assess methylation patterns around transcription start sites (TSSs), genome-wide 5mC signals were aggregated and visualized using deepTools (computeMatrix, ±3 kb from TSS; 50 bp bins), and average profiles were generated.

At the pathway level, genes were ranked by Wald statistic (RNA-seq) or methylation change (Δmethylation). Gene set enrichment analysis (GSEA) was performed using fgsea (v1.36.2) with Gene Ontology Biological Process (GO:BP) gene sets (msigdbr, C5 collection).

Pathway overlap between datasets was assessed using Jaccard index and hypergeometric testing. Concordance in enrichment direction was determined from normalized enrichment scores (NES), and correlations between datasets were assessed using Spearman analysis. Over-representation analysis (ORA) was performed on shared high-confidence genes (h ≥ 0.8), and pathways were ranked by false discovery rate (FDR).

Statistical Analysis

For comparisons of time-course experiment, one-way analysis of variance (ANOVA) followed by the Tukey post-test was used. Group comparisons were performed using Student’s t test to assess differences between PGRN^+/+ and PGRN^−/− mice at 6 and 18 months of age, as well as to compare age-related changes within each genotype (6- vs. 18-month-old PGRN^+/+ or PGRN^−/− mice).

Correlation analyses were performed to assess the relationship between PGRN and p21 expression in human and mouse arterial samples. Associations between variables were evaluated using Spearman rank correlation coefficients (rs), as this nonparametric approach does not assume normal distribution. Correlation coefficients and corresponding two-tailed p values were calculated for each dataset independently, and exact p values are reported. The number of samples analyzed (n) is indicated in the respective figure legends. Statistical significance was defined as p < 0.05.

Concentration–response curves were analyzed using a nonlinear interactive fitting program, creating a curve that depicts the relationship between the concentrations of agonists and the resulting contraction/relaxation. This allows for the evaluation of the maximal effect [Emax, refers to the maximum effect achievable by potassium chloride (KCl)] and pD2 values [represents the negative logarithm of the molar concentration of agonist required to achieve 50% of the maximum effect (EC50)].

All analyzes were performed using GraphPad Prism 8.0. (GraphPad Software Inc., San Diego, CA). The level of significance for all variables was 5%.

RESULTS

Vascular aging is associated with increased vascular PGRN

We first assessed PGRN expression in the aging vasculature. PGRN levels were markedly elevated in mesenteric and coronary arteries from aged mice and humans, alongside increased expression of p21, a key marker of cellular senescence. Consistent with this association, PGRN and p21 expression showed a strong positive correlation across both species (Figure 1A–F). The specificity of the PGRN antibody was confirmed by the absence of signal in arteries from PGRN^−/− mice (Figure 1). Moreover, a time-course analysis of mesenteric arteries from mice aged 4 to 12 months revealed a progressive rise in PGRN expression that closely paralleled the increase in p21 (Figure 1G). This elevation in PGRN was not influenced by sex, as aged female mice and humans showed similar increases. In addition, VSMC exposed to irradiation—a known inducer of senescence—also displayed upregulation of both PGRN and p21 (Figure 1H). Together, these findings strongly suggest that PGRN is associated with vascular aging. However, whether this increase represents a protective or maladaptive response remains to be determined.

Figure 1. Vascular aging is associated with progranulin (PGRN) expression.

(A) p21 and (B) PGRN protein expression in mesenteric arteries from male and female mice at 6 months (adult) and 18 months (aged). (C) p21 and PGRN protein expression correlation from male and female mice at 6 months (adult) and 18 months (aged). (D) p21 and (E) PGRN protein expression in coronary arteries from male and female adult and aged human subjects. (F) p21 and PGRN protein expression correlation from male and female adult and aged human subjects. (G) p21 and PGRN protein expression in mesenteric arteries from male and female mice aged 4–12 months old (mo). (H) p21 and (I) PGRN mRNA expression in vascular smooth muscle cells (VSMCs) exposed or not to irradiation (12 Gy). Data are shown as mean ± SEM, N = 3–8. *P < 0.05 vs. 6-month mice, adult subjects, or control cells. ^#P < 0.05 vs. 4- and 6-month mesenteric arteries. Correlations between PGRN and p21 expression were assessed using Spearman correlation analyses. Correlation coefficients (r) and p values are shown in the respective panels (n = 16).

