Ire1α in Pomc Neurons Is Required for Thermogenesis and Glycemia

Obesity and diabetes remains one of the leading causes of death in developed countries. Importantly, a variety of metabolic abnormalities have been linked to endoplasmic reticulum (ER) stress and the unfolded protein response (UPR) (1–3). In particular, genetically obese mice exhibit ER stress and activation of the UPR in the periphery as well as in the central nervous system (CNS) (4,5). Moreover, nutrient-dependent regulation of the UPR has been suggested to be a major contributor to diet-induced metabolic dysfunction within the periphery as well as the brain (4,6–8).

The inositol-requiring enzyme 1/X-box binding protein 1 (Ire1-Xbp1) pathway is the most conserved of the three branches of the UPR (9–13). Activated Ire1 exhibits endoribonuclease activity, which cleaves its primary target, the mRNA encoding Xbp1, and generates a potent transcription factor that stimulates expression of chaperones and components of the ER-associated protein degradation pathway (9). Although global deletion of Ire1 or Xbp1 in mice produces an embryonic lethal phenotype (14–16), neuronal deletion of Xbp1 creates severe ER stress in the hypothalamus, blocks leptin action, and generates leptin resistance in mice (4). (Note that Xbp1 deletion causes Ire1 hyperactivation, which leads to cell death.) Similarly, chemical activation of ER stress in pro-opiomelanocortin (Pomc) neurons results in acute leptin and insulin resistance via a Ptp1b- and Socs3-dependent mechanism (17). Importantly, constitutive activation of the spliced isoform of Xbp1 (Xbp1s) in Pomc neurons mimics a postprandial state, resulting in improved body weight and decreased glucose and insulin levels (17).

Although driving the Ire1-Xbp1s pathway in arcuate Pomc neurons is sufficient to improve body weight and glucose homeostasis, the requirement of this highly conserved pathway in Pomc neurons to regulate metabolism remains to be established. We therefore tested the hypothesis that Ire1 signaling in Pomc neurons is necessary for proper energy balance and glucose metabolism.

Pomc-cre (25) and Ire1α^fe/fe (20) mice were group housed (1–5 mice per cage) in a barrier facility at 23°C unless otherwise noted. Male mice in a C57Bl/6 background were used for all studies. Mice were provided a Harlan Teklad 2016 chow diet and water ad libitum unless otherwise noted. Mice were placed on a high-fat/high-sucrose (HFHS; Research Diets D12331) diet at 8 weeks of age, if applicable, and HFHS diet was removed and refilled weekly. Mice losing >10% of body weight during the acclimation period for metabolic cage studies (see below) were not studied. Body composition was measured using nuclear magnetic resonance (Bruker minispec).

Experiments were performed in a temperature-controlled room containing 36 TSE metabolic cages maintained by University of Texas Southwestern (UTSW) Animal Resources personnel. One week prior to study, mice were singly housed to acclimate to new housing. Three days prior to study, mice were transported to the room containing the metabolic cages to acclimate to a new environment. HFHS diet, if applicable, was also introduced at the beginning of this acclimation period. After 3 days acclimation, cages were connected to the TSE system for a total of 5 days. Days 2–4 were used for data analyses.

Mice were housed in cold chambers maintained at 6°C by UTSW Animal Resources personnel.

Sixteen-week-old male mice were anesthetized and transcardially perfused with a modified ice-cold artificial cerebrospinal fluid (ACSF) (126 mmol/L NaCl, 2.8 mmol/L KCl, 1.2 mmol/L MgCl₂, 2.5 mmol/L CaCl₂, 1.25 mmol/L NaH₂PO₄, 26 mmol/L NaHCO₃, and 5 mmol/L glucose), in which an equiosmolar amount of sucrose was substituted for NaCl. The mice were then decapitated, and the entire brain was removed and immediately submerged in ice-cold, carbogen-saturated (95% O₂ and 5% CO₂) ACSF. Coronal sections (250 μm) were cut with a Leica VT1000S vibratome and then incubated in oxygenated ACSF with tunicamycin (35 μmol/L) for 6 h. The slices were fixed by 10% neural-buffered formalin for 4 h at room temperature and cryopreserved in 30% sucrose at 4°C overnight. Brain slices were then sectioned into 25-μm coronal sections and stored in cryoprotectant at −20°C until use. Brain sections were rinsed with PBS five times for 5 min and incubated with primary antibody (Ireα, NB100-2323, 1:100 diluted in 3% normal donkey serum, PBS and tween–azide) at 4°C overnight. Sections were then rinsed in PBS and incubated with secondary antibody (1:200 diluted in 3% normal donkey serum, PBS and tween) at room temperature for 2 h. Cells were visualized using a Zeiss microscope by an observer that was blinded to the condition or genotype of the mice.

