Leptin, a hormone secreted by adipocytes, exhibits therapeutic potential for the treatment of type 1 diabetes (T1D). Protein tyrosine phosphatase 1B (PTP1B) is a key enzyme that negatively regulates leptin receptor signaling. Here, the role of PTP1B in the treatment of T1D was investigated using PTP1B-deficient (knockout [KO]) mice and a PTP1B inhibitor. T1D wild-type (WT) mice induced by streptozotocin showed marked hyperglycemia compared with non-T1D WT mice. KO mice displayed significantly improved glucose metabolism equivalent to non-T1D WT mice, whereas peripheral or central administration of leptin partially improved glucose metabolism in T1D WT mice. Peripheral combination therapy of leptin and a PTP1B inhibitor in T1D WT mice improved glucose metabolism to the same level as non-T1D WT mice. Leptin was shown to act on the arcuate nucleus in the hypothalamus to suppress gluconeogenesis in liver and enhance glucose uptake in both brown adipose tissue and soleus muscle through the sympathetic nervous system. These effects were enhanced by PTP1B deficiency. Thus, treatment of T1D with leptin, PTP1B deficiency, or a PTP1B inhibitor was shown to enhance leptin activity in the hypothalamus to improve glucose metabolism. These findings suggest a potential alternative therapy for T1D.

Type 1 diabetes (T1D) is caused by insulin deficiency following destruction of insulin-producing pancreatic β-cells (1). The increased worldwide prevalence of T1D has fostered the development of alternative insulin-independent treatments (2). Insulin dependence is mainly observed in T1D. Treatment options for these patients is limited to insulin because most oral hypoglycemic drugs have no proven efficacy (3). Consequently, insulin remains the first-line drug for the treatment of patients with T1D. However, insulin therapy for T1D can cause adverse events, such as life-threatening hypoglycemia, production of insulin antibodies, or onset of insulin allergy (46). Moreover, insulin therapy places a significant burden on patients in terms of self-administering daily subcutaneous injections. Long-term insulin therapy can cause ectopic fat accumulation, leading to arteriosclerosis and insulin resistance in liver and skeletal muscle (7,8). Consequently, alternative therapeutic strategies for the tratment of T1D are needed.

Leptin is a hormone secreted by adipocytes that acts on neurons within the central nervous system, suppressing food intake and increasing energy expenditure to maintain energy homeostasis (9). Studies have demonstrated that intracerebroventricular (ICV) administration of leptin can normalize glucose metabolism in T1D rodent models (10,11). Specifically, leptin acts on hypothalamic neurons to suppress gluconeogenesis in the liver and enhance glucose uptake in brown adipose tissue (BAT) and skeletal muscle, resulting in lowered blood glucose levels (12,13).

Compared with administration of insulin, leptin treatment is associated with a lower risk of hypoglycemia and inhibits fat synthesis (1316). Like insulin, leptin is effective in the treatment of ketoacidosis, a fatal disease state in T1D (17) and, thus, could be an alternative treatment for T1D. However, peripheral administration of leptin into rodents or patients with T1D has failed to normalize blood glucose levels (11,17,18). Because central administration of leptin to patients in daily clinical practice is problematic, such treatment will necessitate the development of innovative procedures. Here, we focused on protein tyrosine phosphatase 1B (PTP1B), a key regulator of leptin receptor signaling in the hypothalamus.

PTP1B negatively regulates leptin receptor signaling through direct dephosphorylation of Jak2 (19). Moreover, PTP1B is an established metabolic regulator in vivo, a deficiency of which enhances leptin receptor signaling in the hypothalamus. Previous studies have shown that PTP1B deficiency in the whole body, brain, pro-opiomelanocortin (POMC), or agouti-related peptide (AgRP) neurons protect against obesity induced by a high-fat diet through enhanced leptin receptor signaling (2023). We reasoned that inhibition of PTP1B might augment leptin activity in the hypothalamus to normalize blood glucose levels. Under these conditions, peripheral administration of leptin could be effective.

In this study, we determined the role of PTP1B in leptin activity for the treatment of T1D using PTP1B-deficient mice. Combined peripheral administration of leptin and an anti-PTP1B drug was also investigated using T1D wild-type (WT) mouse models.

Mice

All animal procedures were approved by the animal care and use committee of Nagoya University Graduate School of Medicine and performed in accordance with institutional guidelines. Mice were housed in a temperature-controlled facility with a 12-h light/dark cycle and ad libitum access to food and water. Age-matched littermates were used in all experiments. To generate streptozotocin (STZ)-induced hyperglycemia, 10–12-week-old mice received a single intraperitoneal injection of STZ (150 μg/g body weight) (12,24).

Mice With Deletion of PTP1B

Global PTP1B-deficient (PTP1B−/− [knockout (KO)]) mice (22) were produced by intercrossing male and female heterozygotes (25). PTP1B+/+ littermates were used as WT mice. Ptpn1loxP/loxP mice (provided by Dr. Kendra Bence, Pfizer, Cambridge, MA) were generated as described previously (21). POMC-Cre transgene mice express functional Cre-recombinase only in POMC cells (hereafter termed POMC-Cre mice) (26), whereas AgRP-IRES-Cre transgene mice express functional Cre-recombinase only in AgRP cells (hereafter termed AgRP-Cre mice) (27). R26GRR mice (ROSA26 Cre-reporter knockin C57BL/6N mice) exhibit green emission before and red emission after Cre-mediated recombination (28). DNA was extracted from the tail of each experimental mouse at the age of 10 days. DNA was subjected to genotyping analyses by PCR with KOD FX (TOYOBO Biotech) DNA polymerase and a set of oligonucleotide primers. PCR was performed on a SimpliAmp Thermal Cycler (Applied Biosystems). Amplification conditions were as follows: 5-min denaturation at 95°C followed by 30 cycles at 95°C for 30 s, 56°C for 20 s, and 72°C for 60 s with a final extension at 72°C for 7 min. Primer sequences used for genotyping of Ptpn1loxP/loxP, POMC-Cre, AgRP-Cre, and R26GRR mice were as follows: PTP1B forward 5′-TGCTCACTCACCCTGCTACAA-3′, reverse 5′-GAAATGGCTCACTCCTACTGG-3′; POMC-Cre forward 5′-TGGCTCAATGTCCTTCCTGG-3′, reverse 5′-CACATAAGCTGCATCGTTAAG-3′ (to detect WT gene) or 5′-GAGATATCTTTAACCCTGATC-3′ (to detect transgene); AgRP-Cre forward 5′-GCTTCTTCAATGCCTTTTGC-3′, reverse 5′-GTGTGTGGTTCCAGCATGAC-3′ (to detect WT gene) or 5′-AGGAACTGCTTCCTTCACGA-3′ (to detect transgene); and R26GRR forward 5′-AAAGTCGCTCTGAGTTGTTAT-3′, reverse 5′-CTTGTACAGCTCGTCCATGCCGAG-3′. Primer sequences used to detect the occurrence of spurious germline deletions were as follows: Ptpn1 Δ/Δ forward 5′-GTGGTGCCTGCAAGAGAACTGAC-3′, reverse 5′-GAAATGGCTCACTCCTACTGG-3′ and GAPDH (as internal control) forward 5′-AACGACCCCTTCATTGAC-3′, reverse 5′-TCCACGACATACTCAGCAC-3′. All Ptpn1loxP/loxP, POMC-Cre, AgRP-Cre, and R26GRR mice were backcrossed >10 generations onto a C57BL/6J background.

