Restoring Lysosomes in Adipose Tissue Macrophages Mitigates Obesity-Induced Inflammation and Insulin Resistance

Abstract

Adipose tissue macrophages (ATMs) are key mediators of obesity-induced inflammation and insulin resistance. However, the contribution of lysosomal dysfunction to ATM inflammatory activation remains poorly defined. Here, we characterized lysosomal structural and functional alterations in ATMs during obesity and examined whether pharmacological restoration of lysosomal function using 2-hydroxypropyl-β-cyclodextrin (HPβCD) ameliorates metabolic inflammation. In diet-induced obese C57BL/6J male mice, adipose tissue exhibited increased lysosomal abundance, accompanied by reduced cathepsin L+V expression, modestly increased lysosomal acid lipase levels, and decreased expression of transcription factor EB (TFEB), a master regulator of lysosomal biogenesis. Despite expanded lysosomal content, ATMs displayed impaired lysosomal acidification, indicating functional lysosomal dysfunction. Intraperitoneal administration of HPβCD for two weeks significantly improved glucose tolerance and insulin sensitivity without affecting body weight. Flow cytometric analysis revealed reduced pro-inflammatory M1 ATMs and CD8⁺ T lymphocytes in visceral adipose tissue, whereas immune cell populations in subcutaneous adipose tissue, blood, and spleen remained unchanged. In vitro, HPβCD suppressed pro-inflammatory gene expression in both classically and metabolically activated macrophages and attenuated inflammatory responses induced by lysosomal stressors, including bafilomycin A1 and chloroquine, while restoring TFEB expression. Collectively, these findings demonstrate that obesity is associated with lysosomal dysfunction in ATMs and that restoration of lysosomal function alleviates adipose tissue inflammation and metabolic dysfunction, highlighting lysosomal regulation in ATMs as a potential therapeutic target for obesity-associated metabolic diseases.

1. Introduction

Chronic low-grade inflammation in adipose tissue is a defining pathological feature of obesity and a key contributor to the development of insulin resistance and related metabolic disorders [1]. Among the diverse cellular components of adipose tissue, adipose tissue macrophages (ATMs) constitute the dominant immune cells and play a central role in shaping inflammatory tone during obesity [2]. In lean conditions, ATMs largely maintain tissue homeostasis. However, a high-fat diet induces both a rapid expansion and phenotypic reprogramming of ATMs. This remodeling includes a shift from CD11c⁻ resident macrophages toward CD11c⁺ pro-inflammatory macrophages, accompanied by altered lipid handling, cytokine production, and spatial organization around dying adipocytes [3]. These quantitative and qualitative changes in ATMs are closely associated with crown-like structure formation, sustained adipose inflammation, and impaired insulin signaling [4,5,6]. Although ATM heterogeneity and functional plasticity in obesity have been extensively described, the intracellular pathways that govern these changes remain incompletely defined.

Lysosomes are essential regulators of macrophage metabolism and inflammatory responses, integrating lipid degradation, autophagy, and nutrient-sensing pathways [7]. In ATMs, lysosomal activity is initially enhanced during early obesity, reflecting an adaptive response to increased lipid uptake and the need for efficient lipid and debris clearance [8]. However, prolonged exposure to excess lipids, cholesterol, and metabolic stress progressively overwhelms lysosomal capacity, leading to lysosomal stress and functional impairment [9]. Obese ATMs, particularly lipid-associated macrophages localized within crown-like structures, exhibit altered lysosomal enzyme expression, impaired acidification, and defective degradative function [10]. These changes compromise efferocytosis and favor persistent inflammatory signaling, thereby sustaining adipose tissue inflammation. Despite the growing recognition of lysosomal involvement in ATM biology, the question of how lysosomal dysfunction develops during obesity and how it mechanistically contributes to inflammatory activation in ATMs remain unresolved.

Lysosomal homeostasis is transcriptionally coordinated by transcription factor EB (TFEB), a master regulator of lysosomal biogenesis, autophagy, and lipid metabolism [11]. In nutrient-replete and obese conditions, the activation of the mechanistic target of rapamycin complex 1 (mTORC1) restrains TFEB nuclear localization, limiting lysosomal adaptive responses. Suppression of TFEB activity has been linked to impaired autophagy–lysosome flux, inflammasome activation, and sustained pro-inflammatory macrophage polarization [12]. In obesity, lysosomal stress and reduced TFEB signaling may therefore represent a convergent mechanism linking metabolic overload to chronic inflammation in ATMs. While experimental activation of TFEB in macrophages has been shown to improve lysosomal function and dampen inflammatory responses in other disease contexts [13], its functional relevance and therapeutic tractability in obesity-associated adipose inflammation remain to be established.

