Mipep deficiency in adipocytes impairs mitochondrial protein maturation and leads to systemic inflammation and metabolic dysfunctions

Nozaki, Yuka; Kobayashi, Masaki; Fukuoh, Tomoyoshi; Ishimatsu, Mamiko; Narita, Takumi; Taki, Kanari; Hirao, Yuto; Ayabe, Shota; Yokoyama, Miku; Otani, Yuina; Mizunoe, Yuhei; Matsumoto, Mami; Ohno, Nobuhiko; Kaifu, Tomonori; Okazaki, Shogo; Goitsuka, Ryo; Nakagawa, Yoshimi; Shimano, Hitoshi; Iwakura, Yoichiro; Higami, Yoshikazu

doi:10.1038/s41598-025-97307-6

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Published: 14 April 2025

Mipep deficiency in adipocytes impairs mitochondrial protein maturation and leads to systemic inflammation and metabolic dysfunctions

Scientific Reports volume 15, Article number: 12839 (2025) Cite this article

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Abstract

Most mitochondrial proteins encoded in the nuclear genome are synthesized in the cytoplasm. These proteins subsequently undergo maturation through the cleavage of a signal sequence at the N-terminus by one or two mitochondrial signal peptidases, which is essential for their function within mitochondria. The present study demonstrates that adipocyte-specific knockout of one mitochondrial signal peptidase, mitochondrial intermediate peptidase (MIPEP), resulted in disordered mitochondrial proteostasis of MIPEP substrate proteins and their defective maturation. MIPEP deficiency in white and brown adipocytes suppressed the expression of adipocyte differentiation, lipid metabolism, and mitochondrial biogenesis genes. These alterations led to lipoatrophy in white adipose tissue and the whitening of brown adipose tissue. Additionally, it induced an atypical mitochondrial unfolded protein response and local inflammation in white and brown adipose tissue. Furthermore, it induced fatty liver and splenomegaly and caused systemic impairments in glucose metabolism and inflammation. These findings indicate that maturation defects of certain mitochondrial proteins and subsequent proteostasis disorders in white and brown adipocytes cause chronic and systemic inflammatory and metabolic dysfunctions.

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Introduction

Mitochondria play key roles in regulating oxidative phosphorylation (OXPHOS), the TCA cycle, β-oxidation, calcium homeostasis, and apoptosis in living organisms. Therefore, mitochondrial dysfunction is implicated in various pathologies, including metabolic disorders and age-related diseases¹ .

Over 99% of mitochondrial proteins are encoded by nuclear DNA and synthesized in the cytoplasm. Most mitochondrial proteins exist as a precursor form with a mitochondrial targeting signal sequence at the N-terminus. After targeting and entering mitochondria, the targeting signal is cleaved from precursor proteins by mitochondrial signal peptidases, including mitochondrial processing peptidase (MPP) and mitochondrial intermediate peptidase (MIPEP); this leads to the mature protein form existing in the mitochondrial matrix^2,3. Therefore, it could be argued that the maturation of mitochondrial proteins, and more broadly, mitochondrial proteostasis, is regulated by mitochondrial signal peptidase.

White adipose tissue (WAT) predominantly comprises adipocytes, with preadipocytes, mesenchymal cells, and inflammatory cells also present⁴. Although white adipocytes contain few mitochondria, these mitochondria play a pivotal role in WAT function and whole-body metabolism⁵. By contrast, brown adipocytes are rich in mitochondria that contain uncoupling protein 1, which is involved in heat production. Thus, mitochondria in white and brown adipocytes play pivotal roles in WAT and BAT function and whole-body metabolism⁶. Previously, we reported that long-term caloric restriction, which improves metabolism, suppresses age-related pathophysiology, extends lifespan, and enhances mitochondrial biogenesis in WAT via peroxisome proliferator-activated receptor gamma coactivator 1α (PGC-1α) in a sterol regulatory element binding protein (SREBP-1c)-dependent manner⁷. We also found that caloric restriction upregulates Mipep, but not Mpp in a SREBP-1c-dependent manner in WAT^8,9.

Previously, four patients with a Mipep variant (recently designated as combined oxidative phosphorylation deficiency 31) were found to exhibit a systemic disorder involving developmental delay, cardiomyopathy, hypotonia, and infantile lethality¹⁰. In addition, a recent report described a patient with heterozygous Mipep variants who did not present with cardiomyopathy but instead exhibited developmental delay, hypotonia, mild optic neuropathy, and mild ataxia¹¹. Moreover, a functional analysis of patient fibroblasts and HEK293FT cells carrying heterozygous Mipep variants demonstrated that MIPEP deficiency impairs the processing of certain components of the OXPHOS complex and other mitochondrial proteins linked to human diseases¹¹. Thus, alterations in MIPEP expression and function appear to have important cellular and systemic effects.

MIPEP cleaves an octapeptide from the N-terminus of intermediate precursor proteins processed by MPP, thus contributing to the second of two successive cleavage steps^12,13,14,15. Cytochrome c oxidase subunit IV (COX4) and malate dehydrogenase 2 (MDH2) are substrates of MIPEP^16,17. In a previous report, we also demonstrated that a representative mitochondrial protein, sirtuin (SIRT3), was cleaved sequentially by both MPP and MIPEP⁸. Given that SIRT3 activates several mitochondrial enzymes, including superoxide dismutase 2 (SOD2) and isocitrate dehydrogenase 2, via its deacetylase activity^18,19, MIPEP appears to play a pivotal role in mitochondrial functions. Mitochondrial dysfunction results in the activation of a stress response, such as the mitochondrial unfolded protein response (UPR^mt). In mitochondrial dysfunction, the mitochondria-derived retrograde signaling pathway is activated to transmit mitochondrial dysfunction to the nucleus, which results in the induction of the UPR^mt; this, in turn, leads to the transcriptional activation of nuclear-encoded genes including mitochondrial chaperones, proteases, and mitokines^20,21. The UPR^mt is mediated by the expression of transcription factors, including C/EBP homologous protein 10, activating transcription factor 3 (ATF3), ATF4, and ATF5^22,23,24,25. Furthermore, mitochondrial dysfunction is linked to the repression of cytosolic translation through the phosphorylation of eukaryotic translation initiation factor 2 A (eIF2α)²⁶.