To generate POMC or AgRP neuron-specific PTP1B-deficient (P-KO or A-KO, respectively) mice, Ptpn1loxP/loxP mice were crossed with POMC-Cre or AgRP-Cre mice as described previously (21,23). Ptpn1+/loxP POMC-Cre or Ptpn1+/loxP AgRP-Cre mice were then crossed with Ptpn1loxP/loxP mice to yield Ptpn1loxP/loxP POMC-Cre and Ptpn1loxP/loxP AgRP-Cre mice, respectively, along with littermate controls (hereafter termed P-WT or A-WT mice, respectively). Deletion of the Ptp1b allele in P-KO mice was only detected in DNA extracts from the hypothalamus, pituitary, and hindbrain (including the nucleus of the solitary tract) (Supplementary Fig. 5A). Similarly, deletions of the Ptp1b allele in A-KO mice were detected in DNA extracts from the hypothalamus (Supplementary Fig. 5B). In contrast, no recombined alleles were detected in P-WT and A-WT mice (Supplementary Fig. 5A and B).

Detecting Recombination of Floxed Alleles

DNA from tissue samples was genotyped by PCR. Tissue samples of male P-KO mice (cerebral cortex, hypothalamus, pituitary, hindbrain, liver, muscle, white adipose tissue) and male A-KO mice (cerebral cortex, hypothalamus, lung, adrenal gland, liver, muscle, white adipose tissue) at the age of 8 weeks were collected and digested in 50 mmol/L NaOH for 10 min at 95°C. The digestion process was stopped by the addition of 1 mol/L Tris-HCl (pH 8.0). Digested tissue samples were centrifuged (13,000g for 10 min), and the clarified supernatants were recovered. Extracted DNA was subjected to genotyping analyses by PCR as described above.

Immunohistochemistry

STAT3 phosphorylation was detected by immunohistochemistry. For detection of STAT3phosphorylation, 10-week-old male mice were fasted overnight (12 h) and injected intraperitoneally with mouse recombinant leptin (1 μg/g body weight) as described previously (21). Mice were anesthetized 45 min after injection of leptin and perfused with 4% paraformaldehyde in PBS (pH 7.4). After fixation, brains were removed and immersed in the same fixative for 2 h at 4°C. The brains were transferred into PBS containing 10–20% sucrose at 4°C for cryoprotection. Brains were embedded in Tissue-Tek OCT Compound (Sakura Finetek) and stored at −80°C before sectioning. Sections (thickness 20 μm) of brain were cut on a cryostat at −20°C, thawed, and mounted on Superfrost Plus Microscope Slides (Matsunami). The sections were stored at −80°C until immunohistochemistry. Immunohistochemistry of STAT3 phosphorylation was performed as described previously (21). Sections were washed with 1× PBS before and between successive blocking steps with 1.0% H2O2/1.0% NaOH in H2O for 20 min, 0.3% glycine for 10 min, 0.03% sodium dodecyl sulfate for 10 min, and finally 0.2% sodium azide/3% normal goat serum/0.25% Triton X-100 in PBS for 1 h at room temperature. The sections were then incubated with anti-STAT3 phosphorylation antibody (1:100 dilution; Cell Signaling Technology) and diluted in azide blocking solution overnight at 4°C. Sections were washed in 1× PBS and incubated in sodium azide–free blocking solution containing Alexa Fluor 488–conjugated anti-rabbit IgG secondary antibody (1:500; Invitrogen) for 1 h at room temperature. After washing in 1× PBS, sections were placed on slides and air dried, and coverslips were positioned with VECTASHIELD (Vector Laboratories). All fluorescently stained sections were examined with a confocal laser microscope (TiEA1R; Nikon Instech) and viewed using NIS-Elements software (Nikon Instech). Cells labeled for phosphorylated STAT3 were counted bilaterally in a blinded fashion. For analysis, we used four to six mice in each group for the staining, and the mean values of three or four serial sections from each mouse were calculated. The sections included the arcuate nucleus (−1.70 to −1.82 mm from bregma based on coordinates in the brain atlas [29]). The sample numbers in figure legends indicate the number of animals analyzed.

For detection of PTP1B, 10-week-old male mice were anesthetized and perfused with 4% paraformaldehyde in PBS (pH 7.4). After fixation, brain sections were prepared using the same protocol described above. The sections were subjected to antigen retrieval in citrate acid buffer (10 mmol/L sodium citrate, 0.05% v/v Tween 20 [pH 6.0]) at 85°C for 20 min and incubated at room temperature for 1 h in blocking buffer (0.1 mol/L phosphate buffer, 0.2% v/v Triton X-100, 10% BSA [Wako] in PBS) and then overnight at 4°C in goat anti-PTP1B (1/100 dilution, AF3954; R&D Systems) in blocking buffer. After washing with PBS, sections were treated with Alexa Fluor 488–conjugated anti-goat IgG secondary antibody (1:500 dilution; Invitrogen) in blocking buffer for 2 h at room temperature. After washing in 1× PBS, the sections were placed on slides and air dried, and coverslips were positioned with Vectashield (Vector Labs). All fluorescently stained sections were examined with a TiEA1R confocal laser microscope and viewed using NIS-Elements software (Research Resource Identifier SCR_014329).

For immunostaining analysis, P-WT, P-KO, A-WT, and A-KO mice were first crossed with R26GRR mice in which green fluorescence changed to red fluorescence in Cre-recombined cells (28). The tdsRed-positive cells were detected in the arcuate of the hypothalamus in all mice (Supplementary Fig. 5C and D). Immunostaining with PTP1B revealed that PTP1B was expressed in POMC neurons of the hypothalamus in P-WT mice but was rarely detected in P-KO mice (Supplementary Fig. 5C). Similarly, immunostaining with PTP1B revealed that PTP1B was expressed in AgRP neurons of the hypothalamus in A-WT mice but was rarely detected in A-KO mice (Supplementary Fig. 5D).

