
==== Front
J Biol Chem
J Biol Chem
The Journal of Biological Chemistry
0021-9258
1083-351X
American Society for Biochemistry and Molecular Biology

S0021-9258(24)02191-4
10.1016/j.jbc.2024.107690
107690
Research Article
Iron supplementation and iron accumulation promote adipocyte thermogenesis through PGC1α-ATGL–mediated lipolysis
Mai Xudong 12‡
Liu Yifan 2‡
Fan Jigang 1‡
Xiao Lanling 2
Liao Miaomiao 2
Huang Zhipeng 2
Chen Zijian 1
Huang Shaojun 1
Sun Rui 1
Jiang Xiaowan 1
Huang Liujing 3
Sun Jia 1
Xie Liwei xielw@gdim.cn
124∗
Chen Hong chenhong123@smu.edu.cn
1∗
1 Department of Endocrinology and Metabolism, Zhujiang Hospital, Southern Medical University, Guangzhou, China
2 State Key Laboratory of Applied Microbiology Southern China, Guangdong Provincial Key Laboratory of Microbial Culture Collection and Application, Guangdong Open Laboratory of Applied Microbiology, Institute of Microbiology, Guangdong Academy of Sciences, Guangzhou, China
3 Medical Affairs Department, Guangzhou Betrue Technology Co, Ltd, Guangzhou, China
4 College of Life and Health Sciences, Guangdong Industry Polytechnic, Guangzhou, Guangdong, China
∗ For correspondence: Hong Chen; Liwei Xie xielw@gdim.cnchenhong123@smu.edu.cn
‡ These authors contribute equally to this work.

17 8 2024
9 2024
17 8 2024
300 9 1076909 1 2024
24 7 2024
© 2024 The Authors
2024
https://creativecommons.org/licenses/by/4.0/ This is an open access article under the CC BY license (http://creativecommons.org/licenses/by/4.0/).
Iron homeostasis is essential for maintaining metabolic health and iron disorder has been linked to chronic metabolic diseases. Increasing thermogenic capacity in adipose tissue has been considered as a potential approach to regulate energy homeostasis. Both mitochondrial biogenesis and mitochondrial function are iron-dependent and essential for adipocyte thermogenic capacity, but the underlying relationships between iron accumulation and adipose thermogenesis is unclear. Firstly, we confirmed that iron homeostasis and the iron regulatory markers (e.g., Tfr1 and Hfe) are involved in cold-induced thermogenesis in subcutaneous adipose tissues using RNA-seq and bioinformatic analysis. Secondly, an Hfe (Hfe−/−)-deficient mouse model, in which tissues become overloaded with iron, was employed. We found iron accumulation caused by Hfe deficiency enhanced mitochondrial respiratory chain expression in subcutaneous white adipose in vivo and resulted in enhanced tissue thermogenesis with upregulation of PGC-1α and adipose triglyceride lipase, mitochondrial biogenesis and lipolysis. To investigate the thermogenic capacity in vitro, stromal vascular fraction from adipose tissues was isolated, followed with adipogenic differentiation. Primary adipocyte from Hfe−/− mice exhibited higher cellular oxygen consumption, associated with enhanced expression of mitochondrial oxidative respiratory chain protein, while primary adipocytes or stromal vascular fractions from WT mice supplemented with iron citrate) exhibited similar effect in thermogenic capacity. Taken together, these findings indicate iron supplementation and iron accumulation (Hfe deficiency) can regulate adipocyte thermogenic capacity, suggesting a potential role for iron homeostasis in adipose tissues.

Keywords

iron homeostasis
adipose thermogenesis
stromal vascular fractions
HFE
Abbreviations

AMPK adenosine 5′-monophosphate-activated protein kinase

BAT brown adipose tissue

CST Cell Signaling Technology

DAB 3,3′-diaminobenzidine

DEG differentially expressed gene

DMEM Dulbecco's modified Eagle's medium

FAC ferrous ammonium citrate

FBS fetal bovine serum

Fth ferritin heavy chain

iWAT inguinal white adipose tissue

KEGG Kyoto Encyclopedia of Genes and Genomes

RT room temperature

Sf serum transferrin

SVF stromal vascular fraction

TCA tricarboxylic acid

Tfr transferrin receptor

Reviewed by members of the JBC Editorial Board. Edited by Donita C. Brady
==== Body
pmcEnergy expenditure is a fundamental aspect of metabolic balance, and is influenced by body composition, physical activity, and dietary preference (1). Basic thermogenesis or basal heat production, involves energy consumption in both basal metabolism and physiological states, also referred to as resting metabolic rate or resting energy expenditure (1,2). Adaptive thermogenesis, particularly under cold stimulation, reflects an organism's ability to generate heat through metabolic processes in adipocytes, utilizing chemical energy stored in the form of carbohydrates and fats (3). Enhancing adipocyte thermogenic capacity could be a promising approach to counteract metabolic imbalances associated with obesity and type 2 diabetes (4, 5, 6).

Mitochondrial biogenesis and function are central to the maintenance of adipocyte thermogenic capacity, where both iron-sulfur clusters and heme proteins play critical roles during this biological process. These components are instrumental in mitochondrial processes, including the tricarboxylic acid (TCA) cycle and fatty acid β-oxidation (7). Cellular iron regulation is mediated through a coordinated system involving iron transport receptors, storage proteins, pumps, and sensing complexes (8,9). Serum transferrin (Tf), the primary iron carrier, exists in iron-free (Apo-Tf) and iron-bound (Holo-Tf) forms, with Holo-Tf facilitating iron uptake via transferrin receptors (Tfr) (8). The interaction between Tfr1, hemochromatosis protein (HFE), bone morphogenetic protein receptors, and hematopoietin forms a complex that regulates hepcidin expression, a key modulator of systemic iron balance (8).

Previous studies have underscored a close association between iron homeostasis disruption and chronic metabolic diseases like obesity, type 2 diabetes and hepatic fibrosis (10,11). Obese adolescents in some regions, for instance, the prevalence of iron deficiency and iron deficiency anemia is notably higher than normal-weight counterparts, with a negative correlation between transferrin saturation and body mass index (12). Obese children in some regions exhibited a lower serum iron levels and transferrin saturation, alongside elevated serum hepcidin and leptin (13).

The perturbation of iron levels has profound implications for adipose tissue metabolism and plasticity. Our previous study and others found adipose-specific ablation of Tfr1 results in impaired thermogenic capacity in brown or beige adipocytes (14,15). Meanwhile, our previous work also indicated that iron deficiency induced in a model lacking transmembrane serine protease 6, exhibits impaired thermogenesis in beige or brown fat (14). Conversely, increased iron stores, through hepcidin absence, can attenuate insulin resistance and inflammation in adipose tissue induced by a high-fat diets (16). Hfe deficiency, primarily impacting liver iron homeostasis (17), also affects other organs including spleen (18), muscle (19), and adipose tissues (20,21). Studies have shown that iron accumulation affects fatty acid oxidation in muscle tissues of Hfe KO mice (19). However, the effects of increased iron storage due to Hfe deficiency on energy metabolism in adipose tissue remain underexplored. Our study, through transcriptomic analysis and cellular extracellular flux assessments in combination of in vivo and in vitro models, demonstrates that both ferrous ammonium citrate (FAC) supplementation and Hfe deficiency—induced iron storage can regulate thermogenic capacity in adipocytes and stromal vascular fractions (SVFs) of adipose tissue.

