Abstract

   Stimulating adipose tissue thermogenesis has emerged as a promising
   strategy for combating obesity, with uncoupling protein 1 (UCP1)
   playing a central role in this process. However, the mechanisms that
   suppress adipose thermogenesis and energy dissipation in obesity are
   not fully understood. This study identifies mitochondrial carrier
   homolog 2 (MTCH2), an obesity susceptibility gene, as a negative
   regulator of energy homeostasis across flies, rodents, and humans.
   Notably, adipose‐specific MTCH2 depletion in mice protects against
   high‐fat‐diet (HFD)‐induced obesity and metabolic disorders.
   Mechanistically, MTCH2 deficiency promotes energy expenditure by
   stimulating thermogenesis in brown adipose tissue (BAT) and browning of
   subcutaneous white adipose tissue (scWAT), accompanied by upregulated
   UCP1 protein expression, enhanced mitochondrial biogenesis, and
   increased lipolysis in BAT and scWAT. Using integrated RNA sequencing
   and proteomic analyses, this study demonstrates that MTCH2 is a key
   suppressor of thermogenesis by negatively regulating autophagy via
   Bcl‐2‐dependent mechanism. These findings highlight MTCH2's critical
   role in energy homeostasis and reveal a previously unrecognized link
   between MTCH2, thermogenesis, and autophagy in adipose tissue biology,
   positioning MTCH2 as a promising therapeutic target for obesity and
   related metabolic disorders. This study provides new opportunities to
   develop treatments that enhance energy expenditure.

   Keywords: adipose tissue, autophagy, mitochondrial carrier homolog 2
   (MTCH2), obesity, thermogenesis
     __________________________________________________________________

   This study demonstrates that MTCH2 acts as an evolutionarily conserved
   regulator of energy homeostasis across Drosophila, rodents, and humans.
   Adipose‐specific MTCH2 ablation enhances brown/beige adipose
   thermogenesis through Bcl‐2‐dependent autophagy, promoting energy
   expenditure and counteracting obesity‐associated metabolic disorders.
   These findings establish MTCH2 as a pivotal metabolic gatekeeper and
   propose its therapeutic targeting for obesity treatment via adipose
   tissue remodeling.

   graphic file with name ADVS-12-2416598-g003.jpg

1. Introduction

   Obesity, characterized by chronic caloric imbalance where intake
   surpasses expenditure,^[ [58]^1 ^] affects approximately one billion
   individuals worldwide,^[ [59]^2 ^] presenting significant challenges
   for effective weight management. Adipose tissue, traditionally regarded
   as a passive energy reservoir, is now recognized as a dynamic organ
   that plays a crucial role in regulating energy homeostasis.^[ [60]^3 ,
   [61]^4 ^] A key process in this regulation isUCP1‐mediated
   thermogenesis, which dissipates the mitochondrial proton gradient to
   generate heat and increase energy expenditure.^[ [62]^3 , [63]^5 ,
   [64]^6 ^] Several key regulators of UCP1 expression and activity have
   been identified,^[ [65]^3 , [66]^5 , [67]^7 , [68]^8 ^] including PR
   domain zinc finger protein 16 (PRDM16), peroxisome
   proliferator‐activated receptor γ (PPARγ), PPAR co‐activator 1α
   (PGC‐1α), fibroblast growth factor 21 (FGF21), and β‐adrenergic
   signaling pathways. However, the complete molecular mechanisms
   governing thermogenesis in adipose tissue remain incompletely
   understood.

   While genome‐wide association studies (GWAS) have identified numerous
   genetic loci associated with obesity,^[ [69]^9 ,[70] ^10 ^] relatively
   few studies have focused specifically on the regulation of energy
   balance within adipose tissue.^[ [71]^11 , [72]^12 , [73]^13 , [74]^14
   ^] This gap underscores the need for targeted investigations to uncover
   novel adipose‐specific regulators of energy homeostasis. In the
   DiOGenes study, we previously identified 29 genes in abdominal
   subcutaneous adipose tissue associated with weight loss during calorie
   restriction.^[ [75]^15 ^] However, the functional roles of these genes
   in energy regulation remain largely unexplored.

   In this study, we employed Drosophila as a model organism to
   investigate the metabolic roles of these candidate genes, identifying
   several candidate regulators of energy balance. Among them, MTCH2
   emerged as a key regulator due to its established links to obesity and
   energy homeostasis. Through in vivo and in vitro models, coupled with
   comprehensive metabolic phenotyping and integrative proteomic and
   transcriptomic analyses, we demonstrate that MTCH2 acts as a suppressor
   of thermogenesis by negatively regulating autophagy via the Bcl‐2
   pathway. Our findings establish MTCH2 as a critical regulator of
   thermogenesis, offering new insights into its role in energy balance
   and its therapeutic potential in combating obesity and related
   metabolic disorders.

2. Results

2.1. Identification of MTCH2 as a Conserved Negative Regulator of Energy
Homeostasis

   We performed an RNAi screen targeting the 34 Drosophila homologs of 29
   human genes that were revealed in our previous study to have
   differential expression in adipose tissue during calorie restriction.^[
   [76]^15 ^] This screen assessed their impact on fat metabolism in adult
   flies using the Actin‐Gal4 driver (Figure [77]1A). We identified six
   genes that play a critical role in fat metabolism, with their silenced
   homologs (6/34, 17.6%) leading to a reduction in TAG levels
   (Figure [78]1B). Among these, MTCH2 is of particular interest due to
   its established role in the regulation of energy homeostasis.^[ [79]^16
   ^] To validate the RNAi findings, we analyzed Drosophila Mtch
   hypomorphic mutants with a 70% reduction in MTCH2 protein levels
   (Figure [80]S1A, Supporting Information). TAG levels were markedly
   reduced in these mutants (Figure [81]1C). Importantly, TAG levels were
   significantly lower in Mtch mutant flies fed a high‐sugar or high‐fat
   diet (Figure [82]1C), demonstrating resistance to obesogenic diets.

Figure 1.

   Figure 1
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   Identification of MTCH2 as a conserved regulator of energy homeostasis.
   (A) A scheme of functional RNAi knockdown screen of 29 genes associated
   with weight loss using Drosophila melanogaster. (B) Z‐scores of TAG
   levels for each UAS‐RNAi line were presented. Z≥ 1.96 or ≤−1.96 (red
   dotted lines indicate threshold) was regarded as significant hits. (C)
   TAG levels were decreased in Mtch mutant flies fed normal, high sucrose
   or high‐fat diet (n = 5 biological sample, 6 flies per sample). (D)
   Mtch2 mRNA expression was elevated in mice fed a high‐fat diet (DIO
   mice) compared to those mice fed a chow (lean mice) (n = 6). (E,F)
   MTCH2 mRNA expression was lower in the abdominal scWAT of individuals
   with obesity than those without obesity (n = 770 biological samples for
   E, n = 56 for (F). (G–J) A positive correlation existed between MTCH2
   mRNA expression and body fat (G), BMI (H), waist‐to‐hip ratio (WHR)
   (I), and HOMA‐IR (J) in abdominal scWAT of individuals (n = 49
   biological samples for G, n = 770 biological samples for (H–J). (K)
   MTCH2 mRNA expression was significantly reduced in the abdominal scWAT
   of patients with obese after dietary intervention (n = 46 biological
   samples). Data are represented as mean ± SEM. Two‐tailed unpaired
   Student's t‐test (C–F), The Spearman's rank‐order correlation
   coefficient (r) (G–J), and two‐tailed paired t‐test (K) were used. *p
   < 0.05, **p < 0.01, ***p < 0.005, ****p < 0.001.

   To further investigate the role of MTCH2 in adiposity and energy
   homeostasis, we examined Mtch2 expression in adipose tissue and other
   metabolically important tissues in chow and high‐fat diet (HFD, 60%
   fat) ‐fed mice. In mice fed a chow diet, Mtch2 expression was detected
   in all tissues examined including BAT, scWAT, epididymal WAT (eWAT),
   retroperitoneal WAT (rWAT), muscle, and liver, with the highest levels
   found in BAT (Figure [84]S1B, Supporting Information). In HFD‐fed mice,
   Mtch2 mRNA expression was significantly upregulated in both scWAT and
   eWAT compared with chow‐fed mice (Figure [85]1D), suggesting a positive
   relationship between Mtch2 expression and diet‐induced obesity.