Brain Collection for Immunohistochemistry

Male mice (9 weeks of age) were anesthetized and transcardially perfused with a cold fixative containing 4% paraformaldehyde in PBS (pH 7.4). After fixation, brains were removed and immersed in the same fixative for 2 h at 4°C. The brains were transferred into PBS containing 10–20% sucrose at 4°C for cryoprotection. Brains were embedded in Tissue-Tek OCT Compound and stored at −80°C before sectioning. Sections (thickness 20 μm) of brain were cut on a cryostat at −20°C, thawed, and mounted on Superfrost Plus Microscope Slides. The sections were stored at −80°C until immunohistochemistry.

Body Weight and Food Intake

Mice were fed a chow diet (24.9% protein, 4.6% fat, and 70.5% carbohydrate, CE-2; CLEA Japan). Body weight was monitored every 2 days for 10 days from the first week after STZ administration at the beginning of the light cycle (between 9:00 and 10:00 a.m.) when the mice were in a fed state. Two weeks after STZ administration, the amount of food consumed was assessed over a 3-day period using a multifeeder (Shin Factory).

Blood Glucose and Serum Levels of Insulin and Leptin

Blood glucose, serum insulin, and leptin were measured. Blood glucose was assayed every 2 days for 10 days from the first week after STZ administration at the beginning of the light cycle (9:00–10:00 a.m.). Tail vein blood was analyzed using a glucometer that can measure blood glucose up to 1,000 mg/dL (30). Blood was collected on day 10 through submandibular bleeding at the beginning of the light cycle (9:00–10:00 a.m.), and serum was separated by centrifugation. β-Hydroxybutyrate was measured using a FreeStyle Libre continuous glucose monitor and FreeStyle Optium β-Ketone Test Strips (Abbott Diabetes Care). Serum levels of insulin and leptin were measured by ELISA as previously described (31).

Measurement of Serum Glucagon and Corticosterone Levels

Blood was collected through submandibular bleeding. Serum was separated by centrifugation at 3,500g for 10 min. Glucagon and corticosterone levels were measured by ELISA (Funakoshi and Assay Pro).

Glucose and Pyruvate Tolerance Tests

Glucose tolerance tests (GTTs) and pyruvate tolerance tests (PTTs) were performed. For GTTs, mice were fasted for 16 h in the untreated group, for 4 h in the peripherally treated leptin group, and for 2 h in the centrally treated leptin group before intraperitoneal injection of 10% d-glucose at 2 mg/g body weight. Glucose was assayed as previously described (31). In the leptin-treated group, serum insulin levels in tail vein blood samples collected in heparin-coated capillary tubes (HEMATO-CLAD; Drummond) were also measured by ELISA as previously described (31) at 0 and 30 min. For the PTT, mice in the leptin-treated group were fasted for 2 h before intraperitoneal injection of pyruvate dissolved in saline at 2 mg/g body weight (32). Blood was collected as previously described (31,32).

2-Deoxyglucose Uptake

Uptake of 2-deoxyglucose (2DG) was measured in insulin-responsive tissues. For the leptin-treated group, 2DG uptake into interscapular BAT and soleus muscle on day 10 from the first week after STZ administration was measured using a previously described protocol (33). Food was withdrawn for 4 h in the peripherally treated leptin group and 2 h in the centrally treated leptin group before the experiment. Insulin (0.75 units/kg) was injected intraperitoneally 10 min before injection of 2DG. After intraperitoneal injection of 2DG (5 μmol/L), blood glucose was maintained at a constant level by administration of 150 μL of 20% glucose. For measurement of 2DG content, tissue samples (10 mg for soleus muscle and BAT) were homogenized in 500 μL of 10 mmol/L Tris-HCl (pH 8.1), heated to 95°C for 10 min, and then centrifuged at 15,000 rpm for 10 min at 4°C. The supernatants were diluted (20× for soleus muscle, 10× for BAT) with 10 mmol/L Tris-HCl (pH 8.1) and assayed for 2DG content according to the 2DG Uptake Measurement Kit (Funakoshi Co., Inc.). Propranolol (10 mg/kg; Sigma-Aldrich) or saline was injected twice intraperitoneally, 15 min and 3 h before performing this procedure, as previously described (34). The 2DG uptake was also measured in BAT and soleus muscle without using insulin in T1D WT and KO mice. After 12 h fasting, 1 μg of leptin was centrally injected, and 3 h 40 min later, 2DG was administered intraperitoneally. The mice were sacrificed 20 min after administration of 2DG, and BAT and soleus muscle were immediately dissected.

Osmotic Pumps

Leptin was administered either peripherally or centrally using an osmotic pump. In the peripheral procedure, T1D WT and TID KO mice were anesthetized using inhaled isoflurane before being implanted with an osmotic pump (ALZET Micro-Osmotic Pump Model 1002; Alza) that delivered either saline or recombinant mouse leptin (20 μg/day; Amgen) subcutaneously for 10 days. In the central procedure, T1D WT and T1D KO mice were implanted with the pump in a similar manner, which was connected to an ICV cannula to enable the continuous central infusion of leptin (0.25 μg/day) or saline for 10 days. An ICV cannula minipump implantation was performed as previously described (13). In brief, a cannula was implanted into the cerebral lateral ventricle (anterior-posterior −0.50 mm, medial-lateral ±1.3 mm, dorsal-ventral −2.3 mm), and an osmotic pump was implanted subcutaneously through a catheter connected to the cannula for ICV infusion. PTP1B inhibitor was subcutaneously injected once daily with vehicle (5 mg/kg/day, DPM-1001; Glixx Laboratories) (35) or saline every day for 10 days into T1D WT mice, which were implanted with an osmotic pump for delivering leptin (20 μg/day).

Denervation of BAT

Denervation of BAT was performed as described previously (36). Briefly, mice were anesthetized using inhaled isoflurane, and after shaving the area above the region of BAT and sterilizing with 70% ethanol, the skin was opened, and BAT was detached from the underlying muscle layer. The five branches of nerve fibers innervating each BAT lobe were exposed and cut. After severing the nerves, the fat pads were rinsed with sterile 0.9% NaCl solution, and the incision was closed using 5-0 suture (A13-50N3; Akiyama Medical Manufacturing).

Determination of mRNA Levels by Quantitative RT-PCR

Total RNA was extracted from samples using TRIzol (Invitrogen) and the RNeasy Kit (QIAGEN). cDNA was synthesized from 100 ng of total RNA using ReverTra Ace qPCR RT Kit (TOYOBO Biotech). Quantitative RT-PCR (qRT-PCR) was performed using Brilliant III Ultra-Fast SYBR Green QPCR Master Mix (Agilent Technologies), and samples were analyzed using the Stratagene Mx3000P qPCR system. The relative mRNA levels of glucose-6-phosphatase (G6PC) and phosphoenolpyruvate carboxykinase (PEPCK) in liver were assessed by qRT-PCR using GAPDH as an internal control. qRT-PCR was performed and then relative mRNA expression levels calculated using a comparative Ct method as described previously (31). The sequences of primers used for qRT-PCR were as follows: G6PC forward 5′-TGCAAGGGAGAACTGAGCAA-3′, reverse 5′-GGACCAAGGAAGCCACAATG-3′; PEPCK forward 5′-GAGATAGCGGCACAAT-3′, reverse 5′-TTCAGAGACTATGCGGTG-3′; and GAPDH forward 5′-AGGTCGGTGTGAACGGATTTG-3′, reverse 5′-TGTAGACCATGTAGTTGAGGTCA-3′.

