CAPG enhances adipogenesis and inflammatory cytokine expression in adipocytes

ABSTRACT

The expression of CAPG (capping actin protein, gelsolin-like) is upregulated in visceral white adipose tissue of high-fat diet -fed mice; however, its impact on adipocyte functionality remains unclear. We observed upregulated CAPG expression in the epididymal adipose tissue of high-fat diet-fed mice. To investigate the impact of CAPG on adipocyte differentiation and function, we generated a Capg knockdown 3T3-L1 cell line and induced adipogenic differentiation to mature adipocytes. Adipogenesis was assessed via Oil Red O and BODIPY staining, revealing that Capg knockdown markedly suppressed adipogenesis. Western blot analysis demonstrated that CAPG depletion reduced PPARγ expression. Additionally, Western blot analysis revealed that Capg knockdown significantly enhanced lipid utilization in adipocytes. ELISA and qPCR results further demonstrated that Capg knockdown effectively attenuated inflammatory responses in adipocytes. In conclusion, CAPG promotes adipogenesis and inflammatory responses, suggesting that targeted inhibition of CAPG may represent a potential therapeutic strategy for obesity-associated adipose tissue dysfunction.

Introduction

Adipocytes, the primary cellular constituents of adipose tissue, undergo hypertrophy and functional dysregulation during adipose tissue dysfunction. The most prominent metabolic alteration is enhanced lipolysis, resulting in chronic low-level leakage of free fatty acids (FFAs) into systemic circulation and ectopic adipose deposition [1]. Furthermore, hypertrophic adipocytes secrete pro-inflammatory cytokines, promoting immune cell recruitment and activation, thereby amplifying adipose tissue inflammation [2]. Elevated levels of FFAs and pro-inflammatory cytokines collectively contribute to the initiation and progression of atherosclerotic cardiovascular diseases [3].

Capping actin protein, gelsolin like (CAPG), a calcium-sensitive barbed-end capping protein belonging to the gelsolin family, localizes to both the cytoplasm and nucleus [4]. CAPG was initially identified in macrophages and later found to be widely expressed in kidney, lung, and multiple cell types [5]. In the cytoplasm, CAPG interacts with the actin cytoskeleton, precisely regulating actin filament polymerization, organization, and spatial distribution, thereby modulating cell motility and morphology. Within the nucleus, CAPG influences transcriptional activity by regulating actin nucleation and assembly [6]. Hansson et al. demonstrated that high-fat diet (HFD) induces marked upregulation of CAPG expression in epididymal fat pads of wild-type mice, suggesting a potential role in diet-induced adipose dysfunction [7].

Accumulating evidence indicates that CAPG is overexpressed across diverse cancer types, including breast, lung, and colorectal carcinomas, where it functionally promotes tumour cell proliferation, invasion, and metastatic dissemination [8]. Notably, siRNA-mediated Capg knockdown significantly suppresses DU145 cell proliferation, suggesting that CAPG may also influence preadipocyte proliferation [9]. Additionally, emerging evidence indicates that cytoskeletal proteins regulate adipogenesis and nutrient transport [10,11]. However, the role of CAPG in adipocyte differentiation, metabolic function, and inflammatory responses remains unclear and warrants further investigation. To address these questions, we established a Capg-knockdown 3T3-L1 stable cell line using lentiviral-mediated shRNA, aiming to elucidate CAPG’s role in preadipocyte differentiation, metabolism, and inflammatory responses.

Materials and methods

Animals

Twelve 8–10-week-old male C57BL/6J wild-type (WT) mice (Vital River, Beijing) were randomly divided into two groups (n = 6 per group) by random number table method. One group received normal diet (ND: 1022, Beijing HFK Bioscience), while the other was fed high-fat diet (HFD: Research Diet d12492, 60 kcal% fat, Shanghai Yes Service Biotech) for 20 weeks. Mice were maintained at 20–25°C with 40–70% humidity under 12-h light/12-h dark cycles with ad libitum chow and water. Three mice were housed per cage.

Mice were then anesthetized with 2% isoflurane and subsequently euthanized via blood collection. Measurements for different groups were performed in interleaved sessions to neutralize time-of-day effects. Group assignment codes were accessible solely to the study designer. Epididymal white adipose tissue (eWAT) was isolated for further analysis using established protocols [12]. The animal study protocol was approved by the Animal Ethics Committee of Peking University (Approval No. LA2022278), and the study was designed and reported following the ARRIVE guidelines.

