Central α-Klotho Suppresses NPY/AgRP Neuron Activity and Regulates Metabolism in Mice

By electricdiet / July 2, 2020


Introduction

α-Klotho, a well-documented antiaging protein primarily produced in the kidney and choroid plexus (1,2), has recently been observed to have therapeutic potential in rodent models of metabolic disease (36). Studies show α-klotho promotes lipid oxidation, protects pancreatic β-cells from oxidative damage, increases energy expenditure, and facilitates insulin release (37). Furthermore, circulating α-klotho concentrations are decreased in patients with obesity and diabetes (8), suggesting a possible direct role in the pathophysiology of metabolic disorders. Notably, studies have primarily investigated peripheral α-klotho, which neglects the central function of α-klotho due to its impermeability to the blood-brain barrier (9). The few studies investigating centrally circulating α-klotho demonstrate that α-klotho has antioxidative and anti-inflammatory properties (10), is involved in myelination (11), and can be therapeutic in models of hypertension (12). However, the role of central α-klotho in the regulation of metabolism remains unexplored.

Neuropeptide Y/agouti-related peptide (NPY/AgRP)-expressing neurons are located within the arcuate nucleus (ARC) of the hypothalamus and are critical to homeostatic regulation of metabolism. NPY/AgRP neurons sense nutritional changes in the cerebrospinal fluid (CSF) to regulate feeding behavior (13), energy expenditure (14), and glucose metabolism (1517). However, disordered overactivity of these neurons results in phenotypes resembling diabetes and obesity (13,15). Some circulating factors, such as leptin and insulin, also modulate NPY/AgRP neurons (18,19), but in metabolic disease states, signaling of these hormones is disrupted. Therefore, identification of novel regulators of this neuron population could facilitate the development of therapeutic tools for the prevention and treatment of metabolic disease.

Recent studies have identified several fibroblast growth factor (FGF) hormones that activate FGF receptor (FGFR)–phosphatidylinositol 3 kinase (PI3K) signaling to elicit antidiabetic effects and regulate NPY/AgRP neurons (2025). Interestingly, α-klotho serves as a critical scaffolding protein to the FGF23-FGFR complex to promote FGFR activity (26,27). The current study investigates the novel role of central α-klotho in the regulation of NPY/AgRP neurons and whole-body metabolism via an FGFR/PI3K mechanism.

Research Design and Methods

Cell Culture

Cell culture experiments were performed on immortal hypothalamic GT1-7 cells cultured in high-glucose (4.5 mg/dL) DMEM, 10% FBS, and 1% penicillin-streptomycin. Cells were treated with 3.65 mmol/L α-klotho (R&D Systems) (10,11,28), treated with 100 ng/mL FGF23 (R&D Systems) (28), and/or pretreated with 10 nmol/L FGFR1 antagonist PD173074 (Fisher Scientific) (29) or 50 nmol/L PI3K inhibitor wortmannin (Fisher Scientific) (25). All experiments used vehicle-treated cells as controls.

Experimental Animals

C57BL/6 and B6.Tg(NPY-hrGFP)1Lowl/J (NPY-GFP reporter) mice were cared for in accordance with the National Institutes of Health Guide for the Care and Use of Laboratory Animals, and experimental protocols were approved by Institutional Animal Care and Use Committees of East Carolina University. Mice were housed at 20–22°C with a 12-h light-dark cycle.

High-Fat Diet–Induced Obesity

The 6-week-old male C57BL/6 mice were given ad libitum access to a high-fat diet with a kilocalorie composition of 58%, 25%, and 17% of fat, carbohydrate, and protein, respectively, for 10 weeks (D12331; Research Diets, New Brunswick, NJ) before undergoing intracerebroventricular (ICV) cannulation.

ICV Cannulation

Prior to the procedure, mice were given oral analgesic meloxicam and anesthetized with intraperitoneal (i.p.) injection of ketamine and xylazine. Mice were placed on a stereotaxic device, and a midline incision was made on the head. A hole was drilled (1.0 mm lateral, −0.5 mm posterior, 2.5 mm deep to the bregma), and a cannula was placed into the lateral ventricle (Supplementary Fig. 1). Another hole was drilled, and a screw was placed approximately at the ipsilateral lambdoid structure to aid in supporting the cannula in the skull with 3M carboxylate dental cement. Mice recovered for 14 days before immunohistochemical experiments and 7 days before all other experiments. All ICV treatments were administered via Hamilton syringe as 2.0 μL between 6:30 and 7:30 p.m.

The 12-Day ICV Injection Timeline

In high-fat diet–induced obesity (DIO) mice, central administration of either 2.0 μg recombinant α-klotho (R&D Systems) alone, 25.0 μg PD173074 alone, 25.0 μg PD173074 10 min before 2.0 μg α-klotho (20), or vehicle was performed daily. On day 7, glucose tolerance tests (GTTs) or insulin tolerance tests (ITTs) were performed, and on day 12, a body composition analysis was performed using EchoMRI (Echo Medical Systems, Houston, TX). Mice were then euthanized, and tissues were collected. Food intake and body weight data were analyzed from the first 7 days to prevent confounding effects from additional assays.

Single ICV Injection Timeline

In DIO mice, ICV administration of either 2.0 μg recombinant α-klotho (R&D Systems), 10 ng wortmannin, 10 ng wortmannin 1 h before 2.0 μg α-klotho (30), or vehicle was performed the night before a GTT or ITT. Food intake was measured for 4 days after the injection.

Streptozotocin-Induced Diabetes

The 8- to 9-week-old chow-fed male mice underwent ICV cannulation before receiving 3 days i.p. injection of 100 mg/kg streptozotocin (STZ). The dose was determined by pilot studies yielding consistently elevated fed glucose levels between 250 and 600 mg/dL (Supplementary Fig. 2A). At 7 days after STZ injection, mice received 7 days ICV treatment with either 2.0 μg recombinant α-klotho (n = 10) or vehicle (n = 9). Food intake, body weight, and fed glucose levels were monitored daily. To identify the effects of α-klotho treatment on glucose levels independent from food intake, a pair-fed experiment was performed using the same protocols, and on day 7, fasting glucose levels were measured (n = 8–9/group).

Central α-Klotho Inhibition via ICV Anti–α-Klotho Antibody

The 9-week-old male, chow-fed mice were ICV treated with 1.0 μg anti–α-klotho (ab-α-klotho) (R&D Systems) or vehicle (n = 8/group). Treatments were performed on the evening of day 1 and morning of day 2, while subsequent treatments were performed between 6:30 and 7:30 p.m. for 7 days. GTTs were performed on day 3, and on day 7, mice were euthanized, followed by tissue collection for further assays.

Food Intake Measurements

Food intake was measured daily by weighing food (8–9 g) and subtracting from the total food. Bedding was inspected thoroughly for residual bits of food, which were included in measurements. On day 4 in one cohort of DIO mice, food was removed from cages during the light phase and replenished at the beginning of the dark phase (7:30 p.m.). Food intake was measured at 0.5, 1, 1.5, 2, 3, 4, 8, 14, and 24 h after food reintroduction.

GTTs and ITTs

For GTTs, 20% glucose solution (1.0 g/kg body weight) was i.p. injected after an overnight fast, and for ITTs, 0.6 units/kg insulin were i.p. injected after a 4-h fast. Tail blood samples were collected 15, 30, 60, 90, and 120 min after injections for analysis using a glucose meter (ReliOn Prime Blood Glucose Monitoring System; ARKRAY Inc., Kyoto, Japan). Serum was isolated from clotted blood spun at 4°C and 2,000g for 30 min. Insulin levels were quantified using an insulin ELISA kit (Crystal Chem).

Insulin-Stimulated Signaling

On day 12 of ICV treatment, a weight-matched cohort of DIO mice was i.p. injected with 10 units/kg insulin or saline. At 7 min after the injection, hypothalamus, epididymal adipose tissue (eWAT), liver, and hindlimb skeletal muscle were flash frozen for future Western blot analysis.

Immunohistochemistry

For immunofluorescent analysis, mice were intracardially perfused with PBS followed by 10% formalin before immunohistochemistry was performed as described previously (31). Briefly, brains were sliced into 20-μm coronal sections using a freezing microtome (VT1000 S; Leica, Wetzlar, Germany) and incubated overnight in antibody to phosphorylated ERK (1:500; Cell Signaling Technology, Danvers, MA) or cFOS (1:500; Santa Cruz Biotechnology, Santa Cruz, CA), followed by incubation with Alexa-fluorophore secondary antibody for 2 h. Stains were photographed using an optical microscope (DM6000; Leica), followed by blind analysis using ImageJ. At least three anatomically matched images per mouse were quantified.

Western Blot

Western blot was performed as described previously (31). Briefly, equal protein samples were loaded into a 4–20% HCL gel, transferred to a nitrocellulose membrane, and incubated overnight in 1:500–1:1,000 antibody dilutions in 5% milk with Tris-buffered saline with Tween for phosphorylated (p)ATser473, total AKT, pFOXO1ser256, total FOXO1, pERKthr202/thr204 (Cell Signaling Technology), pIRtyr972 (Invitrogen), and total ERK (Santa Cruz). Image J software was used to quantify mean intensity of equal-area sections representing each sample.

Quantitative PCR

Cell and tissue RNA was extracted by Trizol (Thermo Fisher Scientific, Waltham, MA). The expressions of specific mRNA were analyzed using quantitative real-time PCR (RT-qPCR) (Power SYBR Green PCR Master Mix; Applied Biosystems, Foster City, CA). Reactions were performed in triplicate for each sample, while GAPDH was used as a reference gene for normalization.

Patch Clamp Electrophysiology

We conducted cell-attached voltage clamp recordings of NPY/AgRP neurons as described previously (32,33). Briefly, mice were deeply anesthetized by isoflurane followed by intracardial perfusion with chilled N-methyl-d-glucamine solution and sliced into 200- to 300-μm sections. Slices recovered for an hour in HEPES recovery solution, and recordings were conducted in a normal artificial cerebrospinal fluid (aCSF) bath. For whole cell recordings, gigaohm seals were obtained, and the cells were broken into using negative pressure. Data were sampled at 10 kHz. Current clamp recordings were stabilized for repeated firing under the baseline condition. Equivalent length periods (0.5–5 min) were set within each recording during perfusion of aCSF or α-klotho (3.65 mmol/L). Firing rate (Hz) was calculated by dividing the number of events by the number of seconds. Bath application of terodotoxin (TTX) (1.0 μmol/L) prior to α-klotho treatment was used to determine action potential–independent effects on membrane potential. Voltage clamp whole cell recordings were conducted at a −70 mV holding potential and a high KCl intracellular solution (130 mmol/L KCl, 5 mmol/L CaCl2, 10 mmol/L EGTA, 10 mmol/L HEPES, 2 mmol/L magnesium adenosine triphosphate, 0.5 mmol/L sodium guanosine triphosphate, and 5 mmol/L phosphocreatine) was used. For voltage clamp recordings of miniature inhibitory postsynaptic currents (mIPSCs), glutamatergic blockade was induced using NMDA receptor blocker AP5 (50 μmol/L) and AMPA receptor blocker cyanquixaline (10 nmol/L), followed by α-klotho administration, and then currents were abolished with picrotoxin (100 μmol/L) (34).

Statistical Analysis

Unpaired t tests were used for in vivo mouse experiments to compare differences between groups in food intake, body weight, and body composition. To compare pre-post within-group changes over the course of the experiment, paired t tests were performed. To compare differences at GTT or ITT time points, two-way ANOVA with repeated measures for time and Bonferroni corrections for multiple comparisons were used. Unpaired t tests or one-way ANOVA with Tukey correction for multiple comparisons were used in cell culture experiments when appropriate. Paired t tests or repeated-measures ANOVA with Tukey correction for multiple comparisons was used in patch clamp electrophysiology experiments when appropriate. All analyses were performed using GraphPad Prism statistics software, and a P value <0.05 was considered statistically significant.

Data and Availability Statement

The data sets generated during the current study are available from the corresponding author on reasonable request.

Results

Seven Days of Central Administration of α-Klotho Results in Weight Loss, Suppressed Food Intake, and Improved Glucose Regulation in DIO Mice

A 7-day ICV α-klotho treatment in DIO mice significantly reduced body weight (4.9%) compared with vehicle-treated controls (Fig. 1A and B). These changes were, at least in part, due to decreased food intake both daily (14.8%) and after a daytime food restriction (11.4%) (Fig. 1EH). ICV α-klotho treatment also improved glucose clearance and insulin release during a GTT, as well as insulin sensitivity during an ITT (Fig. 1IN).

Figure 1
Figure 1

Seven days of central administration of α-klotho results in weight loss, suppressed food intake, and improved glucose regulation in DIO mice. A: Body weight. B: Changes in body weight. C: Fat mass. D: Lean mass. E: Average daily food intake. F: Timeline of daily food intake. G: Cumulative food intake after a daytime food restriction. H: Timeline of food intake after a daytime food restriction. I: Blood glucose levels during a GTT. J: Area under the curve (AUC) of the GTT. K: Serum insulin levels 30 min into the GTT. L: Blood glucose during an ITT. M: AUC of the ITT. N: Fasting serum insulin. AN: In 17- to 18-week-old male DIO mice after 7 days ICV α-klotho or vehicle injections (n = 8–13/group). Data represented as mean ± SEM. *P < 0.05 vs. ICV control.

A Single ICV α-Klotho Injection Improves Glucose Clearance and Suppresses Food Intake in DIO Mice

To determine if the effects of central α-klotho on glucose metabolism were independent from changes in body weight, a single ICV α-klotho injection was performed in DIO mice the night before a GTT or ITT. Acute ICV α-klotho treatment improved glucose clearance during a GTT (Fig. 2AC) and even decreased food intake the following day (Fig. 2D). Interestingly, acute ICV α-klotho treatment had no effects on insulin sensitivity (Fig. 2EH). These data suggest central α-klotho regulates glucose metabolism independent from changes in body weight and insulin sensitivity. Supporting this hypothesis, 12 days of ICV α-klotho had no effects on insulin-stimulated signaling in hypothalamus, skeletal muscle, eWAT, or liver in weight-matched DIO mice (Supplementary Fig. 3).

Figure 2
Figure 2

Acute central administration of α-klotho improves glucose clearance and suppresses food intake independent of body weight in DIO mice. A: Blood glucose during a GTT. B: Area under the curve (AUC) of the GTT. C: Body weight before the GTT. D: Daily food intake (including overnight fast before GTT on day 1). E: Blood glucose during an ITT. F: AUC of the ITT. G: Food intake the night before the ITT. H: Body weight before the ITT. AH: In 17- to 18-week-old male DIO mice after a single ICV α-klotho or vehicle injection (n = 7–10/group). Data represented as mean ± SEM. *P < 0.05 vs. ICV control.

To begin to investigate alternative peripheral mechanisms through which central α-klotho improves glucose regulation, basal hepatic gene expression was analyzed. Hepatic PEPCK mRNA was significantly reduced (0.75-fold reduction) in DIO mice treated with α-klotho for 12 days, suggesting attenuated hepatic gluconeogenesis, despite no changes in pyruvate kinase, glucose-6-phsophatase, or glucokinase. α-Klotho–treated mice also had reduced hepatic lipid accumulation and upregulated ACC1 and ACC2 mRNA (Supplementary Fig. 4).

Seven Days of Central α-Klotho Administration Attenuates the Progression of Diabetes in STZ-Treated Mice

The therapeutic potential of α-klotho was also investigated in a model of type 1 diabetes induced by STZ treatment. Similar to DIO mice, ICV α-klotho decreased body weight (5.3%), suppressed food intake (27.8%), and attenuated hyperglycemia (20.2% reduction in fed glucose levels) in STZ-treated mice compared with vehicle-treated controls (Fig. 3AG). Even in pair-fed STZ-treated mice, ICV α-klotho attenuated hyperglycemia and trended to improve fasting glucose levels (Fig. 3KM). These data further demonstrate glucoregulatory and anorexic action of central α-klotho in both type 1 and type 2 diabetes models.