Statistical Analyses

The statistical significance of the differences between groups was analyzed by either unpaired t test, two-way ANOVA with repeated measures, or two-way factorial ANOVA followed by Bonferroni post hoc test using SPSS Statistics version 25 (Research and Resource Identifier: SCR_002865; IBM Corporation). Results are expressed as mean ± SEM, and differences were considered significant at P < 0.05.

Data and Resource Availability

The data sets generated and/or analyzed during the current study are available from the corresponding author upon reasonable request. No applicable resources were generated during the current study.

PTP1B Deficiency Enhances the Activity of Leptin Administered Peripherally to Improve Glucose Metabolism in T1D

Global PTP1B-deficient mice (PTP1B−/− [KO]) were generated, and their PTP1B+/+ littermates were used as WT mice. Experimental details are given in the Supplementary Material. Mice were given a single intraperitoneal injection of STZ or vehicle, and 1 week later, blood glucose and body weight were measured at the end of the dark cycle. Hereafter, T1D WT or T1D KO refer to mice after injection of STZ for generating T1D mice. T1D WT mice had blood glucose levels of >400 mg/dL and blood insulin levels of <0.1 ng/mL (Supplementary Fig. 1A and B).

To establish whether PTP1B deficiency enhances leptin activity in T1D to improve glucose metabolism, T1D WT and T1D KO male mice were subcutaneously implanted with osmotic pumps that delivered either saline or recombinant mouse leptin. In accordance with previous reports, the dosage of leptin was 20 μg/day (3739). Systemic leptin treatment decreased blood glucose in T1D WT mice, although values were significantly higher than in control mice (Fig. 1A). By contrast, treatment of T1D KO mice with leptin decreased blood glucose to almost the same levels as control mice (Fig. 1A). Body weight of T1D mice given vehicle was significantly decreased compared with WT mice, but there were no significant differences in body weight between WT mice and leptin-treated T1D mice (Fig. 1B). There were no significant differences in body weight between leptin-treated T1D WT and T1D KO mice (Fig. 1B). Food intake was significantly higher in the T1D group compared with the non-T1D group, while there were significant differences in food intake between T1D mice given vehicle and leptin-treated T1D mice (Fig. 1C). Moreover, there were no significant differences in food intake between leptin-treated T1D WT mice and T1D KO mice (Fig. 1C). Serum leptin concentration was significantly lower in T1D mice given vehicle compared with control mice, while the corresponding concentration in leptin-treated T1D mice was significantly higher than that of control mice (Fig. 1D). Plasma levels of β-hydroxybutyrate in T1D mice given vehicle were significantly higher than in control mice and leptin-treated T1D mice, while there were no significant differences in β-hydroxybutyrate between control mice and leptin-treated T1D mice (Fig. 1E).

Figure 1

PTP1B deficiency enhances the activity of leptin administered peripherally to improve glucose metabolism in T1D. AE: T1D WT and PTP1B-deficient (KO) mice were subcutaneously (SC) treated with vehicle or a high dose of leptin (20 μg/day). Various parameters were then measured 1 week later, including blood glucose, body weight, food intake, serum leptin concentration, and plasma levels of β-hydroxybutyrate. F and G: A GTT was performed 10 days after leptin administration. The AUC of GTT and serum insulin concentrations at 0 and 30 min are shown. H and I: Uptake of 2DG into interscapular BAT and soleus muscle 10 days after leptin administration was also measured. Data are mean ± SE (n = 10–12 mice/group, all data from male mice). The statistical significance of the differences between groups was analyzed by two-way ANOVA with repeated measures (A, B, and GTT in F), two-way factorial ANOVA (CE and G) followed by Bonferroni post hoc test, and unpaired t test (AUC in F, H, and I).

Figure 1

PTP1B deficiency enhances the activity of leptin administered peripherally to improve glucose metabolism in T1D. AE: T1D WT and PTP1B-deficient (KO) mice were subcutaneously (SC) treated with vehicle or a high dose of leptin (20 μg/day). Various parameters were then measured 1 week later, including blood glucose, body weight, food intake, serum leptin concentration, and plasma levels of β-hydroxybutyrate. F and G: A GTT was performed 10 days after leptin administration. The AUC of GTT and serum insulin concentrations at 0 and 30 min are shown. H and I: Uptake of 2DG into interscapular BAT and soleus muscle 10 days after leptin administration was also measured. Data are mean ± SE (n = 10–12 mice/group, all data from male mice). The statistical significance of the differences between groups was analyzed by two-way ANOVA with repeated measures (A, B, and GTT in F), two-way factorial ANOVA (CE and G) followed by Bonferroni post hoc test, and unpaired t test (AUC in F, H, and I).

Close modal

Leptin treatment of T1D KO mice significantly improved glucose tolerance as measured by GTT compared with leptin-treated T1D WT mice, and the area under curve (AUC) of GTT in leptin-treated T1D KO mice was significantly lower than in leptin-treated T1D WT mice (Fig. 1F). Serum insulin levels were <0.1 ng/mL at 0 and 30 min in both leptin-treated T1D WT mice and T1D KO mice (Fig. 1G). To evaluate glucose utilization in peripheral tissues, 2DG uptake was measured in insulin target organs of leptin-treated T1D WT and T1D KO mice. The ratio of 2DG uptake was significantly higher in leptin-treated T1D KO mice compared with leptin-treated T1D WT mice in interscapular BAT and soleus muscle, suggesting that PTP1B deficiency enhanced the effect of leptin in correcting glucose utilization in peripheral tissues (Fig. 1H and I). Additionally, leptin treatment significantly decreased serum glucagon and corticosterone levels in T1D KO and WT mice, but there were no significant differences in both hormone levels between genotypes (Supplementary Fig. 1C and D).