   To evaluate whether MTCH2 expression is associated with adiposity and
   metabolic health in humans, we analyzed publicly available datasets
   ([86]GSE77962, [87]GSE70353, [88]GSE77962) from the Gene Expression
   Omnibus (GEO). Consistent with our results in DIO mice, MTCH2
   expression was significantly elevated in scWAT from obese compared with
   non‐obese individuals (Figure [89]1E,F). Moreover, MTCH2 expression in
   human scWAT was positively correlated with body fat (Figure [90]1G),
   body mass index (BMI) (Figure [91]1H), waist‐to‐hip ratio (WHR)
   (Figure [92]1I), and insulin resistance (Figure [93]1J). Notably, MTCH2
   expression was downregulated following diet‐induced weight loss in
   patients with obesity (Figure [94]1K), implying that Mtch2 levels are
   dependent on the prevailing adiposity and metabolic state.
   Collectively, these findings reveal MTCH2 as a conserved obesity‐linked
   factor in adipose tissue across species, and that elevated MTCH2
   expression in adipose tissue may play a role in the negative regulation
   of energy homeostasis and metabolic dysfunction such as insulin
   resistance.

2.2. Adipose Tissue‐Specific Mtch2 Knockout Prevents HFD‐Induced Obesity and
Metabolic Disorder

   To examine the role of MTCH2 specifically in adipose tissue, we
   generated adipocyte‐specific Mtch2 knockout mice
   (Mtch2^flox/flox,Adipoq‐Cre referred to as Mtch2^AKO hereafter) by
   crossing Mtch2^flox/flox (Mtch2^f/f ) mice with adiponectin‐Cre mice
   (Figure [95]S2A, Supporting Information). The successful deletion of
   mtch2 in adipose tissue of Mtch2^AKO mice was confirmed by the presence
   of a single DNA band at 221 and 204 bp (Figure [96]S2B, Supporting
   Information). Furthermore, Mtch2^AKO mice exhibited a significant
   reduction in Mtch2 mRNA and protein levels in various adipose tissue
   depots, while its levels in non‐adipose tissues, the liver and skeletal
   muscle, remained unchanged (Figure [97]S2C,D, Supporting Information),
   thus confirming the specificity of Mtch2 deletion in adipose tissue.

   To assess Mtch2’s role in energy balance and glucose homeostasis,
   Mtch2^AKO mice and littermate Mtch2^f/f control mice were fed a chow or
   HFD for up to 26 weeks. Body weight (BW) was not different between
   Mtch2^AKO and control mice fed a chow diet (Figure [98]2A; Figure
   [99]S3A,B, Supporting Information), suggesting that adipocyte‐specific
   deletion of Mtch2 has minimal effects on energy balance under normal
   dietary conditions. However, when subjected to high‐fat feeding,
   Mtch2^AKO mice displayed resistance to BW gain, showing a 14.2%
   reduction compared to controls (Figure [100]2B,C; Figure [101]S3C,D,
   Supporting Information). Analysis by magnetic resonance imaging (MRI)
   revealed that the lower BW gain was accompanied by a 29.2% reduction in
   fat mass and fat mass percentage in Mtch2^AKO compared to control mice
   (Figure [102]2D,E; Figure [103]S3E, Supporting Information). Similarly,
   dissected weights of total fat depots, scWAT, and rWAT, were
   significantly reduced in Mtch2^AKO compared to control mice
   (Figure [104]2F). Strikingly, H&E staining revealed a higher proportion
   of relatively small adipocytes in scWAT of Mtch2^AKO compared to
   control mice (Figure [105]2G). Similarly, after HFD feeding, BAT weight
   was reduced by 42.5% in Mtch2^AKO compared to control mice
   (Figure [106]2H). Furthermore, H&E staining and transmission electron
   microscopy (TEM) revealed the presence of smaller adipocytes in BAT
   from Mtch2^AKO compared to control mice (Figure [107]2I). These
   findings indicate that adipocyte expression of Mtch2 is critical for
   the long‐term remodeling of adipose tissues in HFD mice.

Figure 2.

   Figure 2
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   Mtch2^AKO reduces fat depots and hepatic accumulation under HFD. (A)
   Mtch2^AKO did not affect body weight in mice fed chow (n = 8–9). (B,C)
   Body weight was gained less in Mtch2^AKO mice under HFD feeding for 19
   weeks. Representative image (B) and body weight growth curve (n = 10,
   C). (D,E) Mtch2^AKO mice displayed a lower fat composition.
   Representative NMR analysis image (D) and fat mass composition by NMR
   (E). (F,G) Mtch2^AKO mice exhibited fewer fat depots. The dissected
   weight of total WAT, rWAT, scWAT and eWAT (F, n = 10), H&E staining and
   adipocyte size analysis of scWAT (G). (H,I) BAT was reduced in
   Mtch2^AKO mice. Representative image of dissected BAT (H, n = 10), the
   analyses of H&E staining and TEM on BAT and adipocyte size of BAT in
   H&E staining (I). (J–L) Fat accumulation was decreased in Mtch2^AKO
   mice. Representative image of the dissected liver (J, n = 10), hepatic
   TAG levels (K, n = 5), and liver H&E staining (L). (M) Mtch2^AKO mice
   exhibited enhanced insulin sensitivity in mice fed HFD for 18 weeks.
   Insulin tolerance test and quantified in the area under curves
   (n = 8–10). Data are represented as mean ± SEM. Two‐way ANOVA followed
   by Bonferroni's multiple comparisons test (C,M), Two‐tailed unpaired
   t‐test was used (E–K,M), *p < 0.05, **p < 0.01, and ***p < 0.001, ns,
   not significant.

   We next examined the livers of Mtch2^AKO and Mtch2^f/f mice following
   HFD feeding. The liver weight was reduced by 21.3% and appeared less
   pale in Mtch2^AKO mice compared to control mice (Figure [109]2J). This
   was accompanied by a 35.8% decrease in hepatic TAG content
   (Figure [110]2K) and the presence of fewer lipid droplets in the liver
   (Figure [111]2L). Moreover, while aspartate aminotransferase (AST)
   remained unchanged, plasma alanine transaminase (ALT) and the ratio of
   ALT to AST were significantly reduced in Mtch2^AKO mice (Figure
   [112]S3F–H, Supporting Information). This systemic reduction in
   adiposity was associated with improved insulin sensitivity in Mtch2^AKO
   mice compared to controls (Figure [113]2M). The area under the curve
   (AUC) for blood glucose levels from 0 to 120 min during the insulin
   tolerance test was significantly reduced in Mtch2^AKO mice compared to
   control mice (Figure [114]2M). Together, these findings suggest that
   fat tissue‐specific Mtch2 deletion not only protects against
   HFD‐induced fat accumulation in adipose tissue but also reduces liver
   lipid deposition and enhances whole‐body insulin sensitivity in mice.

2.3. Adipose Tissue‐Specific Mtch2 Knockout Increases Energy Expenditure by
Promoting BAT Thermogenesis and scWAT Browning

   To investigate whether the leaner phenotype observed in Mtch2^AKO mice
   under HFD feeding results from changes in energy balance, we measured
   food intake and energy expenditure using metabolic chambers. Mtch2^AKO
   mice exhibited no significant changes in average daily food intake
   (Figure [115]3A) or cumulative energy intake over a 2‐day period
   (Figure [116]3B) compared to their littermate controls, indicating that
   the reduction in BW and fat mass observed in Mtch2^AKO mice was not due
   to altered calorie intake. In contrast, Mtch2^AKO mice exhibited
   increased oxygen (O[2]) consumption (Figure [117]S4A, Supporting
   Information) and carbon dioxide (CO[2]) production (Figure [118]S4B,
   Supporting Information), resulting in a significant increase in overall
   energy expenditure (Figure [119]3C,D). These findings suggest that
   augmented energy expenditure is likely responsible for the reduced BW
   gain and fat accretion in Mtch2^AKO mice. Additionally, there were no
   significant changes in physical activity between the groups (Figure
   [120]S4C, Supporting Information), excluding altered physical movement
   as a potential cause for the increased energy expenditure.

Figure 3.

