Obesity-dependent increase in RalA activity disrupts mitochondrial dynamics in white adipocytes

Mitochondrial dysfunction is a characteristic trait of human and rodent obesity, insulin resistance, and fatty liver disease. Here we report that mitochondria undergo fragmentation and reduced oxidative capacity specifically in inguinal white adipose tissue after feeding mice high fat diet (HFD) by a process dependent on the small GTPase RalA. RalA expression and activity are increased in white adipocytes from mice fed HFD. Targeted deletion of Rala in white adipocytes prevents the obesity-induced fragmentation of mitochondria and produces mice resistant to HFD-induced weight gain via increased fatty acid oxidation. As a result, these mice also exhibit improved glucose tolerance and liver function. In vitro mechanistic studies revealed that RalA suppresses mitochondrial oxidative function in adipocytes by increasing fission through reversing the protein kinase A-catalyzed inhibitory Ser637phosphorylation of the mitochondrial fission protein Drp1. Active RalA recruits protein phosphatase 2A (PP2Aa) to specifically dephosphorylate this inhibitory site on Drp1, activating the protein, thus increasing mitochondrial fission. Adipose tissue expression of the human homolog of Drp1, DNML1, is positively correlated with obesity and insulin resistance in patients. Thus, chronic activation of RalA plays a key role in repressing energy expenditure in obese adipose tissue by shifting the balance of mitochondrial dynamics towards excessive fission, contributing to weight gain and related metabolic dysfunction.

Ral GTPases are members of the Ras superfamily involved in multiple cellular processes. We previously demonstrated that RalA is activated by insulin in adipocytes, and in turn interacts with members of the exocyst complex to target GLUT4 vesicles to the plasma membrane for docking and subsequent fusion, leading to increased glucose uptake [21][22][23] . Insulin activates RalA through inhibitory phosphorylation of the RalGAP complex (RGC) 24 , as well as localization of RGL2, a guanine-nucleotide exchange factor (GEF) for RalA 25 . In vivo activation of RalA through targeted deletion of the RalGAP protein Ralgapb promotes glucose uptake into brown adipose tissue (BAT) 26 , and dramatically improves glucose homeostasis in mice on HFD. Similarly, targeted deletion of Ralgapa1 improves postprandial glucose and lipid disposal into muscle 27 .
We report here that RalA gene and protein expression and activity are increased in adipocytes from obese mice, and further that targeted deletion of Rala in white, but not brown adipocytes, attenuates HFDinduced obesity, due to dramatically increased energy expenditure and mitochondrial oxidative phosphorylation, speci cally in inguinal white adipose tissue (iWAT). These bene cial effects of RalA deletion were driven by a reversal of the increased mitochondrial ssion in white adipocytes induced by feeding mice HFD. In vitro studies revealed that RalA interacts with PP2Aa to promote the dephosphorylation of inhibitory S637 on Drp1, rendering Drp1 active, leading to excessive ssion and mitochondrial fragmentation. Taken together, these data reveal that persistent elevation of RalA in obesity produces mitochondrial dysfunction in white adipocytes, with profound effects on systemic metabolism.