Protein was extracted from brown adipose tissue (BAT) by using RIPA buffer (Boston BioProducts) supplemented with complete protease inhibitor cocktail (Roche). For Western blot analyses, 80 µg protein was subjected to SDS-PAGE under reducing conditions, transferred, and blotted with the anti-UCP1 antibody (ab10983; Abcam).

After an overnight fast, 10- to 14-week-old male mice received intraperitoneal injections of 1.5 g/kg d-glucose. Blood glucose was measured from tail blood using a glucometer at serial time points as indicated in figures. A separate cohort of mice was used for glucose-stimulated insulin secretion (GSIS). Blood was collected at 0 and 30 min after glucose administration. Blood samples were centrifuged, and serum samples were collected from the supernatants. Insulin levels were measured using an ELISA kit (Crystal Chem Inc., Downers Grove, IL) according to the manufacturer’s instruction.

After a 4-h fast to empty the stomach, 10- to 14-week-old male mice received intraperitoneal injections of insulin (1.2 units/kg for chow-fed mice or 1.4 units/kg for high-fat diet [HFD]–fed mice). Blood glucose was measured from tail blood as described above.

After a 6-h fast to empty the stomach, 10- to 14-week-old male mice received intraperitoneal injections of 2 mg/kg pyruvate. Blood glucose was measured from tail blood as described above.

Tissues were dissected and fixed in formalin for 48 h at 4°C followed by 50% ethanol. BAT histology was performed with assistance from the UTSW Histology Core. Inguinal white adipose tissue (iWAT) was performed with assistance from the Harvard Histology Core.

Whole-cell patch-clamp recordings from Pomc-hrGFP (humanized, Renilla reniformis green fluorescent protein) neurons maintained in hypothalamic slice preparations and data analysis were performed as previously described (17,18). In brief, 4- to 16-week-old male mice were anesthetized and transcardially perfused with a modified ice-cold ACSF (described below), in which an equiosmolar amount of sucrose was substituted for NaCl. The mice were then decapitated, and the entire brain was removed and immediately submerged in ice-cold, carbogen-saturated (95% O₂ and 5% CO₂) ACSF (126 mmol/L NaCl, 2.8 mmol/L KCl, 1.2 mmol/L MgCl₂, 2.5 mmol/L CaCl₂, 1.25 mmol/L NaH₂PO₄, 26 mmol/L NaHCO₃, and 5 mmol/L glucose). Coronal sections (250 μm) were cut with a Leica VT1000S vibratome and then incubated in oxygenated ACSF at room temperature for at least 1 h before recording. Slices were transferred to the recording chamber and allowed to equilibrate for 10–20 min before recording. The slices were bathed in oxygenated ACSF (32°C–34°C) at a flow rate of ∼2 mL/min.

The pipette solution for whole-cell recording was modified to include an intracellular dye (Alexa Fluor 594 or Alexa Fluor 350) for whole-cell recording: 120 mmol/L K-gluconate, 10 mmol/L KCl, 10 mmol/L HEPES, 5 mmol/L EGTA, 1 mmol/L CaCl₂, 1 mmol/L MgCl₂, and 2 mmol/L MgATP and either 0.03 mmol/L Alexa Fluor 594 or Alexa Fluor 350 hydrazide dye (pH 7.3). Epifluorescence was briefly used to target fluorescent cells, at which time the light source was switched to infrared differential interference contrast imaging to obtain the whole-cell recording (Zeiss Axioskop FS2 Plus equipped with a fixed stage and a QuantEM:512SC electron-multiplying charge-coupled device camera). Electrophysiological signals were recorded using an Axopatch 700B amplifier (Molecular Devices), low-pass filtered at 2–5 kHz, and analyzed offline on a PC with pCLAMP programs (Molecular Devices). Recording electrodes had resistances of 2.5–5 mol/LΩ when filled with the K-gluconate internal solution. Input resistance was assessed by measuring voltage deflection at the end of the response to a hyperpolarizing rectangular current pulse step (500 ms of −10 to −50 pA).