PGRN deficiency induces a vascular senescence-associated transcriptional and epigenetic program

To investigate the role of PGRN in vascular smooth muscle cell (VSMC) biology, we performed RNA sequencing in VSMCs isolated from PGRN+/+ and PGRN^−/− mice. Loss of PGRN induced extensive transcriptomic remodeling, with 2,863 genes differentially expressed, including 654 genes with a log₂ fold change >1 (Figure 2A).

Figure 2. Progranulin (PGRN) deficiency alters vascular transcriptome and epigenetic signatures.

(A) Volcano plot of differentially expressed genes (DEGs) in vascular smooth muscle cells (VSMCs) from male PGRN+/+ (WT) and PGRN−/− (KO) mice. Genes passing |log₂FC| > 1 and FDR < 0.05 are highlighted, with significance and fold-change thresholds indicated by color. (B, E) Heatmaps showing normalized expression of significant (FDR < 0.05) DEGs that intersected with annotated senescence-related pathways (B) and pathways involved in regulating DNA methylation (E). Columns represent individual WT (blue) and KO (red) replicates; row splits denote functional categories. Colors are reflective of normalized z-scores (−2 to 2), with red indicating higher relative expression and blue indicating lower relative expression. (C) Stacked bar plots summarizing differentially methylated CpG sites between WT and KO mesenteric arteries. Top: Number of high-effect (lower bound of Cohen’s h > 0.8) CpGs assigned to each labeled regulatory category. Bottom: Mean methylation differences (KO − WT, percentage points) for the same categories. Bar color reflects the magnitude of methylation loss in KO, with darker shades indicating larger decreases. (D) Heatmaps depicting the average fraction of modified cytosines (5mC + 5hmC) in mesenteric arteries obtained from wild-type (WT) and progranulin-deficient (KO) mice. Signal is plotted over CpG islands (left) and promoters (right) found to be differentially methylated between the two conditions, sorted by average KO signal. Each row represents an individual promoter or CpG island identified as differentially methylated. For CpG islands, the plotted region is ±4 kb flanking the island center; for promoters the plotted region is −5 kb to +0.5 kb relative to the transcription start site (TSS).

To determine whether these changes reflect a senescence-associated phenotype, we performed an unbiased pathway-based analysis of genes linked to cellular senescence. Differentially expressed genes were enriched across multiple canonical senescence pathways, including regulators of cell cycle arrest (e.g., Cdkn2a, Cdk4/6, Rb1), chromatin remodeling and epigenetic regulation (e.g., histone variants and Ehmt2), DNA damage and stress signaling (MAPK pathways), and SASP-associated transcriptional programs (e.g., Il6, Jun, Ets1) (Figure 2B). These findings indicate that PGRN deficiency activates a broad senescence-associated transcriptional program.

Given the prominent enrichment of chromatin-related pathways, we next examined whether PGRN deficiency alters the epigenetic landscape. Nanopore-based methylation profiling revealed widespread DNA hypomethylation in PGRN^−/− VSMCs, with marked reductions across promoters, CpG islands, enhancers, and other regulatory regions (Figure 2C–D), consistent with widespread epigenomic remodeling associated with cellular senescence.

To determine the relationship between epigenomic and transcriptional changes, we performed integrative analyses of RNA-seq and DNA methylation datasets. At the gene level, overlap between differentially methylated and differentially expressed genes was limited but statistically significant (Jaccard index = 0.041; p = 0.00124), with no strong global correlation between methylation changes and gene expression (Spearman ρ = −0.067, p = 0.422; Supplementary Figure 1). Increasing the inclusion of lower-effect methylation changes increased gene overlap but reduced signal coherence, indicating that high-effect methylation changes are more biologically meaningful. Consistent with this, genes with the strongest methylation differences exhibited coordinated depletion of 5mC signal around transcription start sites in PGRN-deficient cells (Supplementary Figure 1).

