KSRP Ablation Enhances Brown Fat Gene Program in White Adipose Tissue Through Reduced miR-150 Expression

Obesity, caused by positive energy balance leading to expansion of adipocyte mass, increases the risk of diabetes, heart disease, and some forms of cancer. There are two major types of adipose tissues in mammals: white adipose tissue (WAT) and brown adipose tissue (BAT). WAT is the main storage site of excess energy, primarily in the form of triacylglycerol (TG). Conversely, BAT is specialized to dissipate chemical energy for heat generation primarily through uncoupling protein 1 (UCP1). UCP1 uncouples mitochondrial electron transport from ATP synthesis by permeabilizing the inner mitochondrial membrane to allow intermembrane protons to leak back into the mitochondrial matrix (1). The activity of BAT inversely correlates with BMI in adult humans (2,3). Promoting BAT function has a potential to defend against obesity and obesity-associated disorders (4). A distinct type of UCP1-positive adipocytes, designated as beige or brite (brown-in-white) cells, is found sporadically in WAT of adult animals upon exposure to long-term cold or β-adrenergic agonists (5–7). This brown-like transformation of WAT is most prominent in the inguinal subcutaneous depot, whereas epididymal/perigonadal adipose tissue is less susceptible to browning (8), the formation of multilocular UCP1-positive adipocytes. The emergence of these inducible brown-like adipocytes is associated with a protection against obesity and metabolic diseases in rodent models (8–11).

Classic brown adipocytes develop from Myf5-positive precursors through the action of transcriptional regulators PR domain containing 16 (PRDM16) and CCAAT/enhancer binding protein-β (12,13). However, the inducible brown-like adipocytes arise from a non-Myf5 cell lineage. The browning of WAT in rodents can be induced by hormones such as irisin (14) and FGF12 (15), pharmacological activation of peroxisome proliferator–activated receptor (PPAR)-γ (16), and modulation through various transcriptional regulators including PRDM16 (8), PPARGC1α/PPARγ coactivator 1α (17), forkhead box class C2 (9), receptor interacting protein 140 (10,18), transcription intermediary factor 2 (19), pRb, and p107 (20,21). Recently, several microRNAs (miRNAs) were shown to regulate brown fat differentiation and brown-like transformation of WAT. miRNA (miR)-133 inhibits brown adipocyte differentiation by directly repressing Prdm16 expression (22), and also regulates the choice between myogenic and brown adipose determination using multipotent adult skeletal muscle stem cells (satellite cells) that can give rise to both myogenic and brown adipogenic lineages (23). miR-196a is essential for brown adipogenesis of white fat progenitor cells by repressing the expression of Hoxc8 (24). miR-193b and miR-365 are essential for brown fat differentiation by repressing myogenesis as well as by promoting brown adipogenesis (25). miR-155 inhibits brown and beige adipocyte differentiation by repressing CCAAT/enhancer binding protein-β (26).

KH-type splicing regulatory protein (KSRP) is a multifunctional RNA-binding protein involved in post-transcriptional regulation of gene expression. This includes splicing (27), mRNA decay (28), primary miRNA (pri-miRNA) processing (29), and translation (30). KSRP binds the AU-rich elements in the 3′ untranslated regions (UTRs) of inherently unstable mRNAs and promotes their decay by recruiting mRNA decay machineries (28,31). KSRP also interacts with the terminal loops of a subset of miRNA precursors and promotes their maturation in cultured cells (29). However, the in vivo function of KSRP in controlling mRNA decay and pri-miRNA processing, and the associated phenotypes resulting from KSRP deficiency have not been completely established. To do this, we have generated Ksrp-null mice (32). In the current study, we report that Ksrp^−/− mice exhibit decreased fat mass owing to a reduction in TG content. The expression of brown fat–selective and fatty acid oxidation genes is increased in WAT in the absence of KSRP. We also find that high-fat feeding upregulates miR-150 and KSRP in inguinal WAT (iWAT) and the deletion of KSRP significantly decreases adiposity in diet-induced obesity (DIO). Thus, these findings establish KSRP and miR-150 as important regulators for brown-like transformation of iWAT and whole-body adiposity.