To elucidate the role of MIPEP in adipocytes, we generated adipocyte-specific Mipep knockout (aMKO) mice. We examined the impact of MIPEP deficiency on both WAT and BAT, as well as systemic effects caused by adipocyte-specific Mipep knockout.

Results

Characteristics of aMKO mice

We generated Mipep^flox/flox mice as illustrated in Supplementary Fig. 1. Initially, we crossed Mipep^flox/flox mice with CAG-Cre mice, but the resulting systemic Mipep knockout (KO) mice were embryonically lethal. Subsequently, we obtained aMKO mice by crossing Mipep^flox/flox (control) mice with Adiponectin-Cre mice. Both Mipep mRNA expression and MIPEP protein expression were significantly lower in the both WAT and BAT of aMKO mice than in that of control mice (Supplementary Fig. 2A-D). In addition, the body weight of aMKO mice was lower than that of control mice, despite similar food intake, body temperature, and locomotive activity (Supplementary Fig. 2E-I). Furthermore, there were no significant differences in VO₂, VCO₂, respiratory quotient, and energy expenditure between control and aMKO mice (Supplementary Fig. 2J-M).

Computed tomography (CT) analysis revealed that aMKO mice had less adipose mass, particularly in subcutaneous WAT (sWAT) (Fig. 1A). In aMKO mice, gonadal WAT (gWAT) was markedly atrophic (Fig. 1B). Histological analysis revealed that the size of mature white adipocytes remained unaltered. However, small adipocyte clusters and cells exhibiting lipoblast-like characteristics were scattered throughout gWAT, indicating defective differentiation of preadipocytes into mature white adipocytes in aMKO mice (Fig. 1C, D). Electron microscopic analysis showed that mitochondria with numerous cristae in adipocytes exhibited a long oval shape in control mice. However, the shape of mitochondria was short and swollen, and the cristae were absent, in adipocytes (but not in non-adipocytes, as indicated in Fig. 1E-γ) from aMKO mice. Additionally, donut-shaped mitochondria were identified in aMKO mice (Fig. 1E).

In the BAT of aMKO mice, the typical brownish appearance was lost, and more large lipid droplets were observed histologically rather than control mice, indicating that MIPEP deficiency promoted whitening (Fig. 1F, G). Electron microscopy analysis also revealed that lipid droplets were enlarged. Conversely, morphological abnormalities in mitochondria were not as pronounced as in WAT, and the cristae structure appeared largely intact in aMKO mice (Fig. 1H).

Finally, liver and spleen weights were significantly higher in aMKO mice than in control mice; no significant differences were observed in the weights of other tissues, including the heart, kidney, and quadriceps femoris muscle (Table 1).

Table 1 Organ weights of adipose-specific Mipep knockout (aMKO) mice.

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Disordered proteostasis of MIPEP substrates in the WAT and BAT of aMKO mice

Because SIRT3, COX4, and MDH2 are the substrate proteins of MIPEP^8,16,17, we examined the protein expression, enzymatic activity, and mRNA expression of these proteins in gWAT and BAT. The protein level of SIRT3 was unaltered in both the gWAT and BAT of aMKO mice (Fig. 2A, B, G, H). SIRT3 deacetylates SOD2, thereby activating its enzymatic activity¹⁸. The ratio of acetylated SOD2 to total SOD2 was significantly higher in aMKO mice than in control mice, suggesting that the enzymatic activity of SOD2 was diminished in both the gWAT and BAT of aMKO mice (Fig. 2C, I). The protein level of COX4 and its enzymatic activity was significantly lower in both the gWAT and BAT of aMKO mice than in that of control mice (Fig. 2A, B, D, G, H, J). By contrast, the protein level of MDH2 was significantly higher in the gWAT but was unaltered in the BAT of aMKO mice; however, the enzymatic activity of MDH2 was significantly lower in the gWAT but was unaltered in the BAT of aMKO mice (Fig. 2A, B, E, G, H, K). Interestingly, as illustrated in the Western blot image, the band representing MDH2 appeared slightly higher, particularly in the BAT and to a lesser extent in the gWAT, in aMKO mice than in control mice (Fig. 2A, G). Despite the observed changes in the protein levels and enzymatic activities of MIPEP substrates in gWAT and BAT, the mRNA levels of Cox4, Mdh2, and Sod2 remained unaltered in aMKO mice (Fig. 2F, L).

Altered gene expression profiles in the WAT and BAT of aMKO mice

To gain insight into the distinctive features of gWAT and BAT in aMKO mice, we conducted a comparative transcriptome analysis of these tissues using RNA-Seq analysis. Differentially expressed genes were extracted using the edgeR (v4.2.0) program package²⁷. In both gWAT and BAT, pathway enrichment analysis demonstrated that expression of genes in the “immune system and related GO” category was upregulated in aMKO mice compared with control mice, with a more pronounced trend observed in gWAT (Fig. 3A, D). By contrast, the expression of genes in the “lipid metabolism and related GO” category was downregulated in both the gWAT and BAT of aMKO mice compared with control mice, with a more pronounced trend observed in BAT (Fig. 3B, E). In gWAT, the expression of genes associated with the “synapse and related GO” category was downregulated in aMKO mice compared with control mice (Fig. 3B).

Mitochondrial proteostasis disorder might induce the UPR^mt and lead to the expression of mitokines including growth differentiation factor 15 (Gdf15; also known as macrophage inhibitory cytokine-1 [Mic1]) and fibroblast growth factor 21 (Fgf21)^20,21. The biosynthesis of fatty acids and cholesterol is regulated by the transcription factors SREBP-1 and 2 (encoded by Srebf1 and 2, respectively)²⁸. The expression of the mitokines Gdf15 and Fgf21 was upregulated, while that of transcription factors involved in fatty acid synthesis (Srebf1; WAT: logFC − 0.90, p-Value 1.1E-05; BAT: logFC − 0.84, p-Value 7.2E-04) was downregulated in aMKO mice compared with control mice (Fig. 3C, F).