In contrast, integration at the pathway level revealed substantial convergence between datasets. Gene set enrichment analysis identified 118 shared biological pathways between transcriptomic and methylation profiles (Jaccard index = 0.185; p = 1.61 × 10⁻⁸¹), including processes related to vascular development, angiogenesis, cytoskeletal organization, and stress-response pathways such as ferroptosis (Supplementary Figure 2). Notably, many of these pathways exhibited coordinated transcriptional repression alongside methylation changes, suggesting that PGRN deficiency drives structured epigenomic remodeling that converges with transcriptional programs at the level of biological processes rather than individual genes.

Finally, transcriptomic analysis revealed coordinated dysregulation of pathways known to regulate DNA methylation fidelity, including oxidative stress, mitochondrial function, metabolic cofactors, and inflammatory signaling (Figure 2E). Together, these findings support a model in which PGRN deficiency induces a senescence-associated state characterized by global hypomethylation, transcriptional reprogramming, and functional convergence across mitochondrial, epigenetic, and inflammatory pathways.

The senescent phenotype was further validated in mesenteric arteries from 6-month-old PGRN−/− mice, which showed elevated p21 expression (Figure 3A), and in VSMC, where we observed increased SaβG activity and impaired cell cycle progression, with PGRN−/− cells exhibiting G1-phase arrest—a hallmark of cellular senescence (Figures 3B and 3C). In line with our previous work identifying PGRN as a key regulator of mitochondrial quality in VSMCs(15), transcriptomic analysis revealed significant enrichment of genes linked to impaired oxidative phosphorylation in PGRN-deficient cells (Figure 3D). Because mitochondrial dysfunction in oxidative phosphorylation is often coupled to oxidative stress, we next assessed ROS levels. PGRN deletion markedly elevated ROS production in VSMCs, an effect further amplified by antimycin A, a Complex III inhibitor that induces mitochondrial ROS by collapsing the proton gradient and promoting electron leakage (Figure 3E).

Figure 3. Progranulin (PGRN) deficiency induces premature vascular senescence.

(A) p21 protein expression in mesenteric arteries from male 6-month-old PGRN+/+ and PGRN−/− mice. (B) Senescence-associated β-galactosidase (SA-β-gal) accumulation in vascular smooth muscle cells (VSMCs) from PGRN+/+ and PGRN−/− mice. (C) Cell cycle analysis of VSMCs from PGRN+/+ and PGRN−/− mice. (D) Gene Set Enrichment Analysis (GSEA) of the oxidative phosphorylation pathway in VSMCs from male PGRN+/+ and PGRN−/− mice. (E) Reactive oxygen species (ROS) levels measured by lucigenin in VSMCs from male PGRN+/+ and PGRN−/− mice with or without antimycin A treatment (AMA, 20 μM, 30 min). Data are presented as mean ± SEM, N = 3–5. *P < 0.05 vs. PGRN+/+.

Deficiency of PGRN induces vascular dysfunction and a mild vascular inflammation in adult mice

Primarily We evaluated the impact of PGRN deficiency on vascular function, inflammation, remodeling, and renal fibrosis, given the well-established link between vascular dysfunction, cellular senescence, and age-related renal injury, as well as our previous findings of impaired sodium and volume handling in PGRN-deficient mice(14). We found that deficiency in PGRN does not affect the KCl-induced vascular contraction, but it did increase the vascular contractility for PE and TXA2 mimetic (Figures 4A–B). It further induced endothelial dysfunction, characterized by an impaired vasodilation response to ACh, without affecting endothelial Nitric Oxide Synthase (eNOS) expression (Figure 4C), but did not affect the endothelium-independent vasodilation (SNP response) (Figure 4B).

Figure 4. Progranulin (PGRN) deficiency impairs vascular function and promotes inflammation in mesenteric arteries.