Results

Iron supplementation regulates energy consumption in SVFs and primary adipocytes

Investigating the impact of iron supplementation on adipocyte function, SVFs from inguinal white adipose tissue (iWAT) of C57BL6/J mice were isolated and differentiated into primary adipocytes. Prior to the differentiation into primary adipocytes, SVFs cell were treated with FAC at 100 μM. This treatment resulted in significant upregulation of the thermogenic gene Cox8b, along with notable changes in basal respiration, proton leak, and ATP production, pointing to alterations in metabolic activity due to iron supplementation (Fig. 1, A–C). Postdifferentiation, primary adipocytes with FAC-treated exhibited a pronounced increase in thermogenic markers (Ucp1 and PGC1α) and mitochondrial complex proteins (Fig. 1, D and E). This was paralleled by a reduction in lipid droplet size within the adipocytes (Fig. 1F). Further gene expression analysis post-FAC treatment revealed a significant elevation in thermogenic genes (Ucp1, Prdm16, and Cox8b) (Fig. 1G). Additionally, the iron-responsive gene Tfrc was notably downregulated following FAC supplementation, while iron storage (Fth1 and Ftl1) and transport genes (Slc11a2 and Slc40a1) showed varied responses (Fig. 1G). Mitochondrial stress tests conducted on these differentiated primary adipocytes indicated enhanced basal and maximal respiration rates, with no significant alterations in proton leak, ATP production, or spare respiratory capacity (Fig. 1, H and I). These findings underscore the role of iron in modulating thermogenesis and metabolic responses both in predifferentiated SVFs and in differentiated primary adipocytes, highlighting the potential of iron supplementation to influence adipocyte function and energy consumption in vitro.Figure 1 Iron supplementation regulates energy consumption in stromal vascular fractions and primary adipocytes.A, qPCR analysis of iWAT SVF between H2O and FAC 100 μM group (n = 10/group). B, representative oxygen consumption rate of iWAT SVFs in Mito-stress test between control and FAC 100 μM group (n = 5/group). C, assay parameters of the Mito-stress test in iWAT SVFs (n = 5/group), similar result obtained in three independent experiments. D, representative immunoblots of mitochondrial complex, Ucp1, Tfr1, Fth, PGC-1α, and tubulin of iWAT primary adipocytes, an iWAT-cold tissue sample as a positive control for Ucp1 (n = 3/group). E, protein level analysis of the bands (n = 9/group), pooled from three independent experiments. F, representative images of fluorescence staining of iWAT primary adipocytes between control and FAC 100 μM treatment group and lipid droplet analysis (n = 15 ∼ 20 wells/group). G, qPCR analysis of iWAT primary adipocytes between control and FAC group (n = 11 ∼ 12 wells/group). H, representative oxygen consumption rate of iWAT primary adipocytes in Mito-stress test between control and FAC 100 μM group (n = 4 wells/group). I, assay parameters of the Mito-stress test in iWAT primary adipocyte (n = 4 wells/group), similar trends obtained in three independent experiments. Data showed as the mean ± SD. ∗p < 0.05, ∗∗p < 0.01, and ∗∗∗p < 0.001, N.S., no significance. Unpaired student’s t test for two group comparison. FAC, ferrous ammonium citrate; Fth, ferritin heavy chain; iWAT, inguinal white adipose tissue; qPCR, quantitative PCR; SVF, stromal vascular fraction; Tfr, transferrin receptor.

Role of iron in cold-induced thermogenesis in adipose tissue

The effects of cold exposure on iron metabolism in adipose tissues were examined by analyzing brown adipose tissue (BAT) and iWAT from 10 ∼ 15-week-old C57BL6/J mice under room temperature (RT) and cold conditions (Fig. 2, A and B). Cold exposure led to a gradual increase in mRNA levels of thermogenic genes (Ucp1, Ppargc1α, Cox7a1, and Cox8b) in BAT, with a modest increase in Ucp1 protein on the seventh day of cold stimulation (Fig. 2, C and E). Similar trends were observed in iWAT, with increased mRNA and protein levels of thermogenic markers and smaller lipid droplets (Fig. 2, A, D, and F). RNA-seq analysis revealed enrichment in mitochondrial organization, fatty acid metabolism, oxidative phosphorylation, and fatty acid oxidation processes in iWAT postcold exposure (Fig. S1A). Cellular component and molecular function enrichment analyses indicated upregulated gene enrichment in mitochondrial membranes, respiratory chain complexes, and iron-binding molecules (Figs. 3A and S1B). Kyoto Encyclopedia of Genes and Genomes (KEGG) pathway analysis highlighted enhancements in thermogenesis, oxidative phosphorylation, TCA cycle, and adenosine 5′-monophosphate-activated protein kinase (AMPK) signaling pathways in iWAT post cold exposure (Fig. 3B), with iron-sulfur cluster enrichment (Fig. 3C). Similar enrichments were observed in BAT (Fig. S1, C–F). Differential gene expression analysis revealed gradually upregulated Tfr1 and gradually downregulated Hfe in iWAT during cold exposure, indicating a role for iron in cold-induced adipose thermogenesis (Figs. 3, D and E, and S1G).Figure 2 Cold-induced thermogenesis in adipose tissues.A, representative H&E staining images of BAT and iWAT from 10 ∼ 15-week-old WT mice at room temperature and cold challenge. B, representative images of BAT and iWAT from 10 ∼ 15-week-old WT mice in room temperature and cold 7 days. C, Western blot results of tubulin and Ucp1 in BAT and iWAT (D) from 10 ∼ 15-week-old WT mice under room temperature and cold challenge. E, qPCR analysis of BAT and iWAT (F) under room temperature and cold stimulation time points (n = 3 mice/group). Data show as the mean ± SD. ∗p < 0.05, ∗∗p < 0.01, and ∗∗∗p < 0.001, N.S., not significant, compared with RT. Unpaired student’s t test for two group comparison. One-way ANOVA analysis for multiple groups comparison with a bonferroni post hoc analysis. BAT, brown adipose tissue; iWAT, inguinal white adipose tissue; qPCR, quantitative PCR; RT, room temperature.