   Figure 3
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   Mtch2^AKO increases energy expenditure by promoting BAT thermogenesis
   and scWAT browning in HFD‐fed mice. (A,B) Caloric intake was not
   altered in Mtch2^AKO mice. Daily food intake (A, n = 10) and
   accumulative food intake (B, n = 6). (C,D) Energy expenditure was
   enhanced in Mtch2^AKO mice (n = 6). (E,F) Mtch2^AKO mice displayed
   higher surface temperature and core temperature under cold exposure
   (4 °C) at the age of 8 weeks. Representative infra‐red thermal image of
   lumbar back and BAT and qualification of temperatures of them (n = 7
   for E and 4 for F). (G,H) UCP1 protein levels were elevated in BAT and
   scWAT of Mtch2^AKO mice as shown in western blot (G, n = 4) and IHC
   staining of UCP1 (H). Data are represented as mean ± SEM. Two‐tailed
   unpaired t‐test (A,C,E–G) or ANCOVA using body weight as covariate (D)
   was used. *p < 0.05, **p < 0.01, and ***p < 0.001, ns, not significant.

   A key component of energy expenditure involves body temperature
   control, with BAT activation important for maintaining core temperature
   during cold exposure. To explore this, we employed a non‐invasive
   infra‐red camera to measure skin surface temperature^[ [122]^17 ^]
   under cold conditions. We calculated temperatures above the BAT as well
   as over the lumbar spine region (Figure [123]3E). Mtch2^AKO mice
   displayed significantly higher temperatures in both the lumbar spine
   region and BAT under cold exposure compared to control mice
   (Figure [124]3E). Increased body temperatures were also observed in
   Mtch2^AKO mice using the conventional method of rectal probes to
   measure core body temperature over an 8‐h monitoring period
   (Figure [125]3F).

   Given the critical role of UCP1 in BAT thermogenesis and scWAT
   browning,^[ [126]^5 , [127]^18 ^] we examined the UCP1 protein levels
   in BAT and scWAT of HFD‐fed mice. UCP1 levels were significantly
   upregulated in both BAT and scWAT of Mtch2^AKO mice compared to control
   mice (Figure [128]3G). Immunohistochemical staining further revealed
   abundant UCP1 expression in smaller, multilocular adipocytes within the
   BAT of Mtch2^AKO mice (Figure [129]3H). Additionally, clusters of
   UCP1‐positive adipocytes were detected in the scWAT of Mtch2^AKO mice
   (Figure [130]3H), indicating that Mtch2 deletion promotes the
   development of beige fat. Collectively, these results suggest that
   adipose tissue‐specific deletion of Mtch2 enhances energy expenditure
   under HFD condition through the promotion of BAT thermogenesis and the
   conversion of scWAT into energy‐burning beige WAT via upregulation of
   UCP1.

2.4. Adipose Tissue‐Specific Mtch2 Knockout Increases Mitochondrial
Biogenesis and Oxidative Phosphorylation in Adipose Tissues

   We next assessed potential mechanisms for the changes in fat
   thermogenesis in Mtch2^AKO mice by examining mitochondrial features.
   Mtch2 knockout led to a significant increase in mitochondrial DNA
   (mtDNA) copy number in BAT (Figure [131]4A) and scWAT (Figure [132]4B),
   indicating an increase in mitochondrial abundance. Furthermore, TEM
   analysis revealed that BAT adipocytes from Mtch2^AKO mice displayed an
   increased number of mitochondria with more pronounced cristae
   structures compared to control mice (Figure [133]4C). These findings
   uncover a novel regulatory function for Mtch2 in mitochondrial
   biogenesis in BAT and scWAT.

Figure 4.

   Figure 4
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   Mtch2^AKO increases mitochondrial biogenesis and oxidative
   phosphorylation in adipose tissues of HFD‐fed mice. (A,B) The copies of
   mtDNA were increased in BAT (A) and scWAT (B) of Mtch2^AKO mice by qPCR
   qualification (n = 5). (C) The number and structure of mitochondria
   were increased in BAT of Mtch2^AKO by TEM analysis. (D,E) Mitochondrial
   biogenesis was boosted in BAT (D, n = 5) and scWAT (E, n = 4–5) of
   Mtch2^AKO mice by qPCR analysis of typic markers. (F,G) Oxidative
   phosphorylation was enhanced in BAT (F, n = 5) and scWAT (G, n = 4–5)
   of Mtch2^AKO mice by qPCR analysis of typic markers. Data are
   represented as mean ± SEM. Two‐tailed unpaired t‐test was used
   (A,B,D–G). *p < 0.05, **p < 0.01, and ***p < 0.001, ns, not
   significant.

   We next examined changes in the expression of genes involved in
   mitochondrial biogenesis. The expression of Pgc1α, Tfam, and Nrf1 were
   significantly increased in both BAT (Figure [135]4D) and scWAT
   (Figure [136]4E) of Mtch2^AKO mice compared to control mice. We next
   assessed changes in the expression of genes critical for mitochondrial
   oxidative phosphorylation. The expression of Ndufs8, Sdhb, Uqcrc1, and
   Atp5a1 were markedly upregulated in BAT (Figure [137]4F) and scWAT
   (Figure [138]4G) of Mtch2^AKO mice compared to control mice. These
   findings suggest Mtch2 deficiency promotes mitochondrial biogenesis and
   augments mitochondrial function in adipose tissues. Taken together, the
   results suggest that the enhanced thermogenesis and energy expenditure
   observed following Mtch2 deletion is driven by increases in
   mitochondrial abundance, function and uncoupling of the electron
   transport chain.

2.5. Proteomics and Transcriptomics Analyses Reveal that Mtch2 Deficiency
Enhances Thermogenesis and Autophagy

   To gain a comprehensive and unbiased understanding of the underlying
   molecular mechanisms for the thermogenic role of Mtch2 in adipose
   tissues, we employed a multi‐omics approach across flies and rodents,
   namely label‐free quantitative mass spectrometry in Mtch‐mutant flies,
   and RNA sequencing (RNA‐seq) on BAT from HFD‐fed Mtch2^AKO mice. Mass
   spectrometry identified 1036 differentially expressed proteins between
   hypomorphic Mtch mutant and wildtype flies (Figure [139]5A). Ingenuity
   Pathway Analysis (IPA) of differentially expressed proteins showed
   enrichment of autophagy, oxidative phosphorylation, and mitochondrial
   dysfunction pathways (Figure [140]5B). RNA‐seq analysis of BAT from
   Mtch2^AKO and control mice identified 2491 differentially expressed
   genes (Figure [141]5C; Figure [142]S5A, Supporting Information). KEGG
   pathway enrichment analysis identified upregulated genes involved in
   metabolic pathways, thermogenesis, lysosome, and autophagy in Mtch2^AKO
   mice (Figure [143]S5B, Supporting Information). A comparison of fly
   proteomics and BAT transcriptomic analyses in Venn diagrams indicated
   an overlap of 290 upregulated genes (Figure [144]5D). These genes were
   enriched in pathways related to energy metabolism, including energy
   expenditure, autophagy, and lysosome function (Figure [145]5E). This
   suggests that Mtch2 deficiency leads to a coordinated enhancement of
   cellular energy expenditure and metabolic efficiency. Specifically, the
   upregulation of autophagy‐related genes, such as Atg7 in both fly and
   mouse datasets (Figure [146]5E) and Atg5 and numerous other
   autophagy‐related genes in BAT transcriptome (Figure [147]S5C,
   Supporting Information) reveals a potential link between thermogenesis,
   autophagy, and lysosome function. Among the downregulated genes with
   Mtch2 deficiency in the fly proteome and mouse BAT transcriptome, 52
   were common in both datasets, with 12 identified in the MitoCarta
   Inventory^[ [148]^19 ^] (Figure [149]5F). Notably, leucine
   aminopeptidase III (Lap3) and peptidylprolyl isomerase F (Ppif, encodes
   Cyclophilin D, CypD), previously characterized as mitochondrial matrix
   proteins whose localization is facilitated by MTCH2, functioning as a
   insertase^[ [150]^20 ^]‐have been implicated in fat metabolism and
   obesity,^[ [151]^21 , [152]^22 , [153]^23 ^] further supporting the
   MTCH2's critical role in regulating energy homeostasis.

Figure 5.