(A) KCl-induced contraction and (B) concentration–response curves (CRCs) to phenylephrine, U46619, acetylcholine (ACh), and sodium nitroprusside (SNP) in mesenteric arteries from male 6-month-old PGRN+/+ and PGRN^−/− mice. (C) Gene expression of endothelial nitric oxide synthase (eNOS). (D-E) Gene expression of structural and inflammatory markers in mesenteric arteries. (F) Gene set enrichment analysis (GSEA) highlighting collagen-related pathways in VSMCs from PGRN+/+ and PGRN^−/− mice. (G) Arterial wall cross-sectional area, (H) fibrosis score, (I) representative Masson’s trichrome staining, and (J-K) gene expression of structural and inflammatory markers in aortas from male 6-month-old PGRN+/+ and PGRN^−/− mice. Data are shown as mean ± SEM (N = 4–7 per group). *P < 0.05 vs. PGRN+/+.

Transcriptomic analysis of VSMCs from PGRN+/+ and PGRN^−/− mice revealed significant enrichment of extracellular matrix and collagen-related pathways, including collagen fibril assembly, biosynthesis, modification, and crosslinking (Figure 4D). These findings suggested that PGRN deficiency may promote vascular remodeling and fibrosis.

To validate these transcriptomic predictions in vivo, we next assessed markers of vascular phenotype and inflammation in mesenteric arteries. While PGRN deficiency did not alter α-smooth muscle actin (αSMA) or collagen 3α1 (Col3α1) expression, it significantly increased the expression of inflammatory markers, including intercellular adhesion molecule-1 (ICAM-1) and interleukin-1β (IL-1β), with no changes observed in tumor necrosis factor-α (TNF-α) or vascular cell adhesion molecule-1 (VCAM-1) (Figure 4E–F).

Given the established link between vascular dysfunction, inflammation, and end-organ damage, we next evaluated structural remodeling in the aorta and kidney. At 6 months of age, PGRN^−/− mice did not exhibit increased arterial wall cross-sectional area; however, they showed enhanced collagen deposition (Figure 4G–I), increased αSMA expression, and elevated inflammatory markers, including VCAM-1, IL-1β, and TNF-α (Figure 4J–K). These changes suggest that PGRN deficiency promotes early vascular inflammation and fibrotic remodeling, consistent with dysregulated extracellular matrix dynamics.

Furthermore, PGRN−/− mice exhibited increased renal mass, elevated fibrotic markers with no effects on renal inflammation, and higher expression of neutrophil gelatinase-associated lipocalin (NGAL), a marker of kidney disease progression (Supplementary Figure 3A-C).

Elimination of vascular senescent cells improves endothelial function but exacerbates vascular contraction of mesenteric arteries

To investigate the contribution of vascular senescence to vascular dysfunction, we first treated 6-month-old PGRN^−/− mice with the senolytic agent navitoclax (ABT-263) (Figure 5A). This treatment effectively reduced vascular senescence, as evidenced by a significant decrease in mesenteric p21 expression (Figure 5B). Despite this reduction, navitoclax further enhanced vascular contractile responses to KCl (Figure 5C), phenylephrine (PE), and U46619 (Figure 5D). Notably, navitoclax restored endothelial function, as indicated by improved endothelium-dependent vasodilation, without affecting responses to the endothelium-independent vasodilator sodium nitroprusside (SNP) (Figure 5D).

Figure 5. Navitoclax, a senolytic drug, modulates vascular function in progranulin (PGRN)–deficient mice.

(A) Schematic of Navitoclax (ABT-263, 50 mg/kg/day) treatment. (B) p21 protein expression. (C) KCl-induced contractility. (D) Concentration–response curves (CRCs) to phenylephrine, U46619, acetylcholine (ACh), and sodium nitroprusside (SNP). All experiments were performed in mesenteric arteries from male PGRN−/− mice treated with vehicle (5% DMSO, 95% corn oil) or Navitoclax. Data are shown as mean ± SEM, N = 4–5. *P < 0.05 vs. PGRN−/− vehicle.