PTP1B Deficiency Enhances Leptin Activity in the Brain to Improve Glucose Metabolism in T1D

To determine the target organ of leptin, T1D WT and T1D KO male mice were implanted with a pump connected to an ICV cannula to facilitate continuous central infusion of either leptin (0.25 μg/day) or saline. In T1D KO mice, central administration of leptin decreased blood glucose to almost the same levels as control mice (Fig. 2A), whereas peripheral administration of leptin with an osmotic minipump did not alter blood glucose levels in T1D WT and KO mice (Supplementary Fig. 2A). These observations suggest that the target organs of leptin were not peripheral tissues. No significant differences were observed in body weight, food intake, and serum leptin concentration between T1D WT and T1D KO mice (Fig. 2B–D). However, T1D KO mice displayed significantly enhanced glucose tolerance as measured by GTT. Specifically, the AUC of GTT in leptin-treated T1D WT mice was significantly higher than in leptin-treated T1D KO mice (Fig. 2E). Serum insulin levels were <0.1 ng/mL at 0 and 30 min in both leptin-treated T1D WT and T1D KO mice (Fig. 2F). To evaluate gluconeogenesis, leptin-treated T1D WT and T1D KO mice were subjected to a PTT 3 weeks after STZ administration. Leptin-treated T1D KO mice displayed significantly decreased blood glucose compared with leptin-treated T1D WT mice, and the AUC of PTT in leptin-treated T1D KO mice was significantly lower than in leptin-treated T1D WT mice, suggesting that gluconeogenesis was suppressed in the former (Fig. 2G). The mRNA levels of G6PC and PEPCK in liver, which encode enzymes involved in gluconeogenesis, were significantly decreased in leptin-treated T1D KO mice compared with leptin-treated T1D WT mice (Supplementary Fig. 2B and C). In addition, the ratio of 2DG uptake was significantly higher in leptin-treated T1D KO mice compared with leptin-treated T1D WT mice in BAT and soleus muscle, suggesting that glucose utilization was enhanced in leptin-treated T1D KO mice (Fig. 2H and I). There were no significant differences in blood glucose levels and serum insulin levels between genotypes immediately before dissection (Supplementary Fig. 2D and E). Collectively, these results demonstrate that enhanced leptin receptor signaling in the brain elicited by PTP1B deficiency normalizes blood glucose levels through suppressing gluconeogenesis in the liver and enhancing glucose utilization in BAT and soleus muscle.

Figure 2

PTP1B deficiency enhances leptin activity in the brain to improve glucose metabolism in T1D. AD: T1D WT and PTP1B-deficient (KO) mice were given a continuous central infusion of either a low dose of leptin (0.25 μg/day) or saline (0.25 μg/day) for 10 days. Various parameters were then measured, including blood glucose, body weight, food intake, and serum leptin concentration. E and F: GTT was performed 7 days after initiation of central leptin administration. The AUC of GTT and serum insulin concentrations at 0 and 30 min are shown. G: A PTT was performed 10 days after initiating central leptin administration. The blood glucose levels and AUC for GTT tests are shown. H and I: Uptake of 2DG into interscapular BAT and soleus muscle in T1D WT and T1D KO mice was also measured 10 days after initiating the central administration of leptin. Data are mean ± SE (all data from male mice). The statistical significance of the differences between groups was analyzed by two-way ANOVA with repeated measures (A, B, GTT in E, and PTT in G), two-way factorial ANOVA (F) followed by Bonferroni post hoc test, and unpaired t test (C, AUC in E and G, and H and I).

Figure 2

PTP1B deficiency enhances leptin activity in the brain to improve glucose metabolism in T1D. AD: T1D WT and PTP1B-deficient (KO) mice were given a continuous central infusion of either a low dose of leptin (0.25 μg/day) or saline (0.25 μg/day) for 10 days. Various parameters were then measured, including blood glucose, body weight, food intake, and serum leptin concentration. E and F: GTT was performed 7 days after initiation of central leptin administration. The AUC of GTT and serum insulin concentrations at 0 and 30 min are shown. G: A PTT was performed 10 days after initiating central leptin administration. The blood glucose levels and AUC for GTT tests are shown. H and I: Uptake of 2DG into interscapular BAT and soleus muscle in T1D WT and T1D KO mice was also measured 10 days after initiating the central administration of leptin. Data are mean ± SE (all data from male mice). The statistical significance of the differences between groups was analyzed by two-way ANOVA with repeated measures (A, B, GTT in E, and PTT in G), two-way factorial ANOVA (F) followed by Bonferroni post hoc test, and unpaired t test (C, AUC in E and G, and H and I).

Close modal

Cotreatment of an Anti-PTP1B Drug and Leptin Potentially Improves Glucose Metabolism in T1D

To further delineate the contribution of PTP1B with the effect of leptin on glucose homeostasis, pharmacological modulation was performed using a PTP1B inhibitor, DPM-1001. Based on previous reports, the dosage of PTP1B inhibitor was set at 5 mg/kg/day (35). Systemic leptin or PTP1B inhibitor treatment decreased blood glucose in T1D mice compared with T1D mice given vehicle, although blood glucose values were still significantly higher than in control mice (Fig. 3A). By contrast, T1D mice treated with leptin and PTP1B inhibitor (lep + 1bi) displayed markedly decreased blood glucose levels akin to control mice (Fig. 3A). The body weight of T1D mice given vehicle significantly decreased compared with control mice, although there were no significant differences in body weight among control, leptin-treated T1D, PTP1B inhibitor–treated T1D, and lep + 1bi–treated T1D mice (Fig. 3B). Food intake was significantly higher in the T1D group compared with control mice, while there were significant differences in food intake between T1D mice given vehicle and leptin-treated and/or PTP1B inhibitor–treated T1D mice (Fig. 3C). Moreover, there were no significant differences in food intake among leptin-treated, PTP1B inhibitor–treated, and lep + 1bi–treated T1D mice (Fig. 3C). GTT showed that lep + 1bi–treated T1D mice displayed significantly improved glucose tolerance. Specifically, blood glucose levels of lep + 1bi–treated T1D mice were almost the same as those of control mice at each measurement point (Fig. 3D). The AUC of GTT in T1D mice given vehicle was significantly higher than those of leptin-treated, PTP1B inhibitor–treated, lep + 1bi–treated T1D and control mice, whereas AUC in leptin-treated and PTP1B inhibitor–treated T1D mice were significantly higher than those of lep + 1bi–treated T1D and control mice (Fig. 3D). Lep + 1bi–treated T1D and control mice showed no significant differences in AUC (Fig. 3D).

Figure 3

Anti-PTP1B drug treatment enhances leptin activity to improve glucose metabolism in T1D. AC: WT mice and T1D WT mice were subcutaneously (SC) treated with vehicle, PTP1B inhibitor (1bi), leptin, or lep + 1bi for 7 days. Various parameters were then measured, including blood glucose, body weight, and food intake. D: A GTT was performed 7 days after administration of vehicle, 1bi, leptin, or lep + 1bi. The AUC of GTT is shown. Data are mean ± SE (n = 10–12 mice/group, data from all male mice). The statistical significance of the differences between groups was analyzed by two-way ANOVA with repeated measures (A, B, and GTT in D) or two-way factorial ANOVA (C and AUC in D) followed by Bonferroni post hoc test.