Cyclodextrins are cyclic oligosaccharides widely used as pharmaceutical excipients and have recently gained attention for their ability to modulate lysosomal lipid handling. Among the various CD derivatives, 2-hydroxypropyl-β-cyclodextrin (HPβCD) is a hydroxypropyl-modified β-cyclodextrin with high water solubility and low toxicity, and effectively modulates lysosomal lipid trafficking relative to other CD derivatives. HPβCD has been shown to restore lysosomal cholesterol trafficking and improve lysosomal function in lysosomal storage disorders [14]. HPβCD is internalized through endocytic pathways and accumulates within lysosomes, where it facilitates lipid mobilization and enhances degradative capacity [15,16,17]. Beyond inherited lysosomal diseases, HPβCD has been reported to exert anti-inflammatory effects in models of chronic tissue injury [18]. However, its impact on macrophage lysosomal stress and inflammatory activation in obesity has not been examined. In the present study, we systematically characterized lysosomal alterations in ATMs during obesity and tested whether the pharmacological restoration of lysosomal function using HPβCD attenuates ATM-driven adipose inflammation and insulin resistance in diet-induced obese mice and macrophage culture models.

2. Results

2.1. Obesity Is Associated with Lysosomal Dysfunction in Adipose Tissue Macrophages

To determine whether obesity alters lysosomal homeostasis in adipose tissue macrophages (ATMs), we first examined the lysosome-associated protein expression in epididymal white adipose tissue (eWAT) from normal diet (ND)- and high-fat diet (HFD)-fed mice. Protein levels of LAMP1, a lysosomal membrane protein reflecting lysosomal abundance, were significantly increased in eWAT from HFD-fed mice. In contrast, levels of lysosomal proteases cathepsin L+V (CtsL+V) were significantly reduced, while lysosomal acid lipase (LAL) showed an increasing trend that did not reach statistical significance (Figure 1A). Autophagy related marker LC3 II showed an increasing trend while LC3 I remained unchanged, consistent with altered autophagy lysosome dynamics in obese adipose tissue (Figure 1A). Consistent with impaired lysosomal adaptation, the expression of transcription factor EB (TFEB), a master regulator of lysosomal biogenesis and function, was significantly reduced in obese eWAT (Figure 1A).

Figure 1.

Obesity leads to lysosomal dysfunction. (A) Immunoblot analysis of eWAT lysate from ND- and HFD-fed mice. Cathepsin L+V (first), lysosomal acid lipase (second), TFEB (third), LAMP1 (fourth), LC3I/II (fifth), and heat shock protein 90 (HSP90, bottom) protein levels are presented. Relative protein abundance normalized to housekeeping protein HSP90 expression. (B) qRT-PCR analysis of macrophage, pro-inflammatory, lysosomal function, and lipid handling markers in ATM from ND- and HFD-fed-mice. (C) Representative flow cytometry analyses of Lysotracker⁺ ATMs in ND- and HFD-fed mice. (D) Representative LAMP1 immunofluorescence staining of CD11b⁺ cells isolated from ND and HFD-fed mice, showing the overall cellular phenotype and quantitative analysis. (E) Representative comparison of lysotracker⁺ median fluorescence intensity (MFI) in the indicated ATM populations. Error bars represent SEM. * p < 0.05, **** p < 0.001.

To assess lysosomal alterations specifically in ATMs, CD11b⁺ ATMs were isolated and analyzed for lysosomal gene expression and protein abundance. Compared with ND-fed mice, HFD feeding significantly increased the expression of Lamp1, Lipa, and the lipid scavenger receptor Cd36, while reducing the expression of Ctsl, suggesting lysosomal expansion accompanied by compromised degradative capacity in obese ATMs (Figure 1B). Flow cytometric analysis further revealed an increased proportion of LysoTracker⁺ CD45⁺CD64⁺ ATMs in HFD-fed mice (Figure 1C). Consistently, confocal microscopy demonstrated enhanced LAMP1-associated lysosomal signal in CD11b⁺ cells from HFD-fed mice (Figure 1D), supporting increased lysosomal abundance in obese ATMs.

Given the apparent increase in lysosomal content despite the reduced expression of lysosomal enzymes, we next assessed lysosomal function. The median fluorescence intensity (MFI) of LysoTracker staining was significantly reduced in M1 ATMs from HFD-fed mice compared with ND-fed controls (Figure 1E). LysoTracker MFI was also decreased in total ATMs and M2 ATMs by approximately 22% and 19%, respectively. Because lysosomal mass was increased in obese ATMs, the reduced LysoTracker signal indicates impaired lysosomal acidification rather than decreased lysosomal number. Collectively, these data indicate that obesity induces lysosomal expansion in ATMs while concomitantly impairing lysosomal acidification and degradative capacity, consistent with lysosomal stress or functional immaturity in obese ATMs.

2.2. HPβCD Treatment Improves Glucose Tolerance and Insulin Sensitivity in Obese Mice

To examine whether the restoration of lysosomal function affects obesity-associated metabolic dysfunction, HFD-fed mice were treated with the lysosomal activator 2-hydroxypropyl-β-cyclodextrin (HPβCD) or vehicle for 20 days (Figure 2A). Body weights were comparable between groups throughout the treatment period (Figure 2B). HPβCD treatment significantly reduced inguinal white adipose tissue (iWAT) and liver weights, whereas eWAT weight was modestly increased (Figure 2C).

Figure 2.

HPβCD treatment improves obesity-induced glucose intolerance and insulin resistance in obese mice. (A–G) Male C57BL/6 mice were fed a high-fat diet (HFD) for 15 weeks to induce obesity (n = 8 per group). (A) Animal experiment scheme, (B) total body weights, (C) Organ weight-to-body weight ratio. (D) fasting blood glucose, (E) fasting blood serum insulin ELISA, (F) HOMA-Insulin Resistance (IR) analysis, and (G) blood glucose tolerance test. Error bars represent SEM. * p < 0.05, ** p < 0.01, *** p < 0.001.