To determine whether these effects were specific to navitoclax or reflected a broader consequence of reducing senescence burden, we next treated PGRN^−/− mice with a second senolytic agent, fisetin (Figure 6A). Fisetin treatment recapitulated the effects of navitoclax, reducing p21 expression, improving endothelial function, and further increasing vascular contractility (Figure 5B–D).

Figure 6. Fisetin, a senolytic drug, modulates vascular function in progranulin (PGRN)–deficient mice.

(A) Schematic of Fisetin (100 mg/kg/day) treatment. (B) p21 protein expression. (C) KCl-induced contractility. (D) Concentration–response curves (CRCs) to phenylephrine, U46619, acetylcholine (ACh), and sodium nitroprusside (SNP). All experiments were performed in mesenteric arteries from male PGRN−/− mice treated with vehicle (5% DMSO, 95% corn oil) or Fisetin. Data are shown as mean ± SEM, N = 4–6. *P < 0.05 vs. PGRN−/− vehicle.

Together, these findings indicate that senolytic treatment reduces senescence-associated signaling and improves endothelial function, while paradoxically augmenting vascular contractility in PGRN-deficient mice.

Deficiency of PGRN exacerbates aging-induced vascular injury

As expected, aged male PGRN^+/+ mice (18 months old) displayed marked vascular and endothelial dysfunction, accompanied by significant changes in inflammatory, remodeling, and senescence markers in the mesenteric arteries, aorta, and kidneys, compared to adult male PGRN^+/+ mice (6 months old) (Supplementary figures 4 and 5). To determine whether exacerbated senescence could further worsen the vascular phenotype in the absence of PGRN, we compared 18-month-old PGRN^+/+ and PGRN^−/− mice. While PGRN deficiency did not alter KCl-induced vascular contractility (Figure 7A), it significantly enhanced vasoconstrictive responses to PE and U46619 (Figure 7B). In addition, PGRN^−/− mice displayed aggravated endothelial dysfunction (Figure 7B) followed by increased eNOS gene expression (Figure 7C), and impaired vasodilation in response to SNP (Figure 7A).

Figure 7. Progranulin (PGRN) deficiency does not further increase vascular senescence but aggravates age-associated vascular dysfunction.

(A) KCl-induced contraction and (B) concentration–response curves (CRCs) to phenylephrine, U46619, acetylcholine (ACh), and sodium nitroprusside (SNP) in mesenteric arteries from male 18-month-old PGRN+/+ and PGRN^−/− mice. (C) eNOS gene expression and (D) p21 protein expression in mesenteric arteries. The immunoblots were obtained from the same experiment in which all groups were analyzed on a single membrane. (E–F) Gene expression of structural and inflammatory markers in mesenteric arteries. (G) Arterial wall cross-sectional area, (H) fibrosis score, (I) representative Masson’s trichrome staining, and (J–K) gene expression of structural and inflammatory markers in aortas from male 18-month-old PGRN+/+ and PGRN^−/− mice. Data are shown as mean ± SEM, N = 4–6. *P < 0.05 vs. PGRN+/+.

Interestingly, vascular senescence levels—indicated by p21 expression—remained unchanged between aged PGRN^+/+ and PGRN^−/− mice (Figure 7D). Our data suggest that PGRN deficiency may advance the onset of vascular senescence (premature senescence) rather than further increasing senescence burden at advanced age; however, this remains inferential given we primarily assessed p21 and a limited SASP profile. Similarly, markers of vascular inflammation and remodeling in the mesenteric arteries were not further altered by PGRN deficiency (Figure 7E–F).

In the aorta, we observed increased arterial wall cross-sectional area with no difference for fibrosis (Figure 7G–I) followed by no additional changes in inflammatory or structural markers (Figure 7J–K). To determine whether chronic PGRN deficiency promotes age-associated renal injury, we evaluated renal structure and inflammatory markers in kidneys from aged PGRN+/+ and PGRN^−/− mice. PGRN^−/− mice exhibited significantly increased kidney weight relative to body weight compared with aged WT mice, suggesting renal remodeling. Histological analysis using Masson’s trichrome staining revealed increased renal fibrosis in PGRN-deficient kidneys (Supplementary Figure 6A).