Figure 3

Anti-PTP1B drug treatment enhances leptin activity to improve glucose metabolism in T1D. AC: WT mice and T1D WT mice were subcutaneously (SC) treated with vehicle, PTP1B inhibitor (1bi), leptin, or lep + 1bi for 7 days. Various parameters were then measured, including blood glucose, body weight, and food intake. D: A GTT was performed 7 days after administration of vehicle, 1bi, leptin, or lep + 1bi. The AUC of GTT is shown. Data are mean ± SE (n = 10–12 mice/group, data from all male mice). The statistical significance of the differences between groups was analyzed by two-way ANOVA with repeated measures (A, B, and GTT in D) or two-way factorial ANOVA (C and AUC in D) followed by Bonferroni post hoc test.

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In female WT mice, systemic leptin and/or PTP1B inhibitor treatment decreased blood glucose in T1D WT compared with T1D WT mice given vehicle, although blood glucose values in PTP1B inhibitor–treated were significantly higher than those of lep + 1bi–treated T1D and control mice (Supplementary Fig. 3A). There were no significant differences in blood glucose values between leptin-treated and PTP1B inhibitor–treated male mice. Of note, there were no significant differences in blood glucose values among leptin-treated, lep + 1bi–treated T1D, and control mice (Supplementary Fig. 3A). The body weight of T1D mice was significantly decreased compared with control mice, but there were no significant differences in body weight among vehicle-treated, leptin-treated, PTP1B inhibitor–treated, and lep + 1bi–treated T1D mice (Supplementary Fig. 3B). Food intake was significantly higher for given vehicle-treated T1D mice compared with leptin-treated, PTP1B inhibitor–treated, lep + 1bi–treated T1D and control mice, while food intake in leptin-treated and PTP1B inhibitor–treated T1D mice was significantly higher than for control mice (Supplementary Fig. 3C). There were no significant differences in food intake between lep + 1bi–treated T1D and control mice (Supplementary Fig. 3C). GTT showed that lep + 1bi–treated T1D mice displayed significantly improved glucose tolerance. Specifically, the blood glucose levels of lep + 1bi–treated T1D mice were almost the same as that of control mice at each measurement point (Supplementary Fig. 3D). The AUC of GTT in T1D mice given vehicle was significantly higher than leptin-treated, PTP1B inhibitor–treated, and lep + 1bi–treated T1D and control mice, whereas the AUC in leptin-treated and PTP1B inhibitor–treated T1D mice was significantly higher than in lep + 1bi–treated T1D and control mice (Supplementary Fig. 3D). Lep + 1bi–treated T1D and control mice showed no significant differences in AUC (Supplementary Fig. 3D). Taken together, these data demonstrate that cotreatment with an anti-PTP1B drug and leptin can potentially improve glucose metabolism in both male and female T1D mice.

PTP1B Deficiency Enhances Leptin Activity to Increase Glucose Uptake in Peripheral Tissues Through β-Adrenergic Receptor Signaling in the Sympathetic Nervous System

Although PTP1B is expressed throughout the brain, it is highly enriched in the arcuate nucleus of the hypothalamus, which is an important site of leptin action (22). We previously showed that intraperitoneal injection of leptin significantly increased STAT3 phosphorylation in the hypothalamus in PTP1B-deficient compared with WT mice (25). Here, immunohistochemistry was used to verify increased STAT3 phosphorylation in the arcuate nucleus of the hypothalamus in T1D KO mice and T1D WT mice treated with PTP1B inhibitor after peripheral administration of leptin. Moreover, leptin significantly increased STAT3 phosphorylation in the hypothalamus of both T1D KO mice and T1D WT mice treated with PTP1B inhibitor compared with T1D WT mice (Supplementary Fig. 4A–C).

Previous studies showed that leptin in the hypothalamus enhances glucose uptake into specific peripheral tissues through β1- and β2-adrenergic receptor signaling (34,40). Here, we adopted a pharmacological approach to evaluate how β-adrenergic receptor signaling contributes to enhanced glucose uptake. T1D WT and T1D KO mice were fitted with an osmotic pump to deliver recombinant mouse leptin (20 μg/day). The mice were subcutaneously treated with vehicle or propranolol, an inhibitor of β-adrenergic receptor signaling, before measuring 2DG uptake in BAT and soleus muscle. Administration of propranolol significantly decreased the ratio of 2DG uptake in BAT and soleus muscle of T1D WT mice (Fig. 4A and B). In T1D KO mice, the ratio of 2DG uptake was also significantly decreased after propranolol treatment compared with vehicle (Fig. 4A and B). These findings, together with the results shown in Fig. 2, suggest that PTP1B deficiency enhances the action of central leptin to increase glucose uptake into peripheral tissues, at least in part, through β-adrenergic receptor signaling.

Figure 4

PTP1B deficiency enhances leptin activity in the brain to increase glucose uptake in peripheral tissues through β-adrenergic receptor signaling in the sympathetic nervous system. A and B: T1D WT and T1D KO mice were treated with subcutaneous leptin (20 μg/day) without (−) and with (+) propranolol. Uptake of 2DG into interscapular BAT and soleus muscle was measured. C: T1D WT and T1D KO mice were also treated with a central infusion of leptin (0.25 μg/day) without (−) and with (+) denervation of BAT. Uptake of 2DG into BAT was measured. Data are mean ± SE (n = 6–8 mice/group, data from all male mice). The statistical significance of the differences between groups was analyzed by two-way factorial ANOVA followed by Bonferroni post hoc test.

Figure 4

PTP1B deficiency enhances leptin activity in the brain to increase glucose uptake in peripheral tissues through β-adrenergic receptor signaling in the sympathetic nervous system. A and B: T1D WT and T1D KO mice were treated with subcutaneous leptin (20 μg/day) without (−) and with (+) propranolol. Uptake of 2DG into interscapular BAT and soleus muscle was measured. C: T1D WT and T1D KO mice were also treated with a central infusion of leptin (0.25 μg/day) without (−) and with (+) denervation of BAT. Uptake of 2DG into BAT was measured. Data are mean ± SE (n = 6–8 mice/group, data from all male mice). The statistical significance of the differences between groups was analyzed by two-way factorial ANOVA followed by Bonferroni post hoc test.

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To further investigate this mechanism, we denervated the BAT of T1D WT and T1D KO mice before evaluating 2DG uptake. Briefly, nerve fibers innervating each BAT lobe were severed and the mice implanted with a subcutaneous osmotic pump connected to an ICV cannula to enable continuous central infusion of saline for 2 weeks. After a recovery period of 1 week, mice received a single intraperitoneal injection of STZ to generate T1D. One week later, the mice were anesthetized using isoflurane for subcutaneous replacement with another osmotic pump to deliver leptin (0.25 μg/day). After treatment with leptin for 1 week, 2DG uptake was measured. The denervation of BAT significantly decreased the ratio of 2DG uptake in BAT of both genotypes (Fig. 4C), indicating that the absence of innervation of BAT crucially affected glucose metabolism in T1D mice. Moreover, we confirmed 2DG uptake in BAT and soleus muscle without using insulin, which was significantly increased in T1D KO mice compared with T1D WT mice by central leptin injection, and the denervation of BAT significantly decreased 2DG uptake in BAT for both genotypes (Supplementary Fig. 4D and E).