Leptin (100 nmol/L, provided by A.F. Parlow through the National Hormone and Peptide Program, Harbor-UCLA Medical Center, Torrance, CA) or insulin (50 nmol/L, Humulin R 100 units/mL; Eli Lilly and Company) was added to the ACSF for specific experiments. Solutions containing leptin or insulin were typically perfused for 2–4 min. A drug effect was required to be associated temporally with peptide application, and the response had to be stable within a few minutes. A neuron was considered depolarized or hyperpolarized if a change in membrane potential was at least 2 mV in amplitude.

A coronal slice between bregma –1.22 and –2.70 mm was made from age- and weight-matched mice, and then the arcuate nucleus was microdissected with a scalpel under a microscope. Total RNA was extracted from tissues with TRIzol reagent (Invitrogen) according to the manufacturer’s instructions. Total RNA (1 μg) was converted into first-strand cDNA with oligo(dT) primers as described by the manufacturer (Clontech). PCR was performed in an Mx3000P Q-PCR system (Stratagene) with specific primers and SYBR Green PCR Master Mix (Stratagene). The relative abundance of mRNAs was standardized with 18S mRNA as the invariant control.

Experiments were done in conscious, chronically catheterized mice using previously described techniques (17). After a 5-day recovery, food was removed on day of experiment at 0800 h to begin a 5-h fast. [3,4-¹³C₂]glucose (Cambridge Isotopes) was infused beginning at t = –120 min to calculate glucose turnover. Humulin R (2.5 mU/kg/min) was then infused at t = 0 min to induce hyperinsulinemia. Blood samples from the cut tail were taken every 10 min, and dextrose (50%) was infused as needed to maintain target blood glucose levels (∼150 mg/dL).

Statistical analysis was performed using GraphPad. All data were evaluated using a two-tailed Student t test, with a P value of <0.05 considered significant. In all instances, data are presented as mean ± SEM. Body weight curves were compared using a linear regression analysis. Degrees of freedom (DF) for t statistics are marked as t_(DF). For glucose tolerance test, insulin tolerance test, and pyruvate tolerance test (PTT), data were analyzed consistent with current recommendations (19,20).

Pre-established criteria for excluding data points were data two SDs outside the mean or any data obtained from mice that died or lost >10% of body weight due to metabolic cage acclimation or clamp studies.

The downstream target of Ire1α, Xbp1s, improves energy expenditure and glucose metabolism in the periphery as well as the CNS (4,5,17,21,22). In particular, we recently demonstrated that overexpression of Xbp1s in Pomc neurons alone was sufficient to protect against diet-induced obesity as well as glucose and insulin intolerance (17). In order to test the requirement of the Ire1-Xbp1s arm of the UPR in Pomc neurons to regulate body weight, we generated mice that were deficient for Ire1α specifically in Pomc neurons, Pomc-cre::Ire1α^fe/fe mice (23). Fluorescent immunohistochemistry demonstrated that pIre1α can be detected in the arcuate nucleus (Fig. 1). The cells expressing pIre1α include those identified as Pomc neurons as visible by tdtomato expression (Fig. 1A). Notably, Pomc neurons deficient for Ire1α (Pomc neurons from Pomc-cre::Ire1α^fe/fe mice) exhibit a reduced expression of pIre1α (Fig. 1B). Similar data were obtained with quantitative PCR (qPCR) from arcuate punches (Fig. 1C). As expected, Pomc-cre::Ire1α^fe/fe mice displayed diminished mRNA for Xbp1s as well as the putative target genes of Xbp1s (Erdj4 and GalE) in the arcuate nucleus (for Xbp1s: t₍₁₁₎ = 2.507, P < 0.05; for Erdj4: t₍₄₎ = 6.645, P < 0.05; for GalE: t₍₄₎ = 3.891, P < 0.05) (Fig. 1C). However, we failed to detect alterations of ER stress markers and putative downstream targets for Xbp1s outside the arcuate nucleus between control and Pomc-cre::Ire1α^fe/fe mice (Fig. 1D). These data support a Pomc-specific downregulation of Ire1α expression in mice selectively deficient for Ire1α in Pomc-cre::Ire1α^fe/fe mice.