Figure 3 Iron metabolism involved in cold induced adipose thermogenesis process.A, GO analysis of molecular function (MF) of iWAT from 10 ∼ 15-week-old WT mice between room temperature and cold exposure groups. B, KEGG analysis of iWAT from 10 ∼ 15-week-old WT mice between room temperature and cold exposure groups. C, GSEA enrich analysis of iron-sulfur cluster assembly between room temperature and cold exposure groups in iWAT in WT mice. D, volcano map of different expression genes in iWAT between room temperature and cold exposure groups. E, heatmap analysis of iron metabolism–related genes in iWAT between room temperature and cold exposure groups (n = 3 mice/group). GSEA, gene set enrichment analysis; GO, gene ontology; iWAT, inguinal white adipose tissue; KEGG, Kyoto Encyclopedia of Genes and Genomes.

Iron accumulation in Hfe deficiency modulates adipose thermogenesis

To explore the influence of iron accumulation on adipose thermogenesis, Hfe KO mice (Hfe−/−), a model of hereditary hemochromatosis with systemic iron overload, were used (Fig. S2A). Increased iron levels in serum, adipose tissues, and liver were observed in Hfe−/− mice, compared to WT mice (Fig. S2, B and C). Smaller lipid droplets and relative higher expression of thermogenic proteins were noted in iWAT from 30 ∼ 35-week-aged Hfe−/−mice (Figs. 4, A–F and S2, D and E). H&E staining and immunohistochemistry staining indicated these findings (Fig. 4, B and C). Increased ferritin heavy chain (Fth) protein level in adipose tissues of Hfe−/− mice indicated iron accumulation (Fig. 4, D–F). Transcriptome analysis of WT and Hfe−/− mice revealed upregulation of genes associated with oxidative respiratory chain complexes in Hfe−/− mice's iWAT, including Ndufa1, Sdhb, Cox7a1, Cox8b, and Atp5a1 (Fig. S2, F and G). KEGG pathway analysis indicated enrichment in oxidative phosphorylation, thermogenesis, TCA cycle, and fatty acid metabolism in iWAT of Hfe−/− mice (Fig. S2H), with gene set enrichment analysis indicating these findings (Fig. S2, I–K), along with the validation of protein level supporting the enhanced expression of mitochondrial respiratory chain complexes (Fig. S2L). Genes expression analysis also demonstrated an upregulation of Cox8b in Hfe−/− mice's BAT (Fig. S3, A and B).Figure 4 Iron accumulation caused by Hfe deficiency regulate adipose thermogenesis.A, representative images and H&E staining (B) of BAT, iWAT from 30 ∼ 35-weeks-aged WT, and Hfe−/− mice in RT and 7 days cold exposure. C, IHC staining (Ucp1 and PGC-1α) of iWAT from 30 ∼ 35-week-aged WT and Hfe−/− in RT and cold exposure. D, representative Prussian blue-3,3′-diaminobenzidine staining images of BAT, iWAT, liver from 30 ∼ 35-week-aged WT, and Hfe−/− in RT and cold exposure. E, representative protein immunoblots of Ucp1, Tfr1, Fth, PGC1α, and tubulin of BAT and iWAT (F) from 30 ∼ 35-week-aged WT and Hfe−/− in RT and cold exposure (n = 3 mice/group), similar trends obtained in two independent experiments. BAT, brown adipose tissue; IHC, immunohistochemistry; Fth, ferritin heavy chain; iWAT, inguinal white adipose tissue; RT, room temperature; Tfr, transferrin receptor.

Impact of Hfe deficiency and iron supplementation on SVFs and adipocytes in vitro

The effects of Hfe deficiency and iron supplementation on adipocyte thermogenesis were further assessed in vitro using isolated SVFs from WT and Hfe−/− mice. Fluorescent staining revealed smaller lipid droplets in Hfe−/− adipocytes (Fig. 5A). Increased protein expression of thermogenic markers and mitochondrial complexes was observed in Hfe−/− adipocytes (Figs. 5B and S4, A and B). Quantitative PCR analysis supported the enhanced thermogenic capacity in Hfe−/− adipocytes (Fig. 5C). Seahorse XF Mito-stress test results indicated increased maximum oxygen consumption and spare respiratory capacity in Hfe−/− adipocytes (Fig. 5, D–F). Increased mRNA expression of thermogenic and iron homeostasis genes was noted in SVF cells from Hfe−/− mice (Fig. 5G). Adipose triglyceride lipase (ATGL) is one of the important lipases on adipocyte lipolysis (22). In current investigation, we also found Atgl mRNA expression and ATGL protein level were increased in iWAT of Hfe−/− mice (Fig. S4, F–H), while Atgl mRNA expression also increased in Hfe−/− primary adipocytes (Fig. 5C). In addition, we also found BAT primary adipocytes exhibited modest increased protein levels of Ucp1 and PGC-1α in Hfe−/− group, indicating elevated adipose thermogenesis (Fig. S4, D and E). Moreover, stem cell markers in SVFs revealed significant changes post-FAC treatment, indicating a potential of reprogramming of SVFs before differentiation into mature adipocyte (Fig. 5H).Figure 5 Impact of Hfe deficiency and iron supplementation on SVFs and adipocytes in vitro.A, representative images of fluorescence staining and lipid droplets area analysis (n = 13 ∼ 17 wells/group) of iWAT primary adipocytes from WT and Hfe−/−. B, representative immunoblots of Ucp1, PGC-1α, mitochondrial complex, and tubulin of iWAT primary adipocytes from WT and Hfe−/− (n = 3/group), similar results were seen in three independent experiments, iWAT-cold tissues sample as a positive control for Ucp1. C, qPCR analysis of gene expression in iWAT primary adipocyte of WT and Hfe−/− under differentiation for 7 days (n = 9 wells/group). D, experimental scheme of XF Mito-stress test for primary adipocyte. E, oxygen consumption rate of Mito-stress test and (F) Assay parameters of Mito-stress test of iWAT primary adipocyte from WT and Hfe−/− after adipogenic differentiation (n = 19 ∼ 20 wells/group). G, qPCR analysis of mRNA expression in SVF from iWAT between WT and Hfe−/− group (n = 18 ∼ 19 wells/group). H, qPCR analysis of stem cell marker mRNA expression in SVF from iWAT in WT, WT + FAC 100 μM treatment and Hfe−/− group (n = 13 ∼ 14 wells/group), pooled from three independent experiments. Data show as the mean ± SD. ∗p < 0.05, ∗∗p < 0.01, and ∗∗∗p < 0.001, N.S., not significant. Unpaired student’s t test for two group comparison. qPCR, quantitative PCR; FAC, ferrous ammonium citrate; iWAT, inguinal white adipose tissue; SVF, stromal vascular fraction.

Discussion

Iron homeostasis disorder is closely associated with chronic diseases such as obesity and metabolic syndrome (11). However, the underlying relationships between iron accumulation (or iron overloading) and adipose thermogenesis are less investigated. In the present study, we used Hfe−/− mice and WT mice to explore how iron accumulation is linked to heat production in adipose tissues. Employing RNA-seq bioinformatics analysis, animal and primary adipocyte models, we identified the relationship between iron homeostasis genes and adipose thermogenesis. Along with iron accumulation, Hfe deficiency partially modulates adipose thermogenesis and lipid metabolism.