PTP1B in POMC Neurons of the Hypothalamus Regulates Glucose Metabolism in T1D

To investigate the target neurons of leptin in the hypothalamus, P-KO or A-KO mice were generated along with their littermate controls (P-WT or A-WT mice, respectively). Experimental details are given in the Supplementary Material, and the corresponding data are shown in Supplementary Fig. 5A–D.

After generation of T1D mice by injection of STZ, blood glucose and body weight were measured and GTT performed for male T1D P-KO, P-WT, A-KO, and A-WT mice. Systemic leptin treatment decreased blood glucose in T1D P-WT mice, although these values were significantly higher than those of control mice (Fig. 5A). In contrast, leptin treatment of T1D P-KO mice decreased blood glucose to almost the same levels as the control mice (Fig. 5A). The body weight of T1D mice given vehicle were significantly decreased compared with the control mice (Fig. 5B). By contrast, there was no significant difference in body weight between the control mice and leptin-treated T1D mice or between the leptin-treated T1D P-WT and T1D P-KO mice (Fig. 5B). Leptin treatment of T1D P-KO mice significantly improved glucose tolerance as measured by GTT compared with leptin-treated T1D P-WT mice. Specifically, the AUC of GTT in leptin-treated T1D P-KO mice was significantly lower than that of leptin-treated T1D P-WT mice (Fig. 5C). Moreover, systemic leptin treatment significantly increased STAT3 phosphorylation in the hypothalamus of T1D P-KO mice compared with T1D WT mice (Supplementary Fig. 4B and C).

Figure 5

POMC neuron-specific PTP1B deficiency enhances leptin activity to improve glucose metabolism in T1D. AC: P-WT and P-KO mice were subcutaneously (SC) treated with vehicle or a high dose of leptin (20 μg/day). Various parameters were then measured, including blood glucose, body weight, and GTT along with the AUC. DF: A-KO mice were subcutaneously treated with vehicle or a high dose of leptin (20 μg/day). Blood glucose, body weight, and GTT and AUC are shown. Data are mean ± SE (n = 10–12 mice/group, data from all male mice). The statistical significance of the differences between groups was analyzed by two-way factorial ANOVA (A, B, D, E, and GTT in C and F) followed by Bonferroni post hoc test and unpaired t test (AUC in C and F).

Figure 5

POMC neuron-specific PTP1B deficiency enhances leptin activity to improve glucose metabolism in T1D. AC: P-WT and P-KO mice were subcutaneously (SC) treated with vehicle or a high dose of leptin (20 μg/day). Various parameters were then measured, including blood glucose, body weight, and GTT along with the AUC. DF: A-KO mice were subcutaneously treated with vehicle or a high dose of leptin (20 μg/day). Blood glucose, body weight, and GTT and AUC are shown. Data are mean ± SE (n = 10–12 mice/group, data from all male mice). The statistical significance of the differences between groups was analyzed by two-way factorial ANOVA (A, B, D, E, and GTT in C and F) followed by Bonferroni post hoc test and unpaired t test (AUC in C and F).

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Systemic leptin treatment also decreased blood glucose in T1D A-WT and T1D A-KO mice, although these values were significantly higher than those of control mice (Fig. 5D). No significant differences in blood glucose values were observed between genotypes. Moreover, body weights of A-WT and A-KO mice were similar to those of P-WT and P-KO mice (Fig. 5E). Leptin treatment of T1D A-KO mice tended to improve glucose tolerance as measured by GTT compared with that of leptin-treated T1D A-WT mice, although the differences did not reach statistical significance (Fig. 5F).

This study investigated whether enhanced leptin activity in the hypothalamus improves glucose homeostasis in T1D. Our findings demonstrate that 1) peripheral administration of leptin, or central administration of a low dose of leptin, significantly improves glucose metabolism in T1D KO mice compared with T1D WT mice; 2) peripheral combination therapy of leptin and PTP1B inhibitor in T1D WT mice restores glucose metabolism to the same level as control mice; 3) improvements of glucose metabolism are due, at least in part, to β-adrenergic receptor signaling in the sympathetic nervous system; and 4) POMC neurons in the hypothalamus are crucial for lep + 1bi therapy in T1D mice. These findings suggest an alternative therapy for T1D.

Leptin ameliorated hyperglycemia in T1D WT mice more potently when administered centrally than when administered peripherally, consistent with previous reports (11). Central administration of a low dose of leptin markedly improved hyperglycemia in T1D mice, whereas peripheral administration had no such affect. These observations suggest that the target organ of leptin activity is the brain. Previous studies have shown that the effect of leptin in the hypothalamus is crucial for improving glucose metabolism in T1D (12,13,41). Moreover, PTP1B deficiency is reported to enhance hypothalamic leptin receptor signaling by increasing STAT3 phosphorylation (25). The PTP1B inhibitor used in the current study was previously reported to cross the blood-brain barrier, and its peripheral administration alongside leptin increased STAT3 phosphorylation in the hypothalamus (35). Here, we showed enhanced leptin receptor signaling in the hypothalamus of KO mice, which was associated with increased STAT3 phosphorylation due to the lack of PTP1B (Supplementary Fig. 4A–C). P-KO, but not A-KO, mice showed improved glucose metabolism when leptin was administered peripherally under T1D conditions (Fig. 5A and C). Our data are consistent with previous reports (13,42). However, other studies reported the importance of AgRP neurons in mediating the leptin effect (43,44), which is inconsistent with our findings. These discrepancies appear to arise from differences in the genetic model used. Specifically, PTP1B regulates not only leptin receptor signaling but also a variety of intracellular signaling pathways, including inflammatory signals (19,25). The dose of centrally administered leptin was only 0.25 μg/day, <25% of the amount used in previous experiments (11,41). This dosage of leptin improved glucose metabolism in T1D KO mice to almost an equivalent level as control mice but not to T1D WT mice (Fig. 2A and E). Collectively, our results suggest that enhanced leptin receptor signaling in the hypothalamus due to the lack of PTP1B contributed to an improvement of glucose metabolism in T1D upon leptin administration.