Figure 3
Figure 3

Seven days of central α-klotho administration attenuates the progression of diabetes in STZ-treated mice. A: Body weight. B: Changes in body weight. C: Average daily food intake. D: Timeline of food intake. E: Fed blood glucose levels. F: Timeline of fed blood glucose. G: Change in fed blood glucose levels. AG: In 9- to 10-week-old, STZ-treated, ad libitum fed mice after 7 days ICV α-klotho or vehicle injections (n = 9–10/group). H: Body weight. I: Changes in body weight. J: Average daily food intake. K: Fed blood glucose levels. L: Change in fed blood glucose levels. M: Fasting blood glucose levels. HM: In pair-fed, STZ-treated mice (n = 8–9/group). Data represented as mean ± SEM. *P < 0.05 vs. ICV control.

Central α-Klotho Inhibition Impairs Glucose Tolerance

To determine the effects of central α-klotho inhibition on energy and glucose homeostasis, we performed central administration of ab-α-klotho antibody. A 2-day ab-α-klotho treatment significantly impaired glucose tolerance compared with vehicle-treated controls despite similar body weights (Fig. 4AC). There were no differences in liver PEPCK or glucose-6-phsophatase. However, gene expression of glucokinase and pyruvate kinase trended to be lower in ab-α-klotho mice (P = 0.14 and 0.15, respectively) (Supplementary Fig. 5D). Surprisingly, 7 days of ab-α-klotho significantly decreased body weight with no changes in food intake (Fig. 4DG). Taken together with ICV α-klotho experiments, these data suggest a distinct glucoregulatory role of central α-klotho independent of body weight and food intake.

Figure 4
Figure 4

Central α-klotho inhibition impairs glucose tolerance. A: Blood glucose levels during a GTT. B: Area under the curve (AUC) of the GTT. C: Fasting glucose levels. D: Body weight. E: Changes in body weight. F: Daily food intake. G: Timeline of food intake. AG: In 9-week-old chow-fed male mice treated with ab-α-klotho antibody compared with vehicle-treated controls (n = 8/group). Data represented as mean ± SEM. *P < 0.05 vs. ICV control.

α-Klotho Suppresses NPY/AgRP Neuron Activity, at Least in Part, by Enhancing mIPSCs

We next aimed to investigate the effects of α-klotho on NPY/AgRP neurons considering their critical role in energy homeostasis. A single ICV α-klotho injection in NPY-GFP reporter mice before an overnight fast significantly reduced cFOS colocalization with NPY/AgRP neurons by 49.0% (Fig. 5AC). Furthermore, electrophysiological recordings revealed α-klotho treatment decreases NPY/AgRP neuron firing rate and membrane potential (0.79 vs. 0.22 Hz and −52.7 vs. −57.8 mV, respectively) (Fig. 5DF).

Figure 5
Figure 5

α-Klotho suppresses NPY/AgRP neuron activity, at least in part, by enhancing mIPSCs. A: Representative image of cFOS (red) colocalization with NPY/AgRP (green). B: Number of NPY neurons. C: Number of NPY neurons with cFOS colocalization in the ARC. AC: Mice ICV treated with 2.0 μL α-klotho or vehicle before an overnight fast (n = 4 mice/group). DF: Representative cell-attached recording of an NPY/AgRP neuron (D), calculated firing rate (Hz) (E), and membrane potential (mV) (F) during α-klotho administration (n = 8 neurons from four male mice). G and H: Representative current clamp trace of an NPY/AgRP neuron (G) and mean membrane potential (H) induced by TTX or TTX and α-klotho. I: Representative whole cell recording tracers with α-klotho, glutamatergic blockade, and GABA-A receptor antagonist picrotoxin. JM: Mean amplitude (J and K), differences in cumulative probability of mIPSC amplitude (L), and mean frequency (M) of IPSCs under glutamatergic blockade with and without α-klotho treatment (n = 5 neurons from three male mice). Data represented as mean ± SEM. *P < 0.05 vs. aCSF.

To determine if α-klotho’s suppressive effects on NPY/AgRP neurons are due to pre- or postsynaptic events, brain slices were pretreated with TTX (1.0 μmol/L) to block action potentials. In the presence of TTX, α-klotho still decreased membrane potential (−53.4 vs. −58.4 mV), suggesting postsynaptic action of α-klotho on NPY/AgRP neurons (Fig. 5G and H). We also observed α-klotho to increase the magnitude, but not the frequency, of mIPSCs in NPY/AgRP neurons under glutamatergic blockade (25.9 vs. 34.4 pA) (Fig. 5IM), indicating α-klotho is directly antagonizing NPY/AgRP neurons by modulating receptor availability or intracellular signals (35). Overall, these experiments illustrate that α-klotho directly suppresses NPY/AgRP neuron activity by, at least in part, increasing receptor-mediated inhibitory signals.

α-Klotho Induces Cell Signaling, Alters Gene Expression, and Decreases NPY/AgRP Neuron Activity via FGFRs

To investigate the potential of α-klotho to alter cell signaling and gene expression in the hypothalamus, we first used the GT1-7 immortal hypothalamic cell line (36). A 30-min α-klotho treatment increased phosphorylation of ERKthr202/tyr204, AKTser473, and FOXO1ser256 (Supplementary Fig. 6A). Additionally, α-klotho treatment during both overnight and 2 h of serum starvation significantly reduced AgRP mRNA (by 28.5% and 30.3%, respectively), suggesting hormonal action of α-klotho in GT1-7 cells (Supplementary Fig. 6C). To investigate if α-klotho has hormonal action in the hypothalamus in vivo, we performed acute ICV α-klotho administration in healthy, fed mice and observed elevated phosphorylated ERK after 90 min in the ARC compared with vehicle-treated controls (Supplementary Fig. 6B).

Previous studies demonstrate the importance of α-klotho as a scaffolding protein increasing the affinity of FGF23 to FGFR1 (26,27). In hypothalamic GT1-7 cells, 30-min FGF23 (100 ng/mL) treatment had no effects on phosphorylated ERK or AKT (Fig. 6AC), while cotreatment with FGF23 and α-klotho had no synergistic effect compared with α-klotho alone. This suggests, at least in hypothalamic GT1-7 cells, α-klotho is independent of exogenous FGF23-mediated signaling. When cells were pretreated with FGFR1 inhibitor PD173074 (10 nmol/L), α-klotho–mediated cell signaling and suppression of AgRP mRNA were abolished (Fig. 6AD), indicating hypothalamic α-klotho action is dependent on FGFR1 activity. Moreover, immunofluorescent staining of cFOS revealed that ICV pretreatment with PD173074 also inhibited the ability of α-klotho to decrease NPY/AgRP neuron activity in vivo (Fig. 6EG).

Figure 6
Figure 6

α-Klotho–mediated cell signaling and regulation of NPY/AgRP neurons in the hypothalamus is dependent on FGFRs. A: Representative Western blot image. B: Phosphorylation of ERK. C: Phosphorylation of AKT. D: AgRP mRNA expression. AC: In GT1-7 cells treated with α-klotho, FGF23, PD173074, and/or wortmannin (n = 5–10/group). E: Representative image of cFOS (red) colocalization with NPY/AgRP neurons (green). F: Number of NPY neurons. G: Number of NPY neurons colocalized with cFOS. DG: In the ARC of the hypothalamus of mice ICV treated with vehicle, FGFR inhibitor with vehicle, or FGFR inhibitor with α-klotho before an overnight fast (n = 3 mice/group). Data represented as mean ± SEM. *P < 0.05 vs. controls. tAkt, total Akt; tERK, total ERK.

PI3K is a downstream mediator of FGFR1 and is also an important negative regulator of NPY/AgRP neurons (25,37). PI3K inhibition using wortmannin (50 nmol/L) also eliminated α-klotho’s ability to suppress AgRP gene expression (Fig. 6D). Taken together, these data demonstrate the importance of FGFR1/PI3K signaling in hypothalamic α-klotho function.

Seven Days of Central α-Klotho Treatment Suppresses Food Intake and Reduces Body Weight via FGFR and PI3K Signaling in DIO Mice

To determine if the therapeutic effects of α-klotho in DIO mice were dependent on FGFRs, we centrally injected PD173074 to inhibit endogenous FGFR function. Mice receiving α-klotho treatment alone experienced significantly decreased food intake and body weight compared with all groups (Fig. 7AD), while PD173074 treatment alone induced weight gain (Fig. 7A and B). PD173074 treatment blunted α-klotho–mediated reductions in food intake and body weight, suggesting the effects of central α-klotho on energy balance are mediated by FGFR signaling. Surprisingly, both α-klotho with an FGFR inhibitor and α-klotho alone groups experienced improved glucose clearance compared with vehicle-treated controls (Fig. 7E and F), suggesting FGFRs may not be involved in central α-klotho–mediated glucose regulation.

Figure 7
Figure 7

Central inhibition of FGFR1 blunts the therapeutic effects of 7 days α-klotho in DIO mice. A: Body weight. B: Changes in body weight. C: Average daily food intake. D: Timeline of food intake. E: Blood glucose levels during GTT. F: Area under the curve (AUC). AF: In DIO mice receiving 7 days ICV injection with vehicle, α-klotho, FGFR inhibitor, or inhibitor with α-klotho (n = 7–11/group). Data represented as mean ± SEM. *P < 0.05 vs. ICV vehicle.

Similar to FGFR antagonism, central inhibition of PI3K abolished α-klotho’s ability to suppress food intake and improve glucose clearance (Fig. 8). These data indicate that PI3K is critical to α-klotho–mediated regulation of food intake and glucose metabolism. Overall, coupled with in vitro cell signaling experiments, these data demonstrate a novel α-klotho/FGFR/PI3K mechanism in the central regulation of metabolism.

Figure 8
Figure 8

Central inhibition of PI3K negates the therapeutic effects of a single α-klotho injection in DIO mice. A: Timeline of food intake (including overnight fast before GTT on day 1). B: Average 48-h food intake. C: Blood glucose during a GTT. D: Area under the curve (AUC) of the GTT. AD: In DIO mice receiving a single injection with vehicle, α-klotho, wortmannin, or wortmannin with α-klotho (n = 9–10/group). Data represented as mean ± SEM. *P < 0.05.

Discussion

To our knowledge, this is the first study to provide evidence that α-klotho functions as a hypothalamic hormonal agent. ICV α-klotho administration improved glucose regulation, suppressed food intake, and reduced body weight in mouse models of type 1 and 2 diabetes, illustrating the therapeutic potential of central α-klotho in metabolic disease states. Although deeper investigation is required to identify the direct connection between central α-klotho activity and peripheral glucose metabolism, the current study determined that the glucose-lowering effects of α-klotho are independent from insulin sensitivity but, rather, are mediated by augmented insulin secretion during a glucose challenge. However, STZ experiments revealed that some of the glucoregulatory actions of α-klotho are independent of insulin altogether. Basal hepatic PEPCK mRNA was decreased in α-klotho-treated DIO mice, suggesting decreased hepatic glucose output may be an alternative mechanism.

The central and peripheral pools of α-klotho have distinct, independent functions due to α-klotho’s inability to cross the blood-brain barrier (9). While recent publications have demonstrated metabolic roles of α-klotho in the blood, including long-term α-klotho injection improving adiposity in DIO mice (6) and ameliorating diabetic cardiomyopathies in STZ-treated mice (38), the current study identifies distinct differences between central and peripheral α-klotho–mediated metabolic regulation. For example, α-klotho’s effects on food intake and glucose metabolism seem to be mainly via central mechanisms, while peripherally circulating α-klotho regulates gene expression to promote lipid oxidation and energy expenditure (6,38). Notably, whole-body α-klotho knockout and knockdown models have been previously utilized to investigate α-klotho’s functions (1,7), but these approaches do not distinguish between peripheral and central α-klotho function.

ICV ab-α-klotho was used in this study as a novel approach specifically impairing central α-klotho signaling, and as expected, ab-α-klotho treatment impaired glucose clearance. Although central α-klotho concentrations have yet to be quantified in patients with diabetes, past studies show blood α-klotho concentrations to be decreased in some populations with diabetes (8,39). Thus, our data connecting central α-klotho impairment and disordered glucose regulation may provide new insight into the pathophysiology of metabolic disorders.

Contrary to our hypotheses, central α-klotho inhibition resulted in decreased body weight with no differences in food intake. α-Klotho knockout mice also experience weight loss, primarily due to atrophy of metabolically active organs, resulting in premature death (1,7). These findings highlight the complicated and diverse metabolic functions of α-klotho. For example, while evidence from the current study and past literature describes α-klotho as an antidiabetic agent (36,38), overexpression of α-klotho has been shown to elicit insulin resistance (1). Notably, α-klotho–overexpressing mice do not experience hyperglycemia, adiposity, or hyperphagia associated with clinical insulin resistance (1). Moreover, α-klotho is an important negative modulator of insulin and IGF-I signaling to regulate apoptosis and ROS buffering (10,11). The many complex physiological roles of α-klotho may explain the unexpected results in response to central α-klotho inhibition.

The current study identified central α-klotho as a novel antagonist of NPY/AgRP neurons. Considering NPY/AgRP neuron overactivity is associated with disordered feeding, body weight, and glucose regulation (13,40), our data provides encouraging evidence of α-klotho as a potential therapeutic target in metabolic disease prevention. At the present, it is unclear if NPY/AgRP neurons are the primary mediators of central α-klotho’s regulation of metabolism, underscoring the importance of further investigation into the specific neuronal effectors and cell signaling involved. However, the observed ICV α-klotho phenotype has many similarities to the previously described effects of NPY/AgRP neuron inhibition, including suppressed food intake, reduced body weight, improved glucose clearance and insulin release, and decreased hepatic gluconeogenic gene expression (13,1517,41,42).

Similar to findings in studies using hippocampal and oligodendrocyte progenitor cells (10,11,28), α-klotho induced phosphorylation of ERKthr202/tyr204, AKTser473, and FOXO1ser256 in hypothalamic GT1-7 cells—all of which are established signaling molecules involved in downregulating NPY/AgRP gene transcription and activity (18,43). Furthermore, the observed ICV α-klotho phenotype resembles FGFR activation, which also results in suppressed food intake, improved glucose regulation, attenuated NPY/AgRP neuron activity, and decreased liver gluconeogenic gene expression (2024). α-Klotho serves as a nonenzymatic scaffold to increase FGF23 affinity to FGFR1 (27). Thus, we investigated the potential importance of a hypothalamic α-klotho–FGFR1 signaling mechanism. Similar to previous studies in hippocampal cells, our results show that hypothalamic α-klotho–mediated signaling and AgRP mRNA regulation in GT1-7 cells were abolished with pretreatment with FGFR1 antagonist PD173074 (28). Additional experiments determined that PI3K signaling, a downstream mediator of FGFR1 (37) and potent regulator of NPY/AgRP neurons (25), was also required for α-klotho–mediated AgRP mRNA suppression. Future studies should further investigate the possible involvement of a novel α-klotho–FGFR1–PI3K axis in the homeostatic modulation of NPY/AgRP neurons.

We further investigated the involvement of FGFR/PI3K signaling to central α-klotho–mediated regulation of metabolism. Central FGFR or PI3K inhibition blunted ICV α-klotho’s effects on food intake and body weight, while only PI3K inhibition affected α-klotho–mediated glucose regulation. Overall, these data support the hypothesis that central FGFR-PI3K signaling is critical to α-klotho–mediated regulation of metabolism. However, studies investigating the function of central FGFRs in metabolism yield mixed results depending on animal model and experimental approach. ICV PD173074 (FGFR inhibitor) impairs glucose clearance in healthy rats, but it is described as stress related (23,44). ICV PD173074 in DIO mice elicits no phenotype (21,24). Furthermore, antibody-mediated inhibition of FGFR1 in rodents and monkeys increases energy expenditure, decreases food intake, and reduces body weight, while genetic deletion of FGFR1 in NPY/AgRP neurons also results in no metabolic phenotype (4547). Additionally, the specificity for PD173074 in vivo is unclear; thus, it likely has nonspecific antagonism of other FGFRs. Future studies should investigate the specific roles of FGFRs, their isoforms, and their neuronal effectors in central regulation of metabolism by performing selective deletion of FGFRs in specific neurons of mature mice using the inducible Cre-LoxP system.