The metabolic assessment revealed that HPβCD-treated mice exhibited significantly lower fasting blood glucose and serum insulin levels compared with vehicle-treated controls (Figure 2D,E). Accordingly, the homeostasis model assessment of insulin resistance (HOMA-IR) index was reduced by approximately 2.5-fold in HPβCD-treated mice (Figure 2F). Glucose tolerance tests further demonstrated improved glucose clearance in HPβCD-treated mice (Figure 2G). Serum lipid profiling revealed a significant reduction in HDL cholesterol and a modest decrease in total cholesterol, whereas triglycerides, LDL-C, and free fatty acids were comparable between groups (Supplementary Figure S1). In parallel, histological analysis indicated reduced hepatic lipid accumulation in HPβCD-treated mice, accompanied by decreased expression of lipogenic gene Scd1 (Supplementary Figure S2). Collectively, these findings indicate that HPβCD treatment improves glucose tolerance and insulin sensitivity in obese mice, accompanied by adipose tissue redistribution and modest changes in lipid handling, including limited alterations in circulating lipid profiles and hepatic steatosis, independently of body weight changes.

2.3. HPβCD Reduces Pro-Inflammatory ATMs and CD8⁺ T Cells in Visceral Adipose Tissue

We next evaluated whether metabolic improvement following HPβCD treatment was associated with reduced adipose tissue inflammation. A histological analysis of eWAT from vehicle-treated obese mice revealed abundant crown-like structures (CLSs) indicative of macrophage accumulation around dying adipocytes. In contrast, HPβCD-treated mice displayed a marked reduction in CLS number (Figure 3A). Consistent with these findings, immunofluorescence analysis using Caveolin to delineate adipocyte and Mac2 staining as a pro-inflammatory macrophage marker demonstrated reduced macrophage accumulation in adipose tissue from HPβCD treated mice (Supplementary Figure S3).

Figure 3.

HPβCD treatment decreases pro-inflammatory M1 ATMs and ATTs in visceral AT significantly. (A–G) Male C57BL/6 mice were fed an HFD for 15 weeks to induce obesity (n = 8 per group), and then checked adipose tissue macrophages (ATMs) and adipose tissue T lymphocytes (ATTs) from visceral fat were checked. (A) H&E staining (scale bar = 100 μm), (B) representative ATM plots, (C) comparison of the number of M1 and M2-polarized ATMs and dendritic cells (ATDCs) per gram adipose tissue, (D) M1 and M2-polarized ATMs ratio, (E) comparison of the number of adipose tissue CD3⁺ T lymphocytes per gram adipose tissue, (F) comparison of both CD4⁺ and CD8⁺ T cells measured among CD3⁺ T cell population, and (G) qRT-PCR analysis of macrophage, pro-inflammatory, and anti-inflammatory markers in eWAT from both vehicle- and HPβCD-treated HFD mice. Error bars represent SEM. * p < 0.05, ** p < 0.01.

Flow cytometric analysis showed that within the CD45⁺ leukocyte population, CD64⁺CD11c⁺ M1 ATMs predominated in vehicle-treated mice, whereas HPβCD treatment resulted in a relative enrichment of CD64⁺CD11c⁻ M2 ATMs (Figure 3B). Quantitative analysis demonstrated a significant reduction in M1 ATMs, while M2 ATM abundance remained unchanged, leading to a reduced M1/M2 ratio in HPβCD-treated mice (Figure 3C,D). These data indicate that HPβCD attenuates visceral adipose tissue inflammation primarily through the suppression of pro-inflammatory ATMs.

Adipose tissue T lymphocytes (ATTs) were also examined. The total CD3⁺ T cell abundance in eWAT showed a modest, non-significant reduction following HPβCD treatment (Figure 3E). Notably, CD8⁺ T cells were significantly reduced, whereas CD4⁺ T cells showed a decreasing trend (Figure 3F), suggesting a preferential reduction in pro-inflammatory ATTs.

Consistent with these cellular changes, gene expression analysis revealed reduced expression of macrophage markers Emr1 and Itgax in eWAT from HPβCD-treated mice (Figure 3G). While the expression of pro-inflammatory cytokines (Il6, Il12, Nos2) was largely unchanged, the expression of the regulatory T cell marker Foxp3 was increased by approximately two-fold, with modest increases in Il10 and Mgl1. Together, these findings suggest that HPβCD selectively attenuates visceral adipose tissue inflammation by reducing pro-inflammatory ATMs and CD8⁺ ATTs.

2.4. HPβCD-Induced Immunomodulation Is More Prominent to Visceral Adipose Tissue and Does Not Affect Systemic Immunity

To determine whether HPβCD-induced immunomodulation extended beyond visceral adipose tissue, immune cell populations in subcutaneous adipose tissue and peripheral compartments were analyzed. In iWAT, flow cytometry revealed modest reductions in M1 ATMs, M2 ATMs, and adipose tissue dendritic cells (ATDCs) in HPβCD-treated mice; however, the M1/M2 ATM ratio remained unchanged (Figure 4A,B). Total CD3⁺ T cells showed a decreasing trend, while CD4⁺ and CD8⁺ T cell populations were comparable between groups (Figure 4C,D), indicating a weaker effect in subcutaneous adipose tissue.