   Figure 5
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   Mtch2 deficiency promotes autophagy. (A) Volcano plot of differentially
   expressed proteins in Mtch mutant flies (fold change > 1 and p < 0.05).
   (B) Ingenuity pathway analysis (IPA) of up‐regulated proteins in Mtch
   mutant flies. (C) Volcano plot of differentially expressed genes in BAT
   of HFD‐fed Mtch2^AKO mice (fold change > 1 and p < 0.05). (D) Venn
   diagram of the up‐regulated proteins in Mtch mutant flies and the
   up‐regulated genes in BAT of Mtch2^AKO mice. (E) Functional enrichment
   analysis of the overlapped up‐regulated genes in flies and mice. (F)
   Venn diagram of the down‐regulated genes in BAT of Mtch2^AKO mice and
   proteins in Mtch mutant flies (Left). The overlapped proteins
   localizing to the mitochondria were displayed (Right).

2.6. Adipose Tissue‐Specific Mtch2 Knockout Enhances Autophagy to Promote
Lipolysis in Adipose Tissues

   To validate the upregulation of autophagy pathways identified in both
   fly proteomic and mouse BAT RNA‐seq data, we conducted qPCR, western
   blot, and TEM analyses on BAT and scWAT. As expected, HFD‐fed Mtch2^AKO
   mice exhibited significantly increased mRNA expression of key autophagy
   genes, including Atg5, Atg7, Beclin1, and Lc3b, compared to controls
   (Figure [155]6A,B). Activation of autophagy facilitates the conversion
   of LC3I to LC3II and a decline in levels of the autophagy substrate
   p62. Here, we found elevated protein levels of LC3II/LC3I and reduced
   P62 in both BAT (Figure [156]6C) and scWAT (Figure [157]6D) of
   Mtch2^AKO mice compared to control. TEM further revealed that the lack
   of Mtch2 led to a notable increase in the number of autophagosomes and
   autophagolysosomes within BAT (Figure [158]6E), indicating enhanced
   autophagic activity. These results suggest that the autophagy pathway
   is enhanced in adipose tissue with Mtch2 knockout.

Figure 6.

   Figure 6
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   Mtch2^AKO promotes autophagy and lipolysis in the adipose tissues in
   HFD‐fed mice. (A–D) Autophagy was induced in BAT (A,C) and scWAT (B,D)
   of Mtch2^AKO mice by qPCR and western blot analysis of classic marker
   (n = 4–6). (E) The number of autophagosomes and autophagolysosomes was
   increased in BAT of Mtch2^AKO mice by TEM analysis. (F,G) Mtch2
   knockout reduced Bcl‐2 protein levels in BAT (F) and scWAT (G) in 19
   weeks of HFD‐fed mice (n = 6–8). (H–K) Lipolysis was increased in BAT
   (H,J) and scWAT (I,K) of Mtch2^AKO mice by qPCR and western blot
   analysis of lipolytic gene and protein expression (n = 4–5). Data are
   represented as mean ± SEM. Two‐tailed unpaired t‐test was used. *p
   < 0.05, **p < 0.01 and ***p < 0.001, ns, not significant.

   To explore the molecular mechanism underlying this enhanced autophagy,
   we focused on the potential regulators of autophagy. Bcl‐2 is a key
   suppressor of autophagy,^[ [160]^24 , [161]^25 ^] and Mtch2 is reported
   to interact with the Bcl‐2 family of proteins.^[ [162]^26 , [163]^27 ^]
   We investigated the possible involvement of Bcl‐2 in the suppression of
   autophagy by Mtch2 in adipose tissue. We observed a significant
   decrease in Bcl‐2 protein levels in both BAT (Figure [164]6F) and scWAT
   (Figure [165]6G) of Mtch2 ^AKO mice compared with the control. Since
   autophagy has been implicated in promoting lipolysis in adipocytes^[
   [166]^28 , [167]^29 , [168]^30 ^] and lipolysis promotes
   thermogenesis,^[ [169]^3 , [170]^5 , [171]^31 ^] we next examined
   changes in the expression of key lipolytic enzymes in adipose tissue
   following the loss of Mtch2. mRNA levels Atgl, Hsl, and Mgl, were
   significantly elevated in both BAT (Figure [172]6H) and scWAT of
   Mtch2^AKO mice relative to controls (Figure [173]6I). Furthermore, we
   found elevated protein levels of HSL and ATGL in BAT (Figure [174]6J)
   and ATGL in scWAT (Figure [175]6K) of Mtch2^AKO mice compared to
   control mice. These findings suggest that the deletion of Mtch2
   increases lipolysis in adipose tissue. Collectively, these findings
   support the notion that Mtch2 deletion disinhibits autophagy, leading
   to increased lipolysis in adipose tissue, which in turn mediates the
   enhanced thermogenesis and energy expenditure under HFD conditions.

2.7. Mtch2 Suppresses Thermogenesis Through the Bcl‐2‐Autophagy Pathway

   To gain direct functional insights into the influence of Mtch2 on
   thermogenesis and autophagy in adipocytes, we utilized CRISPR/Cas9
   technology to specifically delete Mtch2 in mouse 3T3‐L1 preadipocytes,
   a widely used model system for analysis of adipocyte biology.
   Consistent with in vivo results in adipose tissues, Mtch2 ablation led
   to changes in markers (increased LC3II/LC3I and reduced p62) indicative
   of increased autophagy in 3T3‐L1 preadipocytes (Figure [176]7A). Mtch2
   deletion in mature 3T3‐L1 adipocytes resulted in a reduced lipid
   accumulation as revealed by Oil Red O staining and significantly lower
   levels of the eluted Oil Red O dye (Figure [177]7B). To investigate
   Mtch2's role in UCP1‐dependent thermogenesis, we measured UCP1 protein
   levels in both preadipocytes and mature adipocytes following Mtch2
   ablation. While UCP1 levels remained unchanged in preadipocytes, they
   were significantly increased in mature adipocytes (Figure [178]7C).
   These results with in vitro 3T3‐L1 adipocytes closely resemble the in
   vivo findings, suggesting that the enhancement of thermogenesis in
   mature adipocytes following Mtch2 loss occurs in concert with increased
   UCP1.

Figure 7.

   Figure 7
   [179]Open in a new tab

   Mtch2^AKO promotes thermogenesis through the Bcl‐2‐autophagy pathways.
   (A) Autophagy was increased in 3T3‐L1 preadipocytes following Mtch2
   knockout by CRISPR (n = 4). (B) TAG levels were reduced in 3T3‐L1
   mature adipocytes following Mtch2 knockout by CRISPR‐Cas9 technique
   (n = 3). (C) UCP1 protein levels were elevated in 3T3‐L1 preadipocytes
   and 3T3‐L1 mature adipocytes following Mtch2 knockout by CRISPR
   (n = 4). (D) CQ treatment (50 µm) increased autophagy but reduced UCP1
   protein levels in 3T3‐L1 mature adipocytes following Mtch2 knockout by
   CRISPR (n = 4). (E) MG132 treatment (10 µm) increased Bcl‐2 protein
   levels but reduced UCP1 protein levels in 3T3‐L1 mature adipocytes
   following Mtch2 knockout by CRISPR (n = 3). (F) A summary of MTCH2
   suppresses thermogenesis via Bcl2‐mediated autophagy in adipose
   tissues. MTCH2 functions as a negative regulator of autophagy. MTCH2
   knockout induces Bcl‐2 protein degradation to enhance autophagy,
   leading to increased lipolysis, elevated UCP1 protein levels, and
   promotion of BAT thermogenesis and scWAT browning (Created with
   BioGDP.com). Data are represented as mean ± SEM. Two‐tailed unpaired
   t‐test was used. *p < 0.05, **p < 0.01 and ***p < 0.001, ns, not
   significant.

   We next examined whether changes in autophagic flux contribute to the
   heightened adipose thermogenesis following Mtch2 inhibition. We treated
   control and Mtch2‐deleted mature 3T3‐L1 adipocytes with chloroquine
   (CQ), which inhibits autophagy flux by impairing autophagosome‐lysosome
   fusion.^[ [180]^32 ^] CQ treatment blocked the induction of UCP1
   protein levels in Mtch2‐deleted adipocytes (Figure [181]7D), suggesting
   that Mtch2 deletion promotes adipocyte browning and thermogenesis
   through activation of autophagy.