The generation of Ksrp-null mice on a C57BL/6J congenic background has been described (32). Mice were maintained under a 12-h light/dark cycle at a room temperature of 22°C and were fed a normal chow diet (NCD) or a high-fat diet (HFD) containing 45% kcal from fat (Harlan Laboratories). All experiments were performed using 10- to 16-week-old, age-matched, wild-type and Ksrp^−/− male mice, which were littermate offspring of Ksrp^+/− × Ksrp^+/−. For metabolic analysis, including food intake, activity, and indirect calorimetry, mice were individually housed. Food intake and physical activity were measured using the Comprehensive Lab Animal Monitoring System. VO₂ and VCO₂ were measured by indirect calorimetry. Energy expenditure (EE) was calculated using the equation of EE = (3.941 × VO₂) + (1.106 × VCO₂) and normalized with respect to lean mass. Body fat content and lean mass were measured by dual-energy X-ray absorptiometry. All animal studies were conducted in accordance with guidelines for animal use and care established by the University of Alabama at Birmingham Animal Resource Program and the Institutional Animal Care and Use Committee.

Adipocytes were isolated from epididymal and inguinal fat depots by collagenase digestion as described previously (33). The floating adipocytes were collected and washed with Krebs-Ringer HEPES buffer. Adipocyte number and size were determined as described previously (34). Briefly, aliquots of evenly suspended isolated adipocytes were removed for optical sizing of cell diameter with ImageJ (National Institutes of Health) using a micrometer ruler as a reference for the diameter and for determination of the volume occupied by packed adipocytes using microhematocrit capillary tubes. Adipocyte number was derived by dividing the volume occupied by packed adipocytes with the mean adipocyte volume obtained by optical sizing of the diameter. Lipids were extracted from adipocytes in 1% Triton X-100, and TG content was determined using Infinity reagent (Thermo Scientific).

Total RNA was extracted by TRIzol (Invitrogen). For quantitative real-time RT-PCR analysis, total RNA (1 μg) was reverse transcribed using a mixture of oligo(dT) and random hexamers. Amplification was performed by using an LightCycler 480 and the SYBR Green system (Roche). mRNA levels were normalized to that of β-actin or cyclophilin B mRNAs. The primer sequences are listed in Supplementary Table 1.

RNA was isolated from iWATs of six wild-type and six Ksrp^−/− mice by miRNeasy (Qiagen). Individual RNA of wild-type mice and individual RNA of Ksrp^−/− mice were pooled together, respectively, and subjected to genome-wide miRNA microarray analysis in triplicate, which was performed by Phalanx Biotech. The miRNAs whose signals were fivefold above the background signal (these miRNAs are arbitrarily considered to be expressed in the iWAT) and were elevated by more than twofold in the Ksrp^−/− iWAT were selected for further analysis. For miRNA expression analysis, total RNA was converted into cDNA by miScript II RT Kit (Qiagen) and was subjected to real-time PCR with a specific primer for miR-150 using miScript Primer Assay (Qiagen). miRNA levels were normalized to that of U6 small nuclear RNA.

miRNA target prediction was performed by using the miRWalk database (http://www.umm.uni-heidelberg.de/apps/zmf/mirwalk/).

pri-miRNA processing assays were performed as described previously (29,35). Briefly, a DNA template producing pri-miR-150 was generated by PCR. ³²P-labeled pri-miR-150 was synthesized by in vitro transcription and incubated with total extracts of wild-type and Ksrp^−/− iWATs in a processing buffer containing 100 mmol/L KCH₃COOH, 2 mmol/L Mg(CH₃COOH)₂, 10 mmol/L Tris-Cl (pH 7.6), 2 mmol/L dithiothreitol, 10 mmol/L creatine phosphate, 1 μg of creatine phosphokinase, 1 mmol/L ATP, 0.4 mmol/L guanosine triphosphate, 0.1 mmol/L spermine, and 2 units of RNasin at 37°C. RNA was isolated and subjected to 10% polyacrylamide-urea gel electrophoresis and autoradiography.