Figure 4.

HPβCD-induced decreases in M1 ATMs and ATTs are specific for visceral adipose tissue. (A–D) Male C57BL/6 mice were fed an HFD for 15 weeks to induce obesity (n = 8 per group), and then we performed flow cytometry analysis of ATMs and ATTs from subcutaneous fat. (A) Comparison of M1 and M2-polarized ATMs and ATDCs measured as cell numbers per gram adipose tissue, (B) ratio of M1 and M2-polarized ATMs, (C) comparison of adipose tissue CD3⁺ T cells measured as cell numbers per gram adipose tissue, and (D) comparison of both CD4⁺ and CD8⁺ ATTs measured as cell numbers per gram adipose tissue. Error bars represent SEM.

To assess potential systemic immune effects, circulating leukocyte populations were examined. No differences were observed in blood CD3⁺ T cells, CD115⁺ monocytes, or Ly6G⁺ neutrophils between vehicle- and HPβCD-treated mice (Figure 5A–C). The proportions of inflammatory Ly6C^high and patrolling Ly6C^low monocytes were also unchanged (Figure 5D). Similarly, splenic CD4⁺ and CD8⁺ T cell populations were comparable between groups (Figure 5E). These data indicate that the HPβCD-mediated reduction in adipose tissue inflammation occurs without detectable alterations in systemic immune cell composition.

Figure 5.

HPβCD treatment does not affect blood immune cell population or spleen T cell population in obese mice. (A–E) Male C57BL/6 mice were fed an HFD for 15 weeks to induce obesity (n = 8 per group), and then we performed flow cytometry analysis of immune cell populations from blood and spleen T cell population. (A) Blood lymphocytes population comparison measured among CD3⁺ T cells, (B) blood monocytes population comparison measured among CD115⁺ cells, (C) blood neutrophils population comparison measured among Ly6G⁺ cells, (D) comparison of Ly6C^low and Ly6C^high cell populations measured among CD115⁺ cells, and (E) comparison of Ly6C^high/Ly6C^low ratio between vehicle- and HPβCD-treated mice. Error bars represent SEM.

2.5. HPβCD Suppresses Pro-Inflammatory Macrophage Activation via Restoration of Lysosomal Stress and TFEB Signaling

To determine whether HPβCD directly regulates macrophage activation, bone marrow-derived macrophages (BMDMs) and peritoneal macrophages (pMACs) were stimulated with lipopolysaccharide (LPS), palmitate, or metabolic activation conditions in the presence or absence of HPβCD. As expected, LPS or palmitate robustly induced the expression of M1-associated genes Il1b, Il6, and Nos2 in BMDMs (Figure 6A,B). Co-treatment with HPβCD significantly suppressed the induction of these genes. Similar inhibitory effects were observed in palmitate-stimulated pMACs (Figure 6C). Under metabolic activation conditions, pMACs exhibited increased Il6 and Nos2 expressions, whereas Il1b expression was not induced (Figure 6D). HPβCD treatment significantly attenuated metabolic activation-induced Il6 and Nos2 expression, indicating the suppression of metabolically activated macrophage phenotypes.

Figure 6.

HPβCD decreased M1 macrophage activation and metabolically activated macrophage phenotypes in vitro. (A–D) Characterization of bone marrow derived macrophages (BMDMs) and peritoneal macrophages (pMACs) with HPβCD (1 mM) treatment. (A) qRT-PCR analysis of pro-inflammatory markers and mRNA expression levels in M1-polarized BMDMs induced by palmitate (250 μM) and/or HPβCD treatment for 24 h, (B) qRT-PCR analysis of pro-inflammatory markers and mRNA expression levels in M1-polarized BMDMs induced by LPS (100 ng/mL) and/or HPβCD after treatment for 24 h, (C) qRT-PCR analysis of pro-inflammatory markers and mRNA expression levels in M1-polarized pMACs induced by palmitate (250 μM) and/or HPβCD treatment for 24 h, and (D) qRT-PCR analysis of pro-inflammatory markers and mRNA expression levels in metabolically activated (MMe) pMACs and/or HPβCD treatment for 24 h. Error bars represent SEM. * p < 0.05, *** p < 0.001.

Given the lysosome-activating properties of HPβCD, we next examined the involvement of lysosomal stress and TFEB signaling. Tfeb expression was markedly reduced in palmitate-treated BMDMs and modestly reduced in metabolically activated pMACs; in both settings, HPβCD co-treatment restored Tfeb expression to control or higher levels (Figure 7A). To directly induce lysosomal stress, BMDMs were treated with bafilomycin A1 (Baf) or chloroquine (CQ). Both agents showed a markedly increased expression of pro-inflammatory genes, including Il1b, Il6, Nos2, and Tnfa (Figure 7B,C). HPβCD co-treatment significantly suppressed these responses and restored Tfeb expression under lysosomal stress conditions (Figure 7B,C).

Figure 7.