In addition to FGFR-PI3K signaling, there are likely unknown concurrent mechanisms underlying central α-klotho–mediated metabolic regulation. Other neuron populations, such as proopiomelanocortin (POMC) neurons, which are closely associated with NPY/AgRP neurons, may be involved. Our cell culture and immunohistochemistry data also may suggest ERK as an additional cell signaling mechanism of hypothalamic α-klotho action. ERK signaling is downstream of α-klotho, negatively regulates NPY/AgRP neurons, possibly via Kruppel-like factor 4, and is involved in hypothalamic FGF1- and FGF19-mediated glucose lowering (21,43,48).

To summarize, this study identifies α-klotho as a novel antagonist of NPY/AgRP neurons and demonstrates α-klotho’s importance to central regulation of metabolism via an α-klotho–FGFR1–PI3K signaling axis. Our data revealed central administration of α-klotho to yield various therapeutic effects in models of type 1 and 2 diabetes, including improved glucose regulation, suppressed food intake, and reduced body weight. To our knowledge, this study provides the first evidence of α-klotho as a novel hypothalamic regulator of energy balance and glucose metabolism, thus providing new insight into the pathophysiology of metabolic disease.



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Roasted Zucchini Fries with Parmesan

By electricdiet / June 30, 2020


These roasted zucchini fries with Parmesan are oven-baked to crispy perfection. And you only need a few ingredients to make this delicious summer side dish!

Garlic zucchini fries on an oven pan

Looking for something crispy and easy to serve with your meal?

These roasted zucchini fries with Parmesan are one of my favorite side dishes, especially for the summer! They’re the perfect way to sneak some healthy veggies onto the menu for your next cook out.

And they’re so easy to make. Just slice the zucchini, coat the sticks in Parmesan and garlic pepper, drizzle with olive oil, and throw them in the oven until they’re ready!

I’ve seen a lot of zucchini fry recipes made with bread crumbs to give them crunch. Unfortunately, this adds a lot of carbs and calories.

These zucchini fries, on the other hand, are made with Parmesan to give them that satisfying crispy coating without all the carbs. That means you can enjoy these healthy fries without the guilt!

So whether you have summer zucchini you need to use or just want a delicious and healthy side dish, definitely give this recipe a try!

How to make roasted zucchini fries with Parmesan

These crispy sticks are easy to prep and only take about 20 minutes to bake in the oven.

Step 1: Preheat oven to 425°F. Line a baking sheet with parchment paper.

Step 2: Cut the two zucchini into 16 sticks. To do this, start by trimming the ends of both zucchini. Next, quarter both zucchini lengthwise so you end up with 8 long pieces. Finally, cut each piece in half crosswise to get 16 shorter triangular-shaped pieces.

Step 3: Beat the egg in a shallow bowl. You want to use a bowl that will be large enough to dip the zucchini pieces.

Step 4: Combine the Parmesan and garlic pepper in a second shallow bowl, again ensuring that it’s large enough to dip the zucchini pieces.

Step 5: Take one zucchini strip and dip the cut sides into the egg. Once the cut sides are coated, dip the zucchini into the Parmesan mixture until well-coated.

Step 6: Place the zucchini strip on the baking sheet with the skin side down.

Step 7: Repeat with the remaining zucchini.

Step 8: Once all the zucchini has been coated and placed on the baking sheet, drizzle with olive oil.

Step 9: Place the baking sheet in the oven and roast until the zucchini sticks are golden, about 15 to 20 minutes.

That’s it! Once the exterior is nice and crispy, your zucchini fries are ready to serve.

Cutting zucchini into sticks

Zucchini has recently become popular as a healthy alternative to high-carb and starchy foods like french fries. The trick is to make sure the zucchini turns out crispy and not soggy.

I’ve seen a few recipes for zucchini cooked similar to this one, but it was sliced into rounds instead of sticks.

When I tried that method, the rounds turned out a bit too soggy for my taste.

This triangular shape seemed to get much more crispy. That’s why I prefer to quarter the zucchini and then cut each long piece in half. I also like that they look a bit like house-cut fries this way!

Storing zucchini fries

I recommend serving this dish immediately. The fries will be best when they are hot out of the oven.

If you have leftovers, you can store them covered in the refrigerator for 3-4 days. However, the moisture from the zucchini will make the exterior less crispy than when they’re fresh.

To reheat, I recommend either pan-roasting or sticking them back in the oven. This will help the exterior get crispy again.

Zucchini fries on a wooden plate

Other healthy zucchini recipes

Zucchini is such a versatile vegetable. It’s amazing how many different kinds of recipes you can make with it! Plus, it’s so easy to find in the summertime.

If you’re looking for a few more fun recipes to use up your zucchini, here are some of my favorites that I think you’ll enjoy:

When you’ve tried these zucchini fries, please don’t forget to let me know how you liked it and rate the recipe in the comments below!

Recipe Card

Roasted Zucchini Fries with Parmesan

Roasted Zucchini Fries with Parmesan

These roasted zucchini fries with parmesan are oven-baked to crispy perfection. And you only need a few ingredients to make this delicious summer side dish!

Prep Time:10 minutes

Cook Time:20 minutes

Total Time:30 minutes

Author:Shelby Kinnaird

Servings:4

Instructions

  • Preheat oven to 425°F. Line a baking sheet with parchment paper.

  • Cut the two zucchini into 16 sticks. To do this, start by trimming the ends of both zucchini. Next, quarter both zucchini lengthwise so you end up with 8 long pieces. Finally, cut each piece in half crosswise to get 16 shorter triangular-shaped pieces.

  • Beat the egg in a shallow bowl. You want to use a bowl that will be large enough to dip the zucchini pieces.

  • Combine the Parmesan and garlic pepper in a second shallow bowl, again ensuring that it’s large enough to dip the zucchini pieces.

  • Take one zucchini strip and dip the cut sides into the egg. Once the cut sides are coated, dip the zucchini into the Parmesan mixture until well-coated.

  • Place the zucchini strip on the baking sheet with the skin side down.

  • Repeat with the remaining zucchini.

  • Once all the zucchini has been coated and placed on the baking sheet, drizzle with olive oil.

  • Place the baking sheet in the oven and roast until the zucchini sticks are golden, about 15 to 20 minutes.

Recipe Notes

This recipe is for 4 servings of zucchini fries. If you cut both zucchini into 8 pieces, then one serving will be 4 zucchini fries. This dish is best served immediately. If you have leftovers, they can be stored covered in the refrigerator for 3-4 days. The exterior will lose some of its crispiness, but you can pan roast or bake the fries to reheat and try to re-crisp the outsides.

Nutrition Info Per Serving

Nutrition Facts

Roasted Zucchini Fries with Parmesan

Amount Per Serving (4 pieces)

Calories 127 Calories from Fat 81

% Daily Value*

Fat 9g14%

Saturated Fat 3.5g22%

Trans Fat 0g

Polyunsaturated Fat 2.2g

Monounsaturated Fat 3g

Cholesterol 57.3mg19%

Sodium 297.5mg13%

Potassium 287.6mg8%

Carbohydrates 3.6g1%

Fiber 1g4%

Sugar 2.5g3%

Protein 8.4g17%

Vitamin A 400IU8%

Vitamin C 38mg46%

Calcium 150mg15%

Iron 0.7mg4%

Net carbs 2.6g

* Percent Daily Values are based on a 2000 calorie diet.

Course: Side Dishes

Cuisine: American

Diet: Diabetic, Low Fat

Keyword: low-carb, roasted zucchini fries, zucchini, zucchini fries



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Back on Track – My Bizzy Kitchen

By electricdiet / June 28, 2020


If I were to search my blog and see how many times I’ve said “hit the reset button!” or “the switch is now on!” I would probably have a million dollars and could have retired by now.  But, such is the case.  Since working from home and so close to my kitchen, I’ve been making all the things.  The only problem with that is that I normally only had time on a weeknight pre-COVID to make dinner, do a couple things around the house, watch a 30 minute t.v. show and go to bed.

Now I clock out at 5 and I can easily recipe develop two dishes, taste test and THEN eat dinner.

A week ago Saturday I got on the scale and was like “um, excuse me?”  Last week I walked more, drank water rather than wine, but didn’t go crazy with trying to be perfect.  I was mindful.  And I lost 1.4 pounds.  And it wasn’t that hard.  And I know this.  Yet somehow I still get back on that hamster wheel thinking that the way I used to do things (that got me to 190 pounds) will somehow work in my favor.  But I know that doesn’t work.

I really don’t know what all those numbers mean, but what I realized during the week I first stepped on it, and 7 days later – is that because numbers fluctuate so much, I only stepped on the scale a week later.  My weigh in is normally on Saturday’s and there have been some weeks where I step on the scale on Thursday and see I am down 2.0 pounds and think “well, I guess I can have an extra glass of wine and some pizza!”

So I am being mindful.  Nothing more and nothing less.

+++++++++++

I had a great weekend.  It was really the first full weekend by myself in the house without Hannah, Jacob or my Mom over.  It was . . . quiet.  And weird.  And quiet.  But guess what?  I could listen to music all day long without air pods – that was wonderful!  I also got up early on Saturday morning and made a pot of coffee, did my WW zoom in the kitchen and didn’t have to worry about being too noisy to wake anyone up.

After my WW meeting I hit up the local farmers market for first time this summer.  Farm fresh eggs will forever be worth the $5 a dozen.  Side note:  save these eggs for scrambled eggs or omelets – don’t waste these in a recipe because you won’t tell.

I spoke to a lovely young man at this farmstand – Waypoint Farm.  Each bag of little gem romaine was only $2!  I quickly grabbed two.  He was so passionate about his produce – #love.   I hope to spend more time talking with him next week – it was about to storm as I grabbed my two bags of romaine.

I made another shitty video on my YouTube channel!  This time I showed how to make my KFC chicken nuggets.  So so good.

My new column for The Daily Herald will be posted the week of 4th of July, but my article shows how you can host a BBQ for 8 people on a budget – I made chicken legs, corn salad, grilled potato fries and a strawberry tomato salsa – all for $23.11.  Here is a sneak peak for that article.

It’s funny but since Sunday was just a regular day for me – no father’s day celebrations, I made this dinner for 8 thinking I would just bring it over to my neighbors house.  How rude would that have been to barge in on their father’s day!  So I’ll be having leftovers this week.  Not that I am complaining because this was delicious.

All in all it was a great weekend.  Getting used to this new normal of living by myself.  The longest I’ve lived alone is three weeks before Hannah and Jacob moved in after my husband died in December 2014.  In another week I’ll break that record – ha!

I’ve yet to walk around my house naked, but I have walked from my bedroom to the bathroom in just my underwear and bra, so that’s a start!

Come back tomorrow for “What I Eat in A Day” on #teampurple – it was a delicious day!  Until then, be well.

 



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Apple Cider Vinegar for Diabetes: Does it Help?

By electricdiet / June 26, 2020


Apple cider vinegar — also known as “ACV” — is a trendy ingredient these days. Personally, I love pouring a big splash of it into an ice-cold glass of fruit-flavored seltzer!

But recently, apple cider vinegar is also being touted as a home remedy for people living with diabetes, with claims that it can help control blood sugars and lower A1c levels.

In this article, we’ll discuss the general benefits of apple cider vinegar, what research has determined about its impact on blood sugar levels, precautions to take when adding it to your diet, and who shouldn’t consume apple cider vinegar regularly at all.

Glass bottle of apple cider vinegar in front of basket with apples

First, make sure you buy the right apple cider vinegar

Made by crushing, distilling, and fermenting apples, apple cider vinegar does offer a few generally accepted health benefits.

Those benefits come mostly from “the mother” which is the beneficial bacteria that cause the fermentation process.

“First, yeast is added to apple juice to break down the sugars and turn them into alcohol,” explains Enzymedica. “Then, bacteria is added, which converts the alcohol into acetic acid. This bacteria is what is known as the “mother” because it is the catalyst that gives rise to the vinegar. Many store-bought apple cider vinegars have the mother removed because it gives the vinegar a cloudy appearance, which can lead some customers to believe that the product has gone bad. But this is not the case. In fact, the mother is the healthiest part.”

When purchasing ACV, you should be looking for a product that’s raw, unfiltered, and “comes from the mother.”  All three of these details should be clearly printed on the packaging of your ACV contained in a glass container versus a plastic container.

You can find high-quality ACV in most grocery stores in the baking aisle or online at Trader Joe’s and on Amazon.

*Please note: You should not drink ACV without diluting it in water or another beverage. The high acidity content can damage your teeth, mouth, and throat if consumed regularly without diluting.

General health benefits of drinking apple cider vinegar

Before we discuss how ACV affects blood sugar and A1c for people living with diabetes, let’s look at some of the claims regarding general health benefits.

ACV has been around for a long time

Mentioned several times in the Bible as an “antibiotic,” the problem is that ACV is not actually the cure-all it’s often reported to be — and it’s definitely not going to help you if what you actually need is a legitimate antibiotic.

Again, to be clear, ACV is not an antibiotic.

Instead, there are some basic and generally accepted benefits to drinking a little ACV every day.

Let’s take a look at the facts:

ACV is antimicrobial

ACV is “antimicrobial” which means it’s very similar to alcohol-based hand sanitizers. It helps to prevent the spreading of bacteria, fungi, and some viruses. However, that is not the same as “antibacterial,” which prevents the growth of bacteria.

Extensive research has also found that ACV has a significant direct effect on three specific types of bacteria: E-coli, Staphylococcus aureus, and Candida albicans. However, do not use ACV to treat any of these types of bacteria without consulting your healthcare team.

You can even mix ACV (or standard white vinegar) with a much larger ratio of water to use as an all-natural household cleaning product for your floors, counters, and bathrooms.

ACV contains probiotics…which support healthy digestion

You’ll get the probiotic benefits of ACV is you’re drinking the raw, unfiltered, and “from the mother” version. Like kombucha, ACV contains dozens of beneficial bacteria that support a healthy gut.

When your gut-health is “out of whack” because there is too little of the healthy bacteria and too much of unhealthy bacteria, it can actually affect many parts of your entire well-being.

Gut health has been linked to a variety of health conditions, including:

Apple cider vinegar and diabetes: Can ACV lower blood sugar levels in people with diabetes?

Let’s cut right to the chase: apple cider vinegar has shown to reduce blood sugar levels slightly in people with type 2 diabetes and type 1 diabetes, but the results aren’t going to have a tremendous impact on your A1c from ACV alone.

Instead, the research seems to imply that adding ACV to your many other diabetes management habits can help a little bit. And there’s no question that it could benefit your health in ways unrelated to your blood sugar, too, as explained earlier.

Let’s take a look at some of the most significant research.

In well-managed type 2 diabetes, drinking ACV before bed helps manage morning blood sugars.

In this 2007 study from Arizona, patients with well-managed type 2 diabetes who did not take insulin drank 2 tablespoons of apple cider vinegar with 1 ounce of cheese every night.

The study also included a placebo group of patients with well-managed type 2 diabetes, who drank water instead of ACV.

In the placebo group, morning fasting blood sugar levels were 2 percent lower by the end of the study. In the ACV group, morning fasting blood sugar levels were 4 to 6 percent lower.

The study concluded that ACV can help lower blood sugar levels in those who are engaged in other diabetes management habits.

12 weeks of drinking ACV showed relatively insignificant reductions in A1c levels

This 2018 study from Singapore — involving patients with type 1 and type 2 diabetes — found that while drinking 2 tablespoons of ACV after a meal did demonstrate a slight reduction in post-meal blood sugar levels, the results weren’t remarkably significant.

That being said, blood sugar levels were slightly lower which suggests it can help but it likely can’t replace diabetes medications nor compensate for unhealthy lifestyle habits.

ACV improves insulin sensitivity after high-carbohydrate meals

This 2004 study from Arizona gave patients with type 2 diabetes 20 grams (about 1.5 tablespoons) of ACV with high-carbohydrate meals.

Researchers concluded that consuming vinegar with high-starch meals lowered post-meal blood sugars by increasing a patient’s sensitivity to insulin.

ACV delays gastric emptying and reduces post-meal blood sugar levels

This 2007 study from Sweden focused on patients with type 1 diabetes and gastroparesis.

The results determined that consuming 2 tablespoons of ACV with meals reduced the rate at which the body empties digested food (including glucose) into the bloodstream.