Changes in differentiation and mitochondrial biogenesis biomarkers in the WAT and BAT of aMKO mice

The small adipocyte clusters and lipoblasts observed in the gWAT of aMKO mice suggest that MIPEP deficiency may suppress adipocyte differentiation. As expected, the expression of genes implicated in white adipocyte differentiation including Pparg and Cebpa, and that of the mature adipocyte marker Adipoq, was significantly lower in the gWAT of aMKO mice than in that of control mice (Fig. 4A). In the BAT of aMKO mice, the expression of brown adipocyte differentiation genes including Ucp1, Prdm16, and Cidea, and that of Adipoq, was significantly lower than in that of control mice. However, UCP1 protein expression level was unaltered in BAT of aMKO mice (Supplementary Fig. 3). Consistent with the unaltered UCP1 protein, the body temperature was not changed in aMKO mice (Supplementary Fig. 2H).

Notably, Leptin expression was unaltered in gWAT, but significantly higher in the BAT of aMKO mice compared with control mice (Fig. 4A, C). Because of the slight and marked downregulation of Ppargc1a and Nrf1 expression, respectively, in both gWAT and BAT of aMKO mice, we hypothesized that MIPEP deficiency might suppress mitochondrial biogenesis. Indeed, the amount of mitochondrial DNA (mtDNA), which is reflected by the amount of Nd1 and D-loop, was also significantly lower in both the gWAT and BAT of aMKO mice compared with control mice (Fig. 4B, D).

These results imply that MIPEP deficiency suppresses white adipocyte differentiation in WAT, inhibits brown adipocyte differentiation, and promotes the whitening of BAT. In addition, MIPEP deficiency decreases mitochondrial biogenesis in both WAT and BAT.

Changes in stress response biomarkers in the WAT and BAT of aMKO mice

The UPR^mt increases the expression of mitokines, mitochondrial chaperones, and proteases. mRNA levels of the mitokines Gdf15 and Fgf21 were significantly higher in both the gWAT and BAT of aMKO mice than in that of control mice (Fig. 5A, F). The plasma levels of GDF15 were significantly higher in aMKO mice than in control mice, whereas those of FGF21 were not (Table 2). The mRNA level of Gdf15 was not affected by Mipep KO in other tissues including the liver, kidney, heart, and skeletal muscle (Supplementary Fig. 4A). Thus, we concluded that elevated plasma GDF15 is derived from the gWAT and BAT of aMKO mice. Additionally, there was no change in mRNA expression of Fgf21 in the liver (Supplementary Fig. 4B). In contrast to the observed increase in mitokine expression, the expression of mitochondrial chaperone and protease genes was not higher in aMKO mice, and in some cases, it was even lower than in control mice, particularly in BAT (Fig. 5B, G). Cross-talk between the UPR^mt and the endoplasmic reticulum stress response has been reported; these stress responses are positively regulated by the expression of stress-responsive transcription factor genes such as Chop10, Atf3, Atf4, Arf5, Xbp1 (unspliced, total) and Xbp1s (spliced) and phosphorylation of eIF2a²⁹. The expression of Xbp1s was not changed in gWAT, but it was significantly higher in the BAT of aMKO mice than in that of control mice (Fig. 5C, H). The expression of four stress-responsive transcription factors was higher, three of which were significant, in both the gWAT and BAT of aMKO mice, with a particularly pronounced effect observed in BAT (Fig. 5D, I). In aMKO mice, the ratio of the phosphorylated form of eIF2a to total eIF2 was not changed in gWAT but was significantly higher in BAT compared with control mice (Fig. 5E, J). Therefore, these findings suggest that the stress response induced by MIPEP deficiency is accelerated in BAT rather than WAT.

Table 2 Plasma biochemical data and cyto(mito)kine levels of adipose-specific Mipep knockout (aMKO) mice.

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Inflammation in the WAT of aMKO mice

To gain insight into the enhanced expression of genes classified as “immune system process”-related in the gWAT of aMKO mice, flow cytometry analysis was performed. In the gWAT of aMKO mice, cluster of differentiation (CD)11b⁺ F4/80⁺ cells, which characterize macrophages, were significantly higher than in control mice. Both CD11b⁺ F4/80⁺ cells MHCII^high CD11c^low (dominant macrophages) and MHCII^high CD11c^high (infiltrating macrophages), which infiltrate obese WAT^30,31, were significantly higher in aMKO mice than in control mice (Fig. 6A–E, Supplementary Fig. 5A). In support of the flow cytometry results, the expression of the gene encoding the macrophage marker F4/80 (Adgre1), and the macrophage-related cytokines Mcp1 and Tnfa, was significantly higher in aMKO mice than in control mice (Fig. 6F). These findings suggest that an inflammatory phenotype similar to that observed in obese WAT also occurs in the WAT of aMKO mice^30,31.

Systemic inflammatory and metabolic derangements in aMKO mice

To elucidate the systemic impact of adipose tissue inflammation in aMKO mice, a cytokine array was conducted at 27 weeks of age. The plasma levels of various cytokines were elevated, yet only one cytokine, matrix metalloproteinase-9, was lower in aMKO mice than in control mice (Supplementary Fig. 6A–C).

Spleen weight was significantly higher in aMKO mice than in control mice (Table 1). FACS analysis of splenocytes derived from aMKO mice demonstrated a significantly higher number of monocytes, granulocytes, and macrophages in aMKO mice than in control mice (Fig. 7A). Conversely, the number of T cells was significantly lower in aMKO mice than in control mice, while that of B cells remained unchanged (Fig. 7B).

Basal levels of plasma insulin were significantly higher in aMKO mice than in control mice and plasma glucose was slightly but significantly higher in aMKO mice than in control mice (Table 2). Glucose tolerance test (GTT) and insulin tolerance test (ITT) results revealed mild impairments in glucose and insulin sensitivity in aMKO mice (Fig. 8A, B). Plasma triglycerides were slightly elevated and mild fatty liver was observed in aMKO mice (Table 2; Fig. 8C, D). Although these metabolic derangements appeared not to be severe, they might account for the short lifespan of male mice (Supplementary Fig. 7).