HPβCD inhibits lysosomal stress-induced M1 macrophage activation by enhancing TFEB expression. (A–D) Characterization of BMDMs with either Baf or CQ treatment and with or without HPβCD (1 mM) treatment. (A) qRT-PCR analysis of Tfeb mRNA expression levels in M1-polarized BMDMs induced by palmitate (left) and metabolic condition (right), (B) qRT-PCR analysis of pro-inflammatory markers and Tfeb mRNA expression levels in bafilomycin (100 nM)-treated BMDMs and/or HPβCD treatment for 24 h, (C) qRT-PCR analysis of pro-inflammatory markers and mRNA expression levels in chloroquine (100 μM)-treated BMDMs and/or HPβCD treatment for 24 h, and (D) qRT-PCR analysis of Il1b, Il6, and Nos2 mRNA expression in control and constitutive active TFEB overexpressed (pTFEBΔ30), metabolically activated (MMe), and HPβCD treated BMDMs. (E) Immunoblot analysis of eWAT lysates from vehicle and HPβCD-treated mice. TFEB (top) and β-actin (bottom) protein levels are presented. Relative levels of total protein normalized to b-actin expression. (F) Percentage of CD11b⁺F4/80⁺ macrophages expressing IL-6 and TNF-α, determined by flow cytometry, in visceral adipose tissue of HPβCD- and vehicle-treated HFD-fed mice. Error bars represent SEM. * p < 0.05, **** p < 0.001.

To further probe the TFEB-linked mechanisms, we assessed metabolically activated BMDMs with constitutively active TFEB overexpression, with HPβCD included for comparison (Figure 7D, Supplementary Figure S4). TFEB overexpression recapitulated the anti-inflammatory effects of HPβCD, as Il1b, Il6, and Nos2, failed to increase under metabolic activation (Figure 7D). Notably, HPβCD did not further suppress inflammatory gene expression in TFEB overexpressing macrophages, supporting that the TFEB activation is sufficient to mediate these anti-inflammatory effects in vitro. Consistent with in vitro findings, TFEB protein abundance in visceral adipose tissue was increased approximately 2.5-fold in HPβCD-treated obese mice compared with vehicle controls (Figure 7E). In addition, flow cytometric intracellular cytokine staining demonstrated reduced IL-6 and TNFα in visceral adipose tissue macrophages following HPβCD treatment (Figure 7F). Together, these results demonstrate that HPβCD suppresses pro-inflammatory macrophage activation by alleviating lysosomal stress and restoring TFEB expression, thereby contributing to reduced adipose tissue inflammation.

3. Discussion

Adipose tissue macrophage (ATM)-mediated inflammation is a central driver of obesity-associated metabolic disorders [2,19]. However, the intracellular mechanisms governing pro-inflammatory ATM activation and persistence during obesity remain incompletely understood. Lysosomes have recently emerged as critical regulators of immuno-metabolic signaling in macrophages, integrating lipid handling, nutrient sensing, and inflammatory responses [7,20]. In the present study, we demonstrate that obesity is associated with profound lysosomal dysfunction in ATMs, characterized by increased lysosomal abundance coupled with impaired acidification and reduced degradative capacity. Importantly, pharmacological restoration of lysosomal function using 2-hydroxypropyl-β-cyclodextrin (HPβCD) attenuated visceral adipose tissue inflammation and improved glucose tolerance and insulin sensitivity without altering body weight. These anti-inflammatory effects were observed in both visceral and subcutaneous depots but were more pronounced in visceral adipose tissue, and they were not accompanied by detectable changes in systemic immune cell populations. Together with complementary in vitro data showing the suppression of classically and metabolically activated macrophage inflammatory programs, our findings identify lysosomal dysfunction in ATMs as a key feature of obesity-associated inflammation and highlight lysosome restoration as a potential therapeutic strategy.

Several previous studies have reported obesity-associated alterations in lysosomal pathways within ATMs, although with differing emphases [8,21,22]. Xu et al. [8] described increased lysosomal biogenesis and upregulation of lysosome-related gene expression in obesity, whereas Gabriel et al. [21] emphasized enhanced lysosomal stress under obese conditions. These observations collectively indicate that lysosomal homeostasis is disrupted during obesity, but do not fully resolve the question of how lysosomal alterations relate to macrophage inflammatory activation. Extending these studies, we show that despite increased lysosomal abundance in obese ATMs, as reflected by elevated LAMP1 expression and enhanced lysosomal staining, lysosomal function is compromised, as evidenced by reduced cathepsin expression, impaired acidification, and decreased LysoTracker intensity. In addition, increased LC3 II with unchanged LC3 I together with elevated LAMP1 and reduced cathepsins supports impaired autophagy lysosome flux, consistent with accumulation of autophagosomes and lysosomal structures without adequate degradative capacity. This functional impairment was most pronounced in pro-inflammatory M1 ATMs and was associated with increased lysosomal pH in vivo. Moreover, the experimental induction of lysosomal stress using chloroquine or bafilomycin A1 was sufficient to promote pro-inflammatory gene expression in macrophages in vitro. These findings support a model in which lysosomal biogenesis and lysosomal functionality become uncoupled during chronic overnutrition, contributing to sustained ATM inflammation.