One of our findings is that extracellular iron supplementation increased oxygen consumption and enhanced the thermogenic capacity in SVF and primary adipocytes (Fig. 1). However, it may cause adverse effects with excessive iron intervention in different cell lines (23,24). Conversely, deferoxamine , an iron chelating agent, can suppress adipogenic differentiation and reduce adipocyte thermogenic capacity and downregulate lipolysis (25). This information indicates that appropriate iron level is critical for the regulation of cell mitochondrial function and cell differentiation. Additionally, we classified multiple iron homeostasis genes involved in adipose tissues during cold-induced thermogenesis, paralleling to β3-agonist induced thermogenesis (14). Consistently, our RNA-seq data revealed an upregulation of Tfr1, the gene for Tfr1, during the cold exposure (Fig. 3D), aligning with previous studies showing Tfr1 enrichment in beige adipocytes under β-adrenergic receptor activation (14). However, homeostatic iron regulator (Hfe), known as a pathogenicity gene of hereditary hemochromatosis with multiple tissues excessive storage of iron (17), was identified to be slowly downregulated in iWAT during cold exposure based on the RNA-seq data (Figs. 3E and S1G). Besides, there may be a time lag on Hfe expression level in white adipose tissue, slowly downregulated in β3-agonist (CL-316,243) induced beiging and slowly upregulated upon its withdrawal (26). It is plausible that the expression of Hfe is associate with the thermogenic capacity of adipose tissues and its relationship between Hfe gene expression and thermogenesis in adipose tissues may require additional investigation.

Numerous studies have confirmed that Hfe is an important regulator of iron metabolism (17). Its deletion or mutation can regulate the expression of hepcidin, thereby causing a decline in the negative regulation of iron homeostasis, resulting in a state of higher iron uptake and accumulation in multiple cells and tissues (17,27). Notably, a recent study found that hepcidin (a target protein of Hfe) is necessary for browning capacity in white adipose tissue in mice (28). However, a previous study has reported that iron accumulation in the muscles of Hfe−/− mice and enhanced fatty acid oxygen consumption alongside decreased glucose utilization, linked to an increased expression of carnitine palmitoyl transferase b) and decrease in pyruvate dehydrogenase) enzyme activity (19). Furthermore, increased mitochondrial respiratory capacity in Hfe−/− mice's liver was noted on a normal chow diet, but not on a high-iron diet (27). Indirect calorimetry analysis also indicated a significant increases of heat production and oxygen consumption in Hfe−/− mice with high-fat diet challenge (19). Other than these Hfe−/− studies, it was confirmed that iron accumulation occurred in tissues and serum. Fth, a crucial subunit of ferritin and an important structure for iron storage in cells or tissues (8) and it is essential for mice energy homeostasis and adaptive thermogenesis, which was proved in mice of Fth deficiency (29). This is associated with iron accumulation and high expression of Fth in Hfe−/− mice. As is well known, thermogenic markers in white adipose tissues can be dramatically promoted in cold exposure, compared with RT condition. However, aging is one of the important negative factors in regulating adipose tissues thermogenesis (30), which is also an important reason that aged mice are relatively not tolerant to cold exposure. Previous studies have found an impaired thermogenic capability during cold induced WAT beiging in middle aged WT mice (28-week-old), compared with young mice (8-week-old) (31). Interestingly, along with Fth higher expression in iWAT, we found Ucp1 and PGC-1α protein expression were higher tendency in 30∼35-week-aged Hfe−/− mice, indicating that the iron accumulation status in Hfe−/− mice may play an important role in white adipose tissue thermogenesis and lipid metabolism (Fig. 4). Considering that the iron accumulation level could be a one of the potential factors affecting adipose thermogenesis, it is worth further exploring in the animal models with more server iron overloaded (such as Hjv KO mice, Hfe and Hjv double KO mice (32)) in the future. Notably, the importance of lipolysis in energy homeostasis has been illustrated in numerous studies and the lipase play an important role in adipose lipolysis activity and lipid metabolism (22,33,34). Previous studies have been suggested that both transferrin and iron treatment can contribute to the enhanced lipolytic effect in adipocytes (35). In our study, we speculated that enhanced ATGL expression may contribute to the lipolysis in this iron accumulated status and affect the energy homeostasis (Fig. S4, F–H). SVF and adipocytes are essential parts in adipose tissues, responding to stimuli and metabolic changes (36,37).The SVF derived from adipose tissue contains abundant mesenchymal stem cells with multidirectional differentiation capacity and plasticity (38, 39, 40). Therefore, we isolated SVF from adipose tissues and induced adipogenic differentiation and significant increase in maximal respiration was observed in Hfe−/− primary white adipocytes. This increased oxygen consumption appears to be mainly related to higher expression of Ucp1, PGC-1α, and mitochondrial complex (Fig. 5B).

However, it is also necessary to be noticed that the overall changes of body metabolism in Hfe deficiency is the result of the response of multiple metabolic organs on iron accumulation. Apart from the metabolic changes in adipose tissues and adipocytes, it is also necessary to mention that Hfe deficiency or mutation can increase the risk of some hepatic disease (such as hepatic fibrosis and nonalcoholic fatty liver disease, due to the cell damage and iron overloaded in liver tissue) and glucose metabolic disorders (41, 42, 43). Besides, according to dietary iron supplementation investigations, high-iron diet can induce lower fat mass, higher oxygen consumption (44), and promote fatty acid oxidation (45), but it may also lead to an impaired fasting glucose associated with insulin resistance (46). Therefore, the disadvantage effects of iron accumulation or iron dietary iron supplementation should not be ignored and it is worth more exploration and investigation to precisely regulate iron content and metabolism in different metabolic tissues.

The differentiation and development of iWATs can be traced to a subgroup of adipose vascular endothelial cells, which highly express stem cell markers. Therefore, we speculated that another potential mechanism of iron supplementation promoting cell thermogenesis may be related to cell differentiation. Here, our experiment confirmed iron supplement–enhanced mitochondrial complex genes expression and promoted oxygen consumption rates. The enhanced oxygen consumption metabolic shift was not only found in primary adipocytes but also found in SVF, a population enriched for progenitor cells. Meanwhile, this metabolic shift under iron intervention could be related to higher expressed Cd34 in mice SVF cells (Fig. 5H). Previous studies have showed enhanced proliferative capacity in CD34+ human adipose–derived stem/progenitor cell (47), while human primary adipocyte differentiated from human adipose–derived stem/progenitor cell with high CD34 expression exhibit higher lipid turnover (fatty acid intake, lipidation, and lipolysis) (48). Interestingly, although there is enhanced oxygen consumption in Hfe−/− primary adipocytes and higher expression of thermogenic markers (Cox8b) in Hfe−/− SVF (Fig. 5), no difference was observed in the stem cell markers in Hfe−/− SVF group under normal medium without FAC supplement. This suggests that there is other mechanism involved in regulating energy homeostasis in Hfe−/− SVF group and it may help to identify cell subpopulations of SVF with single cell analysis in future investigation.