Main
White adipocyte-speci c Rala deletion protects mice from high fat diet-induced obesity RNA-seq analysis from isolated mature adipocytes derived from control and HFD-fed mice 28 revealed that Rala expression is signi cantly upregulated in adipocytes from eWAT and iWAT during obesity development, while Ralgapa2 expression is downregulated (Fig. 1a,b). In addition, RalA protein content is increased in mature adipocytes from iWAT of obese mice (Fig. 1c, Extended Data Fig. 1a), accompanied by elevation of RalA-GTP binding (Fig. 1d, Extended Data Fig. 1b). We also observed a trend towards a positive correlation of the expression of the Ral GEF RGL2 in adipose tissue with BMI in a large dataset of obese patients (Extended Data Fig. 1c,d). Together, these observations support the notion that adipocyte RalA activity is constitutively elevated in obesity.
To explore further whether RalA plays a role in glucose homeostasis and energy metabolism, we generated adipocyte-speci c Rala knockout (Rala AKO ) mice by crossing Ralaoxed mice with adiponectin-Cre transgenic mice. Compared to Rala f/f littermates, Rala AKO mice had a greater than 90% decrease of RalA protein in primary adipocytes from WAT and BAT, and an approximately 50% decrease in whole WAT, without changes in liver (Extended Data Fig. 1e). Insulin-stimulated GTP binding of RalA was diminished in WAT of Rala AKO mice compared to control mice, and the same result was observed in primary adipocytes (Extended Data Fig. 1f).
We generated primary white adipocytes by differentiation of iWAT stromal vascular cells from control and KO mice. As previously seen in 3T3-L1 adipocytes 22 , knockout of RalA completely prevented the translocation of GLUT4 from intracellular sites to the plasma membrane in response to insulin (Extended Data Fig. 1g). Moreover, insulin-stimulated glucose uptake in KO cells was signi cantly reduced in knockout cells (Extended Data Fig. 1h).
Adipocyte speci c deletion of Rala had no effect on body weight in chow diet (CD)-fed mice, although these mice displayed a reduction in fat mass and depot weight (Extended Data Fig. 2a-c). Generally, adipocytes from iWAT were considerably smaller than those found in epididymal WAT (eWAT) from mice fed CD. Moreover, Rala AKO mice had smaller adipocytes in iWAT compared to control mice fed with CD, while adipocytes were comparable in eWAT and BAT between the genotypes (Extended Data Fig. 2d).
While Rala AKO mice on chow diet showed no difference in glucose tolerance, there was a slight reduction in insulin tolerance when compared to Rala f/f mice (Extended Data Fig. 2e,f). Insulin levels and HOMA-IR in Rala AKO mice were not different from control mice fed with CD (Extended Data Fig. 2g,h). However, Rala AKO mice gained signi cantly less weight than control littermates when challenged with 60% HFD (Fig. 1e). Rala AKO mice showed a marked reduction of fat mass, with no change in lean body mass ( Fig. 1f). Further analyses revealed that iWAT weight was signi cantly reduced in Rala AKO mice, with no difference in eWAT and BAT (Fig. 1g). HFD increased adipocyte size in all fat depots, but the effect was most pronounced in iWAT; HFD-fed Rala AKO mice displayed a trend towards smaller adipocytes in iWAT compared to control mice, but not in eWAT or BAT (Extended Data Fig. 2d). HFD-fed Rala AKO mice exhibited a marked improvement in glucose tolerance compared to control mice, with no change in insulin tolerance (Fig. 1h,i), but with reduced insulin levels and improved HOMA-IR (Fig. 1j,k). Fasting glucose levels were comparable between the genotypes on either HFD or CD (Extended Data Fig. 2i,j).
To investigate further which adipose tissue depot is responsible for the reduced weight gain in Rala AKO mice fed HFD, we generated BAT-speci c Rala knockout (Rala BKO ) mice by crossing RalA-oxed mice with UCP1-Cre transgenic mice (Extended Data Fig. 2k). Although CD-fed Rala BKO mice showed a reduction in BAT weight, presumably due to reduced glucose uptake, there were no differences in fat mass or depot weight compared with control mice (Extended Data Fig. 2l,m). Glucose and insulin tolerance tests (GTT and ITT) were identical between the genotypes on control diet (Extended Data Fig. 2n,o). Moreover, no differences in body weight, fat mass, tissue weight, GTT, or ITT were observed in HFD-fed Rala BKO mice (Extended Data Fig. 2p-t). These results suggest that speci c Rala deletion in WAT, especially in iWAT, protects mice against obesity.
Loss of RalA in WAT ameliorates HFD-induced hepatic steatosis Since HFD-fed Rala AKO mice showed an improved GTT without markedly altering insulin tolerance, we speculated that the improved glucose handling is due to reduced hepatic glucose production. To test this assumption, we performed a pyruvate tolerance test (PTT) in HFD-fed Rala f/f and Rala AKO mice. Rala AKO mice exhibited substantially lower glucose excursions following pyruvate challenge compared to control mice (Fig. 2a). There was a signi cant downregulation of the hepatic gluconeogenic genes G6pc and Pepck (Fig. 2b). These data suggest that adipose tissue-speci c Rala deletion improved glucose homeostasis partially through reduced hepatic glucose production.
Liver weights and triglyceride (TG) content were signi cantly reduced in HFD-fed Rala AKO mice when compared to control mice (Fig. 2c,d). Both H&E and Oil-Red-O staining indicated less lipid accumulation in the liver of Rala AKO mice (Fig. 2e). In line with histology results, lipogenic genes (Acc, Fasn, Scd1 and Acsl1) were expressed at signi cantly lower levels in the liver of Rala AKO mice (Fig. 2f). However, plasma leptin levels (Fig. 2g) and hepatic expression of genes related to fatty acid oxidation (FAO) (Fig. 2h) were unchanged in Rala AKO mice. In addition, in ammatory (Adgre1) and brosis-related (Col1a1 and Col3a1) genes were expressed at lower levels in livers of Rala AKO mice (Fig. 2i), as were aspartate aminotransferase (AST) and aminotransferase (ALT) activities (Fig. 2j,k). Of note, we did not observe a difference in liver weights in Rala BKO compared to controls fed with HFD (Extended Data Fig. 2r). Together, these observations suggest that WAT-speci c deletion of Rala systemically regulates lipid metabolism to ameliorate liver steatosis and damage in obesity.
RalA de ciency in WAT increases energy expenditure and mitochondrial oxidative phosphorylation To explore why adipose tissue Rala deletion protects mice from HFD-induced hepatic steatosis, weight gain, and glucose intolerance, we investigated energy metabolism in Rala AKO mice with metabolic cage studies. While Rala ablation in adipocytes did not affect energy metabolism and food intake in mice fed CD (Extended Data Fig. 3a-e), HFD-fed Rala AKO mice displayed a signi cant increase in energy expenditure (EE) during the dark phase as determined by ANCOVA using body weight as a covariate (Fig. 3a). Concordantly, oxygen consumption in Rala AKO mice was similarly increased compared to controls (Extended Data Fig. 3f), although there was no difference in respiratory exchange rate (RER), locomotor activity, or food intake between the genotypes (Extended Data Fig. 3g-i). In contrast, Rala BKO mice fed either control or HFD were identical to control littermates in EE, O 2 consumption, RER, locomotor activity, and food intake (Extended Data Fig. 3j-n). These observations demonstrate that Rala de ciency speci cally in WAT increases energy expenditure.
Increased energy expenditure is an indirect re ection of increased mitochondrial oxidative activity. Thus, we assessed the expression of mitochondrial proteins in fat depots. Oxidative phosphorylation (OXPHOS) proteins were markedly increased in iWAT of Rala AKO mice (Fig. 3b,c), but not in eWAT (Extended Data   Fig. 3o,p). Complex I and Complex II levels were modestly increased in BAT of Rala AKO mice (Extended Data Fig. 3q,r). This may occur because of systemic metabolic improvement in Rala AKO mice rather than a cell-autonomous BAT function, since HFD-fed Rala BKO mice did not show an improved metabolic phenotype. In this regard, plasma FFA and TG levels in HFD-fed Rala AKO mice were signi cantly lower (Fig. 3d,e). To test the possible involvement of a generalized browning of iWAT, we also examined thermogenic markers. Ucp1, Cidea, and Prdm16 expression was identical between the genotypes in all three fat depots, indicating that the improvement in energy expenditure in Rala AKO mice did not re ect the development of beige adipose tissue (Extended Data Fig. 3s).
RalA knockout in white adipocytes increases mitochondrial activity and fatty acid oxidation We sought to evaluate further the mechanisms underlying improved energy metabolism in Rala AKO mice, and directly assessed mitochondrial activity in adipocytes. Measurements of basal respiration revealed that oxygen consumption rate (OCR) was signi cantly increased in mitochondria isolated from KO iWAT compared to that from control mice, but was similar in eWAT mitochondria of Rala f/f and Rala AKO mice (Fig. 3f). We also noted that both basal and maximal respiration were signi cantly higher in primary differentiated adipocytes from KO mice, and the difference in maximal respiration was blunted by the addition of the CPT1 inhibitor etomoxir that blocks fatty acid oxidation (FAO) (Fig. 4a, Extended Data Fig. 4a). To investigate directly whether RalA plays a role in controlling FAO, we incubated cells with ( 14 C)labeled palmitic acid (PA) and measured its oxidation to either acid-soluble metabolites (ASM) or CO 2 in WT and KO white adipocytes. In agreement with the OCR results, fatty acid oxidation was signi cantly higher in KO compared to WT adipocytes (Fig. 4b). These data indicate that RalA knockout in WAT increases energy expenditure due to increased mitochondrial oxidation activity.
To ensure that these studies re ected the activity of RalA, we also generated an immortalized preadipocyte line from Rala f/f mice and induced Rala deletion by transducing cells with Cre lentivirus. The Cre recombinase completely ablated RalA in preadipocytes and fully differentiated adipocytes (Extended Data Fig. 4b). BODIPY staining demonstrated that both primary and immortalized preadipocytes from WT and KO mice were fully differentiated. As an orthogonal approach, we performed live cell imaging using the cell permeant uorescent dye, TMRM, to detect mitochondrial membrane potential (MtMP), which re ects electron transport and oxidative phosphorylation in active mitochondria. KO adipocytes exhibited a higher TMRM signal intensity than did their WT counterparts (Fig. 4c, Extended Data Fig. 4c). To specify the ability of TMRM to detect mitochondrial depolarization in active mitochondria, we applied the β3adrenergic receptor agonist CL316,243 (CL) to induce mitochondrial membrane depolarization 29 . The TMRM signal declined quickly after administration of the agonist, which con rms that TMRM stains only active mitochondria (Extended Data Fig. 4d).