Cell culture

The 3T3-L1 cells (mouse embryonic fibroblasts) were purchased from Procell Life Science & Technology Co., Ltd. (Wuhan, China). Cells were cultured in high-glucose DMEM (C11995500BT, Gibco) (4.5 g/L) supplemented with 10% fetal bovine serum (FBS, 10091155, Gibco). Adipogenic differentiation was induced using complete medium containing 0.5 mM 3-isobutyl-1-methylxanthine (IBMX, I5879, Sigma), 1 μM dexamethasone (DXMS, D6950, Solarbio), and 10 μg/mL insulin (I8040, Solarbio) for 48 hours, followed by another 48-hour induction with complete medium containing 10 μg/mL insulin. This induction cycle was repeated once, after which the cells were maintained in complete medium for an additional 48 hours. Overall, the 3T3-L1 cells underwent a 10-day period during adipogenic differentiation induction.

Establishment of stable Capg-knockdown cell lines

The lentiviruses carrying sh-Capg and sh-Ctrl were purchased from GeneChem (Shanghai, China). The 3T3-L1 cells were transfected with lentiviruses at a multiplicity of infection (MOI) of 100. After 48 hours of infection, the lentivirus-containing medium was replaced with complete medium containing 4 μg/mL puromycin (ST551, Beyotime) for selection, resulting in the generation of stable sh-Capg and sh-Ctrl cell lines.

qPCR

Total RNA was extracted from cells using the Freezol Reagent kit (Vazyme, China) according to the manufacturer’s instructions, with RNA concentration and purity assessed using Nanodrop spectrophotometry (Thermofisher, USA). Subsequently, reverse transcription was performed to synthesize cDNA using the HiScript III All-in-one RT SuperMix Perfect for qPCR kit (Vazyme, China). Finally, quantitative PCR (qPCR) reactions were prepared with TB Green® Premix Ex Taq^TM II FAST qPCR kit (Takara, Japan) and performed on a real-time PCR system (Roche, Switzerland). All primers were commercially synthesized by TsingKe Biotech, with specific primer sequences listed in Table 1.

Table 1.

Primer sequences.

Gene Name	Accession number	Forward (5’-3’)	Reverse (5’-3’)
Tubb5	NM_011655	TAGCCATGAGGGAAATCGTGC	TCACCTCCCAGAACTTAGCA
Capg	NM_001271415	GCTGTGTGGCAAAATCTACATC	GATGAAGCCATCAGCCACTT
Il6	NM_031168	TCCGGAGAGGAGACTTCACA	TGCCATTGCACAACTCTTTTC
Mcp1	NM_011333	CCTGCTGCTACTCATTCACCA	TGAGCTTGGTGACAAAAACTAC
Lep	NM_008493	GAGACCCCTGTGTCGGTTC	CTGCGTGTGTGAAATGTCATTG

Western blot

Protein was extracted by lysis buffer at a ratio of 100:1:1 (RIPA: PMSF: phosphatase inhibitor) according to experimental requirements (R0010; P0100; P1260, Solarbio, China). Cryopreserved eWAT (−80°C) was transferred to liquid nitrogen for embrittlement prior to mortar grinding. Cells were scraped from culture plates using clean cell scrapers. The homogenized powder and cells were lysed in pre-prepared lysis buffer on ice for 30 minutes. The lysates were centrifuged at 12,000 g for 15 minutes at 4°C to collect protein supernatants. Protein concentrations were determined using the BCA (23225, Thermofisher, USA) assay. Samples were diluted with 5× loading buffer (P1040, Solarbio, China), boiled, and stored at −20°C. Polyacrylamide gels (WB2302, Biotides, China) were prepared for electrophoresis, and equal amounts of protein samples were loaded into wells for separation under constant voltage. Following electrophoresis, proteins were transferred onto PVDF membranes (GVWP04700, Millipore, USA) and blocked at room temperature for 2 hours. The membranes were then cut into strips according to target protein molecular weights and incubated overnight with appropriately diluted primary antibodies (CAPG, proteintech 10,194–1-AP, China; β-tubulin, proteintech 10,094–1-AP, China; PPARγ, CST, #2430, USA; CD36, MCE, HY-P81793, USA; ATGL, MCE, HY-P80533, USA; GLUT4, MCE, HY-P80495, USA; DDDDK-Tag, ABclonal, AE169PM, China; Adiponectin, CST, #2789, USA; NF-κB P65, YA267, MCE, USA; phospho-NF-κB P65(Ser536), HY-P80839, MCE, USA). The membranes were then incubated with corresponding secondary antibodies (Anti-mouse IgG, HRP-linked Antibody, Proteintech, SA00001–1, China; Anti-rabbit IgG, HRP-linked Antibody, CST, #7074, USA) for 75 minutes. Finally, protein bands were visualized using ECL ultrasensitive luminescence reagent. Quantification of Western blot bands was performed using ImageJ software (1.53k/Java 1.8.0_172, National Institutes of Health, USA).