Discussion

In the present study, adipocyte-specific MIPEP deficiency reduced WAT mass, without significant miniaturization of white adipocytes, and promoted the whitening of BAT in mice. Additionally, MIPEP deficiency enhanced local and systemic inflammation and resulted in systemic metabolic disorders, including impaired glucose and insulin sensitivity and fatty liver. The results indicate that our aMKO mice may be used as a common pathological model of lipoatrophy³². A recent publication described an adipocyte-specific Mipep KO mouse model produced by another research group. They searched public proteome databases to find mitochondrial proteases involved in insulin resistance in adipocytes: the research group identified MIPEP, the expression of which was decreased in multiple insulin resistance models, both in vivo and in vitro. The KO mice that they generated had a lower WAT mass with more small adipocytes, as well as a whitening of BAT, compared with control mice. These observations are almost identical to those of our aMKO mice, except for white adipocyte size. However, their mice displayed an unexpected improvement in an ITT at room temperature, which was no longer evident under thermoneutral conditions (around 30 °C for mice). Therefore, they postulated that the reduction in adiposity and enhancement of insulin sensitivity at room temperature may have been caused by increased thermogenesis, potentially due to elevated skeletal muscle function³³. As described above, both our aMKO mice and their KO mice exhibited similar WAT and BAT phenotypes, yet disparate whole-body metabolism phenotypes. This discrepancy may be attributable to differences in the knockout-targeting locus of the Mipep gene or the flox mouse-generating process.

Lipoatrophy in MIPEP-deficient mice

Genetically engineered lipoatrophic model mice with defects in adipocyte differentiation, adipocyte function, and lipid metabolism have been reviewed previously³⁴. Various molecular mechanisms underlying lipoatrophy have now been uncovered from both mouse and human data. For example, deficiency of peroxisome proliferator-activated receptor-g, a master regulator of adipocyte differentiation, provokes lipoatrophy^35,36,37, while reduced expression of SREBP-1c and PGC-1α contributes to the pathogenesis of antiretroviral-induced lipoatrophy in AIDS patients^38,39. Moreover, mitochondrial function is implicated in the pathogenesis of lipoatrophy³². Indeed, deficiency of mitochondria-related proteins, including caseinolytic protease proteolytic subunit (ClpP), CR6 interacting factor 1 (CRIF1), and mitochondrial transcription factor A (TFAM), cause lipoatrophy^40,41,42,43.

ClpP, which has protease activity, multimerizes with the mitochondrial chaperone and ATPase, ClpX, to form the functional protease ClpXP. Systemic Clpp knockout (ClpP-KO) mice have reduced adiposity and adipocyte size. Paradoxically, however, ClpP deficiency increases whole-body energy expenditure, improves glucose homeostasis, protects against fatty liver, and upregulates mitochondrial biogenesis selectively in WAT. The UPR^mt, including the upregulation of mitochondrial chaperones and proteases, is likely induced in ClpP-KO mice⁴⁰. CRIF1, also known as GADD45GIP1, is a mitochondrial large ribosomal subunit protein that plays a crucial role in the translation of subunits of the OXPHOS system and their insertion into the mitochondrial inner membrane. In adipocyte-specific Crif1 KO (adipo-CRIF-KO) mice, certain components of the OXPHOS system are decreased in WAT, possibly due to defective translation of OXPHOS subunits. Adipo-CRIF-KO mice have mild lipoatrophy, predominantly in epididymal WAT, and a whitening of BAT. In epididymal WAT, they have more small adipocytes; however, they exhibit no whole-body metabolic derangements, including impaired glucose metabolism and fatty liver. Additionally, the UPR^mt, including the upregulation of mitokines, mitochondrial chaperones, and proteases, is induced⁴¹. ClpP and CRIF1 are implicated in the maintenance of mitochondrial proteostasis, as well as in the function of MIPEP. We therefore hypothesized that the induced UPR^mt would manifest similarly in the WAT of systemic ClpP KO mice, adipo-CRIF-KO mice, and aMKO mice. The absence of both ClpP and CRIF1 resulted in the upregulation of mitochondrial chaperones and proteases; however, MIPEP deficiency upregulated the expression of mitokines, including GDF15 and FGF21, but not the expression of chaperones and proteases. These results indicate that MIPEP deficiency is unable to fully induce the previously reported UPR^mt, but is capable of partially activating it. Additionally, ClpP deficiency activated mitochondrial biogenesis, whereas both CRIF1 deficiency and MIPEP deficiency suppressed it. Further analysis is necessary to elucidate the contradictory results observed in ClpP, CRIF1, and MIPEP-deficient models. TFAM plays a role in maintaining the stability of mtDNA during its replication and the transcription of genes that are encoded by mtDNA⁴⁴. Two distinct lines of mice have been reported as adipocyte-specific Tfam KO mice: Tfam^flox/flox mice crossed with aP2-Cre mice and adiponectin-Cre (adipo-TFAM-KO) mice^42,43. Given the observation of off-target effects in adipocyte-specific conditional gene targeting using aP2-Cre mice, the phenotypes of aMKO mice and adipo-TFAM-KO mice were compared. In adipo-TFAM-KO mice, the observed phenotypes included lipoatrophy with normal-sized adipocytes, inflammation in WAT, and a whitening of BAT. The amount and enzymatic activity of specific components of the OXPHOS pathway were also decreased in both WAT and BAT. Moreover, mitochondrial dysfunction in both WAT and BAT resulted in insulin resistance and the development of fatty liver in adipo-TFAM-KO mice⁴³. The UPR^mt was not evaluated in adipo-TFAM-KO mice, but both their adipose tissue and systemic metabolic phenotypes are similar to those we observed in aMKO mice.

The disordered mitochondrial proteostasis and atypical UPR^mt caused by MIPEP deficiency in WAT and BAT

MDH2, COX4, and SIRT3 are substrates of MIPEP^8,16,17; however, to our knowledge, not all MIPEP substrates have been identified, particularly in mammals. Among the three known MIPEP substrates, we observed different effects of MIPEP deficiency in WAT and BAT. MIPEP deficiency resulted in significantly lower COX4 expression in both WAT and BAT, while MDH2 expression was higher only in WAT, and SIRT3 expression was not altered in either WAT or BAT. MIPEP substrate proteins are initially cleaved by MPP; MIPEP subsequently cleaves an octapeptide at the N-terminus to enable the substrate proteins to mature and gain their full physiological function^12,13,14,15. Therefore, the immature forms of MIPEP substrate proteins, as observed in aMKO mice, may be eight amino acids longer and display lower enzymatic activity than their physiologically mature forms. Western blot analysis revealed a shift of MDH2, a MIPEP substrate, to a higher molecular weight, particularly in the BAT of aMKO mice. Based on these observations, we hypothesize that the bands that shifted to a higher molecular weight in the Western blot image indicate an immature form of MDH2. Indeed, MIPEP deficiency reduced the enzymatic activity of three MIPEP substrates in both WAT and BAT. In the mitochondrial matrix, various proteases are localized to maintain tight control of proteostasis. These include three evolutionarily conserved proteases: lon protease homologue, ClpXP, and m-AAA^45,46. It could be that the immature forms of MIPEP substrate proteins are vulnerable or resistant to degradation by these proteases. In the BAT of aMKO mice, we observed an incongruity between the levels of UCP1 mRNA and protein (Fig. 4C and Supplementary Fig. 3). The decrease in UCP1 mRNA could be indicative of the differentiation abnormality of BAT. Furthermore, the findings suggest the possibility that UCP1 may be a substrate of MIPEP. However, to fully understand the proteostasis disorder caused by MIPEP deficiency, a proteomic analysis of the mitochondrial fraction is required, taking into account their degradation by mitochondrial matrix proteases.