We previously reported that lipolysis drives mitochondrial oxidative metabolism in adipocytes 30 . To rule out a possible role for lipolysis as the primary driver of increased oxidative capacity of Rala-KO adipocytes, we performed in vitro and in vivo lipolysis assays. CL robustly stimulated FFA and glycerol release to the same extent in KO and WT immortalized adipocytes, and the molar ratio of FFA to glycerol was approximately 3:1 (Extended Data Fig. 4e,f). Additionally, there was no difference in CL-induced FFA and free glycerol production in Rala f/f and Rala AKO mice (Extended Data Fig. 4g,h). We further tested whether Rala AKO mice are defective in the suppression of FFA release by insulin. Insulin suppressed CLinduced FFA release by approximately 50% in both WT and KO cells (Extended Data Fig. 4e). A single injection of insulin reduced FFA levels in control and Rala AKO mice to the same extent (Extended Data Fig. 4i). Interestingly, KO adipocytes displayed a mild increase in glycerol release in the presence of CL, while Rala AKO mice showed a mild decrease of plasma glycerol levels either in the presence of CL or after fasting (Extended Data Fig. 4f,h,j). Taken together, these results suggest that the absence of RalA in adipocytes enhances mitochondrial oxidative activity without affecting FFA supply.
Targeted Rala knockout protects against obesity-induced mitochondrial ssion in iWAT The increased mitochondrial oxidative activity observed in HFD-fed Rala AKO mice could result from increased mitochondrial biogenesis. Expression of genes related to mitochondrial biogenesis was comparable between the genotypes (Extended Data Fig. 5a,b) in WAT. The activity of AMPK, the master regulator of mitochondrial biogenesis 31,32 , was also comparable between control and Rala AKO mice fed with HFD (Extended Data Fig. 5c-f). In addition to biogenesis, mitochondrial function can also be regulated by dynamic changes in morphology through tightly controlled fusion and ssion events that shape the organelle to comply with energy demands 19,33 . Electron microscopy (EM) revealed that HFD feeding of WT mice induced the appearance of smaller, spherical mitochondria in iWAT (Fig. 4d), consistent with previous reports that mitochondrial function and morphology is impaired in obese adipocytes 34,35 . In agreement with the in vivo metabolic phenotypes, adipocyte Rala deletion did not grossly affect mitochondrial morphology in iWAT of CD-fed mice (Fig. 4d), but the HFD-induced change in mitochondrial morphology was completely prevented in Rala KO iWAT; mitochondria in iWAT from these mice displayed an elongated shape that was indistinguishable from CD-fed mice (Fig. 4d). Indeed, tissue weight ( Fig. 1f), OXPHOS content (Extended Data Fig. 3o,p), and mitochondrial OCR (Fig. 3f) were not affected by RalA deletion in eWAT, corresponding to the observation that the appearance of fragmented mitochondria in this depot was not reversed by RalA KO in HFD mice (Extended Data Fig. 5g). In fact, mitochondria in eWAT do not undergo signi cant fragmentation in response to HFD, possibly because of their already fragmented shape, consistent with the overall anabolic function of visceral adipocytes (Fig. 4d, Extended Data Fig. 5g). We also examined mitochondrial morphology in immortalized adipocytes differentiated from iWAT. As shown in Fig. 4e, mitochondria in KO adipocytes appeared longer than those in WT cells. There was a higher frequency of elongated mitochondria (1.0-1.5 µm) in KO cells (Fig. 4f), and the mean maximal mitochondrial length was signi cantly higher than in WT cells (Fig. 4g).
Inhibition of RalA increases Drp1 S637 phosphorylation in white adipocytes Opa1 and Drp1 have been identi ed as key regulators of mitochondrial fusion and ssion, respectively 36 . Opa1 undergoes proteolytic cleavage to generate long (L-Opa1) and short (S-Opa1) forms that together fuel mitochondrial fusion 37,38 39 . Protein levels of both forms of Opa1 were downregulated in iWAT after HFD feeding (Extended Data Fig. 5h-j); only S-Opa1 was downregulated in eWAT from Rala AKO mice (Extended Data Fig. 5k-m), indicating the likelihood of reduced fusion in KO mice compared to WT littermates. However, the observation of elongated mitochondria in KO mice ( Fig. 4d) suggests that this change in Opa1 processing is likely compensatory. We then focused on Drp1 as a key regulator of ssion.
Interestingly, Drp1 phosphorylation at the anti-ssion S637 site was signi cantly increased in Rala-KO iWAT (Fig. 5a, Extended Data Fig. 6a), whereas Drp1 S637 phosphorylation was comparable between the genotypes in eWAT (Extended Data Fig. 6b,c). To establish whether this effect is cell-autonomous, we examined Drp1 phosphorylation in both immortalized and primary adipocytes. Drp1 S637 phosphorylation is catalyzed by PKA, activated by the b-adrenergic/cAMP pathway 40,41 . Drp1 S637 phosphorylation was triggered by CL after 5 minutes and was maximal after 15 minutes in adipocytes ( Fig. 5b, Extended Data Fig. 6d). Consistent with in vivo results, Rala-KO adipocytes showed a signi cantly higher Drp1 S637 after β-adrenergic stimulation compared to WT cells (Fig. 5c, Extended Data Fig. 6e). We also explored the effect of RalA on Drp1 S637 phosphorylation state using a speci c Ral inhibitor that prevents activation and retains the GTPase in the GDP-bound, inactive state.
Pretreatment with the pan-Ral inhibitor RBC8 26,42 signi cantly increased forskolin-stimulated Drp1 S637 phosphorylation in 3T3-L1 adipocytes (Extended Data Fig. 6f,g). Inhibition of RalA activity with RBC8 also increased forskolin-stimulated Drp1 S637 phosphorylation in the human primary adipocyte cell line (SGBS) (Fig. 5d,e). Thus, RalA speci cally modulates Drp1 S637 phosphorylation downstream of PKA activation across multiple adipocyte cell lines of both murine and human origin. To determine whether RalA in uences CL-induced PKA activation or cAMP breakdown, we measured cAMP production and phosphorylation of hormone sensitive lipase (HSL) in adipocytes. There was no difference in cAMP production between WT and KO primary adipocytes after 5 minutes of CL stimulation (Extended Data Fig. 6h). Similarly, HSL S660 phosphorylation was identical in WT and KO adipocytes (Extended Data Fig. 6i-l).
To examine the relevance of Drp1 as a regulator of metabolism in human obesity, we analyzed microarray data of abdominal subcutaneous WAT from obese and non-obese women. In human subcutaneous WAT, DNM1L (encoding human Drp1 protein) expression was positively correlated with BMI and HOMA ( Fig. 5f,g), and its expression was signi cantly upregulated in obese subjects (Fig. 5h), indicating that increased expression of DNM1L may contribute to mitochondrial dysfunction in obesity. Moreover, bioinformatic analysis of published microarray data (GEO: GSE7053) from 770 human males further con rmed that DNML1 is associated with obesity (Extended Data Fig. 6m-o). Together, these in vivo and in vitro data suggest that upregulated Drp1 activity in adipose tissue may be an important contributor to mitochondrial dysfunction during obesity and further that RalA de ciency protects mitochondria from excessive ssion by increasing Drp1 S637 phosphorylation.
RalA interacts with Drp1 and protein phosphatase 2A, promoting dephosphorylation of Drp1 at S637 To understand the molecular mechanism by which RalA regulates Drp1 S637 phosphorylation, we used proteomics to search for proteins interacting with wildtype (WT), constitutively active (G23V), or dominant negative (S28N) forms of RalA ectopically expressed in liver. Among the binding proteins was protein phosphatase 2A subunit A alpha (PP2Aa), the scaffolding subunit encoded by the Ppp2r1a gene, which preferentially bound to the RalA G23V constitutively active mutant. To con rm these mass spectrometry data, we puri ed RalA WT -Flag protein from HEK293T cells and pulled down PP2Aa from lysates (Fig. 6a).
To determine whether this interaction is dependent on the activation state of the G protein, we coexpressed WT and mutant RalA constructs with PP2Aa in HEK293T cells. As a positive control, the effector Sec5 only bound to active RalA G23V . Similarly, this mutant form of RalA had the highest a nity for PP2Aa (Fig. 6b). We also loaded a RalA-Flag fusion protein in vitro with GTPγS or GDP to evaluate the speci city of effector binding. Both Sec5 and PP2Aa were pulled down by RalA loaded with GTPγS but not with GDP (Fig. 6c). In addition, because PP2Aa and Drp1 did not independently interact (data not shown), we investigated whether RalA directly modi es Drp1 phosphorylation via PP2Aa. When coexpressed, Drp1 and RalA interacted directly with each other, although there was no preference for the activation state of RalA (Extended Data Fig. 7a). Activation of the cAMP/PKA axis by addition of forskolin increased Drp1 S637 phosphorylation, while co-expression of PP2Aa promoted the dephosphorylation of S637 (Fig. 6d), although overexpression of PP2Ab had no effect (Extended Data Fig. 7b). These data suggest that Drp1 is constitutively associated with RalA independent of activation state, and upon activation, RalA recruits PP2Aa to promote the dephosphorylation of Drp1 S637.
To understand further the effects of RalA activation state on Drp1 phosphorylation and mitochondrial function, we transduced immortalized RalA KO cells with RalA WT and RalA G23V lentivirus prior to differentiation into adipocytes. RalA G23V expressing adipocytes showed a robust increase in RalA GTP binding (Fig. 6e), and these cells had signi cantly less Drp1 S637 phosphorylation (Fig. 6f, Extended Data   Fig. 7c). Expression of either RalA WT or RalA G23V signi cantly reduced mitochondrial potential in KO adipocytes (Fig. 6g, Extended Data Fig. 7d). To con rm that this reduction in mitochondrial potential is associated with reduced oxidative function, we performed a seahorse assay. Consistent with results in primary adipocytes, RalA WT and RalA G23V expressing adipocytes displayed reduced basal and maximal oxygen consumption rate (OCR) in comparison to KO adipocytes (Fig. 6h, Extended Data Fig. 7e). In addition, EM revealed that overexpression of WT or constitutively active RalA in adipocytes resulted in fragmented mitochondria, indicating increased ssion compared to RalA KO adipocytes (Fig. 6i).
RalA has previously been reported to promote ssion in proliferating cells, and Rala knockdown led to a long, interconnected mitochondrial network and reduced proliferation 43 . Partially in agreement with this study, we found that RalA de ciency resulted in elongated mitochondria in adipocytes, with increased oxidative phosphorylation that dramatically impacted whole body lipid metabolism. However, unlike the previous study, we did not observe an interaction between RalBP1 and Drp1. Interestingly, total PP2Aa protein levels were increased in Rala KO iWAT compared to control iWAT, without a difference in PP2Ab and PP2Ac content (Extended Data Fig. 7f,g), perhaps re ecting a compensatory pathway. Taken together, our data suggest that obesity drives RalA expression and GTP binding activity, leading to its association with PP2Aa, which in turn recruits the catalytic subunit PP2Ac to dephosphorylate Drp1 S637. We also note that catecholamine resistance, an inherent trait of the obese state 28 , is also expected to lead to reduced PKA-catalyzed S637 phosphorylation. Together, these effects result in constitutive mitochondrial translocation of Drp1 and fragmented mitochondria in adipocytes from obese subjects (Extended Data Fig. 8).