Lysosomal dysfunction in obese ATMs likely arises from the convergence of multiple TFEB-dependent and TFEB-independent regulatory pathways. TFEB is a master transcriptional regulator that coordinates lysosomal biogenesis and autophagy in response to environmental and metabolic cues, and macrophage-specific TFEB activation has been shown to promote lipid catabolism, anti-inflammatory polarization, and improved metabolic outcomes in obese models [11,23,24]. Consistent with these reports, TFEB expression was reduced in obese ATMs and increased following HPβCD treatment in vitro and in adipose tissue in vivo. However, our in vivo data do not directly establish TFEB as the causal mediator of HPβCD effects, and TFEB independent mechanisms may also contribute. TFEB-independent mechanisms that can influence lysosomal function include altered mTORC1–AKT feedback signaling, impaired V-ATPase assembly, lysosomal membrane permeabilization, induction of lysosomal stress markers such as GPNMB, and activation of inflammasome pathways [23]. Lysosomal membrane destabilization and loss of luminal acidity can lead to the leakage of hydrolytic enzymes, impaired autophagic flux, and the amplification of inflammatory signaling independent of transcriptional control [25,26]. Our data are compatible with this integrated model in which obese ATMs exhibit numerically expanded but functionally stressed lysosomes. Future studies using macrophage-specific genetic or pharmacologic TFEB perturbation and direct lysosomal functional assays in vivo will be necessary to define causality and to dissect the relative contributions of these pathways.

Cyclodextrins are well known for their ability to bind membrane lipids and modulate intracellular cholesterol trafficking, thereby influencing lysosomal homeostasis and macrophage function. Previous studies have shown that HPβCD mobilizes lysosomal cholesterol and alters autophagy–lysosome flux, although its effects appear to be context-dependent [15,17]. Early work demonstrated that cholesterol depletion by HPβCD can transiently disrupt autophagic flux yet ultimately restore lysosomal degradative capacity [27], while more recent studies suggest that HPβCD enhances autophagy through TFEB-dependent mechanisms [16]. In line with these findings, our data demonstrate that HPβCD suppresses pro-inflammatory gene expression induced by classical and metabolic activation in macrophages, accompanied by a restoration of TFEB expression. These results support the notion that HPβCD alleviates lysosomal stress and inflammatory activation in macrophages, coinciding with the restoration of lysosome-regulatory programs, while acknowledging that additional TFEB-independent mechanisms may also contribute.

Beyond adipose tissue macrophages, cyclodextrins have been reported to exert broader metabolic effects, including a regression of atherosclerosis through macrophage reprogramming and activation of liver X receptor-dependent cholesterol efflux pathways [28]. Consistent with these observations, we found that HPβCD-treated obese mice exhibited reduced hepatic lipid droplet number and size, along with a decreased expression of lipogenic genes such as Scd1 and Fas (Figure S1). These findings suggest that the modulation of hepatic lipid metabolism may contribute to the overall improvement in metabolic inflammation observed with HPβCD treatment. Importantly, these effects occurred without significant changes in body weight or systemic immune cell composition, indicating a selective impact on tissue-resident metabolic and inflammatory pathways. Thus, improved lysosomal and lipid handling in macrophages and hepatocytes may act in concert to mitigate obesity-associated metabolic dysfunction.

Several limitations of this study should be acknowledged. First, although our data establish a strong association between lysosomal dysfunction and ATM inflammatory activation, causality cannot be definitively inferred. It remains unclear whether lysosomal dysfunction precedes macrophage activation or arises as a consequence of sustained inflammatory stress. Thus, direct macrophage-specific lysosomal functional analyses in vivo are warranted to provide important mechanistic insight. Second, while TFEB restoration correlated with improved lysosomal and inflammatory phenotypes, we did not directly manipulate TFEB genetically, and TFEB-independent pathways likely contribute to the observed effects. Third, HPβCD may exert cell type-specific effects beyond macrophages, particularly in hepatocytes, and this was not fully explored in this study. A detailed assessment of autophagy–lysosome flux, lysosomal membrane integrity, and nutrient-sensing signaling will be necessary to more precisely define the nature of lysosomal dysfunction in obese ATMs. Finally, because only male mice were used, the generalizability of these findings to female mice remains to be determined given known sex-specific differences in adipose inflammation and insulin sensitivity.

In conclusion, our findings demonstrate that, in adipose tissue macrophages, obesity induces a state of lysosomal dysfunction characterized by increased lysosomal abundance but impaired acidification and degradative capacity. The pharmacological restoration of lysosomal function using HPβCD attenuated pro-inflammatory macrophage activation and improved metabolic outcomes in obese mice. These results identify lysosomal regulation in ATMs as a promising therapeutic target and support lysosome-directed strategies as a potential approach for the treatment of obesity-associated metabolic diseases.