Assessment of renal injury and structural markers demonstrated a trend toward increased kidney injury molecule-1 (Kim-1) expression in PGRN^−/− mice, whereas α-smooth muscle actin (αSMA), collagen 3α1 (Col3α1), podocin, and NGAL expression were not significantly altered (Figure XB). Similarly, inflammatory markers including ICAM-1, VCAM-1, IL-1β, and TNF-α were not significantly different between groups ((Supplementary Figure 6B-C). Together, these findings suggest that aging PGRN^−/− mice develop structural renal abnormalities and early kidney injury despite limited changes in inflammatory gene expression at this stage.

DISCUSSION

Our findings identify PGRN as both a marker and regulator of vascular aging. We demonstrate that PGRN expression increases in mouse and human arteries with age, closely paralleling the induction of the senescence marker p21. Loss of PGRN leads to premature activation of vascular senescence pathways, mitochondrial dysfunction, epigenetic remodeling, inflammation, fibrosis, and renal injury—phenotypes that collectively resemble early vascular aging. These observations support a model in which PGRN upregulation represents a compensatory response aimed at preserving vascular integrity during aging.

The increased expression of PGRN in aged arteries and in irradiated VSMCs suggests that PGRN is engaged during vascular stress and may act to counter senescence-associated dysfunction. Whether this upregulation is ultimately protective, maladaptive, or context dependent likely depends on the stage of vascular aging and the balance among mitochondrial dysfunction, inflammation, and chromatin remodeling. Previous studies from our group in vascular biology (14, 15) and others in neurodegeneration(12) and cardiac physiology(19) have shown that PGRN exerts anti-inflammatory, anti-pathological remodeling, and mitochondria and lysosome-stabilizing effects, whereas its deficiency leads to lysosomal dysfunction and neuronal loss. In this context, elevated PGRN during aging may reflect an adaptive attempt to maintain vascular homeostasis under chronic stress, although this compensatory response may become insufficient as aging progresses.

Loss of PGRN in VSMCs induced marked transcriptomic and epigenomic remodeling, with more than 2,000 genes differentially expressed and widespread alterations in DNA methylation patterns. Senescence-associated pathways—including cell cycle regulation, chromatin organization, ubiquitination, DNA damage signaling, and SASP-related pathways—were strongly enriched, supporting the concept that PGRN deficiency disrupts nuclear homeostasis and genomic stability, key hallmarks of aging (27, 28). Importantly, although overlap between differentially expressed genes and differentially methylated regions was limited at the individual gene level, substantial convergence emerged at the pathway level. Shared biological programs included vascular development, extracellular matrix remodeling, cytoskeletal organization, and stress-response pathways. These findings suggest that transcriptional and epigenomic remodeling converge functionally despite incomplete gene-by-gene correspondence. Moreover, depletion of 5mC signal around transcription start sites raises the possibility that altered methylation landscapes contribute to long-term chromatin instability and transcriptional responsiveness in the absence of PGRN.

Our transcriptomic data also highlight several mechanisms that may underlie the reduced DNA methylation observed in PGRN-deficient cells. PGRN−/− VSMCs exhibited clear evidence of oxidative stress—consistent with our previous work demonstrating mitochondrial-driven ROS disruption(15)—and reduced antioxidant capacity, creating a redox environment unfavorable for maintaining cytosine modifications. Under such conditions, ROS can oxidize 5-methylcytosine and related bases, generating lesions that are removed by base-excision repair and replaced with unmodified cytosines(29). This repair-mediated cytosine turnover offers a parsimonious explanation for the broad loss of both methylation and hydroxymethylation in PGRN-deficient cells, independent of an active demethylation program. Given that mitochondrial stress is both a cause and amplifier of premature senescence(30), these redox-driven effects likely link mitochondrial dysfunction to the widespread hypomethylation and accelerated aging phenotype observed in the absence of PGRN.