Ribonucleoprotein immunoprecipitation assays were performed as described previously (35,36). Briefly, cell lysates were immunoprecipitated with Dynabeads (Invitrogen) coated with protein A/protein G and coupled with anti-KSRP serum at 4°C overnight. Pellets were washed four times, and RNA was isolated from the immunocomplexes using miRNeasy, reverse transcribed, and amplified by quantitative PCR (qPCR).

Cells or tissues were lysed in radioimmunoprecipitation assay buffer (0.5% NP-40, 0.5% sodium deoxycholate, 0.1% SDS, 150 mmol/L NaCl, 50 mmol/L Tris-Cl, pH 7.5). Proteins were separated using 7–10% SDS-PAGE, transferred to a polyvinylidene fluoride membrane, and probed with the following antibodies: anti-PRDM16 (R&D Systems), anti-PPARGC1α (Abcam), anti-PPARα (Santa Cruz Biotechnology), anti-KSRP (37), anti-UCP1 (Abcam), anti-β-actin (Abcam), and anti-α-tubulin (Sigma-Aldrich).

Paraffin-embedded sections were incubated with anti-UCP1 antibodies (1:500; Abcam), followed by detection using the Vectastain ABC Kit (Vector Laboratories).

Stromal-vascular fraction (SVF) cells were prepared from iWAT as described previously (33), cultured in DMEM/F12 containing 10% FBS, and induced for differentiation by incubating confluent cells with DMEM/F12 medium containing 10% FBS, 5 mg/mL insulin, 0.5 mmol/L isobutylmethylxanthine, 1 μmol/L dexamethasone, 1 nmol/L triiodothyronine (T3), 125 nmol/L indomethacin, and 1 μmol/L rosiglitazone for 48 h. Cells were maintained in maintenance medium containing 10% FBS, 1 nmol/L T3, and 1 μmol/L rosiglitazone. The medium was replaced every 2 days. For transfection of SVF cells, 80–90% of confluent cells were transfected with miRNA mimics (50 nmol/L), miRNA inhibitors (100 nmol/L), or small interfering RNAs (siRNAs) (60 nmol/L) using HiPerFect (Qiagen). The transfected cells were induced for differentiation the following day and collected for analysis 5 days postinduction.

SVF cells were seeded (15,000 cells/well) in XF24 culture microplates (Seahorse Bioscience) and induced for differentiation. At day 5 of differentiation, the oxygen consumption rate (OCR) was measured using an XF24 analyzer. Basal respiration was assessed in untreated cells, and uncoupled respiration was assessed after the addition of 10 μmol/L oligomycin. Nonmitochondrial respiration was measured after the addition of 4 μmol/L antimycin A and 1 μmol/L rotenone.

Dual luciferase reporters, expressing firefly luciferase containing the 3′ UTRs of Prdm16 and Ppargc1a, and Renilla luciferase, were purchased from GeneCopoeia. A control reporter, pEZX-MT01, was also purchased from GeneCopoeia. The predicted miR-150 target sites were mutated by PCR-mediated site-directed mutagenesis, and mutations were confirmed by DNA sequencing. NIH3T3 cells were transfected with 1 μg of a reporter together with an miRNA mimic (50 nmol/L) using Lipofectamine (Invitrogen). Cells were collected 36–48 h after transfection, and luciferase activities were measured using the Luc-Pair miR Luciferase Assay Kit (GeneCopoeia). Firefly luciferase activity was normalized to internal control Renilla luciferase activity.

miR-150 mimic and inhibitor were purchased from Qiagen. siGENOME SMARTpool siRNAs against Prdm16 and Ppargc1a were purchased from Thermo Scientific. Control miRNA mimic and inhibitor and a control siRNA were purchased from Qiagen.