Discussion
While accumulating evidence suggests that mitochondrial dysfunction is a characteristic trait of obesity in human and rodent adipocytes 16, 34,35,44 , the underlying molecular mechanisms remain unknown. Here, we report a new regulatory axis for the control of mitochondrial morphology and function in the context of obesity involving prolonged activation of the small GTPase RalA. We show that RalA is both induced and activated in white adipocytes after feeding rodents a high fat diet, while the negative regulator of RalA, RalGAP, is downregulated. We also observe a positive correlation of expression of the RalGEF RGL2 with BMI in adipose tissue of humans with obesity, expected to correspond to a chronic increase in RalA activity. The increase in adipocyte RalA mRNA, protein, and activity is associated with mitochondrial dysfunction, characterized by fragmentation and reduced oxidative capacity, speci cally in iWAT. Targeted deletion of RalA in white adipocytes prevents the obesity-dependent fragmentation of mitochondria and produces mice resistant to HFD-induced weight gain via increased energy expenditure. In vitro studies revealed that RalA suppresses mitochondrial oxidative function in adipocytes by increasing ssion through reversing the inhibitory phosphorylation of the mitochondrial ssion protein Drp1. This reduced phosphorylation results from the recruitment of the regulatory subunit of PP2A, which acts as a bona de effector of RalA, leading to the speci c dephosphorylation of the inhibitory Ser 637 residue on Drp1, rendering the protein active. We also note our previous study in which constitutive activation of RalA via adipocyte-speci c KO of Ralgapb produced a signi cant enlargement of white adipocytes and increased adipose tissue mass, even on a control diet 26 . Thus, chronic elevation in RalA activity plays a key role in repressing energy expenditure in obese adipose tissue, contributing to weight gain and related metabolic dysfunction, including glucose intolerance and fatty liver, and may explain in part how energy expenditure is repressed in prolonged obesity 45 .
The observation that adipocyte RalA controls overall systemic metabolism via this mechanism was surprising. We and others previously reported that RalA plays a key role in controlling the tra cking of GLUT4 vesicles in adipocytes and muscle 22,23 . RalA is activated by insulin, mainly by inhibition of its GAP complex through phosphorylation 24,46 , and when activated, RalA interacts with components of the exocyst complex to target GLUT4 vesicles to the plasma membrane for fusion, increasing glucose uptake into fat cells 21 . Indeed, adipocytes treated with a RalA inhibitor 26 or isolated from RalA KO mice showed dramatically reduced GLUT4 translocation to the plasma membrane, with less glucose uptake in response to insulin. Targeted deletion of the scaffolding subunit of the RalGAP complex resulted in constitutive activation of RalA in adipocytes and myocytes, and dramatically improved glucose homeostasis 22,26,46 . However, detailed physiological tracer studies revealed that improvements in glucose disposal in adipocyte-speci c KO mice occurred primarily in brown fat, where glucose uptake was markedly increased 26 . Consistent with these ndings, we observed a small but signi cant reduction in insulin sensitivity in Rala AKO mice on control diet, accompanied by reduced weights of all adipose tissues, likely re ecting less nutrient uptake. However, Rala AKO mice on HFD paradoxically showed improved glucose tolerance and insulin sensitivity. While it remains unclear exactly how these mice overcome the negative effects of RalA deletion on glucose uptake, GLUT4 mRNA and protein levels in WAT are downregulated in obesity 12,47,48 , whereas GLUT1 mRNA and protein levels are increased 49,50 , consistent with our RalGAP KO studies in HFD-fed mice that show little glucose uptake into white fat in response to insulin, but higher basal levels 26 . Thus, it seems likely that improved glucose tolerance in Rala AKO mice occurs because of weight loss and increased energy expenditure.
It was also interesting that liver function was dramatically improved in Rala AKO mice on HFD, with reduced hepatic lipids and gluconeogenesis, as indicated by improvements in pyruvate tolerance. It is well established that WAT plays an important role in regulating whole-body energy metabolism 51 .
Hepatic acetyl-CoA arises from WAT lipolysis to directly promote hepatic gluconeogenesis 52 . The increase in fatty acid oxidation in Rala-KO adipocytes resulted in less circulating FFAs and TGs, likely producing improved liver health and reduced gluconeogenesis.
While the signi cance of the adipose depot speci city of the effects of RalA remains uncertain, we note that adipocytes in visceral, subcutaneous, and brown fat differ in many ways 53,54 . Although RalA was deleted in all adipocytes in Rala AKO mice, mitochondrial function was only improved in iWAT. While there are numerous differences between visceral and inguinal white adipocytes that might explain this, including their response to HFD, one notable issue has to do with inherent mitochondrial morphology.
Upon HFD feeding, adipocytes in iWAT underwent a dramatic size expansion, accompanied by a change in mitochondria from an elongated to a fragmented morphology, re ecting a transition to a largely anabolic state. These changes were not observed in RalA KO mice. Unlike what was observed in iWAT, mitochondria in eWAT display a fragmented morphology even in lean mice, with no change observed after HFD or RalA KO, consistent with the overall energy storage function of this depot even without the anabolic pressure of overnutrition.
Another question concerns the role of RalA in BAT. While BAT tissue weight was reduced in both Rala AKO and Rala BKO mice compared to controls, likely due to reduced glucose uptake, only iWAT adipocytes appear to respond with a change in metabolic activity and mitochondrial morphology. Brown adipocyte mitochondria are morphologically different from those in white adipocytes. These brown adipocyte mitochondria are more numerous and larger than the mitochondria in white adipocytes and contain packed cristae. Comparison of the mitochondria of brown and white adipocytes by proteomic analysis revealed that proteins involved in pathways related to fatty acid metabolism, OXPHOS, and the TCA cycle were highly expressed in BAT compared to WAT 55 . Thus, it seems likely that mitochondria in BAT are subjected to fundamentally different modes of regulation than those in white fat, and the reduced weight of BAT in KO mice can be attributed to reduced glucose uptake.
As mitochondrial function is vital for healthy metabolism, efforts have focused on preventing fragmentation via blocking activity or direct deletion of Drp1 56 . Muscle mitochondrial dysfunction is closely related to excessive Drp1 activity 57 , and elevated Drp1 activating S616 phosphorylation has been found in severely obese human muscle 58,59 . On the other hand, triggering Drp1 S637 phosphorylation has been suggested to increase the uncoupling capacity of FFA in brown adipocytes 29 . In line with this observation, increased S637 phosphorylation was found in BAT after cold exposure 60 . Administration of a Drp1 inhibitor acutely improved muscle insulin sensitivity and systemic glucose tolerance 61,62 . However, the impact of modulating Drp1 levels is complicated and varies between tissues. Targeted deletion of Drp1 in liver reduced hepatic lipid accumulation and body weight in a NAFLD model 63 . Moreover, loss of Drp1 impairs brown adipocyte differentiation and thermogenesis, possibly re ecting the aspects of mitochondrial morphology that are unique to BAT 60,64 . Interestingly, ER stress has been observed in both tissue-speci c Drp1 knockout mice models, which suggests that Drp1 may also regulate ER remodeling 65 . These ndings highlight the likely differences between total ablation of Drp1 activity and changes in its upstream regulatory pathways.