Mitochondrial dysfunction activates the UPR^mt. As stated above, disordered proteostasis was observed in the WAT and BAT of aMKO mice, indicating that the UPR^mt was induced in both adipose tissue types. The UPR^mt induces the expression of mitokine, mitochondrial chaperone, and protease genes²¹. In mice where the UPR^mt was induced in adipose tissue, the expression of mitochondrial stress response genes, including those of mitokines, mitochondrial chaperones, and proteases, was upregulated^41,47,48,49. In both the WAT and BAT of aMKO mice, the gene expression of mitokines, specifically Gdf15 and Fgf21, was increased. However, the expression of mitochondrial chaperone and protease genes was not. These results indicate that the disruption of mitochondrial proteostasis by MIPEP deficiency did not induce a typical UPR^mt response, but rather an atypical response. The elevated expression of Xbp1s and phosphorylation of eIF2α are predominantly elicited by endoplasmic reticulum stress rather than the UPRmt²⁹. In aMKO mice, the expression of Xbp1s was upregulated and phosphorylation of eIF2a was enhanced markedly in BAT, but not in WAT. The stress response in BAT appeared to be more robust than that in WAT, although the ultra-microscopical defects of the mitochondrial structure were not any more severe in BAT than in WAT. As BAT exhibits a higher number of mitochondria per cell than WAT, the retrograde signaling from mitochondria may be more pronounced in BAT than in WAT. However, the molecular mechanisms underlying the atypical UPR^mt we observed and the differential response between WAT and BAT remain to be elucidated; therefore, further analyses are required. As mentioned above, MIPEP deficiency also led to an atypical UPR^mt and an upregulation in the expression of mitokines, including GDF15, but not mitochondrial chaperones and proteases. GDF15 binds to glial-derived neurotrophic factor receptor alpha-like, which is highly expressed in hindbrain neurons, and acts to improve whole-body metabolism^50,51. The precise mechanism by which GDF15 exerts a direct action on peripheral tissues remains to be elucidated, especially as its receptor is not expressed peripherally. However, recent report showed that recombinant GDF15 suppressed C/EBPα mRNA expression and adipose differentiation in human adipose-derived stem cells⁵². Consistent with these data, C/EBPα mRNA levels were decreased in the WAT of aMKO mice (Fig. 4A), the differentiation abnormality in WAT of aMKO mice may be partially due to GDF15. In contrast, GDF15 may induce insulin secretion in beta cells through an as-yet-unknown mechanism^53,54. Beneficial metabolic phenotypes, including polarization of macrophage from M1 to M2 in WAT, suppression of fatty liver, and improvement of ITT, result from the upregulation of GDF15 in adipo-CRIF-KO mice⁴¹. In aMKO mice, despite their severe adipose tissue phenotypes, the pathological changes at a young age were not as severe as expected. Therefore, elevated plasma GDF15 may increase insulin secretion and ameliorate the inflammation and systemic metabolic disturbances caused by adipose tissue dysfunction in aMKO mice.

Conclusion

The present study demonstrates that MIPEP deficiency results in disordered proteostasis in MIPEP substrate proteins due to defects in their maturation. MIPEP deficiency in both white and brown adipocytes resulted in the suppression of adipocyte differentiation, lipid metabolism, and mitochondrial biogenesis genes (Pparg, Srebf1, and Pgc1a/Nrf1, respectively). These changes may contribute to the pathogenesis of lipoatrophy in WAT and the whitening of BAT, resulting in adipose tissue inflammation and systemic inflammatory and metabolic disorders. However, GDF15 upregulation induced by an atypical UPR^mt might attenuate these systemic derangements.

Methods

Animals

All animal experiments and protocols were conducted in accordance with the Fundamental Guidelines for Proper Conduct of Animal Experiments and Related Activities in Academic Research Institutions under the jurisdiction of the Ministry of Education, Culture, Sports, Science and Technology of Japan. They were approved by the Ethics Review Committee for Animal Experimentation at Tokyo University of Science (approval numbers: Y20043, Y21043, Y22037). For generation of Mipep^flox/flox mice, DNA fragments of 2.8 kb (5’ arm), 3.0 kb (medium arm), and 3.1 kb (3’ arm), including exon12, exon13, and exon14, respectively, of the mouse Mipep genomic region were amplified by PCR using KOD FX Neo (Toyobo, Tokyo, Japan) and a bacterial artificial chromosome clone template (RP23-142O16; Advanced Geno Techs Co., Ibaraki, Japan). These fragments were subcloned into the DT-A/conditional KO FW plasmid to prepare the Mipep targeting vector containing loxP-flanked exon13, which encodes zinc-binding sites essential for the enzymatic activity of MIPEP (Supplementary Fig. 1). The linearized targeting vector was introduced into embryonic stem cells derived from C57BL/6 N mice by electroporation. Embryonic stem cell clones with the successfully targeted allele were selected and cocultured with eight-cell embryos derived from ICR mice (obtained from the CLEA Japan) overnight and differentiated into blastocysts. These blastocysts were transferred to pseudopregnant mice to bear chimera mice. The chimeras were crossed with C57BL/6J mice (obtained from the CLEA Japan), and the resulting progeny harbored the targeted allele. Mice with the targeted allele were crossed with CAG-FLPe mice (obtained from RIKEN Bioresource Center) to delete the FRT-flanked PGK-Neo cassette, resulting in Mipep^+/flox mice (Supplementary Fig. 1). Finally, Mipep^+/flox mice were intercrossed to generate homozygous mutant (Mipep^flox/flox) mice, which were subsequently crossed with Adipoq-Cre mice (obtained from the Jackson Laboratory) to prepare aMKO mice. Mipep^flox/flox mice littermates without Adipoq-Cre served as controls. Genotyping of offspring was performed by PCR using KOD FX neo (Toyobo, Osaka, Japan) according to the manufacturer’s protocol. Sequences of primers used for PCR were shown in the Table S1. Mice were maintained under specific-pathogen-free conditions at 23 °C, under a 12-h light/dark cycle in the animal facility at the Faculty of Pharmaceutical Sciences, Tokyo University of Science. They had free access to water and were fed a Charles River Formula-1 diet (21.9% crude protein, 5.4% crude fat, and 2.9% crude fiber; Oriental Yeast, Tokyo, Japan). At 22–27 weeks old, female mice were euthanized under isoflurane anesthesia (Wako, Osaka, Japan) and gWAT, sWAT, BAT, quadriceps femoris muscles, livers, hearts, kidneys, and plasma were collected. These tissues were immediately diced, frozen in liquid nitrogen, and stored at − 80 °C.