All data are presented as the mean ± SEM. Comparisons between two groups were performed using an unpaired two-tailed Student t test. One-way ANOVA, followed by Fisher least significant difference test, was used for multiple comparisons.

Ksrp^−/− mice were lean with a 10% reduction in body weight in males eating a NCD compared with wild-type littermates (Fig. 1A) resulting from a reduction in the mass of distinct fat pads (Fig. 1B). A similar reduction in body weight and fat mass was also observed in female Ksrp^−/− mice (Supplementary Fig. 1A and B). By contrast, no significant difference in the weights of organs, such as liver, spleen, kidney, and heart, was detected between the two groups (Supplementary Fig. 1C). Consistent with these findings, body composition analysis showed a reduction in whole-body fat mass, but not in lean mass, in Ksrp^−/− mice (Supplementary Fig. 1D and E).

Reduction in adipose tissue mass can result from less TG storage, impaired adipocyte differentiation, or both. We isolated primary adipocytes from epididymal WAT (eWAT) and iWAT, and determined the adipocyte number and total TG content. While no difference in adipocyte number was observed (Fig. 1C), TG content was significantly reduced in Ksrp^−/− eWAT and iWAT (Fig. 1D). Consistent with a decrease in TG content, adipocytes of Ksrp^−/− eWAT and iWAT were smaller in size, as revealed by histological analysis (Fig. 1E), and there was a 25% decrease in the mean diameter of adipocytes in both the eWAT and iWAT of Ksrp^−/− mice (Fig. 1F).

To further examine the cause of reduced fat mass, we subjected wild-type and Ksrp^−/− mice to metabolic studies. No significant difference in food consumption and locomotor activity was observed between groups during the light and dark periods (Supplementary Fig. 1F and G). However, Ksrp^−/− mice exhibited a moderate increase in EE as measured by indirect calorimetry during the dark period (Supplementary Fig. 1H). Collectively, these data suggest that the reduced adiposity in Ksrp^−/− mice results from a reduction in TG content in WAT partly due to increased EE, but not reduced food consumption or increased activity.

To determine the molecular mechanism leading to reduced adiposity in Ksrp^−/− mice, we examined the expression of genes involved in adipocyte differentiation and lipid metabolism in WAT. The expression of Pparg, but not of Cebpa, was moderately elevated in Ksrp^−/− eWAT, and the expression of both Cebpa and Pparg was increased in Ksrp^−/− iWAT (Supplementary Fig. 2A). There was no difference in the expression of adipocyte makers such as fatty acid binding protein 4 (Fabp4) and perilipin 1 in either eWAT or iWAT between the two groups (Supplementary Fig. 2A). We examined the expression of genes involved in the pathways of TG synthesis (Gpam, Agpat2, Agpat6, and Dgat2), fatty acid synthesis (Fasn, Acaca/Acc1, and Scd1), and fatty acid uptake (Lpl, Cd36, and Fatp1). While no significant difference in their expression was observed in Ksrp^−/− eWAT, the expression of these genes, except for Dgat2, Lpl, Cd36, and Fatp1, was significantly increased in Ksrp^−/− iWAT (Supplementary Fig. 2B–D). These results suggest that KSRP ablation increases the expression of Cebpa, Pparg, and most of the genes involved in TG synthesis and de novo fatty acid synthesis in iWAT. However, the elevated expression of these genes is not consistent with a reduction in TG content in the WAT of Ksrp^−/− mice.