On a chow diet, Pomc-cre::Ire1α^fe/fe mice displayed similar body weight to littermate controls (Fig. 2A). However when fed an HFD, male Pomc-cre::Ire1α^fe/fe mice exhibit an age-dependent increased body weight compared with wild-type mice (Fig. 2B), which was reflected by increased fat mass (t₍₁₀₎ = 3.171, P < 0.05) (Fig. 2C). Age- and weight-matched Pomc-cre::Ire1α^fe/fe male mice had increased caloric intake and were hypometabolic, as demonstrated by significantly increased food intake and decreased energy expenditure (Fig. 2D–J). Pomc-cre::Ire1α^fe/fe mice were also less sensitive to acute leptin-induced hypophagia when compared with littermate controls at 1 h after refeeding (Fig. 2K).

Several studies have suggested that melanocortin neurons regulate intrascapular BAT (iBAT), which is a key component of adaptive thermogenesis (17,24,25). Components of adaptive “nonshivering” thermogenesis include coordinated neuroendocrine responses to the energy demands of an HFD as well as cold exposure. Even though chow-fed Pomc-cre::Ire1α^fe/fe mice failed to exhibit alterations in body weight, we found that transcripts associated with heat production were reduced in BAT from Pomc-cre::Ire1α^fe/fe mice (Fig. 3A). Similar to previous reports, control mice fed an HFD displayed increased markers of heat production in BAT (Fig. 3A). However, this was not the case in BAT from Pomc-cre::Ire1α^fe/fe littermates (Fig. 3A). Moreover, analyses of BAT histology after 12–24 h at 6°C demonstrated impaired thermogenic responses in Pomc-cre::Ire1α^fe/fe mice (Fig. 3B). In addition, iWAT in cold-exposed Pomc-cre::Ire1α^fe/fe mice failed to show evidence of “browning” based on histology (Fig. 3B) or protein expression (Fig. 3C). Together, these data suggest that Ire1α in Pomc neurons is essential for adaptive thermogenesis including recruitment of BAT and browning of iWAT.

Along with the systemic effects on whole-body energy expenditure and body weight, deficiency of Ire1α in Pomc neurons also leads to alterations in glucose metabolism. In particular, Pomc-cre::Ire1α^fe/fe mice showed impaired glucose tolerance independent of diet when compared with littermate controls (P < 0.05) (Fig. 4A and D). Pomc-cre::Ire1α^fe/fe mice were also insulin intolerant (P < 0.05) (Fig. 4B and E). Although serum insulin was only modestly increased, GSIS was increased 30 min after a glucose load (t₍₂₀₎ = 2.244, P < 0.05) (Supplementary Fig. 1).

Mammals maintain euglycemia within a very tight range, largely dependent upon glucose production and/or secretion from the liver. The livers of chow-fed Pomc-cre::Ire1α^fe/fe mice displayed elevated gluconeogenic markers (for Foxo1: t₍₆₎ = 2.883, P < 0.05; for HNF4α: t₍₆₎ = 5.562, P < 0.05; for Pcx: t₍₆₎ = 3.993, P < 0.05; for G6pc: t₍₆₎ = 4.648, P < 0.05; for Pepck: t₍₆₎ = 3.160, P < 0.05) (Fig. 4G). Moreover, when pyruvate was provided as a fuel source (PTT), chow-fed Pomc-cre::Ire1α^fe/fe mice displayed increased serum glucose levels (Fig. 4C and F), supportive of increased glucose production capacity within the liver.