Moreover, recent study provided evidences that an adipose-liver tissue crosstalk in iron influx for adipose browning mediated by FoxO1-Tgfβ1 signaling (49). Interestingly, we also found Tgfβ1 signaling pathway markers were partially reduced in subcutaneous white adipose tissue in Hfe−/− mice (Fig. S2M). It is plausible that tgfβ1 signaling pathway may also contribute to the thermogenesis in Hfe−/− mice. Due to the impaired adipogenic and lipogenic effect by deferoxamine intervention, it is worth mentioning that recent work also found that iron can organize adipogenic genes expression during 3T3-L1 cell early-stage differentiation, which is mediated by the histone demethylase jumonji domain containing 1A and the DNA demethylase ten-eleven translocation 2 (50). However, considering the differences in adipogenic differentiation capacity between preadipocyte cell lines and progenitor in SVF, it is necessary to further explore the iron-dependent regulation in adipocyte differentiation and thermogenic capacity in future studies.

In summary, our work demonstrated the thermogenic promotion of iron supplementation in primary adipocytes and SVFs. We observed an enhanced thermogenic capacity in adipose tissues and primary adipocytes from the animal model with iron accumulation, suggesting a potential role for iron homeostasis in adipose function.

Limitations of the study

Considering that in animal models of hereditary hemochromatosis (iron accumulation), in addition to Hfe gene knockout, there are several mouse models with Hamp, Hjv, and Slc40a1 (coded iron pump protein) knockout. There may be different iron accumulation level among the mice models in different ages. Therefore, it is worth exploring whether there are differences in metabolic phenotypes of adipose tissues in different model of iron accumulation in future investigation.

Experimental procedures

Experimental animals

The Hfe−/− mice were backcrossed with C57BL6/J mice, reported in the previous studies (51), kindly gifted from Dr Fudi Wang at the Zhejiang University. The mice were housed in specific pathogen-free cages, under a 12-h/12-h light and dark cycle at 23 ± 1 °C. Mice were fed a standard rodent chow diet with free access to food and water. For cold induced thermogenesis time course experiments, 10∼15-week-old male WT and Hfe−/− mice at RT) or cold exposure (5 ± 1 °C) were sacrificed to collect the tissue samples. For middle aged mice cold challenge, 30 ∼ 35-week-old WT and Hfe−/− mice were administered in separate cages at cold exposure (5 ± 1 °C) with sufficient food and water under light and dark cycle (12 h/12 h). After cold exposure, mice were sacrificed to collect the tissue samples for follow-up measurements. Animal experiments were approved by the Animal Care and Use Committee at Guangdong Institute of Microbiology (GT-IACUC201704071) or Animal Experiment Center of Zhujiang Hospital of Southern Medical University (LAEC-2022-004).

Total RNA isolation and quantitative real-time PCR

The total RNA from tissues or cells were extracted with TRIzol reagent (Thermo Fisher Scientific), according to the manufacturer’s instruction. RNA concentration was determined using the NanoDrop absorbance spectroscopy (Thermo Fisher Scientific), followed by reverse transcription to complementary DNA utilizing the 5× All-In-One Master Mix (G490, AbmGood). Complementary DNA was used to determine the gene expression using the SYBR Green Master Mix (A25778, Applied Biosystems) on a QuantStudio 6 Flex Real-Time PCR System (Thermo Fisher Scientific). The gene expression was normalized to the expression of 18S ribosomal RNA. All primer sequences for quantitative PCR were listed in Table S1.

Protein isolation and Western blot analyses

Total protein from tissues or primary cells were extracted with radio immunoprecipitation assay buffer, supplemented with 1× proteasome inhibitor cocktail (Thermo Fisher Scientific). The protein concentration was determined by using bicinchonininc acid protein assay kit (Thermo Fisher Scientific). The protein samples were separated using PAGE and transferred to polyvinylidene fluoride membranes (Merck Millipore) with precooled transfer buffer. Polyvinylidene fluoride membrane with proteins was blocked with 5% (w/v) nonfat milk at RT for 1 h, followed by overnight incubation with primary antibodies: (Tubulin: sc-365791, Santa Cruz); (PGC1α: AB3242, Millipore); (Ucp1:ab10983, Abcam); (total-HSL, #4107, p-HSL(Ser565), #4137, Cell Signaling Technology, CST.); (total-AMPK, #5831, p-AMPK (Thr172), #2523, CST.); (ATGL:#2138, CST.); (Glut4: A7637, Abclonal); (Tfr1: ab84036, Abcam); (Fth: sc-376594 Santa cruz); and (Mito-complex antibody: ab110413, Abcam) on a rocker overnight in 4 °C refrigerator and the secondary antibodies were used for incubation at RT for 1 h. Bands were visualized using enhanced chemiluminescence reagents (Abclonal) using the ChemiDoc Imaging System (Bio-Rad). Protein expressions data were quantified by using ImageJ software (https://imagej.net/).

Tissues and serum iron estimation

Non-heme iron levels of adipose tissues, liver, and serum iron levels were detected following a standard ferrozine assay protocol as described before (14,52). Briefly, tissues were homogenized with ddH2O, followed by the addition of equal volume of 10% trichloroacetic acid in 3 M HCl. After 1 h digestion at 100 °C, 50 μl of each sample and iron standard were incubated at 37 °C for 1 h with equal volume ferrozine working solution (50 μl 100 mM ferrozine, 72 μl 70% mercaptoacetic acid, 2.5 ml 3 M sodium acetate in 5 ml total volume with ddH2O). After incubation, absorbance (A) value at 562 nm was measured by a microplate photometer (Thermo Fisher Scientific). The individual wet tissues weight was used to normalize the iron level.

RNA-seq and bioinformatics analysis

Total RNA samples were sequenced using a BGI-SEQ2500 platform (Beijing Genomics Institute). The high-quality RNA-seq reads were further aligned to the mouse genome (GRCm38/mm10) using HISAT2 and assembled against mouse mRNA annotation using high-throughput sequencing on a high-performance computational system. Differentially expressed genes (DEGs) were analyzed using the DESeq2 (https://bioconductor.org/packages/release/bioc/html/DESeq2.html) package in R. Genes were considered significantly upregulated or downregulated at p-adj < 0.05. Heatmaps were generated using the heatmap package in R based on the raw count of DEGs. Gene ontology analysis was performed using the R package ClusterProfiler for DEGs (upregulated or downregulated). DEGs (p-adj < 0.05) were further analyzed using gene set enrichment analysis. Both upregulated and downregulated genes were functionally categorized using gene ontology and KEGG pathway enrichment analyses.