For patients with gastroparesis, this is actually a disadvantage because they already struggle with significantly delayed and unpredictable digestion — which makes it harder to time and dose insulin.

For patients without gastroparesis, however, this might be helpful. By reducing the rate of gastric emptying, it reduces the post-meal blood sugar spike.

If you decide to add ACV to your diet (and who shouldn’t)

While the results of recent research imply that ACV will have a very modest impact on your blood sugar, there are still plenty of reasons to incorporate it into your daily or weekly diet.

The probiotics alone are remarkably important for the maintenance of your gut’s balance of healthy bacteria.

Remember these three crucial details when consuming ACV:

  • Always dilute it with another beverage (water, seltzer, tea) or by mixing it into your food.
  • Only consume approximately 2 tablespoons per day.
  • Consuming too much ACV can wreak havoc on your teeth, throat, and stomach because of its high-acidity content.

However, in addition to always diluting your ACV with another liquid, there are a few people who shouldn’t drink it all.

You shouldn’t drink ACV if…

If you have any of the following health concerns, talk to your doctor before consuming ACV.

  • You have a history of stomach ulcers
  • You have low potassium levels
  • You have a history of bulimia
  • You have any health or dental conditions in your mouth or throat (discuss with your doctor or dentist first!)

ACV has a lot of subtle but legitimate benefits to offer anyone — including those of us with diabetes. Give it a whirl! And enjoy!



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Vitamin D Receptor Overexpression in β-Cells Ameliorates Diabetes in Mice

By electricdiet / June 24, 2020


Abstract

Vitamin D deficiency has been associated with increased incidence of diabetes, both in humans and in animal models. In addition, an association between vitamin D receptor (VDR) gene polymorphisms and diabetes has also been described. However, the involvement of VDR in the development of diabetes, specifically in pancreatic β-cells, has not been elucidated yet. Here, we aimed to study the role of VDR in β-cells in the pathophysiology of diabetes. Our results indicate that Vdr expression was modulated by glucose in healthy islets and decreased in islets from both type 1 diabetes and type 2 diabetes mouse models. In addition, transgenic mice overexpressing VDR in β-cells were protected against streptozotocin-induced diabetes and presented a preserved β-cell mass and a reduction in islet inflammation. Altogether, these results suggest that sustained VDR levels in β-cells may preserve β-cell mass and β-cell function and protect against diabetes.

Introduction

Vitamin D deficiency has been associated with diabetes, both in individuals with type 1 diabetes (T1D) and insulin resistance/type 2 diabetes (T2D) (13). In addition, a parallel increase in the prevalence of diabetes and in vitamin D deficiency incidence worldwide has been observed, which may result from both deficient sun exposure, involved in vitamin D synthesis by the skin, and inadequate dietary supply (4). Vitamin D exerts its actions mainly through its binding to vitamin D receptor (VDR), and VDR gene polymorphisms have also been associated with the risk of T2D in different ethnic populations (5). Likewise, two single nucleotide polymorphisms in the VDR gene have been associated with T1D (6). This strongly suggests an essential role of the vitamin D/VDR axis in diabetes, although the mechanisms have not yet been elucidated.

VDR belongs to the steroid hormone receptor superfamily, and it is widely expressed in several cell types where it is known to regulate key cellular processes such as proliferation, differentiation, apoptosis, and immunomodulation (7). In addition, genome-wide VDR-binding chromatin immunoprecipitation sequencing (ChIP-seq) data revealed that an important part of target genes is involved in metabolism (810). Moreover, VDR is expressed in a wide variety of immune cells, and vitamin D is a known immunomodulator in both the innate and adaptive arms of the immune system (11). In particular, vitamin D can promote immune tolerance and has immunosuppressive properties (11,12). Vitamin D exposure also causes T cells to change their cytokine production from a proinflammatory to an anti-inflammatory profile (13). It has been hypothesized that vitamin D may have a protective role in diabetes since the immune system is involved in the development of both T1D and T2D (14,15).

VDR is also expressed in several insulin-responsive metabolic tissues, such as the liver, skeletal muscle, or adipose tissue, and it has been reported that vitamin D may improve insulin sensitivity of these tissues (16). Vitamin D may directly increase insulin receptor expression, and thereby enhance insulin stimulation of glucose transport, or indirectly decrease insulin resistance by decreasing inflammatory responses, one of the causes of insulin resistance (16).

Pancreatic islets also express VDR and can metabolize inactive 25-hydroxyvitamin D3 to active 1,25 (OH)2D3 (17). Vitamin D has been reported to exert beneficial effects on glucose tolerance enhancing β-cell function (18,19). Studies in cultured rat islets demonstrated that synthesis and release of insulin may be enhanced by treatment with high doses of vitamin D (20). Moreover, mice with VDR deficiency presented impaired glucose tolerance, defective insulin secretion, and a reduction in insulin mRNA content, suggesting that VDR is a key factor in β-cell function (21). However, the role of VDR, specifically in β-cells, during the development of diabetes remains unknown.

Thus, here we aimed to study the role of VDR in the β-cell in the pathophysiology of diabetes. We found that Vdr expression is decreased in islets from both T1D and T2D mouse models. We also demonstrated that overexpression of VDR in β-cells of transgenic (Tg) mice counteracted experimental diabetes, providing evidence that sustained VDR levels in β-cells may preserve β-cell mass and function and protect against diabetes.

Research Design and Methods

Animals

Female NOD/LtJ and male BKS.Cg-+Leprdb/+Leprdb OlaHsd (db/db), BKS.Cg-m+/+Leprdb/OlaHsd (db/+), and C57Bl6/SJL mice were used. Heterozygous male Tg mice expressing mouse Igf2 under the control of the rat insulin promoter-I (RIP-I) were used (22). Diabetes was induced by streptozotocin (STZ) as previously described (23). All mice were fed ad libitum with a standard chow diet (2018S Teklad Global; Harlan) and maintained under conditions of controlled temperature and light (12-h light/dark cycles). Where stated, mice were fasted for 16 h. Animal care and experimental procedures were approved by the Ethics Committee in Animal and Human Experimentation of the Universitat Autònoma de Barcelona, Bellaterra, Spain.

Generation of Tg Mice

The RIP-I/Vdr chimeric gene was obtained by introduction of a 3.3-kb EcoRV-EclXI fragment containing the entire mouse Vdr cDNA (Open Biosystems INC, Huntsville, AL) (ref. 3710866, GeneBankBC006716) at the EcoRI site in RIP-I/β-globin expression vector (22). This chimeric gene was microinjected into fertilized mouse eggs from a C57BL6/SJL background. The general procedures used for microinjection and detection were as described (24).

Immunohistochemistry and Histopathology

For immunohistochemical detection of VDR, insulin, glucagon, somatostatin, pancreatic polypeptide, TUNEL, and Ki67, pancreas were fixed for 12–24 h in formalin, embedded in paraffin, and sectioned. Sections were then incubated overnight at 4°C with the following antibodies: rat anti-mouse VDR (clone 9A7; Merck KGaA, Darmstadt, Germany), guinea pig anti-porcine insulin (Sigma Chemical, St Louis, MO), rabbit anti-human glucagon (Signet Laboratories, Dedham, MA), rabbit anti-somatostatin (Serotec, Oxford, U.K.), rabbit anti-human pancreatic polypeptide (ICN Biomedicals), and anti-Ki67 (BD Pharmingen). As secondary antibodies, peroxidase-conjugated rabbit anti-guinea pig IgG (Dako, Glostrup, Denmark), biotinylated goat anti-rabbit (Pierce, Rockford, IL), tetramethylrhodamine isothiocyanate (TRITC)-conjugated goat anti-guinea pig (Molecular probes, Leiden, the Netherlands), biotinyilated goat anti-rabbit (Molecular Probes), biotinylated rabbit anti-rat (Dako), and biotinylated horse anti-mouse (Vector Laboratories, Burlingame, CA) antibodies were used. Streptavidin-conjugated Alexa 488 (Molecular Probes) or streptavidin-conjugated Alexa 568 (Molecular Probes) were used as fluorochromes. Images were obtained with a Nikon Eclipse 90i microscope (Nikon, Tokyo, Japan).

Morphometric Analysis

β-Cell and α-cell mass, β-cell replication, and apoptosis determination were performed as previously described (23,25).

Islet Isolation and Culture

Pancreatic islets were isolated as previously described (23) and cultured overnight to recuperate from isolation stress in RPMI-1640 (2.5 mmol/L glucose), supplemented with 1% BSA, 2 mmol/L glutamine, and penicilin/streptomycin at 37°C in an atmosphere of 95% humidified air/5% CO2. To study the effects of glucose on Vdr expression on β-cells, after overnight culture, pools of islets were treated with 2.5 or 9 mmol/L of glucose and 9 mmol/L 2-deoxy-d-glucose. Some of the pools were also cultured with recombinant INS (2 ng/mL) (Sigma). After 8 h of treatment, islets were hand-picked and processed to obtain RNA.

Gene Expression Analysis

For quantitative PCR analysis, total RNA was extracted from isolated islets using Tripure Isolation Reagent (Roche Molecular Biochemicals) and Rneasy Micro Kit (Qiagen, Hilden, Germany). Total RNA (1 µg) was reverse-transcribed for 1 h at 37°C with Transcriptor First Strand cDNA Synthesis Kit (Roche). Quantitative PCR was performed in a Light Cycler 480 II (Roche) using Light Cycler 480 SYBR Green I Master mix (Roche). Values were normalized to Rplp0 as housekeeping. The primers listed in Table 1 (Supplementary Data) were used for murine islets.

Hormone and Metabolite Assays

Blood glucose levels and serum insulin concentrations were measured as previously described (23). A glucose tolerance test was performed as previously described (23). For insulin release determination, glucose (3 g/kg body weight) was injected intraperitoneally, and venous blood from the tail vein was collected at 0, 2, 5, 15, and 30 min in prechilled tubes (Microvette CB 300; SARSTEDT), which was immediately centrifuged to separate plasma and was stored at −20°C. Insulin levels from an insulin release test were measured by ELISA (Crystal Chemical, Chicago, IL).

Statistical Analysis

All values are expressed as the mean ± SEM. Differences between two means were compared by Student t test, and differences between three or more means were analyzed by one-way ANOVA or two-way ANOVA tests using GraphPad Prism software (version 7.00; GraphPad Software). A P value <0.05 was considered statistically significant. Correlations were determined by nonparametric Spearman correlation test using the computer program GraphPad Prism (version 7.00; GraphPad Software).

Data and Resource Availability

To characterize the expression in human β-cells of different genes, such as VDR, INS, or SLC2A2, we used publicly available data sets from Gene Expression Omnibus (GEO) (26) and ArrayExpress databases (27): GSE20966 (28) and E-CBIL-20 (29). For each data set, the publicly available normalized expression levels for each gene were used. When different probes were detecting the expression of the same gene, their genomic locations were determined to assess its relevance. Data were then plotted as relative signal expression.

Results

VDR Expression Is Reduced in Islets From Diabetic Mice

To study the regulation of Vdr gene expression in islets during the diabetic process, Vdr mRNA levels were measured in islets of several diabetic animal models. In islets from nonobese diabetic (NOD) mice (Fig. 1A), the most common type 1 diabetic mouse model, which shows infiltrated islets and β-cell reduction related to glycemia (Supplementary Fig. 1A), Vdr mRNA levels were decreased in hyperglycemic mice compared with normoglycemic littermates (Fig. 1B). This reduction was observed in parallel with a decline in the expression levels of β-cell marker genes, such as insulin (Ins), solute carrier family member 2 (Slc2a2 or Glut2), and glucokinase (Gck) (Fig. 1B). In addition, islet glucagon (Gcg) and uncoupling protein 2 (Ucp2) expression remained unchanged (Fig. 1B). Similarly, in islets from STZ-induced diabetic mice (Fig. 1C), expression of Vdr and β-cell gene markers was also reduced (Fig. 1D). In contrast, Gcg expression was unaltered, and Ucp2 expression presented a clear trend to increase in STZ islets (Fig. 1D). Moreover, Vdr expression was measured in islets from a Tg T2D mouse model in which islets were hyperplasic because of IGF2 overexpression specifically in β-cells (22,25). These Tg mice (Tg-IGF2) develop a prediabetic state with disrupted islet structure, β-cell dysfunction, altered glucose homeostasis, and islet hyperplasia (25) (Supplementary Fig. 1B). Similar to those in the T1D models, islets from prediabetic Tg-IGF2 mice also showed a clear decrease in Vdr expression together with a reduction in β-cell gene markers (Fig. 1E and F). Likewise, a reduction in Vdr expression levels and a decrease in β-cell gene markers were also observed in the well-established T2D model db/db mice, which also displayed unchanged islet Gcg expression and a significant increase of Ucp2 mRNA levels (Fig. 1G and H). Moreover, the correlation between Vdr expression levels and glycemia was examined in T1D and T2D models. Clearly, Vdr expression levels in islets correlated with glucose levels in all analyzed T1D and T2D mice models (Supplementary Fig. 1CF). It is noteworthy that analysis of human islets from healthy and T2D patients from two public available gene expression data set GSE20966 (28) (Fig. 2A) and E-CBIL-20 (29) (Fig. 2B) revealed a nonsignificant trend toward a reduction in the expression levels of VDR and SLC2A2 in islets from T2D patients. These results from different diabetic mouse models and human samples provided evidence that Vdr expression in islets was reduced in conditions of hyperglycemia.

Figure 1
Figure 1

Islet gene expression from diabetic mouse models. A and B: Glycemia, Vdr gene expression analysis, and β-cell gene profile in islets from NOD mice. Blood glucose levels (A) and gene expression in islets (B) from NOD hyperglycemic and NOD normoglycemic female mice (26 weeks old). NOD normoglycemic mice (white bars, n = 7) and NOD hyperglycemic mice (striped bars, n = 9). Results are mean ± SEM. *P < 0.05 and ***P < 0.001 vs. NOD normoglycemic. C and D: Glycemia, Vdr expression analysis, and β-cell gene profile in islets after experimental diabetes induction. Blood glucose levels (C) and gene expression in islets (D) from 3-month-old mice treated with multiple doses of STZ (50 mg/kg body weight) 40 days after treatment. Wt untreated mice (white bars) and Wt STZ-treated mice (blue bars); n = 7 per group. Results are mean ± SEM. *P < 0.05, **P < 0.01, and ***P < 0.001 vs. untreated mice. E and F: Glycemia, Vdr expression analysis and β-cell gene profile in islets from Tg-IGF2 mice. Blood glucose levels (E) and gene expression in islets (F) from hyperglycemic and insulin-resistant Tg mice that overexpress IGF2 specifically in β-cells. Wt mice (white bars) and Tg-IGF2 mice (black bars); n = 6 per group. Results are mean ± SEM. *P < 0.05, **P < 0.01, and ***P < 0.001 vs. Wt. G and H: Glycemia, Vdr gene expression analysis, and β-cell gene profile in islets from db/db mice. Blood glucose levels (G) and gene expression in islets (H) from diabetic db/db and normoglycemic db+/− male mice (19 weeks old). db+/− mice (white bars) and db/db mice (purple bars); n = 8 per group. Results are mean ± SEM. *P < 0.05, **P < 0.01, and ***P < 0.001 vs. db+/−.

Figure 2
Figure 2

Transcriptomic analysis of human β-cells. A: Expression levels of VDR, SLC2A2, and INS in islets from nondiabetic (ND) and T2D patients obtained from the raw data deposited in a Minimum Information About a Microarray Experiment (MIAME) compliant database (GEO accession number: GSE20966); n = 10 per group. B: Expression levels of VDR, SLC2A2, and INS in islets from ND (n = 7) and T2D (n = 5) patients obtained from complete microarray data sets available from ArrayExpress E-CBIL-20.