We next performed hyperinsulinemic-euglycemic clamps to assess whether insulin sensitivity was altered in chow-fed Pomc-cre::Ire1α^fe/fe mice versus wild-type littermates of similar body weight (Fig. 5A). Plasma insulin was elevated similarly in both groups (Fig. 5B) and blood glucose was clamped at target levels (∼150 mg/dL) (Fig. 5C) using an exogenous glucose infusion rate (GIR) (Fig. 5D). The required GIR during the steady-state period (t = 80–120 min) was lower in Pomc-cre::Ire1α^fe/fe mice, indicating impaired insulin sensitivity (Fig. 5D). This was due to impaired insulin-mediated suppression of endogenous glucose appearance (endoR_a) (Fig. 5E). There were no significant differences in insulin-stimulated glucose disappearance (R_d) (Fig. 5F). Together these data suggest that Pomc neurons selectively deficient for Ire1α are linked to impaired insulin sensitivity, which may contribute to hyperglycemia.

Upregulation of the Ire1α-Xbp1s pathway in Pomc neurons mimics a postprandial state and improves metabolism via a Ptp1b/Socs3-dependent mechanism (17). Oppositely, the current study suggests that loss of this pathway impairs energy balance and glucose metabolism. However, whether an inverse ER stress–dependent mechanism is also present remains unclear. Deficiency of Ire1α in Pomc neurons induced Bip, Ptp1b, and Socs3 mRNA within the arcuate nucleus (for Bip: t₍₁₃₎ = 2.241, P < 0.05; for Ptp1b: t₍₁₃₎ = 1.924, P > 0.05; for Socs3: t₍₁₃₎ = 2.375, P < 0.05) (Fig. 6), supporting the hypothesis that Pomc-cre::Ire1α^fe/fe mice display increased basal ER stress.

We previously demonstrated that chemical activation of ER stress (6 h) in the arcuate nucleus results in acute leptin and insulin resistance of arcuate Pomc neurons (17). We hypothesized that the amplified basal ER stress present in the arcuate nucleus of Pomc-cre::Ire1α^fe/fe mice may result in an increased susceptibility to ER stress–induced leptin and insulin resistance. Whole-cell recordings were performed on acute hypothalamic slices containing Pomc-hrGFP neurons within the arcuate nucleus (17,26,27). Similar to previous results (17), leptin depolarized while insulin hyperpolarized subsets of Pomc neurons within the arcuate nucleus (Fig. 7A–F and Fig. 8A). Moreover, Pomc neurons from Pomc-cre::Ire1α^fe/fe mice demonstrated analogous acute cellular responses to leptin and insulin (Fig. 7G and Fig. 8B). Pretreatment of arcuate slices with tunicamycin (30 µmol/L, 1 h) from wild-type Pomc-GFP mice failed to alter the acute leptin (t₍₁₃₎ = 0.6021, P > 0.05) (Fig. 7F) and insulin (t₍₁₄₎ = 1.369, P > 0.05) (Fig. 8B) induced changes in membrane potential. Importantly, pretreatment of arcuate slices with tunicamycin (30 µmol/L, 1 h) from Pomc-cre::Ire1α^fe/fe mice was sufficient to abrogate the ability of leptin (t₍₁₄₎ = 5.708, P < 0.05) (Fig. 7G) or insulin (t₍₁₄₎ = 5.677, P < 0.05) (Fig. 8B) to alter the membrane potential of arcuate Pomc neurons. Pretreatment of tunicamycin alone failed to alter the resting membrane potential and excitability of Pomc neurons (Supplementary Table 1). These data support that Pomc neurons deficient for Ire1α are predisposed to the desensitizing effects of ER stress on acute leptin and insulin signaling.