Materials Animals
RalA-oxed (Rala f/f ) mice were bred with Adiponectin-promoter driven Cre or Ucp1-promoter driven Cre transgenic mice to generate fat depot speci c RalA knockout (Rala AKO or Rala BKO ) mice. All mice have a C57BL/6J background, and all experiments were done using littermates. Male mice were used for in vivo experiments, and female mice were only used for primary preadipocyte isolation. We fed mice with standard chow diet (CD) (Teklad, #7912) or high fat diet (HFD) consisting of 60% calories from fat (Research Diets, #D12492) for 8-12 weeks, starting from 8 weeks old. Mice were housed in a speci c pathogen-free facility with a 12-hr light and 12-hr dark cycle and given free access to food and water. All animal experiments were approved by and followed the guidelines from the Institutional Animal Care and Use Committee (IACUC) at the University of California, San Diego.
Cell lines Primary preadipocytes. The isolation of primary preadipocytes was done as described previously 67 with some modi cation. IWAT from two to three 8-week-old female mice was dissected, minced, and digested in 5 mL 1 mg/mL collagenase (Sigma) for 15 minutes (min) in a 37°C water bath with gentle agitation. HEK 293T cells. HEK 293T cells were cultured in high glucose DMEM-FBS medium. On the same day as seeding, when cells reached 50% con uence, transfection was done as designed using lipofectamine 2000 (Life Technology) following the manufacturer's protocol. Fresh DMEM-FBS medium was added 12-16 hrs after transfection. 48 hrs after transfection, cells reached around 80% con uency and were used for co-immunoprecipitation or pulldown experiments.