Flow cytometry bead-based assay

The cell culture supernatants were collected and diluted to appropriate concentrations, and the levels of cytokines in the supernatants were measured according to the manufacturer’s instructions of the ABplex Mouse Cytokine 15-Plex Assay Kit (RK05203, ABclonal, China).

ELISA

Cell culture supernatants were collected, centrifuged at 2,000 rpm for 5 minutes at room temperature, and interleukin-6 (IL-6) (RK00008, ABclonal, China; SEA079Mu, cloud clone corp, China) and monocyte chemoattractant protein-1 (MCP-1) (887391, Invitrogen, USA) levels were measured in the supernatants using ELISA kits according to the manufacturer’s instructions.

Immunofluorescence protocol

Cells were collected in centrifuge tubes and processed as follows: fixed with 4% paraformaldehyde (G1101, Servicebio, China) for 20 minutes, permeabilized with 0.1% Triton X-100 (T8200, Solarbio, China) for 15 minutes, and blocked with 5% bovine serum albumin (BSA, A8010, Solarbio, China) for 1 hour. The treated cells were then applied to poly-L-lysine-coated glass slides (YA0170, Solarbio, China), with excess blocking buffer aspirated. Primary antibodies were diluted in 5% BSA according to the manufacturer’s recommended ratios, and 100 μL of diluted CAPG and Ki67(28074–1-AP, Proteintech, China) primary antibody was added to each slide. Slides were incubated overnight at 4°C in a humidified chamber. Secondary antibodies (CoraLite594 – conjugated Goat Anti-Rabbit IgG, proteintech, SA00013–4, China) were diluted 1:500 in 5% BSA and stored protected from light. After washing, DAPI (C0065, Solarbio, China) was applied to each slide, followed by coverslip placement. Slides were inverted onto a fluorescence microscope (Keyence, China) stage, with red fluorescence visualized using 594 nm excitation and blue nuclear DAPI signals detected under 405 nm excitation.

BODIPY staining

After removing the culture plate, cells were washed twice with PBS and fixed with 4% paraformaldehyde for 15 minutes at room temperature. Cells were then incubated with 10 μM BODIPY 493/503 working solution (HY-D1614, MCE, USA) for 15 minutes at room temperature under light-protected conditions. After three PBS washes, lipid droplets were visualized under a fluorescence microscope (Keyence, China) using 480 nm excitation.

Oil Red O staining

Following the Oil Red O Staining Kit manufacturer’s protocol (G1262, Solarbio, China), cells cultured in six-well plates were first fixed by adding the provided fixative solution (4% paraformaldehyde) for 20 minutes at room temperature. After fixation, the solution was aspirated and cells were briefly rinsed with 60% isopropanol (I811932, Macklin, China) for 30 seconds. Subsequently, freshly prepared Oil Red O working solution was added for 20 minutes of staining at room temperature under light-protected conditions. The staining solution was then carefully removed, followed by differential destaining with 60% isopropanol until the interstitial matrix became visually clear. Samples underwent 3–5 washes with distilled water before being covered with double-distilled water for immediate bright-field microscopy (Keyence, China) observation at 200× magnification.

The quantitative analysis procedure for Oil Red O staining is as follows: First, images captured by microscopy are uploaded to ImageJ software to calculate the average area of Oil Red O-positive regions per group. Subsequently, the positive areas of all experimental groups were normalized relative to the control group before statistical analysis and graphical presentation.

Quantification of lipid accumulation

Following Oil Red O staining, intracellular lipids were quantitatively extracted by adding 400 μL of absolute isopropanol to each well for 15-minute incubation at room temperature. From each well, 200 μL of the lipid-containing isopropanol solution was carefully transferred to a 96-well microplate. Absorbance measurements were performed using a microplate reader (ThermoFisher, China) at 490 nm with pathlength correction.