VDR Expression Is Controlled by Glucose

To further explore the regulation of Vdr expression by changes in glucose levels, Vdr expression was measured in islets from fed and fasted wild-type (Wt) mice (Fig. 3A). Unexpectedly, our results showed a marked reduction in Vdr mRNA levels in islets from fasted mice compared with islets from fed mice (Fig. 3B). The decrease in Vdr expression in fasted conditions was also parallel to a significant reduction in Ins and Slc2a2 gene expression (Fig. 3B). We next evaluated the effects of changes in glucose concentration on Vdr gene expression in cultured islets. In islets incubated with high glucose, Vdr mRNA levels were increased, whereas this increase was blunted in islets treated with nonmetabolizable 2-deoxy-d-glucose (Fig. 3C), suggesting that glucose metabolization is required to induce Vdr expression. In addition, to examine the effects of insulin on Vdr expression, islets were treated with both insulin and 2-deoxy-d-glucose. Insulin in the presence of nonmetabolizable glucose did not modify either Vdr or Ins mRNA levels (Fig. 3C and D).

Figure 3
Figure 3

Islet gene expression from fed and fasted Wt mice. A and B: Glycemia, Vdr expression analysis, and β-cell gene expression profile in islets from fed and fasted mice. Blood glucose levels (A) and gene expression in islets (B) from mice in fed and overnight-fasted conditions. Fed mice (white bars), fasted mice (orange bars). n = 6 per group. Results are mean ± SEM. *P < 0.05, ***P < 0.001 vs. fed group. C and D: Glucose and insulin effect on Vdr expression in culture islets. C: Vdr expression in islets incubated for 8 h with 2.5 or 9 mmol/L of glucose (Glc), 9 mmol/L of 2-deoxy-d-glucose (2-DG), and both 9 mmol/L 2-DG and 2 ng/mL of insulin (INS). D: Insulin expression in islets incubated for 8 h with 2.5 or 9 mmol/L of Glc, 9 mmol/L of 2-DG, and both 9 mmol/L 2-DG and 2 ng/mL of INS. Results shown represent the data obtained for at least six wells (100 islets per well)/condition and from three independent experiments. Data are expressed as mean ± SEM. A one-way ANOVA with Tukey post hoc analysis was used to determine statistical significance *P < 0.05, ***P < 0.001 vs. 2.5 mmol/L Glc and ##P < 0.01 vs. 9 mmol/L Glc.

Altogether, these results indicate that Vdr gene expression in islets is controlled by glucose, which supports the idea that VDR may play a role in β-cell function.

β-Cell VDR Overexpression Ameliorates Diabetes in Mice

To unravel the role of VDR in β-cells, Tg mice overexpressing VDR, specifically in β-cells, were generated. Three lines of Tg mice (Tg1, Tg3, and Tg4) were obtained that overexpressed murine Vdr under the control of RIP-I. Islets from Tg mice presented higher levels of Vdr mRNAs than Wt littermates (Fig. 4A and Supplementary Fig. 2A). Immunohistochemical analysis revealed that VDR overexpression was detected, specifically in insulin-positive cells (Fig. 4B and Supplementary Fig. 2B). In addition, glycemia and insulinemia remained unchanged in Tg mice when compared with Wt mice (Fig. 4C and D and Supplementary Fig. 2C and D). Since similar results were obtained in the three lines for a number of analyses and to avoid an unnecessary increase in the number of mice studied, Tg3 was selected for a further phenotyping. In accordance with normal glucose and insulin levels, both 4- and 8-month-old Tg mice exhibited normal glucose tolerance (Fig. 4E and Supplementary Fig. 3A). In addition, glucose-stimulated insulin release in 4-month-old mice was not significantly altered (Supplementary Fig. 3B), indicating that VDR overexpression did not modify β-cell function. To evaluate if VDR overexpression affected islet cell mass and distribution, β- and α-cell morphometric analyses were performed. Both 4- and 10-month-old Tg3 mice presented nonsignificant changes in β-cell mass compared with Wt littermates (Fig. 4F and Supplementary Fig. 3C). Immunohistochemical analysis against insulin revealed no differences in islet number (0.12 ± 0.01 and 0.08 ± 0.02 islet number/mg pancreas in Wt and Tg3, respectively) and β-cell distribution in the pancreas between Wt and Tg3 mice (Fig. 4G). Similarly, no alterations in α-cell mass and distribution were observed in 4- and 10-month-old mice (Fig. 4G and Supplementary Fig. 3D and E).

Figure 4
Figure 4

Generation of VDR Tg mice. Tg mice overexpressing murine Vdr cDNA under the control of the RIP-I were obtained. A: Islets Vdr gene expression. Vdr mRNA levels in islets from 2-month-old Wt (white bars) and Tg3 (gray bars) mice; n = 6 per group. Results are mean ± SEM. **P < 0.01 vs. Wt. B: Detection of VDR specifically in β-cell. Immunohistochemical analysis of insulin (red) and VDR (green) in pancreas of 2-month-old mice revealed β-cell–specific VDR expression. Original magnification ×20. C: Fed glycemia in Tg mice. Wt (white bars, n = 15) and Tg3 (gray bars, n = 10) mice. Results are mean ± SEM. D: Serum insulin levels. Insulin concentration was determined in fed conditions by radioimmunoassay. Wt (white bars, n = 15) and Tg3 (gray bars, n = 10) mice. Results are mean ± SEM. E: Glucose tolerance test (1 g/kg glucose), which was performed in 4-month-old Wt mice (white circles) and Tg3 mice (gray squares). F: β-cell mass in VDR Tg mice. β-cell mass was determined in Wt and Tg3 mice pancreas at the age of 4 months. Results are mean ± SEM. n = 4 per group. G: Pancreas immunohistochemical analysis. Immunohistochemical analysis of insulin and Gcg expression in Wt and Tg3 islets from 4-month-old mice. Scale bars, 100 μm.

Next, to examine the effects of VDR overexpression during the diabetic process, Wt and Tg3 mice received five daily consecutive injections of a standard dose of STZ (50 mg/kg) to induce diabetes. After STZ treatment, Wt mice displayed severe hyperglycemia, and 100% of mice developed overt diabetes. However, only 60% of treated Tg mice were diabetic and presented a less severe hyperglycemia at the end of the study (Fig. 5A and Supplementary Fig. 4A and B). In Tg3 mice, glycemia improvement was even more evident in fasting conditions (Fig. 5B). Similar results were obtained when STZ treatment was administered to Tg1 mice (Supplementary Fig. 4C and D). As previously shown, Vdr expression declined in islets from Wt mice after STZ treatment (Fig. 5C). In contrast, although Tg3 mice presented a similar Vdr reduction after STZ treatment, they sustained high Vdr expression levels (Fig. 5C).

Figure 5
Figure 5

Glycemia before and after STZ treatment (5 × 50 mg/kg). A: Evolution of fed glycemia. Glycemia before and after STZ treatment was measured. Wt (blue circles, n = 14) and Tg3 (pink squares, n = 13) mice. A two-way ANOVA with Tukey post hoc analysis was used to determine statistical significance. P < 0.01 was found in time, genotype, and time × genotype. B: Fasted glycemia 60 days after STZ treatment in Wt (white bars, n = 14) and Tg3 (gray bars, n = 13) mice. Results are mean ± SEM. *P < 0.05 vs. Wt. C: Islet Vdr gene expression. Vdr mRNA levels in islets from non-STZ-treated Wt (dotted bars) and Tg3 (striped bars) mice and from STZ-treated Wt (white bars) and Tg3 (dark gray bars) mice. Results are mean ± SEM. ***P < 0.001 vs. Wt. ###P < 0.001 vs. STZ-treated Wt . D: Serum insulin levels. Insulin was determined in fed conditions by radioimmunoassay. Wt (white bars) and Tg3 (gray bars) mice. Results are mean ± SEM. *P < 0.05 vs. Wt. E: β-cell mass in VDR Tg mice after STZ treatment. β-cell mass was determined 4 months after STZ treatment in Wt and Tg3 mice pancreas. Results are mean ± SEM. F: Pancreas immunohistochemical analysis. Immunohistochemical analysis of insulin expression in Wt and Tg3 islets 3 months after STZ treatment. Scale bars, 100 μm.

Immunohistochemical analysis against insulin also clearly revealed that after STZ treatment, Tg mice presented a higher number and area of insulin-positive islets in the pancreas compared with Wt mice, which showed few insulin-positive β-cells (Fig. 5E and F). These results correlated with higher insulin levels in Tg3 compared with Wt mice (Fig. 5D). The observed β-cell mass maintenance was parallel to a lower decrease in mRNA levels of β-cell gene markers such as Ins1, Slc2a2, and GcK after STZ treatment in Tg3 islets (Supplementary Fig. 4E).

The maintenance of pancreatic β-cell mass basically depends on two mechanisms: apoptosis and replication (30). Both mechanisms were explored in mice treated with STZ by morphometric analysis. TUNEL-positive β-cell percentage, an index of apoptosis, was similar in Wt and Tg3 mice, indicating that VDR overexpression did not protect β-cell from apoptosis (Fig. 6A). In contrast, Tg3 islets showed a clear increase in proliferation cell markers Pcna and CdK2 (Fig. 6B and C). In addition, a higher Ki67-positive β-cell percentage, which is a classical β-cell replication index, was detected in Tg3 islets compared with Wt (Fig. 6D and E). Thus, these results indicated that maintenance of the proliferation capacity may be involved in β-cell preservation in Tg3 mice after STZ damage.

Figure 6
Figure 6

β-cell mass in VDR Tg mice. A: β-cell apoptosis analysis. Quantification of apoptotic β-cells. The percentage of apoptotic β-cells was determined in Wt and Tg3 mice 40 days after STZ treatment. B and C: Islet cell cycle gene expression analysis. Pcna (B) and Cdk2 (C) gene expression was analyzed in islets from Wt and Tg3 mice 40 days after STZ treatment. Wt STZ-treated mice (white bars) and Tg3 STZ-treated mice (gray bars). Results are mean ± SEM. **P < 0.01 vs. Wt+STZ. D: Immunohistochemical detection of β-cell replication. Analysis of β-cell replication by double immunostaining with Ki67 replication marker (green) and insulin (red) in Wt and Tg3 islets 40 days after STZ treatment. Original magnification ×20. E: β-cell replication analysis. Quantification of β-cell replication. The percentage of replicative β-cell was determined in Wt and Tg3 mice 40 days after STZ treatment. Results are mean ± SEM. **P < 0.01 vs. Wt+STZ. F: β-cell gene profile analysis in islets after experimental diabetes induction. Gene expression in islets from 3-month-old mice 40 days after treatment with multiple doses of STZ (50 mg/kg body weight). Wt (white-dotted bars) and Tg3 (white-striped bars) non-STZ-treated mice; Wt STZ-treated (white bars) and Tg3 STZ-treated (dark gray bars) mice. Results are mean ± SEM. *P < 0.05, **P < 0.01, ***P < 0.001 vs. Wt, and #P < 0.05 vs. Wt+STZ.

The presence of inflammation was examined in islets by measuring islet proinflammatory and anti-inflammatory markers. Tg3 mice showed higher levels of anti-inflammatory A20 mRNA and a reduction of proinflammatory Tnfa, Tgfb, and H2-Aa mRNA levels related to Wt mice (Fig. 6F). In addition, Ucp2, which has been reported to be an inhibitor of A20 gene expression, was downregulated in Tg3 compared with Wt islets before and after STZ treatment (Supplementary Fig. 4F). Altogether, these data suggested that sustained VDR expression levels protected Tg mice to develop severe hyperglycemia, partially preserving β-cell mass and reducing local inflammation and diabetic consequences.

Since standard doses of STZ may have very toxic consequences (31), the effects of VDR overexpression were also explored in diabetic mice induced by very low doses of STZ (30 mg/kg) (five doses consecutive days) (+STZlow). After STZlow treatment, Wt mice developed hyperglycemia, and about 50% of them became overtly diabetic (Fig. 7A and B). In contrast, Tg3+STZlow mice maintained normoglycemia during all the study with no signs of diabetes (Fig. 7A and B). This improvement was also observed in fasted glycemia, which was clearly lower in Tg3+STZlow mice (Fig. 7C), and in insulin levels that were higher in Tg3+STZlow mice compared with Wt+STZlow mice (Fig. 7D) and were similar to Wt healthy mice (Fig. 4D). In addition, 2 months after STZlow treatment, Tg3+STZlow mice showed an improvement in glucose tolerance compared with Wt+STZlow mice (Fig. 7E). Gene expression analysis of islets at the end of the study revealed a decrease in Vdr expression in Wt+STZlow as observed previously (Figs. 5C and 8A), whereas Tg3+STZlow mice maintained high levels of Vdr, correlating with sustained levels of Ins expression (Fig. 8A). β-cell mass was also higher in Tg3+STZlow mice, accordingly to glucose and insulin values. Likewise, immunohistochemical analysis of the pancreas revealed few insulin-positive islets in Wt+STZlow mice, while Tg3+STZlow mice presented a normal number of insulin-positive islets according to β-cell mass data (Fig. 8B and C).

Figure 7
Figure 7

Glycemia before and after STZlow treatment (5 × 30 mg/kg). A: Evolution of fed glycemia (mean). B: Evolution of individual fed glycemia. Glycemia before and after STZlow treatment was measured. Wt (blue circles n = 9) and Tg3 (pink squares, n = 9) mice. In both A and B, two-way ANOVA with Tukey post hoc analysis was used to determine statistical significance. P < 0.05 (days 8, 21, and 37), P < 0.01 (days 55, 68, 80, 95, and 110) Wt responders vs. Tg3. C: Fasted glycemia. Fasted glycemia 60 days after STZlow treatment in Wt (white bars) and Tg3 (gray bars) mice. *P < 0.05 vs. Wt+STZlow. D: Serum insulin levels. Insulin was determined in fed conditions by radioimmunoassay. Wt (white bars) and Tg3 (gray bars) mice. Results are mean ± SEM. *P < 0.05 vs. Wt+STZlow. E: Glucose tolerance test (1 g/kg glucose), which was performed in Wt mice (white circles, n = 9) and Tg3 mice (gray squares, n = 9) 60 days after STZlow treatment. Results are mean ± SEM. *P < 0.05 vs. Wt+STZlow.

Figure 8
Figure 8

Gene expression analysis. Vdr and Ins gene expression in islets after experimental diabetes induction. A: Gene expression in islets from 3-month-old mice 40 days after treatment with multiple doses of STZ (STZlow) (30 mg/kg body weight). Wt (dotted bars) and Tg3 (striped bars) non-STZlow-treated mice, Wt (white bars) and Tg3 (dark gray bars) STZlow-treated mice. Results are mean ± SEM. A one-way ANOVA with Tukey post hoc analysis was used to determine statistical significance. **P < 0.01 vs. Wt untreated mice, ###P < 0.001 vs. Wt+STZlow-treated mice. B: Pancreas immunohistochemical analysis of β-cell mass in VDR STZ-treated mice. Immunohistochemical analysis of insulin expression in Wt and Tg3 islets 4 months after STZ treatment. Scale bars, 100 μm. C: β-cell mass, which was determined 40 days after STZlow treatment (30 mg/kg body weight) in Wt and Tg3 mice pancreas. Results are mean ± SEM. *P < 0.05 vs. Wt + STZlow.

Overall, these results provide evidence that sustained levels of VDR during the diabetic process revert hyperglycemia onset, improve glucose tolerance, and maintain β-cell mass.

Discussion

The role of vitamin D in the protection against diabetes is now widely accepted, although the underlying mechanisms are mainly unknown. In agreement with a protective role of vitamin D, vitamin D deficiency has been associated with diabetes (1). Similarly, polymorphisms in the Vdr gene, which lead to a significant decrease of mRNA and VDR protein levels, are a risk factor for diabetes development (32). Here, we clearly demonstrate that Vdr expression in mice is downregulated in islets during the development of both T1D and T2D. It is noteworthy that our results obtained from publicly available data sets also showed a trend toward a reduction in VDR expression in human islets from T2D patients. These results were consistent with previous reports showing a reduction in Vdr expression in pancreas from STZ-treated mice and rats (17). In this work, we provided evidence that Vdr reduction also occurs in islets from NOD mice that spontaneously develop diabetes and share many features with human T1D (33). In addition, we observed decreased Vdr levels in two different mouse models of T2D with β-cell hyperplasia (25,34).