Gene analysis in clinical cohorts
The transcriptomics data from abdominal subcutaneous WAT of 30 obese and 26 healthy non-obese women were generated as previously described 72  instructions. RNA quality was checked by Agilent TapeStation. Biological triplicates of isolated 500 ng RNA were used to prepare sequencing libraries using the TruSeq RNA Sample Preparation Kit v2 (Illumina), according to the manufacturer's protocol. Libraries were validated using a 2100 BioAnalyzer (Agilent), then normalized and pooled for sequencing using bar-coded multiplexing at a 90-bp single-end read length on an Illumina HiSeq 4000. Samples were sequenced to a median depth of 14 million reads.

Bioinformatics analysis
For RNA-Seq, sequencing fastq les were generated automatically using the Illumina bcl2fastq2 Conversion Software. Read alignment and junction mapping to genome mm39 (GRCm39) and the mouse Genecode M30 annotation were accomplished using STAR (version 2.7.2b). Known splice junctions from mm10 were supplied to the aligner and de novo junction discovery was also permitted. Differential gene expression analysis and statistical testing were performed using DESeq2. Differentially expressed genes were de ned as having an adjusted P value < 0.05. Raw gene counts were normalized to FPM (fragments per million mapped fragments) using DEseq2. FPM counts were ltered, centered by z-score before gene clustering and heatmap generation using GENE-E (v3.0.215) or GraphPad Prism (8.4.3). For microarray data, gene matrix les were collapsed by Collapse Dataset tool in GSEA (4.3.2) using chip platform (GPL13667) with collapsing mode (Mean_of_probes). Statistical signi cance of differential gene expression was assessed by ComparativeMarkerSelection module (version 11) from GenePattern (https://cloud.genepattern.org/gp/pages/index.jsf).

Gene expression analysis
Tissue RNA was isolated with TRIzol™ reagent in combination with column (PureLink RNA mini, Invitrogen) according to the manufacturer's protocol. cDNA was generated from 1 µg RNA using the cDNA Maxima Reverse Transcription Kit (Thermo Fisher Scienti c). mRNA expression was assessed by realtime PCR using QuantStudio 5 real-time PCR system and SYBR Green PCR master mix (Invitrogen). Gene expression was normalized to Cyclophilin A in murine tissues. Relative mRNA expression levels were calculated using averaged 2 -ΔΔCt values for each biological replicate. Primers are listed in Extended Data Table 1.

Body mass composition
Body mass composition was assessed in non-anesthetized mice by using EchoMRI.

Glucose tolerance test
Mice were fasted for 6 hrs, then intraperitoneally (i.p.) injected with D-[+]-glucose in PBS at a dose of 2 g/kg BW for CD-fed mice or 1.2 g/kg BW for HFD-fed mice. Blood glucose levels were measured before injection and at 15, 30, 60, 90, and 120 min after injection using the Easy Touch glucose monitoring system.

Insulin tolerance test
Mice were fasted for 4 hrs, then i.p. injected with human insulin (Sigma) in saline at a dose of 0.35 U/kg BW for CD-fed mice or 0.6 U/kg BW for HFD-fed mice. Blood glucose levels were measured before injection and at 15, 30, 60, 90, and 120 min after injection using the Easy Touch glucose monitoring system.

Pyruvate tolerance test
Mice were fasted for 16 hrs, then i.p. injected with pyruvate in PBS at a dose of 1.5 g/kg BW for HFD-fed mice. Blood glucose levels were measured before injection and at 15, 30, 60, 90, and 120 min after injection using the Easy Touch glucose monitoring system.

Blood parameters
Whole blood was taken from the facial vein and blood glucose was measured with a glucose meter (Easy Touch) from the tail vein. Plasma was collected after centrifugation at 1,200 x rpm, 4°C for 10 min.