Statistical analysis

All statistical analyses in this study were performed using GraphPad Prism 8 software (GraphPad Software, La Jolla, CA). Quantitative data were first assessed for normality distribution. For comparisons between two groups, two-tailed unpaired t-tests were used for normally distributed data and Mann-Whitney U tests for non-normally distributed data. For multiple group comparisons, one-way analysis of variance (ANOVA) with Bonferroni post-hoc test was applied. A two-tailed p-value <0.05 was considered statistically significant.

Results

High-fat diet and linoleic acid upregulates CAPG expression

The expression level of CAPG in eWAT was significantly higher in HFD-fed mice compared to the ND group (Figure 1(A-B)). To recapitulate CAPG upregulation phenotypes, we treated mature differentiated adipocytes with linoleic acid (LA) at concentrations of 0 μM, 200 μM, 300 μM, and 400 μM, based on established pharmacological ranges reported in the literature. Notably, 400 μM LA upregulated CAPG expression (Figure 1(C-D)). Immunofluorescence analysis revealed CAPG localization in both the cytoplasm and nucleus (Figure 1(E)). However, neither PA treatment alone nor its combination with LA (concentration <200 μM) induced significant changes in CAPG expression levels in adipocytes. (Supplementary Figure S1).

Figure 1.

High-fat diet and LA promotes CAPG expression in adipocytes.

(A) Representative Western blot results of CAPG expression levels in epididymal white adipose tissue (eWAT) from C57BL/6J wild-type mice fed a normal diet (ND) or high-fat diet (HFD). (B) Quantitative comparison of CAPG protein expression in eWAT between ND and HFD groups (n = 6 independent experiments), two-tailed unpaired t-tests. (C) Western blot results of CAPG expression in adipocytes treated with increasing concentrations of linoleic acid (LA). (D) Quantitative comparison of CAPG protein levels following LA treatment at different concentrations (n = 4 independent experiments). (E) Fluorescence microscopy images (400× magnification) of mature adipocytes treated with 0 μM or 400 μM LA. Nuclei stained blue (DAPI), CAPG-positive regions stained red; scale bar = 100 μm. LA, linoleic acid; ns, not statistically significant, one-way ANOVA.

CAPG expression progressively decreases during adipocyte differentiation

We collected cells on days 4, 8, and 10 of induced differentiation, respectively. As shown in Figure 2, CAPG expression gradually decreased during adipogenic differentiation of 3T3-L1 cells, concomitant with increasing PPARγ protein levels.

Figure 2.

Downregulation of CAPG during adipogenic differentiation of 3T3-L1 cells.

(A) Representative Western Blot of CAPG expression levels on Days 4, 8, and 10 of adipocyte differentiation (n = 3 independent experiments). (B-C) Quantitative analysis of CAPG and PPARγ protein levels at indicated time points (n = 3 independent experiments), one-way ANOVA.

CAPG knockdown inhibits adipocyte differentiation

3T3-L1 preadipocytes were transduced with either control shRNA (sh-Ctrl) or three distinct sh-Capg lentiviral constructs, all of which achieved stable Capg knockdown (Figure 3(A)). For subsequent experiments, 3T3-L1 cells transfected with shRNA-1 were selected. The specific nucleotide sequence is provided in the Supplementary Table S1. Following adipogenic induction, mature differentiated sh-Ctrl and sh-Capg adipocytes were treated with vehicle control (NC) or 400 μM LA for 48 hours. Oil Red O and BODIPY staining revealed that while LA treatment markedly enhanced lipid accumulation, Capg knockdown significantly attenuated adipogenesis (Figure 3(B-F)). Western blot analysis demonstrated LA-mediated upregulation of PPARγ expression, which was substantially inhibited by Capg knockdown (Figure 3(G-J)), collectively establishing the essential role of CAPG in adipocyte differentiation.

Figure 3.

CAPG knockdown suppresses adipocyte differentiation.