Altogether, our results suggested that Vdr downregulation in β-cells may be a common feature of diabetes and that this reduction, along with a reduction in the expression of other β-cell markers, may be the result of β-cell loss/dedifferentation in diabetic islets, one of the common features of both T1D and T2D. VDR decrease was also associated with hyperglycemia, the major hallmark of diabetes. Indeed, Vdr expression was negatively correlated to circulating glucose levels in all the diabetic models studied here. Unexpectedly, we also showed that Vdr expression was decreased when circulating glucose levels were physiologically low, i.e., during fasting. Our results obtained from an in vitro study in cultured islets further indicated that Vdr expression was stimulated by glucose. Thus, although we cannot discard that decreased Vdr expression was due to β-cell loss during diabetes, our results may also be explained by the fact that diabetes is associated with low intracellular glucose levels resulting from decreased glucose uptake and a decrease in glucose sensing players, such as GLUT2 in β-cells (35).

Likewise, in fasted conditions, β-cells present similar low intracellular glucose. Thus, we can hypothesize that despite hyperglycemia, the low intracellular glucose levels may be responsible for Vdr downregulation. Accordingly, in islets from all of the diabetic mouse models analyzed, a reduction in expression of genes involved in glucose uptake in β-cells, the glucose transporter Slc2a2 and the glucose-phosphorylating enzyme Gck, was parallel to Vdr downregulation. In human islets from T2D patients, a parallel nonsignificant trend to decrease of SLC2A2 and VDR expression has also been observed. Thus, in humans, decreased glucose uptake in β-cells may also lead to a reduction in VDR expression. However, the mechanisms of regulation of β-cell VDR expression in humans remain to be elucidated. In addition, although our results suggest for the first time, that glucose metabolism may modulate positively Vdr expression, further studies are necessary to carefully explore this issue. It is also noteworthy that Vdr expression in islets declined in fasted compared with fed state, corroborating the importance of glucose metabolism for physiological regulation of Vdr expression. Accordingly, it has been reported recently that fasting-induced transcription factors repress vitamin D bioactivation (36). Thus, our findings reveal that both physiological acute changes in glucose concentration and pathophysiological disruption of glucose uptake alter Vdr gene expression in islets and may be responsible for Vdr decreased expression during diabetes.

Our results also support that VDR deficiency may have an impact on β-cell function alterations during diabetes. Results obtained from mice lacking functional Vdr indicate that VDR loss leads to a reduction in insulin mRNA levels and a deficit in insulin secretion (21). In accordance with this, we observed a decrease in insulin gene expression levels parallel to Vdr reduction in islets from diabetic mice. Moreover, treatment with vitamin D increased Ins1 mRNA expression in control animals, but this effect was lost in STZ-treated mice, possibly because of a decrease in Vdr expression in islets (17). These results suggest that VDR may be involved in Ins1 transcription in mouse pancreas (17). However, so far, no vitamin D response elements have been identified in the human or mouse insulin gene promoters. VDR-deficient mice are glucose intolerant, probably due in part to defects of the β-cell function, but the absence of VDR in other tissues may also contribute to their phenotype (21).

In this study, we found that Tg mice overexpressing VDR specifically in β-cells were resistant to the development of STZ-induced diabetes. Our results show a clear protective effect of sustained VDR levels in β-cell in front of STZ damage. It has been described that multiple doses of STZ induce diabetes through an increase in inflammatory cytokines, β-cell functional defects, and finally β-cell loss (37). Tg mice overexpressing VDR presented a preservation of β-cell mass, at least in part by maintaining β-cell replication capacity in front of STZ damage. In agreement with these results, it has recently been reported that VDR activation leads to an induction of β-cell replication (38). In addition, VDR overexpression partially protected Tg mice from islet inflammation, reducing the expression of inflammatory markers in islets. This reduction in inflammation and the subsequent protection against diabetes may have been mediated by VDR, as suggested by the fact that vitamin D, or its nonhypercalcemic analog, significantly inhibits insulitis and prevents or delays the onset of diabetes in NOD mice (13,39,40). In addition, the activation of VDR in human β-like cells is able to counteract the inflammatory response induced by cytokines and maintain β-cell functionality (38). In contrast, knockdown of VDR led to an increased cytokine-induced cell death in human β-like cells and to a reduction in the expression of key β-cell genes in cytokine-treated rat β-cells INS1 (38). All of these data suggest that VDR may reduce the inflammatory milieu in islets and therefore maintain β-cell mass and function resulting in a protection against diabetes development. Therefore, we demonstrated that sustained VDR levels in β-cells may protect against diabetes-induced damages by increasing β-cell functional target genes, decreasing inflammation and maintaining β-cell proliferation capacity.

Data from animal studies show that although vitamin D administration may delay the onset of diabetes in NOD mice, no or limited benefit has been observed by the administration of a vitamin D analog on β-cell damage after STZ treatment in mice (38). Interestingly, pharmacologically induced VDR signaling by a synthetic ligand in combination with a VDR-downstream modulator is able to partially restore β-cell function and glucose homeostasis in various T2D and T1D mouse models. Nevertheless, these effects were lost when the VDR synthetic ligand was used alone (38). In light of our results, this lack of effectiveness of vitamin D analogs or VDR ligands may be due to reduced Vdr expression levels in diabetic mice.

In humans, clinical data of effectiveness of vitamin D supplements are controversial. Although some data reveal beneficious effects of vitamin D supplementation on glucose metabolism (41,42), others report no effect of vitamin D supplements (43,44). The results of supplementation may be influenced by factors such as baseline vitamin D status, variability in dosages and forms of vitamin D used, duration of the intervention, or heterogeneity of the patients (45). It has also been suggested that genetic variation in VDR may influence metabolic effects of vitamin D supplementation in T2D (46). Our data clearly point out that variations in VDR expression in diabetic patients may also influence the outcomes of vitamin D supplementation. Therefore, a better knowledge of the VDR regulation in diabetes is necessary to define appropriate strategies of supplementation to improve glycemic control and metabolic alterations.

Although various studies described genetic variations in the VDR gene as a risk factor for T1D and T2D development, contradictory results about the association between VDR single nucleotide polymorphisms and T2D have been reported (4749). While genetic variations may not be responsible for diabetes development, a pathophysiological decrease in VDR expression may lead to a loss of protection against β-cell damage. As evidenced here, VDR expression maintenance might be essential to counteract β-cell damage in the diabetic process and to protect against diabetes development. Thus, future strategies for treatment of diabetes should be based on a better knowledge of mechanisms underlying VDR downregulation during diabetes and address restoration of VDR levels.

Article Information

Acknowledgments. The authors thank Malcolm Watford (Rutgers University, New Brunswick, New Jersey) and Ivet Elias (Universitat Autònoma de Barcelona, Bellaterra, Spain), for helpful discussion and Aina Bonet, Marc Leal, Albert Ribera, Marta Moya, Jennifer Barrero, and Lidia Hernández (Universitat Autònoma de Barcelona, Bellaterra, Spain) for technical support.

Funding. This work was supported by grants from FEDER/Ministerio de Ciencia, Innovación y Universidades – Agencia Estatal de Investigación (SAF2017-86166-R), Generalitat de Catalunya (ICREA Academia Award to F.B.), Spain. G.E. received a predoctoral fellowship from Generalitat de Catalunya, Spain.

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

Author Contributions. M.Mor. and L.V. designed experiments, generated reagents, performed experiments, and wrote and edited the manuscript. S.F. designed experiments and wrote and edited the manuscript. C.M. generated reagents, performed experiments, and contributed to discussions. G.E., T.F., M.Mol., and E.C. generated reagents and performed experiments. J.R. analyzed human data and contributed to discussion. A.P. generated transgenic mice and contributed to discussion. N.T. designed experiments and contributed to discussions. F.B. and A.C. designed experiments and wrote and edited the manuscript. A.C. is the guarantor of this work and, as such, had full access to all the data in the study and takes responsibility for the integrity of the data and the accuracy of the data analysis.

  • Received August 2, 2019.
  • Accepted February 14, 2020.



Sell Unused Diabetic Strips Today!

Low-Carb General Tso’s Chicken – Diabetic Foodie

By electricdiet / June 22, 2020


Satisfy your takeout cravings without the unnecessary carbs! You’ll love the sweet flavors and crispy texture of this low-carb General Tso’s chicken.

Low-Carb General Tso's Chicken on a plate with steamed broccoli

Who doesn’t love General Tso’s chicken? This sweet and crispy dish is a westernized version of Chinese food, but it’s one of the most popular carry-out dishes.

Unfortunately, it’s not very heart-healthy or diabetic-friendly. The chicken is usually deep-fried and then coated in a sticky-sweet sauce. This means that it’s packed with calories, carbs, sugar, and sodium.

So when I saw this low-carb General Tso’s chicken in Ching’s Everyday Easy Chinese by Ching-He Huang, I knew I had to try it!

Ching’s recipe is quick and easy to make, and the result is absolutely delicious. It’s less sweet than restaurant versions, which I liked. It allowed some of the other flavors to shine.

It’s also a bit spicy, but not overpowering. Just make sure you don’t eat the chiles!

Her recipe does call for yellow bean sauce, which I couldn’t find at any local markets. But the glossary in the back of her book says yellow bean sauce is “made from fermented yellow soy beans, dark brown sugar and rice wine.” Once I read that, I decided hoisin sauce would be a decent substitute.

If you’ve been missing Chinese carryout, definitely give this healthier General Tso’s chicken a try!

How to make low-carb General Tso’s chicken

This dish comes together with just a bit of prep. Then, simply add everything to your pan and cook!

You can see how i make the recipe in this short video or follow the step-by-step instructions below.

Step 1: In a small bowl, combine the ingredients for the sauce. Mix well and set aside.

Step 2: Place the cubed chicken in a medium bowl and season with salt and white pepper. Add the cornstarch, mix well, then set aside.

Step 3: Heat a wok or large pan over high heat. Once hot, add the peanut oil.

Step 4: Add the garlic and red chiles, stirring for a few seconds, then add the chicken and stir-fry for a couple of minutes.

Step 5: When the chicken starts to turn opaque, add the sherry and cook for 2-3 more minutes.

Step 6: Add the sauce and bring the mixture to a boil.

Step 7: Reduce the heat to medium and continue to simmer until the chicken is cooked through and the sauce has thickened, another minute or two.

Step 8: Turn off the heat, discard the red chiles, and stir in the scallions to serve.

You can choose to leave the red chiles if you want a pop of color on your plate, just be sure not to eat them!

Did you know that General Tso’s chicken in a typical Chinese restaurant has about 850 calories, 40g fat, and 60g carbs?!

This recipe, on the other hand, has about 275 calories, 9 grams of fat, and 18g carbs per serving. How much better is that?

The whole recipe is for 2 servings, so even if you ate the whole thing, you’d still be better off than ordering from a restaurant!

I do recommend using low-sodium tamari or soy sauce to minimize the sodium in this dish. I prefer tamari to traditional soy sauce because it seems less salty to me and it contains no wheat.

If you’re really trying to lower your sodium, you could also reduce or omit the salt used to season the chicken at the beginning.

Looking for some tasty sides to serve with your General Tso’s chicken?

You can never go wrong adding steamed veggies to your plate. I love eating this dish with a side of broccoli or edamame!

The broccoli is also great for sopping up any leftover sauce.

Storage

This tasty chicken dish is best served right away. However, if you have leftovers, you can store them in an airtight container in the refrigerator for 3-4 days.

If you know you won’t eat both servings, I would recommend leaving the scallions off the portion you plan to save.

Scallions are much better fresh, so if you can, don’t add them until right before you serve this dish!

Other healthy chicken recipes

Chicken breast is such a great protein. Not only is it super lean and healthy, but there are also so many ways to cook it! If you’re looking for a few more chicken breast recipes, here are some of my favorites:

When you’ve tried this dish, please don’t forget to let me know how you liked it and rate the recipe in the comments below!

Recipe Card

Low-Carb General Tso's Chicken

Low-Carb General Tso’s Chicken

Satisfy your takeout cravings without the unnecessary carbs! You’ll love the sweet flavors and crispy texture of this low-carb General Tso’s chicken.

Prep Time:5 minutes

Cook Time:5 minutes

Total Time:10 minutes

Author:Shelby Kinnaird

Servings:2

Instructions

  • In a small bowl, combine the ingredients for the sauce. Mix well and set aside.

  • Place the cubed chicken in a medium bowl and season with salt and white pepper. Add the cornstarch, mix well, then set aside.

  • Heat a wok or large pan over high heat. Once hot, add the peanut oil.

  • Add the garlic and red chiles, stirring for a few seconds, then add the chicken and stir-fry for a couple of minutes.

  • When the chicken starts to turn opaque, add the sherry and cook for 2-3 more minutes.

  • Add the sauce and bring the mixture to a boil.

  • Reduce the heat to medium and continue to simmer until the chicken is cooked through and the sauce has thickened, another minute or two.

  • Turn off the heat, discard the red chiles, and stir in the scallions to serve.

Recipe Notes

This recipe is for 2 servings of General Tso’s chicken. Do not eat the red chiles.  Leftovers can be stored covered in the refrigerator for 3-4 days.

Nutrition Info Per Serving

Nutrition Facts

Low-Carb General Tso’s Chicken

Amount Per Serving (1 serving)

Calories 313 Calories from Fat 74

% Daily Value*

Fat 8.2g13%

Saturated Fat 1.2g8%

Trans Fat 0g

Polyunsaturated Fat 2.2g

Monounsaturated Fat 3.1g

Cholesterol 86.7mg29%

Sodium 990.2mg43%

Potassium 463.8mg13%

Carbohydrates 18.3g6%

Fiber 1g4%

Sugar 10.4g12%

Protein 33.6g67%

Vitamin A 0IU0%

Vitamin C 0mg0%

Calcium 0mg0%

Iron 0mg0%

Net carbs 17.3g

* Percent Daily Values are based on a 2000 calorie diet.

Course: Main Course

Cuisine: Chinese

Diet: Diabetic

Keyword: easy dinner recipes, General Tso’s Chicken, low-carb, low-carb chicken



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Pork and Mushroom Wontons – My Bizzy Kitchen

By electricdiet / June 20, 2020


I love when I can make one dish and morph it into a few things later in the week, like these pork and mushroom wontons.

I love wontons so much because the possibilities are endless.  Have some protein and veggies?  Wrap it up in a wonton.  I bought 1/2 a pound of ground pork from Mariano’s – if you stick to chicken or beef, please try ground pork – so much flavor!

These are the wonton wrappers I use.  They seem to be a tiny bit thicker and easier to shape without tearing.  You can find wonton wrappers in the produce section of most grocery stores.  At Mariano’s, they are on the top shelf right above the packaged mushrooms.

If you don’t know how to shape wontons, check out youtube.  You can basically find out how to do anything on youtube.  I am a visual learner and it took me just two tries to get the perfect wonton.

You’re welcome for the no bra and tank top. 😀

One of the biggest things I’ve learned in cooking is: TASTE YOUR FOOD AS YOU GO.  Yes, I am shouting because I was the queen of following a cookbook recipe and not tasting a damn thing until I plated it for dinner.  So many times I under-seasoned, or over-seasoned, or wish I would have added more spice – you get the idea.

Especially when making a big batch of something (this wonton recipe below makes 44 wontons!) always cook off a tiny bit and give it taste.  First go around I was missing both salt, a bit of acid and heat.  I sprinkled in a bit more salt, a tiny bit of rice wine vinegar and some sambal oelek to the filling.  My second test – perfect!

How pretty is that?!

The filling will actually keep in your fridge for a few days, so if you don’t have the time or help to make 44 wontons at once, just make as many as you want.  I made 7 wontons a serving, but you do you.  The first photo above, I brought water to a boil, and then turned the heat down and boiled at a simmer for 3 minutes.   

I posted the first picture of the boiled wonton, and the immediate response was “hey Biz, how would you air fry those?”  Um, I had no idea!  The next day I formed the wontons above (not boiled) and sprayed with avocado oil spray and air fried at 360 for 6 minutes.  Crunchy, delicious and tasted like the best appetizer you could have at a restaurant.

The day after I made the wontons, I decided to make wonton soup for dinner.  The broth recipe will be in the recipe box below, but holy balls was this packed with flavor.  Sautéed mushrooms and garlic were added to spicy broth, then add in the pork wontons and a drizzle of sweet chili sauce – #swoon.  I could probably eat that for lunch every day and not get sick of it.