   Reduced Bcl‐2 protein levels were associated with increased autophagy
   in MTCH2‐deficient adipose cells (Figure [182]6F,G). However, Bcl‐2
   mRNA expression remained unchanged in BAT and scWAT of Mtch2^AKO mice
   compared to control mice (Figure [183]S6A, Supporting Information),
   suggesting that Mtch2 regulates Bcl‐2 protein levels via
   post‐transcriptional mechanisms, potentially involving pathways of
   protein stabilization or degradation processes. Co‐immunoprecipitation
   (Co‐IP) assay further revealed no direct interaction between Bcl‐2 and
   MTCH2 (Figure [184]S6B, Supporting Information). MTCH2 is known to
   function as a membrane protein insertase that impacts cellular
   proteostasis,^[ [185]^33 , [186]^34 , [187]^35 ^] we examined whether
   the lack of MTCH2 enhances autophagy by promoting Bcl‐2 protein
   degradation in mature 3T1‐L1 adipocytes. To test this, we used the
   proteasome inhibitor MG132^[ [188]^36 ^] to block Bcl‐2 protein
   degradation. Remarkably, MG132 treatment restored Bcl‐2 protein levels
   and completely blocked the induction of autophagy in MTCH2‐deficient
   adipocytes, as evidenced by reduced LC3II and increased p62 levels
   (Figure [189]7E). In concert, MG132 treatment blocked the induction of
   UCP1 protein levels in MTCH2‐deficient adipocytes (Figure [190]7E).

   Collectively, these findings reveal a previously unrecognized
   regulatory mechanism whereby MTCH2 represses thermogenesis in obesity.
   MTCH2 knockout in adipocytes results in Bcl‐2 protein degradation and
   disinhibition of autophagy, which leads to upregulation of UCP1 and
   increased thermogenesis (Figure [191]7F).

3. Discussion

   Our research identifies a novel role for MTCH2 as a crucial regulator
   of adipose tissue biology through its control of mitochondrial
   adaptation and thermogenesis during obesity. Adipose‐specific knockout
   of MTCH2 protected against diet‐induced obesity and metabolic
   disorders, as evidenced by reduced fat deposition in adipose tissues
   and the liver, along with improved insulin sensitivity. These
   beneficial effects are mainly driven by enhanced energy expenditure,
   increased lipolysis, elevated BAT thermogenesis, and extensive being of
   scWAT linked to upregulation of UCP1. Mechanistically, adipose Mtch2
   deletion increases mitochondrial biogenesis, and oxidative
   phosphorylation, primarily through heightened autophagic activity via
   the Bcl‐2 pathway. Our findings uncover MTCH2 as a key suppressor of
   adipose thermogenesis and establish a previously unrecognized link
   between the Mtch2‐Bcl‐2‐autophagy pathways in the regulation of
   thermogenesis and energy homeostasis. Thus, our studies suggest that
   the upregulation of adipose MTCH2 contributes to increased adiposity,
   disruption of thermogenesis, and metabolic disorder in obesity.

   Mtch2 is enriched in WAT and its expression is elevated in WAT in
   obesity,^[ [192]^37 ^] suggesting a role in promoting adiposity and
   regulating lipid homeostasis. We found that mice lacking Mtch2
   specifically in adipose tissues gained less body weight and fat mass
   with noticeably smaller adipocytes when fed an HFD. These results
   provide the first direct in vivo evidence that Mtch2 expression in
   adipose tissue is required for fat accumulation during dietary lipid
   oversupply. This was extended in vitro, where Mtch2 deletion by
   CRISPR/Cas9 in mature 3T3‐L1 adipocytes led to reduced lipid
   accumulation, consistent with previous findings.^[ [193]^38 , [194]^39
   ^] Interestingly, prior studies suggest an obesogenic role of Mtch2 in
   other peripheral metabolically active tissues, with Mtch2 deletion in
   muscle^[ [195]^16 ^] and liver^[ [196]^38 ^] leading to reduced fat
   accumulation under HFD conditions. Thus, it is possible that targeting
   Mtch2 in multiple peripheral tissues may synergistically reduce lipid
   accumulation under HFD conditions.

   Obesity arises from a chronic imbalance between food intake and energy
   expenditure. Our research found that adipose‐specific Mtch2 deletion
   did not affect food intake under either chow diet or HFD conditions.
   This finding is consistent with results from mice with liver‐specific
   Mtch2 modification.^[ [197]^38 ^] However, in contrast, increased food
   intake was observed in mice with Mtch2 deletion in either the forebrain
   (chow diet),^[ [198]^40 ^] or muscle (chow diet and HFD).^[ [199]^16 ^]
   Given that appetite and food intake are primarily controlled by the
   brain, particularly the hypothalamus,^[ [200]^41 , [201]^42 ^] it is
   likely that central Mtch2 plays a distinct role in controlling feeding
   behavior compared to peripheral Mtch2. The reason for increased food
   consumption in muscle‐specific Mtch2 deletion is unclear, but it may be
   a secondary effect stemming from muscle alterations, which could signal
   the hypothalamus to influence feeding. The complex regulation of
   feeding by Mtch2 warrants further investigation, particularly within
   the hypothalamus circuits that control feeding. Additionally, human
   studies have linked MTCH2 gene variants to emotional eating,^[ [202]^43
   ^] indicating MTCH2‐ particularly in the central nervous system – may
   play a role in integrating emotional states with feeding behavior.

   The current study reveals that MTCH2 acts as a suppressor of energy
   expenditure in adipose tissue. A previous study also shows that MTCH2
   control energy demand and expenditure to fuel anabolism during
   adipogenesis.^[ [203]^44 ^] The deletion of adipose Mtch2 increases
   whole‐body energy expenditure, largely through increased BAT
   thermogenesis and scWAT browning, driven by UCP1 upregulation. Although
   the mechanisms may differ, muscle‐specific Mtch2 knockout^[ [204]^16 ^]
   also increases energy expenditure in association with augmented
   mitochondrial biogenesis and oxidative phosphorylation in muscle. In
   our study, the enhanced BAT thermogenesis and scWAT beige observed with
   adipose deletion of Mtch2 was linked to greater heat production,
   increased whole‐body energy expenditure, and improved hepatic steatosis
   and insulin resistance under energy‐excess conditions. One limitation
   in our study is not assessing BAT activation by PET‐CT as previously
   described,^[ [205]^45 ^] though we employed multiple alternative
   methods instead. Moreover, the extensive upregulation of genes in BAT
   enriched in pathways related to energy metabolism, such as
   thermogenesis, autophagy, and lysosome function, indicates a
   coordinated response to enhance cellular energy expenditure and
   metabolic efficiency. The previously reported roles of Mtch2, as an
   outer mitochondrial membrane protein gatekeeper^[ [206]^34 ^] involved
   in mitochondrial dynamics and metabolism, including fusion, elongation,
   and regulation of gene transcription,^[ [207]^35 , [208]^46 , [209]^47
   ^] highlight its importance in mitochondrial function and energy
   homeostasis. Additionally, Mtch2 has been implicated in promoting
   adipogenesis^[ [210]^39 ^] and inhibiting fat oxidation,^[ [211]^38 ^]
   further emphasizing its multifaceted roles in regulating lipid
   homeostasis. Together, our findings provide new insights into the
   complex regulation of lipid homeostasis by Mtch2, highlighting its
   potential as a target for metabolic intervention.

   Our study reports an unidentified link between Mtch2, autophagy, and
   thermogenesis. Autophagy has been shown to be critical in lipid
   metabolism^[ [212]^48 ^] and thermogenesis.^[ [213]^49 , [214]^50 ,
   [215]^51 ^] Additionally, autophagy regulates the initiation of
   lipolysis or lipophagy,^[ [216]^52 ^] and its activation promotes
   lipolysis and fatty acid oxidation.^[ [217]^50 , [218]^53 ^] In this
   study, Mtch2 knockout activated autophagy, leading to increased
   lipolysis, upregulated UCP1 levels, and enhanced thermogenesis. Mouse
   RNA‐seq and fly proteomics analyses revealed marked upregulation of
   autophagy and lysosome pathways following Mtch2 deletion, which may
   promote cellular turnover and improve metabolic function by clearing
   damaged organelles and proteins. Overall, the changes point to an
   adaptation favoring increased energy utilization and cellular renewal,
   leading to mitigation of metabolic dysfunction in the context of
   overnutrition.