4. Materials and Methods

4.1. Animal Studies and Metabolic Assessments

Male C57BL/6J mice were obtained from Orient Bio (Seongnam, Republic of Korea) and housed under specific pathogen-free conditions with a 12 h light/dark cycle and ad libitum access to food and water. At six weeks of age, male mice were randomly assigned by an investigator blinded to group allocation and fed either a normal diet (ND; 4.5% fat; PMI Nutrition International, St. Paul, MN, USA) or a high-fat diet (HFD; Research Diets, New Brunswick, NJ, USA, #D12492,). Body weight was recorded weekly. After 15 weeks of HFD feeding, mice were administered intraperitoneal (IP) injections of 2-hydroxypropyl-β-cyclodextrin (HPβCD; 4000 mg/kg body weight; Sigma, St. Louis, MO, USA) or saline every other day for a total of 10 injections. A dosage of 4000 mg/kg of HPβCD was carefully selected based on prior in vivo studies in lysosomal disease models for its efficacy and low toxicity. The dietary intervention used the cage as the experimental unit, whereas HPβCD IP injection used the individual mouse. Following the ninth injection, glucose tolerance tests (GTTs) were performed after a 6 h fast. Mice were IP-injected with D-glucose (0.7 g/kg body weight), and blood glucose levels were measured at 0, 15, 30, 45, 90, and 120 min using a glucometer. For tissue collection, mice were fasted for 16 h, anesthetized with isoflurane inhalation, and euthanized by cervical dislocation. All animal procedures were approved by the Institutional Animal Care and Use Committee of Soonchunhyang University (SCH18-0020).

4.2. Isolation of Stromal Vascular Cells and CD11b⁺ Macrophages

Stromal vascular cells (SVCs) were isolated from white adipose tissue (WAT). Excised tissue was mechanically minced and digested in buffer containing 0.5% BSA, 20 mM HEPES, 50% HBSS, and 50% PBS supplemented with type II collagenase (1 mg/mL; Sigma-Aldrich, St. Louis, MO, USA, #C2139) for 30 min at 37 °C with gentle agitation. Digested tissue was filtered through a 100 μm mesh and centrifuged at 500× g for 5 min at 4 °C. The resulting SVC pellet was subjected to red blood cell lysis (eBioscience, San Diego, CA, USA, #00-4333-57) for 5 min at room temperature, washed, and counted. CD11b⁺ macrophages were purified from SVCs using magnetic-activated cell sorting (MACS) with CD11b MicroBeads (Miltenyi Biotec, Bergisch Gladbach, Germany, #130-049-601), according to the manufacturer’s instructions.

For flow cytometric analysis, cells were incubated with Fc block for 10 min at 4 °C and stained with fluorochrome-conjugated antibodies. For lysosomal labeling, LysoTracker (50 nM; Thermo Fisher Scientific, Waltham, MA, USA, #L12492) was added after antibody staining and incubated for 10 min on ice. Cells were washed three times, and data were acquired using a BD FACSCanto II (BD Biosciences, Franklin Lakes, NJ, USA) and analyzed with FlowJo software v10 (Tree Star). All antibodies are listed in Table S1.

4.3. Bone Marrow-Derived and Peritoneal Macrophage Culture

Bone marrow cells were harvested from femurs and tibias of wild-type mice and differentiated into bone marrow-derived macrophages (BMDMs) in high-glucose Dulbecco’s modified Eagle’s medium (DMEM; Corning, Manassas, VA, USA, #10-013-CV), supplemented with 10% fetal bovine serum (FBS; Corning, #35-015-CV), 1% penicillin–streptomycin (P/S; Corning, #30-002-CI) and 20% L929-conditioned medium for 6 days, with medium replacement every 2 days. Peritoneal macrophages (pMACs) were elicited by intraperitoneal injection of 2 mL thioglycolate. After 5 days, pMACs were collected by peritoneal lavage and cultured in RPMI 1640 (Corning, #10-040-CVR) supplemented with 10% FBS and 1% P/S. Non-adherent cells were removed after 4 h, and fresh medium was added.

BMDMs and pMACs were seeded and treated as indicated. Classical M1 activation was induced by treatment with lipopolysaccharide (LPS; 100 ng/mL) or palmitate (250 μM) for 24 h. Metabolically activated macrophages (MMe) were generated using glucose (30 mM), insulin (10 nM), and palmitate (0.4 mM) for 24 h. Lysosomal stress was induced using bafilomycin A1 or chloroquine, with or without concurrent treatment with HPβCD (1 mM).

For the TFEB overexpression study, BMDMs were transfected with mock (pEGFP) or pEGFP–N1-Δ30-TFEB (Addgene, Watertown, MA, USA, #44445) to achieve constitutively active TFEB overexpression. Similarly, TFEB overexpressed BMDMs were metabolically activated using (30 mM), insulin (10 nM), and palmitate (0.4 mM) for 24 h and treated with or without HPβCD (1 mM).

4.4. Gene Expression Analysis

Total RNA was extracted using TRIzol^® Reagent (Thermo Fisher Scientific #15596018). cDNA was synthesized using a High-Capacity cDNA Reverse Transcription Kit (Applied Biosystems, Foster City, CA, USA). Quantitative PCR was performed using PowerUp™ SYBR™ Green Master Mix (Applied Biosystems, #A25742) on a QuantStudio™ 1 system. Relative gene expression was calculated using the 2^−ΔΔCt method, with β2-microglobulin (B2m) as the internal control for macrophages and 18S rRNA for tissue samples. All primers are listed in Table S2.

4.5. Immunoblotting

Tissues were homogenized in RIPA buffer containing protease inhibitors using a TissueLyser II (Qiagen, Venlo, The Netherlands). Protein lysates were separated by SDS–PAGE and transferred to nitrocellulose membranes. Membranes were incubated with primary antibodies overnight at 4 °C, followed by HRP-conjugated secondary antibodies. Signals were detected using ECL Prime reagent (Cytiva, Marlborough, MA, USA) and visualized with an Agfa CP1000 system. All antibodies are listed in Table S1.