Print

Pork and Mushroom Wontons

These pork and mushroom wontons could really be filled with anything you have in your fridge – wonton wrappers are so versatile and cheap!  I can buy a 50 count wonton package for $1.99 at Mariano’s – found on the top shelf near the packaged mushrooms.

  • Author: Biz
  • Prep Time: depends on how many you make!
  • Cook Time: 3 minutes boiled, 6 minutes air fried
  • Total Time: 44 minute
  • Yield: 44 wontons 1x

Scale

Ingredients

44 wonton skins ( used @friedasproduce wrappers found in the produce section @marianosmarket)
1/2 pound ground pork
4 ounces mushrooms
2 cloves garlic
1 cup chopped spinach
1 cup grated carrots
1/2 teaspoon Five Spice seasoning
1 teaspoon szechuan peppercorns, crushed
1 teaspoon mirin
1 teaspoon soy sauce
1 tablespoon sambal oelek
1/2 teaspoon salt
For the dipping sauce:
1 teaspoon sambal oelek
1 tablespoon rice wine vinegar
1 tablespoon sweet chili sauce

Instructions

Saute the mushrooms and garlic for 4-5 minutes. While that cooks, make the dipping sauce. Let mushroom mixture cool and chop. Add remaining ingredients and use one teaspoon of filling per wonton. Fold in half like an envelope, then, wet the corners of the folded end and bring together and crimp. Bring water to boil, then lower temp and boil the wontons for 3 minutes.

To add the wontons to soup:

Wonton Broth For Two:

4 cups chicken broth
1 teaspoon chicken base
1 teaspoon sambal oelek (crushed red pepper is a good substitute)
1/2 teaspoon szechuan peppercorns, crushed
1 tablespoon mirin
1 tablespoon rice wine vinegar
1/2 cup mushrooms that are sauteed with 1 teaspoon minced garlic

In a skillet, saute mushrooms and garlic for 5-6 minutes. Roughly chop and set aside. Add the chicken broth through rice wine vinegar and cook for 10 minutes over medium low heat. Add in the garlic mushrooms.

To serve: put 1/2 cup chopped baby spinach in your bowl, add in 2 cups of the wonton broth and add your steamed wontons to the bowl. Enjoy!

Notes

To make a whole batch, place all the wontons on a baking sheet, freeze for 30 minutes and store in a zip top bag. Frozen would take probably 5 minutes to boil. Serve with a drizzle of dipping sauce and veggies on the side.

No matter which WW plan you are on, each wonton is .8 points – or if you want to round up to 1, you do you!  

I hope you give these a try.  Leave me a comment and let me know if you would prefer the steamed or fried version!  It’s pretty much a tie for me, but I now have thoughts on making fried crab rangoon wontons – stay tuned for that!





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10 Low-Carb Muffin Recipes | Diabetes Strong

By electricdiet / June 18, 2020


These low-carb muffin recipes are super delicious, easy to make, and perfect for a healthy breakfast or snack on the go!

10 low-carb muffin recipes pin

Why low-carb muffins?

Everybody loves a delicious muffin. Unfortunately, the amount of sugar and carbs you find in traditional muffins makes them more like a dessert than a healthy breakfast.

If you want to be able to enjoy a muffin for a quick morning or afternoon bite without the guilt, the key is to find low-carb muffin recipes!

These healthier recipes use low carb sweeteners and flours that won’t cause a spike in blood sugar. That way, you can enjoy these tasty treats any time.

Diabetes-friendly muffin recipes

If you’ve been looking for some great diabetic or keto muffin recipes, the good news is that there are tons to choose from!

Whether you’re looking for something simple like chocolate or an exciting new flavor combo like raspberry pumpkin, you’ll find plenty of inspiration.

Give these healthy and low-carb muffin recipes a try and enjoy these tasty treats without the guilt!

With so many delicious options, you can try a new flavor of low-carb muffins every week!

More low-carb recipe roundups

Want to check out more of my favorite recipes? Here are a few collections I think you’ll enjoy:

If you try any of these recipes, don’t forget to leave a comment below and let me know how you liked them!



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Cinnamon Rolls with Biscuits Make Easy Cinnamon Rolls Recipe with Kids

By electricdiet / June 16, 2020


Cinnamon Rolls with Biscuits – Easy Cinnamon Rolls Recipe Fun For Kids

Cinnamon Rolls with biscuits make favorite easy cinnamon rolls recipe. In fact, you probably have these four ingredients for Easy Cinnamon Rolls from KITCHEN 101 cookbook already in your panty to easily make cinnamon roll biscuits. You can whip up this simple cinnamon roll breakfast and the kids love to help make them.  Watch and you will see if you involve the kids, usually they will quickly eat it.

Cinnamon Rolls with Biscuits Best Easy Cinnamon Rolls Recipe!

You might have an easy cinnamon rolls recipe but you will really like these cinnamon rolls with biscuits.  First, you can be creative with the recipe and add nuts, dried fruit or whatever extra ingredients you like.  Also, believe it or not, this is a diabetic recipe.  In KITCHEN 101, all recipes have nutritional information and have a “D” to highlight all diabetic recipes.  There are different size biscuit cans but the analysis is based on the 10-count can of biscuits.  Also, look for whole wheat biscuits to use.

Easy Cinnamon Rolls
Keep canned biscuits handy to make this fast and fabulous treat for breakfast or a snack.

    Servings10 rolls

    Ingredients

    • 1


      can refrigerated biscuits or whole wheat biscuits10-biscuit

    • 2tablespoons


      butter

    • 1tablespoon


      sugar

    • 1teaspoon


      ground cinnamon

    • 1/4cup


      chopped pecansoptional

    Instructions
    1. Preheat oven 425°F. Coat 15x10x1-inch baking sheet with nonstick cooking spray.

    2. Flatten each biscuit with your hand or rolling pin. Spread each biscuit with butter.

    3. In small bowl, combine sugar and cinnamon together. Sprinkle cinnamon mixture on top of butter; sprinkle with pecans, if desired.

    4. Roll up each biscuit like a cigar and form a circle by putting the ends together. Bake 8-10 minutes or until golden brown.

    Recipe Notes

    Calories 76, Calories from Fat 35%, Fat 3g, Saturated Fat 1g, Cholesterol 6mg, Sodium 210mg, Carbohydrates 11g, Dietary Fiber 0g, Total Sugars 3g, Protein 1g, Dietary Exchanges: 1 starch, 1/2 fat

    Terrific Tip: These are freezer-friendly so keep in freezer to have to pop out for a quick snack.

    Nutrition Nugget: Light, low-fat meals, especially breakfast foods seem to be the best tolerated of all foods while you are having chemotherapy.

    Another Fun Biscuit Recipe Bunny Biscuits

    Another fun kid biscuit recipe is for Holly’s popular Bunny Biscuits. Don’t save them only for Easter. They are fun and so good to eat year round!

    Get All of Holly’s Healthy Easy Cookbooks

    The post Cinnamon Rolls with Biscuits Make Easy Cinnamon Rolls Recipe with Kids appeared first on The Healthy Cooking Blog.



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    Hyperuricemia Predisposes to the Onset of Diabetes via Promoting Pancreatic β-Cell Death in Uricase-Deficient Male Mice

    By electricdiet / June 14, 2020


    Introduction

    Hyperuricemia (HU), a metabolic condition characterized by elevated serum urate (SU), is a primary cause of gout. Epidemiological studies have demonstrated the rapid increasing prevalence of HU worldwide in the last decades (13). Although some recent studies have highlighted the connection between SU and glucose homeostasis and indicated that each 1 mg/dL increase in SU was accompanied by a 17% increase in the risk of type 2 diabetes (4,5), there is no conclusion of a causal relationship between HU and diabetes. The Mendelian randomization study by Sluijs et al. (6) did not support a causal effect of circulating urate on diabetes onset and thus concluded that urate-lowering therapies (ULTs) may therefore not be beneficial in reducing diabetes risk. Therefore, whether or how SU is involved in disrupting glucose homeostasis remains elusive. Moreover, the investigation of the possible relationship between urate and glucose metabolism has been hindered by the lack of an appropriate animal model.

    The etiology of diabetes is multifactorial, including insulin resistance, defective insulin secretion, and loss of β-cell mass through β-cell apoptosis. Adequate insulin secretion from pancreatic β-cells is necessary to maintain blood glucose homeostasis. Notably, the apoptosis and the subsequent loss of function of pancreatic β-cells is the major contributor to the progression of diabetes. Although the relationship between HU and diabetes has been implicated, the molecular underpinnings of diabetic β-cell apoptosis promoted by HU remain poorly understood. One piece of evidences from Jia et al. (7) demonstrated that soluble urate can directly cause β-cell death and dysfunction by activation of the nuclear factor-κB–inducible nitric oxide synthase–nitric oxide (NF-κB–iNOS–NO) signal axis in vitro. Thus, understanding the mechanisms of β-cell survival in basal or stress conditions associated with HU is imperative for creating new strategies to prevent and manage diabetes, especially for HU individuals.

    Uricase (Uox) expressed in rodents can further degrade uric acid into allantoin (8), which has hindered the establishment of suitable rodent models for HU (9). We previously established a spontaneous HU mouse model with Uox gene deficiency that is characterized by long-term stable SU levels (7–9 mg/dL) (10). This animal model lays a foundation for further glycometabolism investigations. In the current study, we address the following questions: 1) whether the Uox-KO mouse develops spontaneous glucose abnormities (such as insulin resistance, compromised β-cell functions) and even diabetes, 2) whether HU imposes stresses on glucose phenotypes or pancreatic β-cells with additional high-fat diet (HFD) and/or streptozotocin (STZ) stimulation in the Uox gene-deficiency mouse model, and 3) how urate works in mouse isolated islets if we have evidence that urate attacks pancreatic β-cells.

    Research Design and Methods

    Animals

    Uox knockout (KO) mice and their wild-type (WT) counterparts (C57BL/6J background) were generated as previously described (10). Briefly, a region of 28 base pairs in exon 3 of the Uox gene was deleted using the transcription activator-like effector nuclease technique. The animals were maintained in a temperature-controlled room (22°C), with humidity at 55%, and on a 12-h light-dark cycle (lights on from 7:00 a.m. to 7:00 p.m.) under specific pathogen-free conditions. After a 2-week acclimation period, 8-week-old mice were randomly assigned to two groups (1:1) and fed with an HFD (45% total fat, 35% protein, and 20% carbohydrate) or a regular chow diet ad libitum with free access to water for 20 weeks.

    To provoke the potential role of urate in animals, we fed mice the HFD (45% fat, 35% protein, and 20% carbohydrate) for 1 week and then injected them with multiple low-dose STZ (MLD-STZ, 40 mg/kg body wt i.p.) daily for 5 consecutive days. Specifically, STZ was freshly dissolved in 0.1 mol/L citrate buffer (pH 4.5). For comparison, mice were administered STZ and fed a normal diet. Random-fed (9:00–10:00 a.m.) blood glucose levels were determined by glucometer (ACCU-CHEK Inform; Roche Pharmaceuticals). To evaluate the effect of ULT, pegloticase (also known as Krystexxa or Puricase), purchased from Horizon Pharma, was administrated to mice by tail vein injection at a dose of 0.5 mg/kg, with the first injection on the first day of the MLD-STZ intervention, and the second injection was on the 10th day of the MLD-STZ intervention.

    Male mice were used for all studies shown, and the age of mice is indicated in the figures. This study was approved by the Affiliated Hospital of Qingdao University Animal Research Ethics Committee.

    Blood Biochemistry

    Mice were fasted overnight before serum biochemical tests. Blood was collected from the outer canthus the next morning. SU levels were measured immediately from the serum of anesthetized breathing mice (11), using an automatic biochemical analyzer (Toshiba, Tokyo, Japan). Serum creatinine levels and lipid profiles, including total cholesterol (TC), triglycerides (TG), and HDL and LDL cholesterol were assessed by the automatic biochemical analyzer (Toshiba) as well. Blood glucose levels were monitored by tail bleeding with a glucometer (ACCU-CHEK Inform). Diabetes was defined as a random blood glucose of ≥16.7 mmol/L (12).

    Glucose Tolerance Test, Insulin Tolerance Test, and Glucose-Stimulated Insulin Secretion

    Mice were fasted for 8 h before a glucose tolerance test (GTT) or insulin tolerance test (ITT) and then injected with d-glucose at 2 g/kg body wt i.p. or 1 unit/kg insulin (Humulin R; Eli Lilly). Blood was collected at predetermined times (0, 15, 30, 60, and 120 min) after the glucose injection, and the blood glucose levels were determined using a glucometer (ACCU-CHEK Inform).

    Glucose-stimulated insulin secretion (GSIS) testing in vivo and in vitro was performed. Briefly, mice were fasted for 8 h and injected with 2 g/kg d-glucose i.p., and serum insulin levels were tested at 0, 15, 30, and 60 min for in vivo GSIS. For in vitro GSIS, isolated islets were purified and harvested by handpicking under a stereomicroscope as described below. The islets (10 per well) were seeded in 24-well plates and then cultured in complete RPMI 1640 (Gibco, Life Technologies) with 10% FBS (HyClone; GE Healthcare, Little Chalfont, U.K.) in 5% CO2 at 37°C overnight. After incubation for 1 h in glucose-free Krebs buffer (115 mmol/L NaCl, 4.7 mmol/L KCl, 1.2 mmol/L MgSO4, 1.2 mmol/L KH2PO4, 20 mmol/L NaHCO3, 16 mmol/L HEPES, 2.56 mmol/L CaCl2, and 0.2% BSA), the islets were treated for 1 h in Krebs buffer with low (3.3 mmol/L) or high (16.7 mmol/L) concentrations of glucose. After treatment, the supernatants were obtained for determination of insulin concentration with an ultrasensitive ELISA kit (ALPCO Diagnostics, Salem, NH).

    Pathology Analysis

    Mice were sacrificed to extract their pancreas for the pathology analysis. The pancreas was immediately dissected, weighed, and fixed in 10% formalin on ice for 30 min, followed by paraffin embedding of 5-μm serial sections. Tissue serial sections were stained by hematoxylin-eosin (HE) and separately incubated with anti-insulin (1:200) (ab7842; Abcam) rabbit polyclonal antibodies for immunohistochemistry. Primary islet apoptosis was analyzed by the terminal deoxynucleotidyl TUNEL technique according to the manufacturer’s instructions (catalog no. 11684795910; Roche). The samples were stained with DAPI to visualize total cells. Pancreatic sections were also immunostained with anti-insulin antibody to identify β-cells. β-Cell fractional area was determined by quantifying the percentage of insulin-positive pancreas area as a total of the full pancreas area for each section, followed by averaging of six sections per mouse. Images were analyzed using the Pannoramic Digital Slide Scanner. The insulin-positive areas were analyzed by Image-Pro Plus 6 software. β-Cell mass was calculated by multiplying the β-cell fractional area with the initial pancreatic wet weight.

    The kidneys were removed, fixed in 10% formalin, embedded in paraffin, and then cut into 4-μm sections. The sections were used for HE, periodic acid Schiff–methenamine (PASM) staining, and periodic acid Schiff (PAS) staining. The percentage of tubule injuries was assessed by scoring six renal cortical tubule sections in randomly selected fields for each group. The glomerulosclerosis index (GSI) was calculated as [(1 × N1) + (2 × N2) + (3 × N3) + (4 × N4)]/(N0 + N1 + N2 + N3 + N4), where Nx was the number of glomeruli with each given score (0 for normal glomeruli, 1 for up to 25% involvement, 2 for up to 50% involvement, 3 for up to 75% involvement, and 4 for >75% sclerosis). The average GSI was analyzed based on three given PAS staining sections for each group. Renal crystal sections were obtained from absolute ethanol-fixed kidneys to detect urate crystals under polarized light. The urate crystal areas in each group were calculated by the Image-Pro Plus 6.0 system.