(A) Representative Western blot results of CAPG expression in 3T3-L1 cells transduced with sh-Ctrl or three independent sh-Capg lentiviruses. (B) Oil Red O staining of sh-Ctrl and sh-Capg cells treated with vehicle control (NC) or 400 μM linoleic acid (LA) for 48 hours on day 10 of adipogenic differentiation. (C) Quantitative comparison of lipid content across the four experimental groups (n = 3 independent experiments). (D) BODIPY staining images (200× magnification; scale bar = 100 μm). (E) Comparative Oil Red O staining under bright-field microscopy (200× magnification; scale bar = 100 μm). (F) Quantification of Oil Red O-positive areas across groups (n = 3 independent experiments). (G) Western blot analysis of CAPG and PPARγ expression in the four groups (n = 3 independent experiments). (H-J) Quantitative comparison of CAPG and PPARγ protein levels from Western blot results (n = 3 independent experiments), one-way ANOVA.

To provide mechanistic evidence for CAPG’s role in adipocyte differentiation, we generated stable Capg-overexpressing (OE) 3T3-L1 cells. Employing the same induction protocol, 3T3-L1 cells were differentiated into adipocytes. Western blotting demonstrated upregulated adipogenic marker PPARγ, confirming CAPG’s pro-adipogenic function (Supplementary Figure S2).

CAPG knockdown suppresses 3T3-L1 cell proliferation

Through Ki67 immunofluorescence staining and CCK-8 assays performed on sh-Ctrl and sh-Capg cells, we demonstrated that Capg knockdown significantly reduced nuclear Ki67 protein expression and inhibited proliferation in 3T3-L1 cells (Figure 4).

Figure 4.

CAPG knockdown suppresses preadipocyte proliferation.

(A) Representative Ki67 immunofluorescence images in sh-Ctrl vs. sh-Capg groups (200× magnification). (B) Quantitative analysis of Ki67-positive cells (n = 3 independent experiments). (C) CCK-8 viability at 24 hours in sh-Ctrl vs. sh-Capg groups (n = 5 independent experiments). (D) CCK-8 viability at 48 hours in sh-Ctrl vs. sh-Capg groups (n = 5 independent experiments), one-way ANOVA.

CAPG knockdown enhances lipid metabolism

CAPG knockdown significantly increased the expression of CD36, a key membrane protein mediating exogenous fatty acid uptake in adipocytes, thereby promoting cellular fatty acid absorption (Figure 5(A-B)). GLUT4 is the primary regulator of adipocyte glucose uptake. LA treatment markedly suppressed the expression of GLUT4, while CAPG knockdown elevated GLUT4 levels enhancing glucose utilization and de novo fatty acid synthesis (Figure 5(A,C)). Furthermore, CAPG knockdown upregulated ATGL protein expression regardless of LA treatment (Figure 5(A,D)). Surprisingly, CAPG OE in adipocytes also elevated the expression levels of ATGL and CD36 in these cells (Supplementary Figure S2B-H).

Figure 5.

CAPG knockdown promotes lipid metabolism.

(A) Western blot results of CD36, ATGL, CAPG, and GLUT4 protein levels in sh-Ctrl and sh-Capg cells treated with vehicle control (NC) or 400 μM linoleic acid (LA). (B-D) Quantitative comparison of CD36, GLUT4, and ATGL protein expression across experimental groups (n = 3 independent experiments), one-way ANOVA.

CAPG knockdown suppresses adipocyte inflammatory responses

As shown in Figure 6(A-D), LA treatment significantly increased Capg mRNA levels in mature adipocytes, while sh-Capg lentiviral transduction effectively downregulated Capg expression. Subsequent analysis of inflammatory factors revealed that LA treatment markedly elevated Il6 and Mcp1 mRNA expression (Figure 6(A-C)), whereas Capg knockdown significantly reduced Mcp1 levels in control groups and attenuated LA-induced Lep mRNA expression (Figure 6(D)).

Figure 6.

CAPG knockdown suppresses adipocyte inflammatory responses.

(A-D) mRNA expression levels of Capg, Il6, Mcp1, and Lep in sh-Ctrl and sh-Capg adipocytes treated with vehicle control (NC) or 400 μM linoleic acid (LA) for 48 hours (n = 3 independent experiments). (E-I) Comparison of pro-inflammatory cytokine concentrations (IL-6, TNF-α, MCP-1, GM-CSF, IL-12 P40) in cell supernatants from sh-Ctrl and sh-Capg adipocytes following NC or LA treatment for 48 hours (n = 3 independent experiments), one-way ANOVA.