CT imaging

We validated a CT method using a third-generation CT scanner, LaTheta LCT-200 (Hitachi-Aloka, Tokyo, Japan). The tube voltage was set to 50 kV and the current was constant at 0.5 mA. Animals were scanned in a 48 mm-wide specimen holder with a resolution of 96 μm per pixel. For all scans, the same number of views (436) was used, which was the number of data points collected during a single 360° rotation around the object. In accordance with the manufacturer’s recommendations, the estimated radiation exposure of the mice was kept below 40 mSv. In pilot experiments, optimal scanning conditions were evaluated for each tissue; the final conditions were set at a − 550 to − 140 Hounsfield unit density range, 192 μm slice thickness, and a 600 μm slice pitch. Visceral and subcutaneous WAT were distinguished according to whether they were located inside (visceral) or outside (subcutaneous) of the body wall muscles such as the abdominal and thoracic wall muscles. The WAT mass was measured by Latheta software (v3.30) (Hitachi-Aloka).

Hematoxylin and Eosin staining and the size distribution of adipocytes

gWAT, BAT, and liver samples were fixed with 10% formalin solution for 24 h and embedded in paraffin. Section (5 μm thick) were cut from each tissue block, deparaffinized, dehydrated, and stained with hematoxylin and eosin to assess morphology. Images were acquired with an All-in-one fluorescence microscope (BZ-X800, KEYENCE, Osaka, Japan) using a 10 × objective lens. The size distribution of adipocytes from WAT was automatically calculated using a hybrid cell count application (v1.1.2.4) (BZ-H4C, KEYENCE, Osaka, Japan).

Scanning electron microscopy

Mice were anesthetized and perfused with buffered 4% paraformaldehyde (pH 7.4), and the peritesticular adipose tissues were removed and immersed in buffered 4% paraformaldehyde and 2.5% glutaraldehyde over a few nights at 4 °C. The tissues were cut into small pieces, and en bloc metal staining was performed as reported previously with some modifications⁵⁵. Briefly, tissues were washed with PBS, before being treated with 2% OsO4 in 1.5% K4[Fe(CN)6] for 1 h on ice. This was followed by filtered 1% thiocarbohydrazide for 20 min and 2% OsO4 for 30 min at room temperature. Tissues were then treated with uranyl acetate overnight at 4 °C, and lead aspartate solution at 50 °C for 2 h. Each of these treatments was followed by washing five times with double distilled water for 10–15 min. Tissues were dehydrated in a graded ethanol series (60%, 80%, 90%, and 95%, 5 min each); incubated sequentially with acetone dehydrated using a molecular sieve, a 1:1 mixture of resin and acetone, and 100% resin; and embedded in Durcupan (Sigma, MO, USA). The samples were placed in a mold with conductive resin containing 5% Ketjen black in Durcupan and cured at 60 °C for 5 nights. Cured resin blocks were trimmed and mounted on aluminum rivets with conductive glue (CR-2800, Kaken Tech, Osaka, Japan). The surfaces of the trimmed samples were treated with gold sputtering to increase conductivity and imaged in a field emission scanning electron microscope (Sigma from Carl Zeiss, Oberkochen, Germany) equipped with 3View, a system with a built-in ultramicrotome and a back-scattered electron detector (Gatan, CA, USA).

Immunoblotting

The preparation of cell lysates and immunoblotting were performed according to our previously reported methods⁵⁶. Briefly, cells were lysed with SDS sample buffer, boiled for 5 min, and sonicated. Lysates were subjected to SDS/PAGE (15 µg protein per well), and separated proteins were transferred onto nitrocellulose membranes. Membranes were blocked with blocking solution (2.5% skim milk, 0.25% bovine serum albumin in tris-buffered saline with Tween-20 [25 mM Tris-HCl pH 7.4, 140 mM NaCl, 2.5 mM KCl, 0.1% Tween-20]) for 1 h at room temperature and then probed with appropriate primary antibodies overnight at 4°C. The list of primary antibodies is shown in the Table S2. Subsequently, membranes were incubated with appropriate secondary antibodies for 1 h at room temperature (horseradish peroxidase-conjugated F(ab’)2 fragment of goat anti-mouse IgG or anti-rabbit IgG from Jackson Immuno Research [West Grove, PA, USA]). Antibody-bound proteins were visualized using ImmunoStar LD Reagent from Wako (Osaka, Japan) and the ChemiDoc™ Touch; data were analyzed using Image Lab software (v6.1) (Bio-Rad, CA, USA). Specific signal intensities were normalized to those of Coomassie Brilliant Blue (Wako, Osaka, Japan) or Ponceau S (Beacle, Inc., Kyoto, Japan) staining. The original raw blots are included in the source data files.