In parallel, several pathways that support methylation fidelity—including metabolic cofactors, mitochondrial stability, and inflammatory signaling(31, 32)—were also disrupted, creating conditions in which both maintenance methylation and TET-dependent cytosine modification are likely impaired. Because cellular senescence is fundamentally associated with extensive epigenetic remodeling, including context-dependent alterations in DNA methylation patterns(33), the coordinated dysregulation of these processes in PGRN-deficient cells provides a unifying framework. Although the precise mechanism remains uncertain, the combined transcriptomic and methylation patterns are consistent with a model in which stress-driven cytosine turnover and impaired methylation maintenance together contribute to the widespread CpG hypomethylation observed in the absence of PGRN.

Adult PGRN^−/− mice exhibited early vascular dysfunction characterized by enhanced vasoconstriction and impaired endothelial-dependent relaxation. These functional abnormalities were accompanied by increased inflammatory signaling and extracellular matrix remodeling, including elevated ICAM-1 and IL-1β expression, enrichment of collagen-related pathways, increased collagen deposition, and higher αSMA expression. Together, these findings indicate that PGRN is required to preserve endothelial function and restrain vascular inflammation and fibrotic remodeling. Notably, our integrated omics analyses identified activation of collagen biosynthesis and extracellular matrix pathways in PGRN-deficient VSMCs, extending previous observations and providing new mechanistic insight into how loss of PGRN contributes to vascular fibrosis. Consistent with these findings, previous studies have shown that although alterations in PGRN levels are often associated with fibrotic remodeling(34), the broader literature suggests that PGRN functions predominantly as an anti-fibrotic factor across several organs—including skin(35), liver(36), heart(37), and lung(38)—with effects that remain highly context-dependent. Our findings extend this protective role to the vasculature.

End-organ analyses further demonstrated increased kidney mass, fibrosis, and NGAL expression in aged PGRN-deficient mice, suggesting that vascular dysfunction and remodeling extend to the renal compartment. These findings are consistent with previous studies demonstrating protective roles for PGRN in renal injury models, including diabetes and ischemia/reperfusion injury (39, 40). Because the kidney is highly susceptible to age-related vascular dysfunction and microvascular injury, these results support the concept that vascular senescence contributes to progressive cardiorenal damage during aging. Importantly, our previous work demonstrated altered sodium and water handling in young PGRN-deficient mice prior to overt renal injury(14), supporting the possibility that vascular dysfunction precedes and contributes to subsequent renal pathology.

To dissect the contribution of vascular senescence to these phenotypes, we treated PGRN−/− mice with the senolytic agent navitoclax. Senolytic therapy effectively lowered p21 expression and improved endothelial-dependent vasodilation, confirming a detrimental role of endothelial senescence in impaired relaxation. Surprisingly, however, navitoclax also further increased vascular contractility. Importantly, these findings were recapitulated using a second senolytic agent, fisetin, indicating that this phenotype is not specific to navitoclax. These divergent effects highlight that senescent cells within the vascular wall do not exert uniform functional consequences. Our findings raise the possibility that senescent VSMCs may influence vascular contractile tone; however, this interpretation is hypothesis-generating and requires direct experimental validation. Consistent with this possibility, recent evidence indicates that senescent VSMCs exhibit reduced mechanical contractility(41), which may act as an intrinsic brake on vasoconstriction during vascular disease progression. Because navitoclax is not cell-type–specific, it likely eliminated senescent endothelial cells, VSMCs, and potentially perivascular cells, making the net effect a composite of different cellular contributions. This complexity may help explain the mixed vascular outcomes reported with senolytic therapies in both preclinical and clinical studies.

Although fisetin reproduced the major vascular effects observed with navitoclax, suggesting that these findings are not solely attributable to navitoclax-specific off-target actions, we cannot exclude Although fisetin reproduced the major vascular effects observed with navitoclax, suggesting that these findings are not solely attributable to navitoclax-specific off-target actions, including thrombocytopenia(42) due to Bcl-2 inhibition. We also did not assess potential navitoclax-induced vascular apoptosis or activation of the caspase-3 pathway(43), which represents an additional limitation given that senolytic agents can trigger apoptosis of stressed or damaged cells. Although we did not evaluate platelet counts or apoptotic markers systematically, these unmeasured effects should be considered when interpreting the functional consequences of senolytic treatment.