4.6. Histology and Immunofluorescence

WATs were fixed in 10% formalin, paraffin-embedded, sectioned, and stained with hematoxylin and eosin (H&E). Images were acquired using a Leica DMi8 microscope. For immunofluorescence, isolated CD11b⁺ cells (~20,000 cells) were cyto-spun onto glass slides, fixed with 2% PFA, permeabilized, blocked, and incubated with anti-LAMP1 (CD107a; Thermo Fisher #14-1071-81) overnight at 4 °C. Slides were incubated with fluorescent secondary antibodies, counterstained with DAPI, and imaged using a LSM 710 confocal microscope (Zeiss, Oberkochen, Germany). All antibodies are listed in Table S1.

4.7. Serum Insulin Measurement

Serum insulin levels were quantified using an ultra-sensitive mouse insulin ELISA kit (Miobs, Tsurumi-ku, Japan, #3101) according to the manufacturer’s protocol.

4.8. Statistical Analysis

Data are presented as mean ± SEM. Statistical analyses were performed using GraphPad Prism 8. Comparisons between two groups were conducted using unpaired Student’s t-tests, while multiple-group comparisons were analyzed by one-way ANOVA with Tukey’s post hoc test. Statistical significance was defined as p < 0.05.

Abbreviations

The following abbreviations are used in this manuscript:

ATM(s)	Adipose tissue macrophage(s)
CD11c (ITGAX)	Integrin alpha X
CLS(s)	Crown-like structure(s)
TFEB	Transcription factor EB
mTORC1	Mechanistic target of rapamycin complex 1
HPβCD	2-hydroxypropyl-β-cyclodextrin
eWAT	Epididymal white adipose tissue
ND	Normal diet
HFD	High-fat diet
LAMP1	Lysosomal-associated membrane protein 1
CTSL	Cathepsin L
CTSV	Cathepsin V
LAL/LIPA	Lysosomal acid lipase
HSP90	Heat shock protein 90
CD11b	Cluster of differentiation 11b
CD36	Fatty acid translocase/scavenger receptor
qRT-PCR	Quantitative real-time polymerase chain reaction
CD45	Cluster of differentiation 45
CD64	Cluster of differentiation 64
LysoTracker	Lysosomal acidity-sensitive fluorescent dye
MFI	Median fluorescence intensity
LysoSensor	Lysosomal pH-sensitive fluorescent probe
iWAT	Inguinal white adipose tissue
HOMA-IR	Homeostasis model assessment of insulin resistance
M1	Classically activated macrophage
M2	Alternatively activated macrophage
ATT(s)	Adipose tissue T lymphocyte(s)
CD3	Cluster of differentiation 3
CD4	Cluster of differentiation 4
CD8	Cluster of differentiation 8
Emr1 (F4/80)	EGF-like module-containing mucin-like hormone receptor-like 1
Il6	Interleukin-6
Il12	Interleukin-12
Nos2	Nitric oxide synthase 2
Foxp3	Forkhead box P3
Il10	Interleukin-10
Mgl1	Macrophage galactose-type lectin 1
ATDC(s)	Adipose tissue dendritic cell(s)
Ly6G	Lymphocyte antigen 6 complex, locus G
CD115	Colony-stimulating factor 1 receptor
Ly6C	Lymphocyte antigen 6 complex, locus C
BMDM(s)	Bone marrow-derived macrophage(s)
pMAC(s)	Peritoneal macrophage(s)
LPS	Lipopolysaccharide
MMe	Metabolically activated macrophage
Baf	Bafilomycin A1
CQ	Chloroquine
V-ATPase	Vacuolar-type H+-ATPase
AKT	Protein kinase B
GPNMB	Glycoprotein non-metastatic melanoma protein B
Scd1	Stearoyl-CoA desaturase 1
Fas/Fasn	Fatty acid synthase
SEM	Standard error of the mean

Supplementary Materials

The following supporting information can be downloaded at https://www.mdpi.com/article/10.3390/ijms27041755/s1.

Author Contributions

J.C.; Formal analysis, Investigation, Writing—original draft preparation; E.B.; Formal analysis, Investigation, Writing—original draft preparation, Writing—review and editing; S.S.; Methodology, Validation, Visualization; X.R.Y.; Formal analysis; S.C.; Conceptualization, Resources, Funding acquisition; K.W.C.; Conceptualization, Resources, Data curation, Supervision, Project administration, Funding acquisition, Writing—review and editing. All authors have read and agreed to the published version of the manuscript.

Institutional Review Board Statement

The animal study protocol was approved by the Institutional Animal Care and Use Committee (IACUC) of Soonchunhyang University (Approval No. SCH18-0020, approved on 6 October 2018).

Informed Consent Statement

Not applicable.

Data Availability Statement

Data is contained within the article or Supplementary Material.

Conflicts of Interest

The authors declare no conflicts of interest.

Funding Statement

This study was supported by the National Research Foundation of Korea (NRF-2022R1A2C2006224, NRF-RS2023-00222784), Korean Society for the Study of Obesity (KSSO202401) and Korean Diabetes Association Grant (S.C., 2017F-4).