COX activity assay

WAT and BAT samples were homogenized in homogenization buffer containing 50 mM Tris-HCl (pH7.4), 150 mM NaCl, 5mM EDTA, 1% phosphatase inhibitor cocktail (Wako, Osaka, Japan), 1% protease inhibitor cocktail (Wako, Osaka, Japan), 1% Triton-X100 and 0.05% sodium deoxycholate, then centrifuged ay 12,000 × g for 10 min at 4 °C before the supernatant was collected. Protein concentration in each lysate was determined using a Pierce™ BCA Protein Assay Kit (Thermo Fisher Scientific, IL, USA), according to the manufacturer’s protocol. COX activity was measured by monitoring changes in the absorbance of reduced cytochrome c measured at 550 nm. Reduced cytochrome c is generated by adding sodium ascorbate to 10 mg/mL cytochrome c prior to ultrafiltration. Kinetic measurements at 550 nm started immediately after adding tissue extracts to reduced cytochrome c at a ratio of 1:4; measurements were taken every 10 s for 10 min using an Infinite M200 pro microplate reader (TECAN, Zurich, Switzerland). The concentration of reduced cytochrome c was 25 µM in WAT and 100 µM in BAT.

MDH2 activity assay

MDH2 activity was measured in WAT and BAT extracts using a commercially available assay according to the manufacturer’s instructions (MDH2 Activity Assay, Abcam, Cambridge, UK). One unit of MDH2 activity was expressed as the change in absorbance per minute per amount of sample per MDH2 unit (ΔmOD/min/MDH2 unit). The MDH2 unit indicates the relative quantity of MDH2 as determined by Western blotting.

Quantitative real-time PCR

Total RNA was extracted using ISOGEN II (Nippon gene, Toyama, Japan) and reverse transcription was performed using ReverTra Ace^® qPCR RT Master Mix (Toyobo, Osaka, Japan). Quantitative real-time PCR was performed using the CFX Connect™ Real-Time System (Bio-Rad, Hercules, CA, USA) and Thunderbird SYBR qPCR mix (Toyobo), according to manufacturer protocols. Sequences of primers used for PCR are shown in the Table S3. Rps18 was used as the housekeeping gene.

RNA extraction and RNA-seq

Total RNA was extracted from WAT and BAT from 30-week-old aMKO mice using the Relie Prep™ RNA Tissue Miniprep System (Promega, WI, USA) according to the manufacturer’s protocol. The concentration, purity, and integrity number of the RNA were measured using the Agilent 2100 Bioanalyzer (Agilent, CA, USA) by Novogene Co., Ltd. (Beijing, China). RNA samples with an integrity number greater than 7.3 were used for downstream analysis. Library preparation and RNA sequencing were done by Novogene Co., Ltd. (Beijing, China). The RNA-seq reads were aligned to the mouse reference genome GRCm39 was calculated using STAR (v2.711a) (https://github.com/alexdobin/STAR) two-pass alignment^57,58. Transcripts per million were calculated and gene expression levels were quantified using featureCounts from the Subread package (v2.0.6) (https://subread.sourceforge.net/)⁵⁹. Differentially expressed genes were calculated using the edgeR (v4.2.0) (https://bioconductor.org/packages/release/bioc/html/edgeR.html) program package²⁷.

Measurement of mitochondrial DNA

DNA was isolated and subjected to quantitative PCR using the primers shown in the Table S3. Quantitative real-time PCR was performed using the CFX Connect™ Real-Time System (Bio-Rad, Hercules, CA, USA) and Thunderbird SYBR qPCR mix (Toyobo), according to manufacturer protocols. Relative mitochondrial DNA copy number is represented by the amplification ratio of Nd1 or D-loop. β-Globin and Hprt1 were used as housekeeping genes in WAT and BAT, respectively.

Cell preparation and flow cytometry

Single-cell suspensions from the spleen were prepared as previously described^60,61. For stromal cell preparation from WAT, whole WAT was cut and minced in DMEM medium containing 0.2% type II collagenase using scissors (Worthington, Lakewood, NJ, USA) and incubated for 60 min at 37℃ with shaking every 10 min. Cell debris was removed by filtering ruptured tissues through a 70 μm nylon mesh. All samples were treated with ammonium-chloride-potassium buffer (0.15 M NH4Cl, 1.0 mM KHCO3, 0.1 mM Na2-EDTA [pH 7.3]), filtered through nylon mesh, and suspended in magnetic-activated cell sorting buffer (1% penicillin-streptomycin, 2% fetal bovine serum, 0.4% 0.5 M EDTA in PBS); all buffers used for staining were azide-free. Single-cell suspensions were counted and blocked with an anti-CD16/32 (2.4G2) antibody. Single-cell suspensions from the indicated organs were stained with the indicated antibodies; data were collected on FACS flow cytometers (BD Bioscience, NJ, USA) and analyzed using FlowJo software (v10.10.0) (TreeStar, Ashland, OR; https://www.flowjo.com/). Antibodies used for flow cytometry and sorting are listed in the Table S2.

GTTs and ITTs

GTTs and ITTs were performed in control and aMKO mice 21–24 weeks old. Prior to the GTT and ITT, mice were fasted for 24 h and 8 h, respectively. D-glucose (1.0 kg/body weight, Wako) and insulin (1.0 U/kg body weight, Wako) were injected intraperitoneally for the GTT and ITT, respectively. Next, serial blood sampling from the tail vein was performed at 0, 15, 30, 60, and 120 min after injection. Blood glucose levels were measured using an Accu-Chek^® Aviva blood glucose meter (Roche, Basel, Switzerland).

Measurement of triglyceride in the liver

Liver pieces were homogenized with Solution I (chloroform: methanol 1:1) and mixed with a quarter volume of 1 M NaCl. The homogenates were centrifuged for 10 min at 1100 × g at 4 °C, and their lower layers were collected and dried into lipid pellets. The pellets were dissolved in Solution II (tert-butanol: methanol: triton-114 3:1:1) as extracted fat samples. The amount of triglyceride in samples was measured using the LabAssay™ triglyceride kit (Wako, Osaka, Japan) according to the manufacturer’s protocol.

Plasma biochemical analysis

Plasma, non-esterified fatty acids, cholesterol, glucose, adiponectin and insulin levels were measured using the LabAssay™ triglyceride, LabAssay™ NEFA, LabAssay™ Cholesterol, Autokit Glucose, LBIS Mouse/Rat HMW Adiponectin ELISA, and LBIS Mouse Insulin ELISA (U-type) kits (Wako, Osaka, Japan). Plasma Leptin, GDF15, and FGF21 levels were measured using the Mouse/Rat Leptin Quantikine ELISA Kit, Mouse/Rat GDF15 Quantikine ELISA Kit, and Mouse/Rat FGF21 Quantikine ELISA Kit (R&D Systems, MN, USA), respectively. All assays were performed according to manufacturer protocols.