To further investigate the role of CAPG in cytokine production, we collected media from all four experimental groups and quantified inflammatory cytokine concentrations using multiplex flow cytometry. LA treatment substantially increased secretion of MCP-1, granulocyte-macrophage colony-stimulating factor (GM-CSF), and IL-12 P40 (Figure 6(G-I)). Conversely, Capg knockdown significantly decreased TNF-α and MCP-1 levels in NC-treated adipocytes (Figure 6(F,G)) as well as reduced LA-stimulated IL-6, MCP-1, and IL-12 P40 production (Figure 6(E,G,I)), demonstrating the critical involvement of CAPG in mediating adipocyte inflammatory responses. Furthermore, when Capg is overexpressed, the expression levels of IL-6 and MCP-1 are significantly elevated (Supplementary Figure S2 I-J).

To rule out secondary reductions in inflammatory factors, we examined the expression levels of specific proteins in the NF-κB signalling pathway. Our results revealed that Capg knockdown significantly suppressed both phosphorylated P65 and the phosphorylated-to-total P65 ratio, indicating that the CAPG-mediated reduction in inflammatory factors may be associated with inhibition of the NF-κB signalling pathway (Supplementary Figure S3).

Discussion

Adipose tissue possesses remarkable plasticity, which is frequently impaired in obesity, leading to diminished nutrient-buffering capacity and subsequent development of insulin resistance and metabolic disorders. Genetic and pharmacological studies have demonstrated that systemic metabolic dysfunction arises not from adipocyte hypertrophy perse, but rather from the loss of adipose tissue ‘expandability’ [13]. In the present study, we systematically investigated the impact of CAPG on adipocyte dysfunction, revealing four pivotal findings: (1) HFD and LA upregulates CAPG expression in adipocytes; (2) knockdown of Capg effectively suppresses adipocyte differentiation and preadipocyte proliferation; (3) Capg knockdown enhances lipid metabolism; and (4) Capg knockdown in adipocytes substantially inhibits the release of pro-inflammatory cytokines, thereby attenuating chronic low-grade inflammation associated with metabolic dysfunction.

We observed that CAPG expression progressively decreases during adipocyte differentiation. However, in adipose tissue from HFD-fed mice, CAPG expression was elevated. To determine the regulatory mechanisms underlying these opposing changes in adipocytes, we treated adipocytes with distinct types of fatty acids. When treating mature adipocytes with fatty acids, only 400 μM LA induced CAPG upregulation. Neither saturated fatty acids alone nor their combination with unsaturated fatty acids increased CAPG levels. The mechanism underlying this differential response remains unclear. Similarly, Krautbauer et al. reported that LA – but not PA – elevates Galectin-3 (which primarily functions to clear advanced glycation end products) expression in adipocytes [14]. It should be emphasized that 400 μM LA exceeds physiological fatty acid concentrations; therefore, whether CAPG elevation in adipocytes is directly induced by LA or mediated through secondary mechanisms warrants further investigation.

Adipose tissue expansion is primarily achieved through two distinct mechanisms: adipocyte hyperplasia (increase in cell number via preadipocyte differentiation) and adipocyte hypertrophy (increase in cell volume through lipid accumulation) [15,16]. The white adipose tissue undergoes expansion via both hypertrophy and hyperplasia when subjected to HFD. During the early stages of obesity, adipose tissue expansion is predominantly driven by adipocyte hypertrophy until cells reach their maximal volume capacity, at which point they trigger the generation of new adipocytes [17,18]. However, emerging evidence indicates that adipocyte proliferation may commence prior to cells attaining their maximal lipid storage capacity [19]. In this study, we demonstrate that CAPG promotes preadipocyte proliferation and differentiation, thereby increasing adipocyte numbers.