In aged mice, PGRN deficiency aggravated endothelial dysfunction and vascular hypercontractility without further increasing p21 expression or amplifying inflammatory or remodeling markers. This pattern suggests premature, rather than progressive, vascular senescence: PGRN^−/− mice appear to reach a senescent threshold earlier in life, after which senescence burden does not substantially increase with chronological aging. Our data support a model in which PGRN deficiency accelerates the onset of vascular senescence and dysfunction, leading to an early plateau rather than progressive worsening with age; however, this remains inferential given we primarily assessed p21 and a limited SASP profile. This trajectory differs from findings in cardiac tissue, where Zhu et al. (2020)(19) reported that aged PGRN-deficient mice exhibit further accumulation of senescence markers and worsened cardiac hypertrophy. Several factors may account for this discrepancy. Senescence is highly tissue-specific, and the heart and vasculature differ markedly in cellular composition, metabolic demand, and regenerative capacity. Thus, cardiac cells may continue to accrue senescent burden with age, whereas vascular cells—particularly VSMCs—may reach a senescence plateau earlier. Additionally, the prior study assessed a broader panel of senescence markers, while our analysis focused primarily on p21 and selected SASP components, which may capture different dimensions of the senescence program. Distinct inflammatory and mitochondrial stress environments in cardiac versus vascular tissue may further influence how PGRN deficiency shapes aging trajectories. Together, these considerations highlight that PGRN-dependent aging phenotypes are organ-specific and that PGRN loss may drive premature vascular senescence without progressive expansion of senescent burden in late life. Nonetheless, aged PGRN^−/− mice exhibited increased aortic cross-sectional area and worsened renal injury, indicating that early-life vascular dysfunction predisposes to exacerbated age-related cardiorenal pathology.

Several limitations of this study should be acknowledged. First, although our findings support a protective role for PGRN in vascular aging, direct gain-of-function experiments in aged vessels were not performed. Second, while p21 expression, SA-β-gal activity, transcriptomic signatures, and epigenetic remodeling strongly support a senescence-associated phenotype, additional senescence markers such as γH2AX, and telomere dysfunction were not systematically evaluated. Third, although our integrative transcriptomic and methylation analyses revealed coordinated pathway-level remodeling, we did not directly assess chromatin accessibility, histone modifications, or methylation-dependent transcriptional regulation. Fourth, the precise mechanisms underlying vascular dysfunction and injury were not directly investigated. In particular, we did not assess the relative contributions of nitric oxide bioavailability, prostanoid signaling, or immune cell infiltration within vascular tissues. In addition, because this study used a global PGRN knockout model, we cannot distinguish the relative contributions of endothelial versus VSMC-specific PGRN deficiency. Future studies using vascular cell-specific knockout models are currently underway to address these questions. Finally, arterial stiffness measurements such as pulse wave velocity were not performed and will be important for defining the biomechanical consequences of PGRN deficiency during aging.

In conclusion, our findings establish PGRN as a mechanistic regulator of vascular aging. PGRN deficiency induces premature vascular senescence and disrupts mitochondrial, epigenetic, inflammatory, and structural homeostasis, leading to early vascular dysfunction, remodeling, and renal injury. Over time, these alterations recapitulate key features of physiological vascular aging but occur earlier and with greater severity. Importantly, the divergent effects of senolytic therapy reveal that endothelial and VSMC senescence may exert distinct functional consequences within the vasculature. Given the ongoing clinical development of PGRN-targeted therapies for frontotemporal dementia, these findings raise important considerations regarding vascular and cardiorenal health in individuals with impaired PGRN signaling. Collectively, our study identifies PGRN as both a biomarker and potential therapeutic target in age-associated vascular disease.

Supplementary Material

SUPPLEMENTAL MATERIAL

Supplemental Tables S1-S3: DOI. https://doi.org/10.6084/m9.figshare.32220750

Supplemental Figs. S1-S6: DOI. https://doi.org/10.6084/m9.figshare.32220465