   Mechanistically, Bcl‐2 is a well‐known repressor of autophagy^[
   [219]^24 , [220]^54 ^] and a regulator of mitochondrial function.^[
   [221]^26 , [222]^27 , [223]^55 ^] Elevated Bcl‐2 expression has also
   been linked to increased adiposity and body weight in mice.^[ [224]^56
   ^] In this study, Mtch2 knockout mitigated obesity and reduced Bcl‐2
   protein levels, suggesting that Bcl‐2 may mediate metabolic benefit of
   Mtch2 ablation via autophagy. Previous studies demonstrate that Bcl‐2
   downregulation induces autophagy without compromising the mitochondrial
   functions,^[ [225]^57 ^] whereas Bcl‐2 upregulation^[ [226]^58 ^] or
   impaired degradation^[ [227]^59 , [228]^60 ^] suppresses autophagy. Our
   in vivo and in vitro findings reveal that while Mtch2 protein does not
   directly interact with Bcl‐2 protein, it post‐transcriptionally
   regulate Bcl‐2 protein stability, likely through stabilization or
   degradation mechanisms. These results broaden the understanding of how
   Mtch2 impacts cellular proteostasis,^[ [229]^34 , [230]^35 ^]
   particularly in maintaining Bcl‐2 protein levels to suppress autophagy,
   lipolysis, and energy expenditure in obesity. This highlights the
   critical role of Mtch2 in adipose tissue metabolism and energy
   homeostasis through mitochondrial and autophagic pathways. Notably,
   phosphorylation of Bcl‐2 at residues T69, S70, and S87 inhibits its
   anti‐autophagic function;^[ [231]^24 , [232]^61 ^] however, whether
   Mtch2 impacts Bcl‐2 phosphorylation remains unexplored. Further studies
   are needed to elucidate the precise mechanism by which Mtch2 controls
   Bcl‐2‐mediated autophagy thereby influencing thermogenesis and overall
   energy expenditure.

   Obesity is linked to cognitive decline,^[ [233]^62 ^] and MTCH2
   variants have been associated with conditions such as Alzheimer's
   disease^[ [234]^63 ^] and emotional eating.^[ [235]^43 ^] That MTCH2
   may be a critical player in neural cell biology was suggested in a
   study demonstrating mitochondrial dysfunction and impaired cognitive
   function in mice deficient in forebrain Mtch2.^[ [236]^40 ^] More work
   is necessary to elucidate the role of MTCH2 in both obesity and its
   related neurodegeneration. In addition, previous studies have shown a
   gender difference in MTCH2 expression, with protein levels
   significantly upregulated in women but not men with obesity.^[ [237]^37
   ^] Future work is required to explore this gender difference,
   particularly in female mouse models. Importantly from a therapeutic
   perspective, pharmacological or gene‐based interventions aimed at
   modulating MTCH2 activity could offer promising strategies for treating
   metabolic disorders such as obesity and insulin resistance. However,
   due to its extensive distribution and complexity, careful evaluation of
   the systemic effects of MTCH2 manipulation is needed to avoid
   unintended consequences in tissues where MTCH2 may play a protective
   role.

   In conclusion, our study identifies adipose MTCH2 as a key regulator of
   energy expenditure, with broad implications for understanding obesity
   and developing new treatments. By elucidating its role in modulating
   energy expenditure, lipid metabolism, and autophagy, we provide a solid
   foundation for the development of novel therapeutic strategies aimed at
   targeting MTCH2. Future research will continue to advance our
   understanding of Mtch2's mechanisms and potential to improve metabolic
   health.

4. Experimental Section

Flies

   All Drosophila melanogaster stocks were cultivated on a conventional
   medium comprising agar, sugar, and yeast, and were reared in an
   incubator at a constant temperature of 25 °C under a 12‐h light/dark
   period. The Actin‐Gal4 driver line was sourced from the Bloomington
   Drosophila Stock Center. The RNAi fly lines utilized in this study were
   obtained from the Vienna Drosophila Resource Center (VDRC) and listed
   in Table [238]S1 (Supporting Information). The flies were nurtured on a
   standard medium that included 1% agar, 3.6% yeast, 2% yellow corn meal,
   5.4% sugar, and 3% molasses, and were also subjected to a 25 °C
   incubation environment with a 12‐h light/dark cycle. Following
   eclosion, adult male flies, aged 4–7 days post‐hatch, were selected for
   experimentation under various dietary conditions. For the experimental
   groups, flies were subjected to distinct diets. The normal diet
   consisted of 1% agar, 3.6% yeast, and 5.4% sucrose. The high sucrose
   diet was formulated with 1% agar, 3.6% yeast, and an elevated sucrose
   concentration of 16.2%. In contrast, the high‐fat diet contained 15%
   coconut oil, complemented by 1% agar, 3.6% yeast, and 5.4% sucrose.

Mice

   Heterozygous Mtch2^flox/+ mice were generated using CRISPR/Cas9
   technology and obtained from Cyagen Biological Technology Co., Ltd.
   (Suzhou, China) on a C57BL/6J background. Homozygous Mtch2^flox/flox
   mice were subsequently produced by intercrossing heterozygous pairs. To
   generate adipose tissue‐specific Mtch2 knockout mice, Mtch2^flox/flox
   mice were bred with mice expressing Cre recombinase under the control
   of the Adiponectin promoter. Genotyping was performed by PCR, with
   primer sequences listed in Table [239]S2 (Supporting Information).
   Littermate mice of the same sex and age were utilized as controls for
   all experiments. All mice were maintained in an environment with
   ambient room temperature (23 ± 2 °C) and a 12‐h light/dark cycle, with
   unrestricted access to water and food, unless otherwise indicated. The
   experimental procedures involving mice were conducted by an
   Institutional Animal Care and Use Committee (IACUC)‐approved protocol
   from Sun Yat‐sen University, with the specific approval number
   SYSU‐IACUC‐2022‐000298.

Cell and Cell Culture

   Human embryonic kidney cell line HEK293T and the 3T3‐L1 preadipocyte
   cell line were both procured from the American Type Culture Collection
   (ATCC). The cells were maintained in Dulbecco's Modified Eagle Medium
   (DMEM, Life Technologies) supplemented with 10% (v/v) Fetal bovine
   serum (FBS) and 1% penicillin‐streptomycin. Cultivation was conducted
   at 37 °C in an atmosphere containing 5% CO[2].

Functional Analyses of Drosophila

   Initiating crosses by mating ten virgin Actin‐Gal4 females with 5–7
   young males of either UAS‐RNAi or control w1118 strains to induce
   RNAi‐mediated whole‐body knockdown of the target genes. Upon eclosion
   of the F1 progeny, male flies were isolated and, at 4–7 days of age,
   assigned to either a normal diet or a high sucrose diet for 10 days.
   Body weight was ascertained using an analytical balance. To assess
   triglyceride levels, a group of ten male flies was weighed, homogenized
   in 200 µL of ice‐cold PBST (phosphate‐buffered saline with 0.05% Tween
   20), and sonicated. Subsequently, 800 µL of ice‐cold PBST was added to
   the homogenate and mixed thoroughly. A 50 µL aliquot of this mixture
   was utilized to quantify triglycerides with the Roche triglycerides
   assay kit, following the manufacturer's guidelines. Triglyceride
   measurements were normalized relative to body weight. The data were
   standardized to a Z‐score employing the formula: 𝑍 = (𝑋−𝜇)/𝜎, where 𝑋
   is the individual's value, 𝜇 is the control group mean, and σ is the
   standard deviation of the control group. The threshold for statistical
   significance was set at a 95% confidence interval for a two‐tailed
   test, equating to a Z‐score threshold of ≥ 1.96 or ≤ −1.96.^[ [240]^64
   ^] Target genes identified as hits were validated through three
   independent screening assays for confirmation.

Protein Mass Spectrometry

   Protein mass spectrometry analysis was conducted following the methods
   outlined in a previous report.^[ [241]^65 ^] In summary, the proboscis
   or the entire body of the specimen was extracted and homogenized in a
   lysis buffer containing 6 m urea, 2 m thiourea, 25 mm TEAB, and 0.1%
   SDS. The homogenate was then subjected to sonication, followed by
   centrifugation at 15 000 g for 10 min at ambient temperature. The
   resulting supernatant was subjected to precipitation overnight with
   pre‐chilled acetone. The precipitated protein pellets were subsequently
   resuspended in a buffer comprising 6 m urea, 2 m thiourea, and 25 mm
   TEAB. Protein reduction was performed using 10 mm DTT at 30 °C for
   60 min, followed by alkylation with 25 mm IAA for 30 min at room
   temperature in darkness. The protein mixture was then enzymatically
   digested with LysC (enzyme‐to‐substrate ratio of 1:100) at 30 °C for
   2 h. Post‐digestion, the protein solution was diluted 1:5 with 25 mm
   TEAB and further digested with trypsin (enzyme‐to‐substrate ratio of
   1:50) at 37 °C for an extended period. The resultant peptide mixture
   underwent desalting via stage‐tips and was subsequently lyophilized
   using vacuum centrifugation in preparation for total proteome analysis.