Hepatic lipid TG measurement
Frozen liver tissue (50-100 mg) was homogenized in 1 mL of PBS. 800 uL of lysates were added to 4 mL extraction buffer. After thoroughly rotating for 30 min at room temperature, the lipid phase was separated from the aqueous phase by centrifuging at 3,000 x rpm for 20 min. 0.2 mL of lipid fraction in the organic phase was collected and transferred to a 1.5 mL tube to dry under nitrogen stream in the fume hood. 0.2 mL of 2% Triton X-100 solution was used to solubilize the lipids. Triglyceride levels were determined by using the In nity ™ Triglycerides kit (Thermo Fisher). Lipid amount was normalized to liver lysate protein amount.

Indirect calorimetric measurements
For metabolic cage study, mice were individually housed in Promethion metabolic cages maintained at 22°C under a 12-hr light/12-hr dark cycle. Prior to the experiment, mice were adapted to the metabolic cages for 2 days. The monitoring system records and calculates food intake, locomotor activity, oxygen consumption, carbon dioxide production, respiratory exchange ratio (RER), and energy expenditure (EE).
Mice were provided with free access to water and food during the whole measurement. The data were exported with ExpeData software (SABLE SYSTEMS) and EE was analyzed using ANCOVA with body weight as a covariate by web-based CalR tool 75 . Respiration measurement Intact Cells. The cellular OCR was measured using an eXF96 Extracellular Flux Analyzer and analyzed by Agilent Seahorse Wave Software (Seahorse Bioscience). Prior to assay, 2,500 primary preadipocytes were seeded into XF96 microplates. Two days after reaching full con uency, adipogenic differentiation was initiated using a protocol mentioned above. Once fully differentiated, adipocyte culture medium was changed to assay medium containing 25 mM glucose, 1 mM pyruvate and 2 mM L-Glutamine and 0.5 mM carnitine without phenol red or sodium bicarbonate for 3 hrs. Prior to the measurement, cells were incubated in a CO 2 -free incubator for 15 min. Basal rates of respiration were measured in assay medium and followed with sequential injections of oligomycin (2 µM), FCCP (0.5 µM), and Rotenone with Antimycin A (each 0.5 µM). Oxygen consumption values were normalized to protein content. Isolated mitochondrial. Isolation of mitochondrial from HFD-fed mice, and the OCR with 2.5 ug isolated mitochondrial was performed as previously described 30 . Fatty acid oxidation assay Fatty acid oxidation (FAO) assay was modi ed from a previous described protocol 76 . Fully differentiated primary adipocytes in 24-well plates were incubated in 0.5 mL DMEM per well containing 1 mM carnitine and 0.5 µCi/well [ 14 C]-palmitic acid for 60 min at 37°C. Afterwards, 360 µL medium was collected and added to 40 µL 10% BSA in a 1.5 mL tube with a lter paper in the cap. 200 µL 1 M perchloric acid was added to the tube, and the cap was immediately closed tightly and incubated at room temperature. After 1 hr, captured CO 2 and acid-soluble metabolites (ASM) were used to measure radioactivity. The cells were lysed in NaOH/SDS buffer (0.3 N/0.1%) to measure protein concentration. FAO rates were normalized to protein content.

Confocal microscope imaging
Live cell. Fully differentiated adipocytes were cultured in a glass bottom dish (Cellvis) and incubated in phenol red-free DMEM (imaging medium) with 100 nM TMRM (Thermo Fisher) for 30 min to indicate mitochondrial membrane potential, and BODIPY 493/503 ( nal 5 ug/mL, Life Technology) was added to label lipid droplets for the last 15 min. Cells were then washed three times with imaging medium. Live cell images were obtained with Nikon A1R confocal with 100 x or 60 x oil immersion objective. For time-lapse imaging, pictures were taken every 10 min.
Fixed cell. Fully differentiated primary adipocytes were cultured in a glass bottom chamber (Lab-Tek). On the day of the experiment, cells were serum starved for 3 hrs and treated with 100 nM insulin. After 15 min, medium was removed and cells were xed with ice-cold methanol and incubated at -20°C for 10 min.
Cells were then washed twice with PBS and blocked with 10% goat serum in PBS with 0.1% Triton X-100 at room temperature for 30 min. After blocking, cells were incubated with primary antibody at 4°C overnight and secondary antibody for 1 hr at room temperature. Cells were washed three times with PBS before imaging with Nikon A1R confocal microscope using 100x oil immersion objective.

Lipolysis
In vitro. Fully differentiated primary adipocytes in a 24-well plate were serum starved in lipolysis medium (2% BSA-phenol red-free DMEM) for 3 hrs. For insulin treatment, 100 nM insulin was added to cells for 30 min starting at the 2.5th hour of starvation. After starvation, medium was replaced with 0.5 mL fresh lipolysis medium with vehicle, 1 µM CL, 100 nM insulin, or in combination. Medium was collected after 1 hr incubation at 37°C. Released free fatty acids and free glycerol levels were measured using 100 µL medium with NEFA kit (WAKO) and Free Glycerol Reagent (Sigma) according to the manufacturer's protocol.
In vivo. CD-fed mice were used for in vivo lipolysis. For CL-induced lipolysis, ad libitum fed mice were intraperitoneally (i.p.) injected with PBS or CL (1 mg/kg) for 60 min. Circulating free fatty acids and free glycerol levels were measured using 2 uL plasma with NEFA kit (WAKO) and Free Glycerol Reagent (Sigma). For insulin-suppressed lipolysis, overnight fasted mice were i.p. injected with insulin (0.5 U/kg) for 60 min. Circulating free fatty acids and free glycerol levels were measured at indicated conditions. Electron microscopy Adipose tissue. Dissected adipose tissue was immediately xed with 2-3 drops of xative buffer (2% paraformaldehyde, 2.5% glutaraldehyde in 0.15 M sodium cacodylate buffer pH 7.4). Fat tissues were gently removed and xed at room temperature. After 2 hrs incubation, tissues were further cut into around 1 mm 3 cubes and immersed in xative buffer overnight at 4°C. Tissue cubes were post xed in 1% osmium 0.15 M sodium cacodylate buffer (SC buffer) for 1-2 hrs on ice, followed by ve 10-min washes in 0.15 M SC buffer, then rinsed in ddH 2 O on ice. Washed tissues were stained with 2% uranyl acetate for 1-2 hrs at 4°C then dehydrated in an ethanol series (50%, 70%, 90%, 100%, 100%, 10min each time) and dried in acetone for 15 min at room temperature. Dried tissues were in ltrated with 50%:50% Acetone:Durcupan for 1 hr or longer at room temperature then changed to 100% Durcupan overnight. The next day, embedded tissues in Durcupan were placed in a 60°C oven for 36 to 48 hrs. Ultrathin sections (60 nm) were cut on a Leica microtome with a Diamond knife and then post-stained with both uranyl acetate and lead. Images were obtained by using a Jeol 1400 plus Transmission Electron Microscope equipped with a Gatan digital camera.
Immortalized cells. Fully differentiated cells in a 6-well plate were quickly xed with 2% glutaraldehyde in 0.1 M sodium cacodylate buffer (pH 7.4) at room temperature for 15 min then incubated at 4°C for 15 min. Afterwards, cells were scraped down and pelleted by centrifugation. Cell pellets were post xed in 1% OsO4 in 0.1 M sodium cacodylate buffer for 1 hr on ice. The cells were stained all at once with 2% uranyl acetate for 1 hr on ice, then dehydrated in graded series of ethanol (50-100%) while remaining on ice.
The cells were then subjected to one wash with 100% ethanol and two washes with acetone (10 min each) and embedded with Durcupan. Sections were cut at 60 nm on a Leica UCT ultramicrotome and picked up on 300 mesh copper grids. Sections were post-stained with 2% uranyl acetate for 5 min and Sato's lead stain for 1 min. Images were obtained by using a Jeol 1400 plus Transmission Electron Microscope equipped with a Gatan digital camera.