Although the requirement of Ire1α in specific organs and tissues remains incompletely understood, it is of particular interest that deficiency of Ire1α in Pomc neurons has effects on body weight and glucose homeostasis. In particular, we demonstrate that selective deficiency of Ire1α in Pomc neurons (Pomc-cre::Ire1α^fe/fe mice) increased sensitivity to diet-induced obesity in mice. The increased body weight was dependent upon a hypometabolic phenotype (decreased energy expenditure and heat production) and increased food intake. Notably, these responses are phenotypic signatures of impaired leptin action within Pomc neurons (28–31). These data were supported by the reduced efficacy of leptin-induced hypophagia as well as the increased susceptibility of ER stress–induced acute leptin and insulin resistance in mice with Pomc-specific deficiency of Ire1α. The hypometabolic phenotype was also accompanied by decreased markers for thermogenesis in BAT, which suggests that Ire1α deficiency in Pomc neurons impairs compensatory thermogenesis in response to an HFD and may be a contributing factor in increased body weight. Similar to Xbp1s activity in the liver or Pomc neurons (17,22), we also found that deficiency of Ire1α in Pomc neurons alone was sufficient to impair blood glucose and insulin levels (e.g., impaired glucose/insulin tolerance, increased gluconeogenic expression/PTT response, increased GSIS, and reduced hepatic insulin sensitivity) independent of changes in body weight. On a cellular level, Pomc-cre::Ire1α^fe/fe mice exhibited increased basal ER stress in the arcuate nucleus, contributing to an amplified sensitivity to ER stress–dependent leptin and insulin resistance. Together, these data support a model in which loss of Ire1α in Pomc neurons accelerates HFD-induced obesity while at the same time impairing glucose homeostasis.

This study reinforces an important relationship between Ire1α-Xbp1s and Ptp1b/Socs3 in the ER stress–induced acute leptin and insulin resistance of arcuate Pomc neurons. In particular, both Socs3 and Ptp1b are increased within the hypothalamus in a state of excess nutrition or obesity (32–36). Moreover, there is a fundamental role of ER stress in regulating leptin and insulin signaling in hypothalamic Pomc neurons via a Ptp1b/Socs3-dependent mechanism (17). In particular, transcripts of both Ptp1b and Socs3 were also lowered in mice with constitutive activation of Xbp1s in Pomc neurons (17). Pomc neurons selectively deficient for either Ptp1b or Socs3 also demonstrated improved acute leptin and insulin signaling even in the presence of strong activators of ER stress (17). These data were supported in the current study by the inverse finding that Pomc-specific deficiency of Ire1α stimulates Ptp1b and Socs3 in the arcuate nucleus of the hypothalamus. This also correlated with a decreased sensitivity to pharmacological leptin-induced hypophagia as well as at the cellular level an increased susceptibility to ER stress–dependent acute leptin and insulin resistance. These data support a Ptp1b/Socs3-dependent, ER stress–induced acute leptin and insulin resistance in the absence of Ire1-Xbp1 activation.

Another salient finding is the suggestion that the Ire1α-Xbp1s pathway in Pomc neurons is required and sufficient (17) for diet- and cold-induced thermogenesis in iBAT and “browning” of iWAT, respectively. This is topical given that activation of functional iBAT depots in adult humans (37–40) as well as induction of thermogenic programs in WAT (41,42) has recently emerged as a potential therapeutic approach in the treatment of obesity and diabetes (43). However, it is important to clarify that our data cannot firmly establish that defects in iBAT and/or ability to “brown” iWAT contribute to altered energy expenditure, body weight, and/or glycemia in mice. In fact, mice deficient for Ire1α in Pomc neurons exhibited decreased thermogenic markers in BAT independent of changes in body weight on a chow diet. However, when exposed to an HFD, the defects in thermogenesis of BAT were greatly exaggerated. Moreover, transgenic overexpression of the Ire1α-Xbp1s pathway in Pomc neurons increased thermogenesis of BAT and iWAT concomitant with a protection against diet-induced obesity independent of changes in food intake (17). Thus, although it is difficult to determine the independent contribution of energy intake versus energy expenditure resulting in weight gain, the Ire1α-Xbp1s pathway in Pomc neurons consistently modifies thermogenic programs that may underlie deficits associated with obesity.