Cytokine array

Serum cytokines were measured in control and aMKO mice using the Proteome Profiler Mouse XL Cytokine Array (R&D Systems, MN, USA), according to the manufacturer’s protocol. Dot intensity was quantified using Image J software (v1.53k) from NIH (MD, USA; https://imagej.net/ij/index.html).

Statistical analyses

The significance of differences between the two groups was analyzed using the Student’s t-test. Statistical calculations were performed using GraphPad Prism version 10.1.2 (324) and version 8.4.1 (676) (GraphPad Software, USA; http://www.graphpad.com).

Data availability

Raw fastq files and unprocessed aligned data can be accessed free through the Gene Expression Omnibus (accession number GSE282147). In this study, Mipep^flox/flox mice were generated. The mice are available upon request.

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Acknowledgements

The authors are grateful to all members of the Laboratory of Molecular Pathology and Metabolic Disease (Faculty of Pharmaceutical Sciences, Tokyo University of Science) for useful discussions. We also thank members of the Animal Center (Tokyo University of Science) for their contributions to animal care, Mses. Atsuko Imai and Nobuko Hattori (National Institute for Physiological Sciences) and Ms. Megumi Yatabe (Jichi Medical University) for their technical assistance, and Advanced Bioimaging Support (ABiS) supported by Grant-in-Aid for Transformative Research Areas (No. 22H04926). We thank Philippa Gunn, D.Phil., from Edanz (https://jp.edanz.com/ac) for editing a draft of this manuscript.

Funding

This work was supported in part by Grants-in-Aid for Scientific Research (B) (No. 17H02179, 20H04130), Fostering Joint International Research (B) (No. 20KK0228), Grant-in-Aid for Young Scientists (B) (No. 17K13231), and Grant-in-Aid for Early-Career Scientists (No. 20K19686, 22K17814) from the Japan Society for the Promotion of Science, the Uehara Memorial Foundation (No. K19-170), and Academic Research Grants of Natural Sciences Division from the 61th Mishima Kaiun Memorial Foundation. The funders had no role in study design, data collection and interpretation, or the decision to submit the work for publication.

Author information

Yuka Nozaki and Masaki Kobayashi contributed equally to this work.

Authors and Affiliations

Laboratory of Molecular Pathology and Metabolic Disease, Faculty of Pharmaceutical Sciences, Tokyo University of Science, Chiba, Japan
Yuka Nozaki, Masaki Kobayashi, Tomoyoshi Fukuoh, Mamiko Ishimatsu, Takumi Narita, Kanari Taki, Yuto Hirao, Shota Ayabe, Miku Yokoyama, Yuina Otani, Yuhei Mizunoe & Yoshikazu Higami
Department of Nutrition and Food Science, Graduate School of Humanities and Sciences, Ochanomizu University, Tokyo, Japan
Masaki Kobayashi
Institute for Human Life Science, Ochanomizu University, Tokyo, Japan
Masaki Kobayashi
Section of Electron Microscopy, Supportive Center for Brain Research, National Institute for Physiological Sciences, Okazaki, Japan
Mami Matsumoto
Department of Developmental and Regenerative Neurobiology, Institute of Brain Science, Nagoya City University Graduate School of Medical Sciences, Nagoya, 467-8601, Japan
Mami Matsumoto
Department of Anatomy, Division of Histology and Cell Biology, School of Medicine, Jichi Medical University, Shimotsuke, Japan
Nobuhiko Ohno
Division of Ultrastructural Research, National Institute for Physiological Sciences, Okazaki, Japan
Nobuhiko Ohno
Division of Immunology, Faculty of Medicine, Tohoku Medical and Pharmaceutical University, Sendai, Japan
Tomonori Kaifu
Research Institute for Biomedical Sciences (RIBS), Tokyo University of Science, Noda, Japan
Shogo Okazaki, Ryo Goitsuka, Yoichiro Iwakura & Yoshikazu Higami
Division of Complex Biosystem Research, Department of Research and Development, Institute of Natural Medicine, University of Toyama, Toyama, Japan
Yoshimi Nakagawa
Department of Endocrinology and Metabolism, Institute of Medicine, University of Tsukuba, Ibaraki, Japan
Hitoshi Shimano

Authors

Yuka Nozaki
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Masaki Kobayashi
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Tomoyoshi Fukuoh
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Mamiko Ishimatsu
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Takumi Narita
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Kanari Taki
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Yuto Hirao
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Shota Ayabe
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Miku Yokoyama
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Yuina Otani
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Yuhei Mizunoe
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Mami Matsumoto
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Nobuhiko Ohno
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Tomonori Kaifu
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Shogo Okazaki
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Ryo Goitsuka
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Yoshimi Nakagawa
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Hitoshi Shimano
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Yoichiro Iwakura
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Yoshikazu Higami
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Contributions

Y.H. supervised the project. Y.N., M.K. and Y.H. developed the scientific concept of the project and wrote main manuscript text. T.N., T.K., S.O., R.G., Y.N., H.S. and Y.I. contributed to creation of aMKO mice. Y.N., M.K., T.F., M.I., K.T., Y.H., S.A., M.Y., Y.O., Y.M., M.M. and N.O. performed the experiments and data analysis. T.F. prepared figure 1 A-D, F and Figs. 2, 3, 4, 5, 6 and 7. M.M. and N.O. prepared Fig. 1E and G. All authors reviewed the manuscript and agreed to its submission.

Corresponding author

Correspondence to Yoshikazu Higami.

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Ethics

The experimental procedures and reporting of this study were conducted in accordance with the ARRIVE guidelines. All animal experiments were conducted in accordance with ethical regulations for animal research and were approved by the Ethics Review Committee for Animal Experimentation at Tokyo University of Science (Y20043, Y21043, Y22037).

Competing interests

The authors declare no competing interests.

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Nozaki, Y., Kobayashi, M., Fukuoh, T. et al. Mipep deficiency in adipocytes impairs mitochondrial protein maturation and leads to systemic inflammation and metabolic dysfunctions. Sci Rep 15, 12839 (2025). https://doi.org/10.1038/s41598-025-97307-6

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Received: 19 December 2024
Accepted: 03 April 2025
Published: 14 April 2025
DOI: https://doi.org/10.1038/s41598-025-97307-6