We also observed that genes selectively expressed in BAT, including Ucp1, Cidea, Cox8b, and Dio2, were significantly upregulated in the iWAT of Ksrp^−/− mice (Fig. 2A). The fold increase in brown fat–selective genes was much higher than that of TG synthesis and fatty acid synthesis genes (compare Fig. 2A with Supplementary Fig. 2). The expression of some brown fat genes was also moderately increased in Ksrp^−/− eWAT, although their expression was much less than that in iWAT (Fig. 2A). In addition to brown fat markers, the expression of genes encoding brown fat transcriptional regulators, including Ppargc1a and Ppara, but not Prdm16, was also elevated in the iWAT and eWAT of Ksrp^−/− mice (Fig. 2B). By contrast, no difference in the expression of these genes was observed between wild-type BAT and Ksrp^−/− BAT (Supplementary Fig. 3A and B). The lack of an alteration in brown fat gene expression in Ksrp^−/− BAT is likely due to a lower KSRP expression in BAT compared with iWAT (Supplementary Fig. 3C). Protein levels of PRDM16 (while Prdm16 mRNA was not elevated), PPARGC1α, and PPARα were significantly increased in Ksrp^−/− iWAT (Fig. 2C). Consistent with the elevated expression of PPARGC1α and PPARα, important regulators of fatty acid oxidation, the expression of Cpt1b, Acadm, and Acadl was also significantly increased in Ksrp^−/− iWAT and eWAT (Fig. 2D). We also detected elevated levels of UCP1 protein and increased numbers of UCP1-positive adipocytes in Ksrp^−/− iWAT, which contained clusters of multilocular cells (Supplementary Fig. 4A and B). Thus, the deletion of KSRP leads to brown-like transformation of iWAT that is characterized by the enhanced expression of brown fat–selective and fatty acid oxidation genes.

KSRP regulates the maturation of some miRNAs by facilitating pri-miRNA processing. To determine whether the impaired expression of miRNAs plays a role in enhancing brown fat gene expression in Ksrp^−/− iWAT, we subjected RNA samples to miRNA microarray analysis and only selected miRNAs whose expression was reduced in the absence of KSRP. We identified only one miRNA, miR-150, whose expression was reduced by more than twofold in Ksrp^−/− iWAT (data not shown; see Research Design and Methods). The expression of miR-150 was higher in the iWAT than in the eWAT and BAT of wild-type mice, and there was a threefold reduction in Ksrp^−/− iWAT (Fig. 3A). Conversely, pri-miR-150 levels in Ksrp^−/− iWAT were higher than that in wild-type iWAT (Fig. 3B), suggesting that the reduction in miR-150 levels was likely due to impaired pri-mR-150 processing. To determine the role of KSRP in pri-miR-150 processing, we performed ribonucleoprotein immunoprecipitation assays and observed that KSRP physically associated with pri-miR-150 in wild-type iWAT extracts (Fig. 3C). In contrast, pri-miR-34a was not associated with KSRP (Fig. 3C). The residual signals of pri-miR-34a in the anti-KSRP immunoprecipitates were likely due to a nonspecific interaction of anti-KSRP serum with pri-miR-34a. We carried out in vitro pri-miRNA processing assays and found that production of pre-miR-150 was significantly reduced using the cell lysate of Ksrp^−/− iWAT (Fig. 3D). The addition of recombinant KSRP to Ksrp^−/− lysate restored pre-miR-150 production (Fig. 3E). By contrast, processing of a control pri-miR-23b was equivalent between wild-type and Ksrp^−/− lysates (Fig. 3F). These data indicate that decreased miR-150 expression in Ksrp^−/− iWAT is indeed due to a reduction in pri-miR-150 processing.

To determine whether the increased brown-like transformation is cell autonomous, we isolated SVF cells of iWAT that are able to express brown fat–selective genes upon differentiation to adipocytes (8). After differentiation, the expression of Ucp1, Cidea, Cox8b, Ppargc1a, and Ppara, but not of Prdm16, was elevated in Ksrp^−/− SVF cells (Fig. 4A and B). By contrast, no difference in the expression of adipocyte markers common to white and brown adipocytes, such as Fabp4 and Adipoq, and a brown adipocyte–selective marker, Elovl3, was detected between wild-type and Ksrp^−/− SVF cells (Fig. 4C). The expression of PRDM16, PPARGC1α, and PPARα was elevated in differentiated Ksrp^−/− SVF cells (Fig. 4D). We examined miR-150 expression and observed a fivefold increase upon differentiation of wild-type SVF cells; we also observed that this increase was completely blunted in Ksrp^−/− SVF cells (Fig. 4E). Furthermore, pri-miR-150 levels were decreased in differentiated wild-type SVF cells, but those levels were not altered in differentiated Ksrp^−/− SVF cells compared with undifferentiated cells (Fig. 4E). These data indicate that the induction of miR-150 upon adipocyte differentiation occurs primarily through pri-miR-150 processing due to an increase in KSRP levels (Fig. 4D), and that the elevated expression of brown fat–selective genes and decreased miR-150 expression in the absence of KSRP can be reproduced in adipocytes derived from SVF cells of iWAT.