The current study also highlights an important transient regulation of leptin and potentially insulin signaling within arcuate Pomc neurons. In particular, leptin signaling within Pomc neurons is necessary and sufficient to regulate proper energy and glucose homeostasis (28,29). Although insulin signaling alone in Pomc neurons minimally alters energy balance or glucose metabolism, loss of both leptin and insulin signaling results in larger glycemic dysregulation than loss of either receptor alone (44). Chemical activation of ER stress abrogates leptin and insulin signaling within the periphery as well as the CNS (including Pomc neurons) (17,21,45). Also, increased basal ER stress as occurs with loss of Ire1α in Pomc neurons accelerated leptin and insulin resistance. The acute gating of these signaling pathways (both leptin and insulin via a mechanism similar to a rheostat) may contribute to alterations in energy balance and systemic glucose metabolism and insulin sensitivity. Importantly, both Ire1α and Xbp1s are regulated depending on various metabolic states (17,22,46). Thus, this may highlight an important conserved mechanism that responds to fluctuating energy demands, independent of altered leptin and insulin levels. It is important to note that in addition to the role of Ire1α in the regulation of Xbp1s activity, Ire1α has additional endonuclease and intrinsic kinase activity that ultimately facilitates recovery from ER stress (47–50). Although the current study supports a model demonstrating a requirement of the Ire1α-Xbp1s pathway in the protection from obesity and diabetes, we cannot exclude the possibility that additional activities of Ire1α (endonuclease or intrinsic kinase activity) may contribute to the metabolic phenotypes observed in the current study. Moreover, the age dependence of the body weight phenotype as well as the tunicamycin-dependent blunting of acute leptin and insulin signaling in arcuate Pomc neurons may suggest the requirement of a “trigger” in order to elicit changes in metabolism. Several triggers could be sufficient to drive these processes, including aging and HFD (both of which can precipitate hypothalamic ER stress).

Overall, these findings extend the physiologically important role of ER stress and the UPR in Pomc neurons to regulate sensitivity to humoral signals, ultimately contributing to diet-induced obesity and diabetes. Notably, Ptp1b and Socs3 are reciprocally regulated in response to altered Ire1α-Xbp1s signaling, providing a potential molecular link between ER stress and leptin/insulin resistance. Together, these data highlight multiple molecular targets that may contribute to obesity and diabetes.

Acknowledgments. The authors thank Dr. Joel K. Elmquist (Division of Hypothalamic Research, Department of Internal Medicine, UTSW Medical Center) for kindly providing the Pomc-cre and Pomc-hrGFP mice.

Funding. This work was supported by grants to T.Y. (China Scholarship Council 201406280111), J.S. (National Natural Science Foundation of China 81300689 and 81670783), X.K. (National Institute of Diabetes and Digestive and Kidney Diseases [NIDDK] 5K99DK106550), E.D.B. (NIDDK R01 DK109408), R.J.K. (NIDDK R37 DK042394, R24 DK110973, R01 DK103825, and R01 DK103185), T.L. (American Heart Association 14SDG20370016), and K.W.W. (NIDDK R01 DK100699). This work was also supported by NIDDK PL1 grant DK081182 and National Institutes of Health grant UL1RR024923.

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

Author Contributions. T.Y., Z.D., and J.S. designed and performed all experiments except immunoblotting, analyzed the data, and wrote the manuscript. Y.G. performed electrophysiological experiments, analyzed data, and wrote the manuscript. X.K. analyzed gene expression in adipose depots, performed immunoblotting in BAT, analyzed the data, and reviewed the manuscript. Y.H., Z.H., and Y.X. assisted in performing experiments. Y.C., K.-j.Y., R.-t.W., H.G., H.C., and X.L. designed experiments and edited the manuscript. B.G.F. performed hyperinsulinemic-euglycemic clamps and analyzed the data. E.D.B. designed and performed hyperinsulinemic-euglycemic clamp experiments, analyzed the data, and reviewed the manuscript. R.J.K. designed and developed the IRE1α^fe/fe mouse and edited the manuscript. J.Y. and T.L. supervised development of the mouse models, designed experiments, and edited the manuscript. K.W.W. conceived and designed the study, supervised development of the mouse models, designed experiments, and edited the manuscript. K.W.W. 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.