cAMP measurement
To induce cAMP production, fully differentiated primary adipocytes were stimulated with 1 µM CL for 5 min. Cells were then immediately lysed in lysis buffer (0.1 N HCL) and cAMP levels were measured with the Direct cAMP Enzyme Immunoassay kit (Sigma) according to the manufacturer's protocol. min then cleared by centrifugation. Protein concentrations were measured with the DC protein assay (Bio-Rad) and 0.5-1 mg protein was used for incubation at 4°C with 20 µL GST-Δ RalBP1 agarose beads (Millipore) for 45 min or 20 µL ANTI-FLAG™ M2 A nity gel (Sigma) overnight. After incubation, beads were washed three times with RalA buffer and boiled at 65°C in 2X SDS buffer for 10 min.
Cell lysates were rotated for 15 min at 4°C and cleared by centrifugation for 15 min at 17,000 x g at 4°C.
Flag-RalA WT lysates were incubated with 20 µL ANTI-FLAG™ M2 A nity gel (Sigma) at 4°C. After 2 hrs rotation, the empty M2 or Flag-RalA WT beads were washed three times with lysis buffer then incubated with GFP-PP2Aa lysates at 4°C overnight. The next day, beads were washed three times with washing buffer (25 mM Tris-HCl, 40 mM NaCl, 30 mM MgCl 2 , 0.5% NP-40, EDTA-free protease inhibitor) and boiled in 2X SDS buffer at 65°C for 10 min. For GTPγS and GDP loading to Flag-RalA WT beads, washed beads were rinsed with loading buffer (20 mM Tris, 50 mM NaCl, 1 mM DTT, 2 mM EDTA) then incubated with 2 mM GTPγS or 200 µM GDP in loading buffer for 1 hr at 25°C with 600 x rpm agitation. After loading, 10 mM MgCl 2 was added to stop the loading, and loaded beads were incubated with GFP-PP2Aa lysates as above.
Co-immunoprecipitation. For the co-IP experiment, it is critical to harvest cells at around 70-80% con uency. Co-transfected cells were washed twice with ice-cold TBS and lysed in 0.5 mL lysis buffer (the same as above) or Drp1 buffer (25 mM Tris, 50 mM NaCl, 0.5 mM MgCl 2 , 10% glycerol, 0.5% NP-40, EDTA-free protease inhibitor). Lysates were cleared by centrifugation and protein concentrations were measured with BCA (Pierce). 0.5-1 mg protein was used for incubation with 20 µL ANTI-FLAG™ M2 A nity gel (Sigma) at 4°C. After overnight gentle rotation, beads were washed three times with washing buffer (the same as above) or Drp1 wash buffer (25 mM Tris, 50 mM NaCl, 0.5 mM MgCl 2 , 0.1% NP-40, EDTA-free protease inhibitor) and boiled in 2X SDS buffer at 65°C for 10 min.

Statistics and reproducibility
Statistical analyses were performed using GraphPad Prism (8.4.3). All data in bar graphs are shown as mean ± SEM. N represents the number of biological replicates. All experiments were performed at least 3 times independently. For comparison between two groups, datasets were analyzed by Two-tailed unpaired Student's T-test. For experiments with a two-factorial design, multiple comparisons were analyzed by two-way ANOVA to determine the statistical signi cance between groups based on one variable. Differences in EE were calculated with CalR using ANCOVA with body weight as a covariate. The signi cance of the correlations between gene expression with BMI and HOMA values were calculated using Spearman's correlation test. Values of p < 0.05 were considered as signi cantly different.

Schematics
Schematics were prepared using Adobe Illustrator or Biorender.com with publication permissions.

Declarations
Data availability RNA-Seq data reported in this paper have been deposited in NCBI SRA database (BioProject PRJNA727566). Human study data were deposited in NCBI gene expression omnibus (GEO) with the accession number GSE25402 and retrieved from GEO (GSE70353).

Code availability
Adipocyte size from CD-fed mice was assayed using Cell Pro ler with an in-housed modi ed pipeline described previously 30 , code will be made freely available upon request. RalA de ciency in WAT increases energy expenditure and mitochondrial oxidative phosphorylation. a, Regression plot of energy expenditure (EE) measured in HFD-fed Rala f/f and Rala AKO mice (n = 5-8) during dark phase. ANCOVA test was performed using body weight (BW) as a covariate, group effect p = 0.0391. b, c, Immunoblot (b) and quanti cation (c) of OXPHOS proteins in iWAT of HFD mice (n = 10-13). d, e, Plasma non-esteri ed fatty acid (NEFA, d) and TG (e) levels in HFD-fed Rala f/f and Rala AKO mice (n = 10-13). f, Basal oxygen consumption rate (OCR) in mitochondria measured by Seahorse. Mitochondrial fractions were isolated from primary mature adipocytes in iWAT or eWAT of HFD-fed Rala f/f and Rala AKO mice (n = 4-5). The data (c-f) are shown as the mean ± SEM, *p < 0.05, **p < 0.01, ***p < 0.001 by unpaired t-test (c-f).