Mass Spectrometry Analysis and Data Processing

   MS‐based total proteome analysis was performed on an Easy nLC‐1000
   UHPLC coupled to a Q‐Exactive in positive polarity mode as in the
   previous report.^[ [242]^65 ^] The in‐house packed C18AQ column (75 µm
   × 40 cm, 1.9 µm particles) separated the peptides using a 360‐min
   gradient of 10–35% acetonitrile (ACN) containing 0.1% formic acid (FA)
   at a flow rate of 200 nL min^−1 and a column temperature of 55 °C. The
   MS1 scan encompassed a range of 300–1750 m/z with a resolution of
   70 000, AGC target of 3e6, and injection time of 100 ms. Data‐dependent
   MS/MS acquisition subsequently targeted the top 20 ions using HCD
   fragmentation with a resolution of 17 500, AGC target of 5e5, injection
   time of 60 ms, normalized collision energy (NCE) of 25, and an
   isolation window of 2.0 m/z. Raw data processing employed MaxQuant
   (v1.5.3.25) against a UniProt Drosophila database. Minor adjustments
   were made to the default settings: oxidation of methionine and
   N‐terminal protein acetylation were designated as variable
   modifications, while cysteine carbamidomethylation was set as a fixed
   modification. Re‐quantification, second peptide searches, and
   match‐between‐runs features were enabled. Peptide spectral match (PSM)
   and protein false discovery rate (FDR) were both set to 1%. The
   “Label‐free Quantification (LFQ) method” was employed for protein
   quantification and normalization. Bioinformatics analysis, primarily
   conducted using the LIMMA package within the R programming environment,
   aimed to identify differentially expressed proteins. Functional
   annotation was performed using DAVID v6.8 beta
   ([243]https://david‐d.ncifcrf.gov/).

Gene Analysis in Clinical Cohorts

   The initial transcriptomics data were derived from abdominal
   subcutaneous white adipose tissue (WAT) biopsies of 25 male and 28
   female individuals, generated as previously reported. These
   transcriptome profiles were acquired using the Affymetrix Human Gene
   1.1 ST Array, and the data have been deposited in the NCBI Gene
   Expression Omnibus (GEO) under the accession code [244]GSE77962.^[
   [245]^66 ^] Subsequent transcriptome profiles were sourced from
   subcutaneous WAT biopsies of 770 male participants in the METSIM study,
   utilizing the Affymetrix Human Genome U219 Array, with the dataset
   archived under accession code [246]GSE70353.^[ [247]^67 ^] Additional
   transcriptomics data were procured from abdominal subcutaneous adipose
   needle biopsies of 56 women, employing the Affymetrix Human Gene 1.0 ST
   Array, and registered with the accession code [248]GSE25401.^[ [249]^68
   ^] In these analyses, obesity was defined by a BMI exceeding 30 kg m^−
   ^2.

Body Weight

   Male mice were fed either a normal chow diet (NCD) from four weeks of
   age or a high‐fat diet (HFD; Research Diets, D12492, 60% kcal from fat,
   20% kcal from carbohydrate, 20% kcal from protein, 5.42 kcal g^−1)
   starting at nine weeks of age for the indicated experimental periods.
   Body weight was monitored weekly at a consistent time point throughout
   the study.

Intraperitoneal Glucose Tolerance Test (IPGTT) and Insulin Tolerance Test
(ITT)

   IPGTT and ITT were performed at the indicated time points. Mice were
   fasted for 15 h (18:00‐09:00) before IPGTT or for 6 h (09:00‐15:00)
   before ITT. Glucose (2.0 g kg^−1 body weight) or insulin (0.75 U kg^−1
   body weight) was administered intraperitoneally, and blood glucose
   levels were measured using a digital glucometer (Yuwell, Jiangsu Yuyue
   Medical Equipment & Supply Co., LTD, China) at baseline and 15, 30‐,
   60‐, 90‐, and 120‐min post‐injection.

Food Intake

   Spontaneous food intake was measured after a 24‐h acclimation period in
   individual cages with ad libitum food access, as previously
   described.^[ [250]^1 ^] Equal food quantities were provided to each
   mouse at the start of the measurement period. Food remnants and
   spillage were collected and weighed after 24 h. Food intake was
   calculated by subtracting the combined weight of remnants and spillage
   from the initial food amount.

Core Temperature and Surface Temperature

   Core body temperature was measured rectally in conscious mice at
   specified time points following cold exposure (4 °C) using an
   electronic thermometer (FT3400, DEJR, China). Surface temperature was
   assessed using thermal imaging with a Fluke Ti480U infrared thermal
   camera (Fluke Corporation, USA) after 2 h of cold exposure (4 °C).

Energy Metabolism by Indirect Calorimetry

   Mice were acclimatized to Promethion Metabolic Systems (Sable Systems
   International, North Las Vegas, NV, USA) for 24 h before data
   collection. Oxygen consumption, carbon dioxide production, respiratory
   exchange ratio, energy expenditure, and locomotor activity were
   continuously monitored for 48 h. Oxygen consumption, carbon dioxide
   production, and energy expenditure were normalized to body weight as
   previously described.^[ [251]^1 , [252]^69 ^]

Body Composition

   Body composition, including fat and lean mass, was assessed in mice
   using nuclear magnetic resonance (NMR) spectroscopy (Small Animal Body
   Composition Analysis and Imaging System, MesoQMR23‐060H‐I, Suzhou
   Niumag Corporation, Shanghai, China) according to the manufacturer's
   instructions.

Tissue Collection

   Mice were euthanized by intraperitoneal injection of pentobarbital
   sodium (50 mg kg^−1). Following euthanasia, adipose tissues (BAT,
   scWAT, eWAT, and rWAT) and liver were carefully excised, weighed,
   snap‐frozen in liquid nitrogen, and stored at −80 °C for subsequent
   analysis.

Hepatic Triglyceride Measurements

   ≈20 mg of snap‐frozen tissue was homogenized in 180 µL of ice‐cold
   absolute ethanol and subsequently centrifuged at 2500 rpm for 10 min at
   4 °C. A 2.5 µL aliquot of the supernatant was used to quantify hepatic
   triglycerides using the GOP/PAP method according to the manufacturer's
   instructions.

Plasma Parameters Measurement

   Plasma ALT and AST were determined using a Beckman CX5 automated
   biochemical analyzer (Beckman Coulter, Inc., United States) according
   to standard operating procedures of the Guangdong Engineering &
   Technology Research Center for Disease‐Model Animals, Sun Yat‐sen
   University.

RNA Extraction and qPCR

   Total RNA was extracted from tissues or cells using TRIzol reagent.
   Subsequently, 1 µg of RNA was reverse transcribed into cDNA using a
   PrimeScript RT reagent kit according to the manufacturer's protocol.
   Real‐time quantitative PCR (qPCR) was performed using SYBR Green dye on
   a QuantStudio 5 system (Applied Biosystems, Thermo Fisher Scientific).
   Gene expression levels were normalized to the housekeeping gene, 18S
   ribosomal RNA (Rn18S),^[ [253]^70 ^] using the ΔΔCt method and
   expressed as fold change relative to controls. Primer sequences are
   listed in Table [254]S2 (Supporting Information).

RNA‐Sequencing and Data Analysis

   RNA sequencing was outsourced to BerryGenomics. Before library
   preparation, RNA quality from BAT was assessed using an Agilent
   TapeStation. Sequencing libraries were constructed using the TruSeq RNA
   Sample Preparation Kit v2 (Illumina) according to the manufacturer's
   instructions. Library quality and fragment size were validated using a
   2100 Bioanalyzer (Agilent). Normalized libraries were then pooled for
   sequencing on an Illumina NovaSeq 6000 platform using barcoded
   multiplexing for single‐end reads of 150 bp. RNA sequencing data
   analysis was performed using OmicShare tools, a freely accessible
   online platform^[ [255]^71 ^] ([256]https://www.omicshare.com/tools).