Target prediction analysis indicated that Prdm16 and Ppargc1a are targets of miR-150. To determine the molecular mechanism leading to enhanced expression of brown fat genes and whether it is due to a reduction in miR-150 expression, we manipulated miR-150 levels in SVF cells of iWAT. The inhibition of miR-150 expression in wild-type SVF cells increased mRNA levels of Ucp1, Cidea, Cox8b, Ppargc1a, and Ppara, but not of Prdm16 (Fig. 5A and B). Conversely, ectopic miR-150 expression in Ksrp^−/− SVF cells markedly repressed the expression of Ucp1, Cidea, Cox8b, Ppargc1a, and Ppara, but not of Prdm16 (Fig. 5A and B). By contrast, neither reduction nor overexpression of miR-150 changed the expression of Fabp4 and Adipoq (Fig. 5C). The inhibition of miR-150 expression in wild-type SVF cells moderately enhanced the expression of PRDM16 and PPARGC1α, and, conversely, the overexpression of miR-150 in Ksrp^−/− SVF cells caused a marked decrease in the protein levels (Fig. 5D).

To correlate the brown fat gene program with mitochondrial functions, we examined mitochondrial aspiration in differentiated SVF cells. Ksrp^−/− adipocytes exhibited higher OCRs for basal and uncoupled (treated with oligomycin) aspirations, but not for nonmitochondrial aspiration (treated with antimycin A and rotenone), than wild-type adipocytes (Fig. 5E, left panel). Importantly, the treatment of Ksrp^−/− adipocytes with a miR-150 mimic decreased basal and uncoupled aspirations (Fig. 5E, right panel). These data indicate that KSRP deficiency increases mitochondrial respiration activity because of reduced miR-150 expression.

To demonstrate that miR-150 can directly target Prdm16 and Ppargc1a mRNAs, we cloned their 3′ UTRs into a dual luciferase reporter. We subcloned two overlapping fragments from each 3′ UTR owing to their large size (>3.5 kb). Fragment (F) 2 of Prdm16 contains a predicted miR-150 site, and both fragments (F1 and F2) of Ppargc1a contain one predicted site (Fig. 6A). The expression of miR-150 decreased the expression of luciferase reporters harboring F2, but not F1, of Prdm16, and F2, but not F1, of Ppargc1a compared with a control miRNA (Fig. 6B). By contrast, miR-150 expression did not alter the expression of a control reporter (MT01) without an insertion (Fig. 6B). Mutations of the miR-150 target sites in F2 of Prdm16 and F2 of Ppargc1a resisted inhibition by miR-150 (Fig. 6B). These data indicate that miR-150 directly regulates the expression of PRDM16 and PPARGC1α, leading to changes in the expression of their downstream target genes, such as Ucp1, Cidea, Cox8b, and Ppara, and suggest that the absence of KSRP leads to derepression of Prdm16 and Ppargc1a through a decrease in miR-150 expression.

To determine whether the increase in brown fat–selective gene expression in the absence of KSRP is due to increased expression of PRDM16 and PPARGC1α, we downregulated their expression by siRNAs in SVF cells. The downregulation of PRDM16 in wild-type and Ksrp^−/− SVF cells decreased the expression of Ucp1, Cidea, Cox8b, Ppargc1a, and Ppara. PPARGC1α knockdown decreased the expression of Ucp1, Cidea, Cox8b, and Ppara, but not of Prdm16, in wild-type and Ksrp^−/− SVF cells (Supplementary Fig. 5A and B). By contrast, the downregulation of PRDM16 or PPARGC1α did not alter the expression of Fabp4, Adipoq, and Elovl3 (Supplementary Fig. 5C), indicating that adipocyte differentiation was not affected under these conditions. These data strongly indicate that the increase in the brown fat gene program in the absence of KSRP is indeed due to increased expression of PRDM16 and PPARGC1α.