SVF isolation and primary cell culture

The SVF of BAT and iWAT were isolated as previously described (53). Adipose tissue from 5∼7-week-old mice was carefully separated into enzyme working solution (1.5 mg/ml collagenase type II, 2.4 U/ml dispase II, and 10 mm CaCl2, filtered through a 0.22 μm filter), scissors cut to pieces. Digestion was carried out in a water bath at 37 °C for 30 min, and the mixture was observed every 10 min. The digested tissue fragments were filtered through a 40 μm sterile filter, neutralized in medium containing 10% fetal bovine serum (FBS), and centrifuged (1000×g, 10 min) to precipitate cells as SVF. The cells were resuspended in complete medium (Dulbecco's modified Eagle's medium [DMEM] supplemented with 10% FBS, 1% Penicillin, Streptomycin) and seeded into 24-well plates overlaid with collagen. After 5 h, the cells were rinsed with complete medium (DMEM, 10% FBS, 1% P.S) to remove tissue debris and red blood cells. Cultures were incubated in a 5% CO2 incubator at 37 °C. After cells 100% adherence, they were induced adipocyte differentiation. BAT primary cell culture protocol: 3 days in induction medium, a fresh complete DMEM-high glucose medium with 3-isobutyl-1-methylxanthine, 0.5 mM (Sigma: I-5879); indomethacin, 125 μM (Sigma: I7378); dexamethosone, 2 μg/ml, (Sigma: D1756); insulin, 1 μM (Solarbio, I8830); T3, 1 nM (Sigma: T2877); rosiglitazone, 1 μM (Sigma: R2408), along with differentiation medium (insulin, 1 μM, Solarbio I8830; T3, 1 nM, Sigma: T-2877; rosiglitazone, 1 μM) for 7 days culture. iWAT primary cell culture protocol: induction medium supplemented with 3-isobutyl-1-methylxanthine 0.5 mM; indomethacin, 125 μM; dexamethosone, 2 μg/ml; rosiglitazone, 1 μM, culture for 3 days, and cells induced differentiation with differentiation medium (insulin, 1 μM and T3, 1 nM) for another 7 days. FAC treatment was performed in differentiation medium for 7 days. Bodipy staining and 4′,6-diamidino-2-phenylindole staining were performed by using lipid probes (Thermo Fisher Scientific, D-3834) and Hoechst reagent (Invitrogen, H1399) before cell harvested. Images were captured by using fluorescence microscopes (Invitrogen EVOS), and adipocyte lipid droplet sizes were analyzed by using ImageJ. The analysis was performed with minor adjustments as previously described (54). Briefly, after setting the scale of the images, 8 bit images were converted into binary images and watershed for lipid object separation. The size (μm2) of lipid droplets were displayed by ImageJ and presented in GraphPad prism 8.0 (https://www.graphpad.com) with relative frequency.

Oxygen consumption rate assays

After 3 days of induction, the primary cells were digested with 0.25% trypsin and seeded into seahorse V28 well plates (∼20,000 cells/well), and white SVF cells from WAT were seeded in seahorse V7-PS cell plates. Oxygen consumption rate assays were performed on the seventh day of primary adipocyte differentiation or on the third day of SVF cells with FAC treatment. Preheat the machine in advance and hydrate the probe plate overnight. Cell culture plates were rinsed three times with working assay solution (pH 7.4) (1 mM glucose, 2 mM glutamine, and 10 mM pyruvate). Finally, 500 μl working solution was added per well. The cells were incubated in a CO2-free incubator at 37 °C for 50 min. The hydrated probe plate was added with drugs (oligomycin 1.5 μM, carbonyl cyanide 4-(trifluoromethoxy)phenylhydrazone 4.0 μM, antimycin/rotenone 1.0 μM), and then put into the cell plate to run the XF Cell Mito Stress Test program. Data were collected and normalized with cell protein level in Seahorse Software Wave Desktop (V2.6).

H&E staining

Fresh tissues were immersed in 10% formalin for 48 h, dehydrated, and made into paraffin-embedded sections with a thickness of 4 μm. For H&E staining, paraffin wax was melted (65 °C, 40 min), sections were deparaffinized by immersing in xylene (three times for 5 min each), and xylene was rinsed with alcohol series. The tissues were stained with hematoxylin for 3 min, differentiated in 0.3% hydrochloric alcohol for 1 ∼ 2 s, and blued with ammonia solutions. The sections were immersed in eosin, dehydrated with alcohol series, dried with xylene, and mounted with mounting medium (Macklin, 96949-21-2). Images were captured by microscope system (Hitachi) or scanned on automatic digital slice scanning system (Guangzhou Betrue Technology Co, Ltd).

Immunohistochemistry

Paraffin slices with tissue sections at 4 μm thickness were deparaffinized in xylene and hydrated in alcohol series, followed with ddH2O rinsed. Soak the slides in Tris EDTA buffer in a boiling water bath for 1 h for antigen retrieval. After cooling to RT, the slides were washed with PBS two times for 5 min, followed with 0.5% Triton X-100 permeabilization for 10 min. Incubate at RT with 3% H2O2 for 10 min to eliminate endogenous peroxidase activity and wash with ddH2O three times. Sections were blocked in 3% bovine serum albumin containing 5% goat serum for 1 h, followed with primary antibody incubation (Ucp1, 1:500, abcam; PGC1α, 1:500, millipore) at 4 °C overnight. After incubation, the sections were washed with PBS for three times and incubated secondary antibody (1:500) in 3% bovine serum albumin containing 5% goat serum for 1 h at RT. The sections were incubated with 3,3′-diaminobenzidine (DAB) reagent for 10 ∼ 12 min. Restained the nucleus for 3 min with hematoxylin, followed with differentiator to remove excess dye. After water washing, bluing with ammonia solution for 10 min. The sections were dehydrated and cleared with alcohol series and xylene. Sections were mounted with mounting reagent and images captured and scanned as described above.

Prussian blue-DAB staining

Prussian blue-DAB staining was as performed as described (55). Paraffin slices dewaxed, rehydrated with standard procedures, and incubated in 1% H2O2 in methanol for 10 min. After ddH2O washing, sections were immersed in 2% HCL and 2% potassium ferrocyanide at ratio of 1:1 for 30 min. Then sections were washed with ddH2O and immersed in 0.05% DAB in PBS pH 7.4 for 10 min. Sections were immersed in 0.044% H2O2 with 0.05% DAB in PBS pH 7.4 for another 10 min and washed with ddH2O. Then sections were dehydrated in alcohol series and dried with xylene, followed by mounting and images were captured and scanned.

Statistical analysis

Data were presented as the mean ± SD. Statistical analysis was performed in GraphPad prism 8.0 by using the unpaired Student’s t test for two group comparison. One-way ANOVA analysis was used for multiple groups comparison, followed by a Bonferroni post hoc analysis. p values is shown as ∗p < 0.05, ∗∗p < 0.01, and ∗∗∗p < 0.001 for showing differences and the NS stands for not significant (p > 0.05).

Data availability

RNA-seq sequencing data were uploaded at National Microbiology Data Center (NMDC: http://nmdc.cn/) and requests for data should be approved by H. C. (chenhong123@smu.edu.cn) and L. X. (xielw@gdim.cn). Further information and requests for resources and reagents should be directed to and will be fulfilled by the contact, H. C. (chenhong123@smu.edu.cn) and L. X. (xielw@gdim.cn).