Mtch2 mRNA Expression in Mice

   Twelve‐week‐old male wide‐type C57BL/6J mice were euthanized, and the
   tissues were carefully excised to extract RNA. Adipose tissues of obese
   mice and same‐aged lean mice were obtained from the previous
   research.^[ [257]^1 ^] Mtch2 mRNA expression was analyzed by qPCR.

Mitochondrial DNA (mtDNA) Copy Number Analysis

   Relative mtDNA copy number analysis was performed as previous report.^[
   [258]^72 ^] Briefly, genomic DNA from adipose tissue was extracted
   using a TIANamp Genomic DNA Kit. Quantitative PCR was performed using
   QuantStudio 5 (Applied Biosystems, Thermo Fisher Scientific) to
   quantify mtDNA and nuclear 18S rRNA. The ΔΔCt method was used to
   calculate the relative mtDNA copy number, normalized to 18S rRNA.
   Primer sequences are listed in Table [259]S3 (Supporting Information).

Western Blot

   Tissue lysates were prepared using RIPA buffer (Hangzhou FuDe
   Biological Technology Co., LTD, China) supplemented with protease and
   phosphatase inhibitors (including 1 mm PMSF). Total protein lysates
   were boiled with a loading buffer containing 10% SDS‐PAGE. Soluble
   proteins (30 ug) were separated on a 10% or 12.5% SDS polyacrylamide
   gel and transferred onto a 0.2 µm PVDF membrane (Immobilon‐PSQ, Merck).
   After blocking with 5% non‐fat milk in TBST buffer (containing 0.1%
   Tween 20 in 1xTBS) for 1 h at room temperature, membranes were
   incubated with primary antibodies overnight at 4 °C. Membranes were
   then rinsed three times for 10 min each time in TBST and were incubated
   with an HRP‐conjugated secondary antibody for 1 h at room temperature.
   Antibody α‐tubulin (Ray antibody Cat#RM2007), β‐actin (Servicebio
   Cat#GB15003), UCP1 (Abcam Cat#ab209483), HSL (ABclonal Cat#A15686),
   ATGL (ABclonal Cat#A5126), LC3A/B (Cell Signaling Technology
   Cat#12741), SQSTM1/p62 (Servicebio Cat#GB11531), Bcl‐2(ABclonal
   Cat#A19693), MTCH2(Absin Cat#abs143485), Goat anti‐rabbit IgG HRP (Ray
   Antibody Cat#RM3001) and Goat anti‐mouse IgG HRP (Ray Antibody
   Cat#RM3002) were used. Immunoblotted bands were detected using enhanced
   chemiluminescence (ECL) reagent, imaged by a BG‐gdsAUTO 720 Imaging
   system (Baygene, Beijing), and quantified by densitometry using Fiji
   software (NIH). The intensity of proteins of interest was normalized to
   the intensity of the loading control.

Co‐Immunoprecipitation (Co‐IP) Assays

   Co‐IP was performed using Immunoprecipitation Kit (Beyotime, Cat
   #P2179S) according to the manufacturer's protocol. 3T3L1 cells were
   washed with PBS for three times and lysed on ice for 30 min. One mg
   cellular proteins were then incubated with 1 µg primary antibodies at
   4 °C overnight on the roller. 20 µL Protein A/G‐Magnetic Beads were
   later added to capture antigen‐antibody complex at 4 °C for 2 h.
   Subsequently, the samples were washed for three times with magnetic
   grate, suspended with loading buffer, and boiled at 95 °C for 5 min.
   The supernatant was collected for Western blot analysis with 20 µL
   supernatant per well.

Histological Analysis and Immunohistochemistry

   Adipose tissues and liver tissues were fixed in 10% neutral buffered
   formalin, embedded in paraffin, and sectioned at a 4 µm thickness.
   Hematoxylin and eosin (H&E) staining of the resulting sections was
   performed according to the manufacturer's instructions. Adipocyte size
   was analyzed by Fiji software (NIH) as previously reported.^[ [260]^17
   ^] Immunohistochemical staining of the resulting sections for UCP1 was
   performed with a rabbit monoclonal antibody (Abcam Cat#ab209483) at
   1:2000 dilution. Antigen retrieval was conducted using an EDTA
   solution. The images were captured using a Nikon microscope.

Transmission Electron Microscope

   BAT tissues from 19‐week HFD‐fed mice were dissected into 1 mm^3 cubes
   and fixed in 2.5% glutaraldehyde prepared in sodium cacodylate buffer
   (2 h at room temperature, followed by 12 h at 4 °C). Subsequent
   processing involved post‐fixation in 1% osmium tetroxide, staining with
   2% uranyl acetate, dehydration in an ethanol series (30–100%), and
   embedding in EMbed‐812 resin. Ultrathin sections (50–60 nm) were
   stained with uranyl acetate and lead citrate before imaging on a
   Hitachi H600 electron microscope (Hitachi, Japan) at 120 kV.

Mtch2 Knockout in 3T3‐L1 Cells by CRISPR/Cas 9

   3T3‐L1 cells were infected with a second‐generation lentiviral system
   packaged in 293T cells to generate Mtch2 knockout (KO) cells as
   previous report.^[ [261]^73 ^] Lentiviral vectors expressing sgRNA
   targeting Mtch2 (sgRNA 1#: TACCGGGAAGAAGGCATCGT; sgRNA 2#: CTTTCA
   CGTACATGAGCGGC) were constructed in pLentiCRISPRv2 and transduced into
   3T3‐L1 cells. Puromycin selection was used to enrich KO cells, which
   were subsequently validated by Western blot.

Adipocyte Differentiation

   3T3‐L1 preadipocytes were induced to differentiate two days
   post‐confluence (day 0). The differentiation medium contained insulin
   (5 µg mL^−1), dexamethasone (1 µm), 3‐isobutyl‐1‐methylxanthine
   (0.5 mm), indomethacin (125 nm), triiodothyronine (1 nm), and
   rosiglitazone (1 µm). From day 2, the medium was replaced with a
   maintenance medium containing insulin (5 µg mL^−1), triiodothyronine (1
   nm), and rosiglitazone (1 µm), which was changed every other day. Fully
   differentiated adipocytes were harvested on day 6.

Oil Red O Staining

   Oil Red O staining was performed as previously reported.^[ [262]^74 ^]
   Oil Red O (0.5% (mg/v) in isopropanol) was diluted with water (3:2),
   and then filtered through a 0.45 mm filter. The differentiated
   adipocytes were washed three times with PBS and fixed for 30 min with
   4% (v/v) paraformaldehyde (PFA), then the fixed cells were incubated
   with filtered Oil Red O for 1 h at room temperature to visualize lipid
   droplets. After being washed with PBS, 100% isopropanol was added as an
   extraction solution to extract the staining dye of cells. The
   absorbance of the extracted dye was measured at 510 nm using a BioTek
   Epoch2 microplate reader.

Statistical Analysis

   Data are presented as Mean ± S.E.M. Statistical analyses were performed
   using GraphPad Prism 9. Unpaired t‐tests, one‐way ANOVA with Bonferroni
   post hoc test, or two‐way ANOVA with Bonferroni post hoc test were used
   as appropriate. Differences were considered significant at *p < 0.05,
   **p < 0.01, ***p < 0.005, or ****p < 0.001.

Conflict of Interest

   The authors declare no conflict of interest.

Author Contributions

   X.Y.Z., Q.P.W., and G.G.N. conceptualized the study. X.Y.Z. and Q.P.W.
   developed the methodology. X.Y.Z., B.C.Z., H.L.L., Y.L., B.W., and
   A.Q.L. conducted the investigation. X.Y.Z., B.C.Z., H.L.L., and Q.P.W.
   performed the validation. X.Y.Z., A.Q.L., J.X.M., and Q.P.W. created
   the visualization. X.Y.Z. and Q.P.W. wrote the original draft. X.Y.Z.,
   T.S.Z., H.X.Y.H., J.S., D.C., N.G., J.H., J.X.M., D.R.L., Y.Y.D.,
   Y.C.S., G.G.N., and Q.P.W. reviewed and edited the manuscript. Q.P.W.
   and Y.C.S. acquired funding. Q.P.W. and G.G.N. provided resources.
   G.G.N., Y.C.S., and Q.P.W. supervised the study.

Supporting information

   Supporting Information
   [263]ADVS-12-2416598-s001.doc^ (1.2MB, doc)

Acknowledgements