The glucose-lowering effects were not due to insulin receptor signaling because insulin concentrations in T1D mice were undetectable when fed (Supplementary Fig. 1B) or at 0 and 30 min during GTT (Figs. 1G and 2F). The amount of food ingested was significantly elevated in the T1D group compared with the non-T1D group (Fig. 3C), as reported previously (37). There were no significant differences in food intake between T1D KO and T1D WT mice independent of leptin treatment (Figs. 1C and 2C). In addition, body weight did not differ between T1D KO and T1D WT mice independent of leptin treatment (Figs. 1B and 2B). Collectively, our results suggest that the improvement in glucose metabolism of PTP1B KO mice was due to enhanced leptin receptor signaling in the hypothalamus, independent of the effects of insulin receptor signaling, or changes in food intake or body weight.

We conclude that PTP1B deficiency enhances leptin activity in the hypothalamus, thereby suppressing gluconeogenesis in the liver (Fig. 2G and Supplementary Fig. 2B and C) and promoting glucose uptake in BAT and soleus muscle (Figs. 1H and I and 2H and I). Several independent lines of evidence indicate that glucose metabolism through the hypothalamus is involved in the activation of the sympathetic nervous system (34,40). Here, glucose uptake in BAT and soleus muscle elicited by leptin treatment significantly diminished upon administration of propranolol (a β-blocker) as reported previously (34,40) (Fig. 4A and B). However, in this regard, some reports have cast doubt on the β-receptor–mediated effects (13,45). The apparent discrepancy is likely to arise from differences in the drug and mouse model used in the respective studies. Further investigations are required to fully resolve this issue. Here, glucose uptake in BAT following leptin treatment was significantly diminished by denervating BAT (34,40) (Fig. 4C). Collectively, our data suggest that upon increased leptin receptor signaling due to the lack of PTP1B, leptin enhances glucose uptake in BAT and soleus muscle through the sympathetic nervous system.

In this study, serum leptin levels increased two- to threefold after administration of leptin (Fig. 1D). However, no significant differences in food intake or body weight were observed between T1D WT and T1D KO mice. We believe that this increase in serum leptin levels is within an acceptable range for clinical use considering that serum leptin levels of obese people are 5–10-fold higher than in healthy individuals (4648). A previous study reported that ICV administration of 10 μg of leptin resulted in an increase of mean arterial pressure (49). In the current study, only 0.25 μg of leptin was used for central injection (Fig. 2A), which was sufficient to improve glucose metabolism in T1D under PTP1B deficiency. We found that the leptin-treated group lost less weight compared with the group not treated with leptin in our T1D model (Figs. 1B and 2B). Elevated levels of β-hydroxybutyrate in T1D WT mice became normalized to the control level by combined administration of leptin and PTP1B inhibitor (Fig. 1E), suggesting that glucose was appropriately metabolized. An anti-PTP1B drug has been successfully tested in phase II clinical trials as a therapeutic agent for patients with type 2 diabetes (50). These findings indicate that combination therapy of the pharmacological inhibition of PTP1B and leptin may be valuable in treating patients with T1D.

Although hyperglycemia in patients with T1D cannot be normalized by peripheral administration of leptin alone (18), it was confirmed in this study that glucose metabolism could be normalized by combined use of a leptin enhancer (PTP1B inhibitor). We consider this to be the most significant finding from our study. Coadministration of leptin and anti-PTP1B drugs may be useful for lean patients with T1D with poor glycemic control. In such cases, administration of these drugs may reduce the risk of ketoacidosis as well as the dosage and/or frequency of insulin treatment. Several parameters still need to be established (e.g., appropriate BMI, PTP1B inhibitor/leptin dosage during insulin treatment).

A scheme outlining the proposed mechanism of leptin is shown in Fig. 6. Our findings suggest that targeting leptin-sensitive pathways could be a therapeutic option for T1D. Moreover, anti-PTP1B drugs might display therapeutic utility for patients with and without T1D.

Figure 6

Scheme outlining the proposed mechanism by which leptin acts on the brain to decrease gluconeogenesis in liver and improve glucose utilization in BAT and soleus muscle through an autonomic nervous pathway. The mechanism is shown for T1D mice (A) or under PTP1B deficiency or use of a PTP1B inhibitor (B). The activity of leptin is enhanced in the brain, resulting in improved glucose metabolism in T1D mice (B).

Figure 6

Scheme outlining the proposed mechanism by which leptin acts on the brain to decrease gluconeogenesis in liver and improve glucose utilization in BAT and soleus muscle through an autonomic nervous pathway. The mechanism is shown for T1D mice (A) or under PTP1B deficiency or use of a PTP1B inhibitor (B). The activity of leptin is enhanced in the brain, resulting in improved glucose metabolism in T1D mice (B).

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This article contains supplementary material online at https://doi.org/10.2337/figshare.20103176.

Acknowledgments. The authors thank Michiko Yamada (Department of Endocrinology and Diabetes, Nagoya University Graduate School of Medicine, Nagoya, Japan) for helpful technical assistance.

Funding. This work was supported by a Grant-in-Aid for Young Scientists from the Japanese Society for Promotion of Science (2618K16225) and a grant from the Japan IDDM Network.

Duality of Interest. Y.I. received grants from Sanwa Kagaku Kenkyusho, Kowa Pharmaceutical, Kureha Corporation, and Nipro Corporation and personal fees from Astellas Pharma and Ono Pharmaceutical Company. T.O. received personal fees from MSD K.K. H.T. received grants from MSD K.K. S.I. received personal fees from Ono Pharmaceutical Company, Bristol-Myers Squibb, MSD K.K., and Chugai Pharmaceutical Co., Ltd. H.A. received grants from Ono Pharmaceutical Company, MSD K.K., and Chugai Pharmaceutical Co. Ltd. and personal fees from Ono Pharmaceutical Company, Bristol-Myers Squibb, and MSD K.K. No other potential conflicts of interest relevant to this article were reported.

Author Contributions. Y.I. and R.B. designed the research, analyzed the data, and prepared the manuscript. Y.I., R.S., H.Y., K.T., A.M., M.S., T.T., H.Ko., H.Ki., and R.B. performed in vivo studies and supervised the study. Y.I. and T.T. performed biochemical and immunohistochemical experiments. T.K., D.H., S.I., H.S., T.O., and H.A. supervised the study and helped to prepare the manuscript. All authors read and approved the final manuscript. R.B. 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.

Prior Presentation. Parts of this study were presented in poster and oral form at the 90th Annual Meeting of the Japan Endocrine Society, Kobe, Japan, 24–26 November 2017; 61st Annual Meeting of the Japan Diabetes Society, Tokyo, Japan, 24–26 May 2018; 45th Annual Meeting of Japan Neuroendocrine Society, Okinawa, Japan, 30 June–3 July 2022; and 48th Annual Meeting of Society for Neuroscience, San Diego, CA, 3–7 November 2018. Per the request of the Japan IDDM Network, part of the research outline was announced at a press conference, published in Japanese newspapers (Asahi, Chunichi, Mainichi, and Yomiuri), and reported in the media (NHK, tv-Aichi, and Kyodo).

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