Supporting information

This article contains supporting information (14, 49).

Conflict of interest

The authors declare that they have no conflicts of interest with the contents of this article.

Supporting information

Supporting information figure legends

Supporting information table

Figure S1

Figure S2

Figure S3

Figure S4

Author contributions

X. M. and L. X. methodology; X. M. and L. X. formal analysis; X. M., J. F., L. X., Z. H., S. H., R. S., X. J., L. H., J. S., and L. X. data curation; Y. L., L. X., and H. C. conceptualization; L. X. writing–review and editing; L. X. and H. C. supervision; L. X. resources; L. X. project administration; L. X. and H. C. funding acquisition; X. M. and Y. L. validation; Y. L. visualization; X. M., J. F., M. L., Z. H., Z. C., S. H., R. S., X. J., L. H., and H. C. investigation.

Funding and additional information

This work was supported by the 10.13039/501100001809 Natural Science Foundation of China (NSFC: 82072436 to L. X., 82170859 , 82370864 to H. C.) and the 10.13039/501100003453 Natural Science Foundation of Guangdong Province General Project (2021A1515010785 ).
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References

1 Westerterp K.R. Feingold K.R. Anawalt B. Blackman M.R. Boyce A. Chrousos G. Corpas E. Control of Energy Expenditure in Humans 2000 Endotext South Dartmouth (MA)
2 Heymsfield S.B. Smith B. Dahle J. Kennedy S. Fearnbach N. Thomas D.M. Resting energy expenditure: from cellular to whole-body level, a mechanistic historical perspective Obesity (Silver Spring) 29 2021 500 511 33624441
3 Brychta R.J. Chen K.Y. Cold-induced thermogenesis in humans Eur. J. Clin. Nutr. 71 2017 345 352 27876809
4 Cheng L. Wang J. Dai H. Duan Y. An Y. Shi L. Brown and beige adipose tissue: a novel therapeutic strategy for obesity and type 2 diabetes mellitus Adipocyte 10 2021 48 65 33403891
5 Li Y. Wang D. Ping X. Zhang Y. Zhang T. Wang L. Local hyperthermia therapy induces browning of white fat and treats obesity Cell 185 2022 949 966.e919 35247329
6 Yin X. Chen Y. Ruze R. Xu R. Song J. Wang C. The evolving view of thermogenic fat and its implications in cancer and metabolic diseases Signal. Transduct. Target Ther. 7 2022 324 36114195
7 Vercellino I. Sazanov L.A. The assembly, regulation and function of the mitochondrial respiratory chain Nat. Rev. Mol. Cell Biol. 23 2022 141 161 34621061
8 Kawabata H. Transferrin and transferrin receptors update Free Radic. Biol. Med. 133 2019 46 54 29969719
9 Venkataramani V. Iron homeostasis and metabolism: two sides of a coin Adv. Exp. Med. Biol. 1301 2021 25 40 34370286
10 Harrison A.V. Lorenzo F.R. McClain D.A. Iron and the pathophysiology of diabetes Annu. Rev. Physiol. 85 2023 339 362 36137277
11 Gonzalez-Dominguez A. Visiedo-Garcia F.M. Dominguez-Riscart J. Gonzalez-Dominguez R. Mateos R.M. Lechuga-Sancho A.M. Iron metabolism in obesity and metabolic syndrome Int. J. Mol. Sci. 21 2020 5529 32752277
12 Manios Y. Moschonis G. Chrousos G.P. Lionis C. Mougios V. Kantilafti M. The double burden of obesity and iron deficiency on children and adolescents in Greece: the Healthy Growth Study J. Hum. Nutr. Diet 26 2013 470 478 23279448
13 del Giudice E.M. Santoro N. Amato A. Brienza C. Calabro P. Wiegerinck E.T. Hepcidin in obese children as a potential mediator of the association between obesity and iron deficiency J. Clin. Endocrinol. Metab. 94 2009 5102 5107 19850683
14 Li J. Pan X. Pan G. Song Z. He Y. Zhang S. Transferrin receptor 1 regulates thermogenic capacity and cell fate in Brown/beige adipocytes Adv. Sci. (Weinh) 7 2020 1903366
15 Qiu J. Zhang Z. Wang S. Chen Y. Liu C. Xu S. Transferrin receptor functionally marks thermogenic adipocytes Front. Cell Dev. Biol. 8 2020 572459
16 James J.V. Varghese J. John N.M. Deschemin J.C. Vaulont S. McKie A.T. Insulin resistance and adipose tissue inflammation induced by a high-fat diet are attenuated in the absence of hepcidin J. Nutr. Biochem. 111 2023 109175
17 Barton J.C. Edwards C.Q. Acton R.T. HFE gene: structure, function, mutations, and associated iron abnormalities Gene 574 2015 179 192 26456104
18 Martins R. Silva B. Proenca D. Faustino P. Differential HFE gene expression is regulated by alternative splicing in human tissues PLoS One 6 2011 e17542
19 Huang J. Jones D. Luo B. Sanderson M. Soto J. Abel E.D. Iron overload and diabetes risk: a shift from glucose to Fatty Acid oxidation and increased hepatic glucose production in a mouse model of hereditary hemochromatosis Diabetes 60 2011 80 87 20876715
20 Yue F. Cheng Y. Breschi A. Vierstra J. Wu W. Ryba T. A comparative encyclopedia of DNA elements in the mouse genome Nature 515 2014 355 364 25409824
21 Fagerberg L. Hallstrom B.M. Oksvold P. Kampf C. Djureinovic D. Odeberg J. Analysis of the human tissue-specific expression by genome-wide integration of transcriptomics and antibody-based proteomics Mol. Cell Proteomics 13 2014 397 406 24309898
22 Cho C.H. Patel S. Rajbhandari P. Adipose tissue lipid metabolism: lipolysis Curr. Opin. Genet. Dev. 83 2023 102114
23 Choi E.J. Jeon C.H. Lee I.K. Ferric ammonium citrate upregulates PD-L1 expression through generation of reactive oxygen species J. Immunol. Res. 2022 2022 6284124
24 Joffin N. Gliniak C.M. Funcke J.B. Paschoal V.A. Crewe C. Chen S. Adipose tissue macrophages exert systemic metabolic control by manipulating local iron concentrations Nat. Metab. 4 2022 1474 1494 36329217
25 Higashida K. Takeuchi N. Inoue S. Hashimoto T. Nakai N. Iron deficiency attenuates catecholamine-stimulated lipolysis via downregulation of lipolysis-related proteins and glucose utilization in 3T3-L1 adipocytes Mol. Med. Rep. 21 2020 1383 1389 32016466