We examined the expression of miR-150 and Ksrp in a model of DIO, and found elevated expression of miR-150 and Ksrp in the iWAT of wild-type mice on an HFD and reduced miR-150 levels in HFD-fed Ksrp^−/− iWAT compared with HFD-fed wild-type iWAT (Fig. 7A). HFD feeding also increased mRNA levels of Prdm16, Ppargc1a, Fabp4, Ucp1, Cox8b, Cidea, Dio2, Cpt1b, and Acadl in wild-type iWAT (Fig. 7A–D), and levels of Ucp1, Cox8b, Cidea, Cpt1b, and Acadl were further upregulated in HFD-fed Ksrp^−/− mice (Fig. 7B–D). We also detected an increase in KSRP and PPARGC1α, but not of PRDM16, in wild-type mice fed an HFD compared with a NCD, and a further upregulation of PPARGC1α and PRDM16 in HFD-fed Ksrp^−/− mice (Fig. 7E). Since the mRNA levels of Prdm16 and Ppargc1a were not elevated in HFD-fed Ksrp^−/− mice compared with HFD-fed wild-type mice, we suggest that the increase in protein levels was likely due to increased translation through reduced miR-150 levels. More importantly, body weight and fad pad weights were reduced in HFD-fed Ksrp^−/− mice (Fig. 7F and G). These data suggest a model in which elevated expression of both KSRP and miR-150 is likely permissive for the development of obesity, and in which the absence of KSRP promotes a reduction in adiposity in DIO by enhancing brown-like transformation of iWAT through the increased expression of PPARGC1α and PRDM16.

This study shows that targeted deletion of KSRP enhances brown fat–selective gene expression in iWAT through the elevated expression of PRDM16, PPARGC1α, and PPARα, which are important regulators for the thermogenic program in BAT and for brown-like remodeling in WAT. We demonstrated that KSRP is involved in the processing of pri-miR-150 in iWAT and its absence results in downregulation of miR-150. Mechanistically, miR-150 directly represses Prdm16 and Ppargc1a expression. Thus, reduction in miR-150 levels in Ksrp^−/− iWAT leads to elevated expression of PRDM16 and PPARGC1α, thereby increasing the brown fat gene program. In addition, we also found increased expression of fatty acid oxidation genes in Ksrp^−/− iWAT, likely due to the upregulation of PPARGC1α and PPARα, which are critical regulators for enhancing fatty acid use (38–40). The expression of brown fat–selective and fatty acid oxidation genes in the eWAT of Ksrp^−/− mice was also increased. By contrast, KSRP ablation did not alter the brown fat gene program in BAT, likely due to a lower expression of KSRP and miR-150 in BAT. These findings suggest that KSRP is a critical factor for balancing energy storage and EE in white adipocytes, and its absence favors EE, partly through post-transcriptional regulation of miR-150 expression.

Targeted deletion of miR-150 leads to B-cell expansion and an enhanced humoral immune response due to c-Myb upregulation (41). The body weight and fat mass of Mir150^−/− mice compared with wild-type mice were not reported. Consistent with the observation with HFD feeding (Fig. 7A), miR-150 was also reported to be upregulated in subcutaneous adipose tissue of obese human subjects (42). While Ppargc1a was predicted to be a target of miR-150, its levels were not altered in obese adipose tissue (42). Using adipocytes derived from SVF cells of iWAT, we demonstrate that ectopic expression of miR-150 attenuates Prdm16, Ppargc1a, and Ppara expression, and, conversely, that the inhibition of miR-150 expression increases their expression. Although Prdm16 and Ppargc1a are directly targeted by miR-150, Ppara does not contain any predicted target site. The regulation of Ppara expression by miR-150 is likely indirect due to the altered expression of Prdm16 and Ppargc1a (see below). Our results point to miR-150 as a negative regulator of inducible brown-like adipocytes in iWAT by repressing Prdm16 and Ppargc1a.