CAPG knockdown promotes lipid metabolism in adipocytes. CD36, a high-affinity receptor for high-density lipoprotein, low-density lipoprotein, and very-low-density lipoprotein on adipocyte membranes, mediates exogenous fatty acid uptake [20]. CAPG depletion increases CD36 expression on adipocyte surfaces, thereby enhancing exogenous fatty acid uptake and utilization to drive lipogenesis. Previous studies have shown that CD36 expression is transcriptionally regulated by PPARγ, with elevated PPARγ levels promoting CD36 upregulation [21]. In this study, upon CAPG overexpression, the levels of PPARγ, ATGL, and CD36 all increased. However, CAPG knockdown suppressed PPARγ expression yet paradoxically increased CD36 levels (Supplementary Figure S4), likely due to direct cytoskeletal protein-mediated regulation of CD36 [22]. Nevertheless, the precise mechanisms warrant further in-depth investigation. ATGL, the rate-limiting enzyme in triglyceride hydrolysis, was significantly upregulated upon CAPG knockdown. CAPG knockdown ultimately enhances net lipid synthesis, promoting adipocyte lipid storage [23]. Additionally, GLUT4 facilitates glucose uptake and transport to fuel de novo fatty acid synthesis [24]. CAPG knockdown also elevates GLUT4 expression in adipocytes.

Chronic inflammation represents a hallmark feature of dysfunctional adipose tissue [3], wherein hypertrophic adipocytes secrete pro-inflammatory cytokines that orchestrate systemic inflammatory cascades [25]. Our study demonstrates that genetic knockdown of CAPG significantly attenuates the release of key inflammatory mediators from adipocytes. NF-κB represents a canonical signalling pathway implicated in adipocyte inflammation [26], which modulates adipose tissue inflammatory responses by regulating the expression of downstream adipokines, chemokines, and inflammatory factors. Ma et al. reported the correlation between CAPG and this signalling pathway [27]. Utilizing mass spectrometry-based immunoprecipitation, they purified and characterized CAPG protein complexes in the human acute myeloid leukaemia cell line THP-1. This analysis identified CAPG-interacting proteins – including RPL4, ZFP91, and CCAR2—all established activators of the NF-κB signalling pathway. Similarly, our study demonstrates that CAPG knockdown inhibits the NF-κB pathway by suppressing P65 phosphorylation, which may be associated with reduced inflammatory factors in adipocytes.

MCP-1, a critical regulator of macrophage activation and recruitment, exhibits elevated expression in insulin-resistant white adipose tissue of obese mice, where it promotes endothelial dysfunction, leukocyte transmigration, and subsequent atherosclerotic progression [28]. Similarly, other cytokines including IL-6 and TNF-α exacerbate local and systemic inflammation through immunocyte activation, thereby contributing to obesity-associated vasculopathies [29]. Our study also revealed that CAPG knockdown suppresses leptin secretion, a multifunctional adipokine that modulates diverse biological processes including inflammatory responses, immune function, and energy homoeostasis through binding to leptin receptors [30]. These findings collectively establish CAPG as a pathogenic amplifier in adipocyte-driven inflammation.

This study has several limitations. First, we investigated the effects of CAPG on adipocyte function solely by altering its expression, without exploring the mechanisms underlying CAPG-mediated changes in adipocyte functionality. Second, while this study investigated CAPG’s impact on adipocyte function through cellular-level interventions targeting CAPG expression, it did not establish adipose-tissue-specific CAPG loss-of-function models. Third, we observed that LA treatment increased CAPG expression in adipocytes, whereas PA treatment showed no significant effect on CAPG levels. The underlying mechanism for this differential regulation of CAPG expression by distinct fatty acid types remains unclear and warrants further investigation. Future studies could explore the impact of CAPG on the structure and function of mature adipocytes by intervening with its expression in adipose tissue of mouse models.

Conclusion

Our findings demonstrate that HFD and linoleic acid promotes the upregulation of CAPG expression in adipocytes. Genetic Capg knockdown suppresses adipogenesis and attenuates inflammatory responses, while enhancing lipid metabolism, suggesting that targeting CAPG may represent a novel therapeutic strategy for mitigating obesity-associated adipose tissue inflammation and metabolic disorders.

Funding Statement

This study was supported by the grants from the National Key R&D Program of China [No. 2020YFC2003000 and 2020YFC2003001], the National Natural Science Foundation of China [No. 82170890] and National High Level Hospital Clinical Research Funding [BJ-2024-219].

Disclosure statement

No potential conflict of interest was reported by the author(s).

Data availability statement

The raw data generated in this study have been deposited in the Figshare repository and are publicly available at 10.6084/m9.figshare.29045045.

Ethical statement

All mouse studies were approved by the Animal Ethics Committee of Peking University (Approval No. LA2022278), in full compliance with the Declaration of Helsinki and relevant biomedical research ethics guidelines.

Supplementary material

Supplemental data for this article can be accessed online at https://doi.org/10.1080/21623945.2025.2549456