26 Yook J.S. You M. Kim J. Toney A.M. Fan R. Puniya B.L. Essential role of systemic iron mobilization and redistribution for adaptive thermogenesis through HIF2-alpha/hepcidin axis Proc. Natl. Acad. Sci. U. S. A. 118 2021 e2109186118
27 Fischer C. Volani C. Komlodi T. Seifert M. Demetz E. Valente de Souza L. Dietary iron overload and Hfe(-/-) related hemochromatosis alter hepatic mitochondrial function Antioxidants (Basel) 10 2021 1818 34829689
28 Deschemin J.C. Ransy C. Bouillaud F. Chung S. Galy B. Peyssonnaux C. Hepcidin deficiency in mice impairs white adipose tissue browning possibly due to a defect in de novo adipogenesis Sci. Rep. 13 2023 12794
29 Blankenhaus B. Braza F. Martins R. Bastos-Amador P. Gonzalez-Garcia I. Carlos A.R. Ferritin regulates organismal energy balance and thermogenesis Mol. Metab. 24 2019 64 79 30954544
30 Silva G.D.N. Amato A.A. Thermogenic adipose tissue aging: mechanisms and implications Front. Cell Dev. Biol. 10 2022 955612
31 Fu W. Liu Y. Sun C. Yin H. Transient p53 inhibition sensitizes aged white adipose tissue for beige adipocyte recruitment by blocking mitophagy FASEB J. 33 2019 844 856 30052487
32 Wu Q. Wang H. An P. Tao Y. Deng J. Zhang Z. HJV and HFE play distinct roles in regulating hepcidin Antioxid. Redox Signal. 22 2015 1325 1336 25608116
33 Lettieri Barbato D. Aquilano K. Baldelli S. Cannata S.M. Bernardini S. Rotilio G. Proline oxidase-adipose triglyceride lipase pathway restrains adipose cell death and tissue inflammation Cell Death Differ. 21 2014 113 123 24096872
34 Khan S.A. Sathyanarayan A. Mashek M.T. Ong K.T. Wollaston-Hayden E.E. Mashek D.G. ATGL-catalyzed lipolysis regulates SIRT1 to control PGC-1alpha/PPAR-alpha signaling Diabetes 64 2015 418 426 25614670
35 Rumberger J.M. Peters T. Jr. Burrington C. Green A. Transferrin and iron contribute to the lipolytic effect of serum in isolated adipocytes Diabetes 53 2004 2535 2541 15448081
36 Yang Loureiro Z. Solivan-Rivera J. Corvera S. Adipocyte heterogeneity underlying adipose tissue functions Endocrinology 163 2022 bqab138 34223880
37 Mahmoud M. Abdel-Rasheed M. Influence of type 2 diabetes and obesity on adipose mesenchymal stem/stromal cell immunoregulation Cell Tissue Res. 394 2023 33 53 37462786
38 Wang T. Sharma A.K. Wolfrum C. Novel insights into adipose tissue heterogeneity Rev. Endocr. Metab. Disord. 23 2022 5 12 34935088
39 Shinde A.B. Nunn E.R. Wilson G.A. Chvasta M.T. Pinette J.A. Myers J.W. Inhibition of nucleotide biosynthesis disrupts lipid accumulation and adipogenesis J. Biol. Chem. 299 2023 104635
40 Yang X. Liu Q. Li Y. Tang Q. Wu T. Chen L. The diabetes medication canagliflozin promotes mitochondrial remodelling of adipocyte via the AMPK-Sirt1-Pgc-1alpha signalling pathway Adipocyte 9 2020 484 494 32835596
41 Tan T.C. Crawford D.H. Jaskowski L.A. Murphy T.M. Heritage M.L. Subramaniam V.N. Altered lipid metabolism in Hfe-knockout mice promotes severe NAFLD and early fibrosis Am. J. Physiol. Gastrointest. Liver Physiol. 301 2011 G865 G876 21817060
42 Sun Z. Pan X. Tian A. Surakka I. Wang T. Jiao X. Genetic variants in HFE are associated with non-alcoholic fatty liver disease in lean individuals JHEP Rep. 5 2023 100744
43 Barton J.C. Barton J.C. Adams P.C. Acton R.T. Undiagnosed diabetes and impaired fasting glucose in HFE C282Y homozygotes and HFE wild-type controls in the HEIRS Study BMJ Open Diabetes Res. Care 4 2016 e000278
44 Gabrielsen J.S. Gao Y. Simcox J.A. Huang J. Thorup D. Jones D. Adipocyte iron regulates adiponectin and insulin sensitivity J. Clin. Invest. 122 2012 3529 3540 22996660
45 Kim M. Kim Y.H. Min S. Lee S.M. High iron exposure from the fetal stage to adulthood in mice alters lipid metabolism Nutrients 14 2022 2451 35745181
46 Dongiovanni P. Ruscica M. Rametta R. Recalcati S. Steffani L. Gatti S. Dietary iron overload induces visceral adipose tissue insulin resistance Am. J. Pathol. 182 2013 2254 2263 23578384
47 Suga H. Matsumoto D. Eto H. Inoue K. Aoi N. Kato H. Functional implications of CD34 expression in human adipose-derived stem/progenitor cells Stem Cells Dev. 18 2009 1201 1210 19226222
48 Raajendiran A. Ooi G. Bayliss J. O'Brien P.E. Schittenhelm R.B. Clark A.K. Identification of metabolically distinct adipocyte progenitor cells in human adipose tissues Cell Rep. 27 2019 1528 1540.e1527 31042478
49 Shi L. Tao Z. Zheng L. Yang J. Hu X. Scott K. FoxO1 regulates adipose transdifferentiation and iron influx by mediating Tgfbeta1 signaling pathway Redox Biol. 63 2023 102727
50 Suzuki T. Komatsu T. Shibata H. Tanioka A. Vargas D. Kawabata-Iwakawa R. Crucial role of iron in epigenetic rewriting during adipocyte differentiation mediated by JMJD1A and TET2 activity Nucleic Acids Res. 51 2023 6120 6142 37158274
51 Liu J. Wu X. Wang H. Wei J. Wu Q. Wang X. HFE inhibits type I IFNs signaling by targeting the SQSTM1-mediated MAVS autophagic degradation Autophagy 17 2021 1962 1977 32746697
52 Ding H. Chen S. Pan X. Dai X. Pan G. Li Z. Transferrin receptor 1 ablation in satellite cells impedes skeletal muscle regeneration through activation of ferroptosis J. Cachexia Sarcopenia Muscle 12 2021 746 768 33955709
53 Aune U.L. Ruiz L. Kajimura S. Isolation and differentiation of stromal vascular cells to beige/brite cells J. Vis. Exp. 2013 50191 10.3791/50191 23568137
54 Sanchez-Ceinos J. Guzman-Ruiz R. Rangel-Zuniga O.A. Lopez-Alcala J. Moreno-Cano E. Del Rio-Moreno M. Impaired mRNA splicing and proteostasis in preadipocytes in obesity-related metabolic disease Elife 10 2021 e65996 34545810
55 Moos T. Mollgard K. A sensitive post-DAB enhancement technique for demonstration of iron in the central nervous system Histochemistry 99 1993 471 475 7691783