PRDM16 is a determinant of brown fat lineage and is able to induce browning of subcutaneous WAT (8,12,13). PPARGC1α is recognized as a critical regulator of thermogenesis and oxidative metabolism (43). PPARα is also critical for the expression of brown fat and fatty acid oxidation genes in BAT (44). We found that PRDM16 knockdown in SVF cells attenuated the expression of Ppargc1a, Ppara, and brown fat markers, and that PPARGC1α knockdown decreased the expression of Ppara, but not of Prdm16, and brown fat markers (Supplementary Fig. 5). These results strongly suggest that the enhanced brown-like transformation in Ksrp^−/− iWAT at least in part results from the elevated expression of Prdm16, Ppargc1a, and Ppara. These data are also consistent with previous studies showing that PRDM16 lines upstream of PPARGC1α and PPARα (8,25,45), and that PRDM16 and PPARGC1α are positive regulators for brown fat gene expression (8,17). While it was shown that PPARα activates Ppargc1a expression through coactivation by PRDM16 (46), our data suggest that PPARGC1α also regulates Ppara expression. Thus, there is a mutual regulation between PPARGC1α and PPARα.

While Ksrp^−/− mice have reduced fat mass and increased EE, it will be of interest to see whether the enhanced brown fat gene program in iWAT is the sole contribution to these phenotypes. Using mice with both KSRP ablation and adipose-specific overexpression of miR-150 should provide an answer. Furthermore, since our mouse model is a global KSRP knockout, other effects on lipid metabolism in metabolic tissues, such as muscle and liver, may also contribute to the observed leanness. Further studies using adipose-specific KSRP knock-out mice should reveal additional functions of KSRP in controlling whole-body adiposity and lipid metabolism. Nevertheless, our observations in Ksrp^−/− mice are consistent with a large body of previous studies (8–11,21,24,47–52) revealing that increasing the browning of WAT shows resistance to DIO and improved glucose metabolism. In summary, this work demonstrates the in vivo role of KSRP in post-transcriptional regulation of miRNA expression to control brown-like remodeling of iWAT, and suggests that modulation of KSRP-dependent pri-miR-150 processing could potentially lead to therapeutics for obesity and metabolic disorders.

Acknowledgments. The authors thank Dr. Maria S. Johnson, University of Alabama at Birmingham, Animal Models Core, for conducting body composition analysis and indirect calorimetry; Dr. Martin Young, University of Alabama at Birmingham, for metabolic analysis using the Comprehensive Lab Animal Monitoring System; Dr. Chih-Hsuan Wang, Auburn University, for biostatistics analysis; and Dr. Douglas Moellering, University of Alabama at Birmingham-Comprehensive Diabetes Center, Core Diabetes Research Center Redox Biology Core (supported by National Institutes of Health grant P60 DK079626), for advice on and design of the measurement of oxygen consumption rates.

Funding. This work was supported by grants from Ministero della Salute (RF-2010-2306205) and the Association for International Cancer Research (#10-0527) to R.G., and by National Institutes of Health grant GM068758 (C.-Y.C.).

Duality of Interest. No potential conflicts of interest relevant to this article were reported.

Author Contributions. C.-F.C. helped to design and performed the experiments, analyzed the data, and wrote the manuscript. Y.-Y.L., H.-K.W., X.Z., M.G., P.B., and R.G. performed the experiments and analyzed the data. W.T.G. helped to conceive the project. C.-Y.C. helped to conceive the project and to design the experiments. C.-Y.C. is the guarantor of this work and, as such, had full access to all the data in the study and takes responsibility for the integrity of the data and the accuracy of the data analysis.