    Islets Isolation

    Pancreatic islets were isolated from mice with digestive enzyme (13,14). Briefly, a mouse was euthanized with CO2, followed by cervical dislocation, and then placed supine under a stereomicroscope, with the abdomen cleaned with 75% ethanol. A laparotomy was performed by cutting the skin and the muscular tissue of the thorax with a V-incision from the pubic region up to the diaphragm to expose the abdominal cavity. The common bile duct close to the duodenum was ligated for the retrograde puncture of the common bile duct, followed by a slow perfusion of 3 mL prechilled collagenase IV (catalog no. C5138S; Sigma-Aldrich) at 0.5 mg/mL concentration (dissolved in Hanks’ balanced salt solution) to fully expand the pancreatic body and tail. The pancreas was then excised and digested at 37°C for 11 min. Islets were purified in Histopaque-1077 (catalog no. 10771; Sigma-Aldrich) by vortexing gently for several seconds.

    Microarrays

    RNA Isolation and Quantification

    Isolated islets from mice were prepared for the following RNA extraction. Briefly, total RNA was extracted and purified using RNeasy Micro Kit (catalog no. 74004; QIAGEN), following the manufacturer’s instructions, and checked for a registrant identification number to inspect RNA integration by an Agilent Bioanalyzer 2100 (Agilent Technologies). Only those samples with a 260 nm–to–280 nm ratio between 1.8 and 2.1, a 28S-to-18S ratio within 1.5 and 2.0, and gel electrophoresis that showed clear 28S and 18S bands and a weak 5S strip were further processed.

    Library Preparation

    Total RNA samples (100–1,000 ng) were polyA enriched, reverse-transcribed into double-stranded cDNA, and then labeled by Low Input Quick Amp Labeling Kit (catalog no. 5190-2305; One-Color; Agilent Technologies) following the manufacturer’s instructions. Each slide was hybridized with 600 ng Cy3-labeled cRNA using the Gene Expression Hybridization Kit (catalog no. 5188-5242; Agilent Technologies) in a hybridization oven. Microarray was performed on Agilent platform using Agilent SurePrint G3 Mouse Gene Expression Microarray 8 × 60K chips (Agilent Technologies).

    Data Analysis

    Data were extracted with Feature Extraction 10.7 software (Agilent Technologies). We used a cutoff of normalized array values (log2-transformed values >2.0 or <0.5) for islet tissue transcripts. Raw data were normalized by quantile algorithm. Limma and agilp packages were loaded in R 3.4.1 software.

    Statistical Analysis

    All experimental statistical analyses were performed using GraphPad Prism 8 software (GraphPad). For two-group comparisons, the Student t test was used. For multiple comparisons, one-way ANOVA, followed by a Dunnett test, was used to compare each group versus a vehicle-treated group. Data are presented as mean ± SEM. Differences with P < 0.05 were considered statistically significant.

    Results

    Uox-KO Mice Develop Glucose Intolerance but Not Diabetes

    In our previous study, we successfully established the mouse model with Uox gene (which is mainly expressed in liver) (Supplementary Fig. 1) deficiency to mimic human HU (10). Shown in Fig. 1A, SU levels in Uox-KO male mice were dramatically increased compared with WT mice (P < 0.001) and stabilized at elevated levels (7–9 mg/dL) from 8 to 56 weeks of age. No difference in body weight was observed between age-matched Uox-KO and WT mice (Fig. 1B). HU did not alter fasting blood glucose or plasma insulin levels even with aging (from 8 to 56 weeks) (Fig. 1C and D). As for lipid profiles, no difference was shown in TC or HDL and LDL cholesterol between Uox-KO males and their WT counterparts, whereas TG were significantly lower in Uox-KO mice compared with WT mice at each age point (P < 0.001 at 8 weeks, 24 weeks, and 56 weeks) (Fig. 1E–H).

    Figure 1
    Figure 1

    General characteristics of Uox-KO male mice and WT counterparts stratified by age. A: SU levels were measured at the age of 8, 24, and 56 weeks in KO and WT mice (n = 9). B: Body weights were evaluated accordingly (n = 8). Fasting blood glucose (C) and fasting plasma insulin (D) were detected (n = 5–8). Lipid profiles, including TC (E), TG (F), LDL cholesterol (G), and HDL cholesterol (H) were determined in 8-, 24‐, and 56-week-old mice (n = 6–8). ***P < 0.001. Data are expressed as mean ± SEM.

    GTT and ITT were performed in mice to further assess the impact of HU on glucose homeostasis. In basal condition, Uox-KO males showed a significant impairment in glucose tolerance, indicated by remarkable elevated blood glucose concentrations at 15 and 30 min after 2 g/kg glucose administration at 8 and 24 weeks of age (Fig. 2A and B) and at 30 min when they were 56 weeks old (Fig. 2C). ITT revealed no insulin resistance in Uox-KO males, even with prolonged HU stress (Fig. 2D–F). These data show Uox-KO male mice do not develop diabetes spontaneously though with aging.

    Figure 2
    Figure 2

    HU impairs glucose tolerance in Uox-KO male mice. GTT were performed in Uox-KO mice and WT controls with a 2 g/kg glucose i.p. injection after 8 h fasting at the age of 8 weeks (A), 24 weeks (B), and 56 weeks (C), separately (n = 6). DF: Insulin (1 unit/kg) was administrated i.p. after 8 h fasting for ITT in 8-, 24‐, and 56-week-old mice (n = 6). *P < 0.05, **P < 0.01. Data are expressed as mean ± SEM.

    HU Does Not Impair Insulin Sensitivity in HFD-Fed Mice

    We fed Uox-KO male mice and the control WT littermates with the HFD or the normal chow diet, starting at 8 weeks of age, and continued for 20 weeks. Both genotypes displayed comparable SU levels and circulating lipids (including TC and HDL and LDL cholesterol) before and after the experimental feeding (Table 1). The HFD elevated TC, TG, and LDL cholesterol and decreased HDL cholesterol in both genotypes (Table 1). After 20 weeks of exposure to the HFD, the WT mice showed increased body weight compared with the Uox-KO mice (Table 1). Although plasma insulin levels displayed a significant elevation in HFD-fed WT mice compared with chow diet–fed mice, the HFD-fed Uox-KO mice showed a dramatic reduction in plasma insulin levels compared with the HFD-fed WT mice (Table 1). This result indicates a failure of compensatory insulin production by pancreatic β-cells in Uox-KO mice.

    Table 1

    Body weight and serum biochemical profiles of Uox-KO and WT mice after 20 weeks on an HFD or normal chow diet

    We also performed GTT and quantified the area under the curve (AUC) that integrates the values from 0 to 120 min of GTT. After 20 weeks of being fed with the chow diet or HFD, blood glucose in WT and Uox-KO male mice was restored to normal concentrations at 120 min after the glucose challenge (Fig. 3A), despite that a higher peak at the 30-min point was observed in HFD-fed Uox-KO mice. However, after 20 weeks of being fed the HFD, both genotypes exhibited a retarded glucose clearance compared with mice fed the normal chow diet (Fig. 3A). Remarkably, the blood glucose levels in HFD-fed Uox-KO mice were sustained at a dramatically higher level over the whole course of GTT compared with HFD-fed WT controls (Fig. 3A), resulting in a significant increase in the AUC of the GTT (Fig. 3B) and indicative of severe glucose intolerance.

    Figure 3
    Figure 3

    HU strengthens glucose intolerance with HFD stimuli. A: GTTs were used for chow diet– and HFD-fed 8-week-old Uox-KO male mice and WT controls (n = 8). B: AUC integrated from the period of 0–120 min was calculated based on the GTT curves. ITTs were monitored (C), and corresponding AUCs were determined (8-week-old mice, n = 8) (D). E: Plasma insulin levels were measured with plasma collected at 0, 15, 30, and 60 min after glucose injection (2 g/kg i.p.) in 8-week-old Uox-KO mice and WT mice (n = 6 in each group). F: Supernatant insulin levels were determined after in vitro GSIS with isolated pancreatic β-cells (n = 6 in each group). *P < 0.05, **P < 0.01, ***P < 0.001. Data are expressed as mean ± SEM.

    The development of glucose dysmetabolism is ultimately attributable to impaired insulin action or insufficient insulin production, or both. To clarify whether insulin resistance accounts for the glycol-metabolic disorders in HFD-fed Uox-KO mice, we used an ITT. Compared with the chow diet, the disappearance of glucose after the insulin challenge was comparable by the HFD (Fig. 3C), displaying no insulin resistance in diet-induced Uox-KO male mice. The AUC of the ITT was comparable between HFD-fed WT and Uox-KO mice as well (Fig. 3D), suggesting an equivalent degree of insulin resistance in the two genotypes.

    Islet function was determined by in vivo and in vitro GSIS. Similar to the results of Uox-KO mice islet function in our previous study (10), HFD-fed Uox-KO mice secreted a significantly lower amount of insulin at 15 and 30 min (Fig. 3E), consistent with the GTT results above. Although isolated islets derived from HFD-fed Uox-KO mice secreted a similar amount of insulin as HFD-fed WT islets at low glucose concentration (3.3 mmol/L), insulin secretion was decreased by 22.6% in HFD-fed Uox-KO islets at a high glucose concentration (16.7 mmol/L), as shown in Fig. 3F. Collectively, these data suggest that HU impairs GSIS in HFD-fed Uox-deficient males, contributing to increased glucose intolerance.

    HU Induces Diabetes With External MLD-STZ Stimulation

    Given HU compromised β-cell functions and impaired glucose tolerance, we suppose urate worsens hyperglycemia in an established model of diabetes induced by HFD + MLD-STZ (40 mg/kg/day for 5 days). Pegloticase, an approved recombinant porcine-like uricase drug, was indicated for the treatment of HU mice. As indicated by Fig. 4A, the random glucose levels in STZ-induced diabetic Uox-KO (STZ-KO) mice elevated markedly on day 7, and then severe hyperglycemia developed, which over time resulted in uncontrolled diabetes (day 0 vs. day 7, P < 0.01; day 0 vs. day 8–20, P < 0.001), whereas the STZ-induced diabetic WT (STZ-WT) mice showed stable glucose levels on the first 8 days (P > 0.05) and were significantly elevated afterward (P < 0.01). Consistent with the previous reports, a low dose of STZ enhanced random glucose, and 23.1% of the mice developed diabetes. However, 87.5% of the STZ-KO mice exhibited average random blood glucose levels >16.7 mmol/L on day 11 and afterward (Fig. 4B). ULT lowered the diabetes incidence, although without a significant difference, and ultimately reached 75.0% in STZ-KO mice (Fig. 4B). Furthermore, ULT delayed the diabetes onset from day 7 to day 10 (Fig. 4B). Therefore, MLD-STZ stimuli could accelerate the development of diabetes, and ULT can only partially reverse this deterioration in STZ-KO males.

    Figure 4
    Figure 4

    HU induces diabetes with external MLD-STZ stimulation. A: Random blood glucose levels were monitored daily after administration of MLD-STZ (40 mg/kg/day for 5 days) for 20 consecutive days (8-week-old mice, n = 13, 8, and 8 for STZ-WT mice, STZ-KO mice, and pegloticase therapy ULT-KO mice separately, respectively). B: The diabetes incidence in 8-week-old mice was calculated by percentage (n = 13, 8, and 8 in each group, respectively). Diabetes was defined as random blood glucose ≥16.7 mmol/L. P trend value is presented as *P < 0.05, **P < 0.01, ***P < 0.001. Data are expressed as mean ± SEM.

    HU-Related Diabetes Is Manifested by Increased β-Cell Apoptosis

    For determination of whether impaired glucose intolerance was caused by a reduction in β-cell number and islet mass, we extracted mice pancreas tissues for further analysis. No apparent histological lesions, stained with HE or with antibodies against insulin, were detected in the pancreatic islets of the Uox-KO males and controls (Fig. 5A and B). Pancreatic sections were coimmunostained with anti-insulin antibody to visualize β-cells. Compared with the control groups, pancreatic insulin content was lower in STZ-KO mice than in control mice (Fig. 5A and B). Accordingly, relative β-cell mass, determined by insulin immunoreactivity, was 62% lower in STZ-KO mice (Fig. 5A and B). To determine whether cell death contributes to reduced β-cell mass in STZ-KO mice, we measured islet apoptosis by TUNEL staining. The number of TUNEL-positive β-cells was higher in STZ-KO mice than in control mice (Fig. 5C and D), tending to a 42% increase (Fig. 5C and D). This finding suggests HU promotes not only the loss of β-cell mass but also apoptosis of the pancreatic β-cell in STZ-KO mice. These morphological alterations are consistent with the changes in plasma insulin levels, indicating that β-cell apoptosis chiefly contributes to the onset of diabetes in STZ-KO males.

    Figure 5
    Figure 5

    HU causes pancreatic β-cell death under MLD-STZ stimuli. A: Pancreatic sections were histologically stained for HE and immunohistochemically (IHC) stained for insulin, respectively (8-week-old mice, n = 6). B: Total areas of pancreatic tissues and insulin-positive cells were traced manually and determined by counting 10 islets or more in 6 sections per mouse (8-week-old mice, n = 6) in each group. β-Cell mass was analyzed, and results are shown in multiplying the β-cell ratio (insulin-positive areas–to–total area) with the initial pancreatic wet weight. C: Insulin-positive cell apoptosis was analyzed by the terminal deoxynucleotidyl TUNEL (8-week-old mice, n = 6). D: Double-positive ratio of insulin and TUNEL was measured in each group (8-week-old mice, n = 6). Scale bars = 50 μm. **P < 0.01, ***P < 0.001. Data are expressed as mean ± SEM.

    We next asked whether ULT would be able to reverse the pancreatic β-cell apoptosis with MLD-STZ in HU mice. Shown in Fig. 6A, the β-cell masses were significantly smaller in STZ-KO males than in STZ-WT controls (P < 0.001). However, the ULT did not reverse the loss of β-cell mass (STZ-KO vs. ULT-KO, P > 0.05) (Fig. 6A). MLD-STZ exaggerated the islet β-cell death compared between STZ-KO and STZ-WT mice, indicated by TUNEL staining (Fig. 6B). Consistently, the number of TUNEL-positive β-cells was significantly higher in STZ-KO mice than in their controls (Fig. 6C). However, no significant decreases in TUNEL-positive β-cell signals (Fig. 6B) or analysis numbers (STZ-KO vs. ULT-KO, P > 0.05) (Fig. 6C) were detected after 21 days of ULT in STZ-induced Uox-KO mice. Thus, our findings did not support the benefits of ULT on β-cell vitality.

    Figure 6
    Figure 6

    ULT reversed few pancreatic β-cell deaths stimulated by MLD-STZ. Mice were sacrificed on the 21st day after the first MLD-STZ (40 mg/kg/day for 5 days) injection. A: β-Cell mass was calculated by multiplying the β-cell ratio (insulin-positive areas–to–total area) with the initial pancreatic wet weight (11-week-old mice, n = 3). B: Insulin-positive cell apoptosis was analyzed by the terminal deoxynucleotidyl TUNEL (11-week-old mice, n = 3). C: Double-positive ratio of insulin and TUNEL was measured in each group (11-week-old mice, n = 3). Scale bars = 50 μm. **P < 0.01, ***P < 0.001. Data are expressed as mean ± SEM.

    ULT Ameliorates Tubulointerstitial Injury in Diabetes

    To evaluate renal changes after STZ stimulation and ULT intervention, we did kidney functional and histological experiments in both genotypes. SU levels were significantly higher in STZ-induced Uox-KO male mice than in their WT controls (P < 0.001) (Fig. 7A). The same results were shown in serum creatinine levels, which are indicators of renal function (P < 0.001) (Fig. 7B). ULT increased renal function in STZ-KO males, displaying significant decreases in SU and creatinine levels (P < 0.001) (Fig. 7A and B).

    Figure 7
    Figure 7

    Renal function and pathological changes were evaluated in male 11-week-old mice. A: SU levels were measured at the end of experiments (n = 13 for WT and n = 8 for Uox-KO mice in each group). B: Serum creatinine levels, indicators of renal function, were determined by the automatic biochemical analyzer (n = 13 for WT and n = 8 for Uox-KO mice in each group). C: Renal pathohistology was detected by HE staining, PASM staining, and PAS staining (n = 6). Deposits of renal urate crystals were investigated under polarized light (n = 3). D: Quantification of tubulointerstitial injury. Quantitative analysis for tubulointerstitial injury was assessed by scoring six renal cortical tubule sections (original magnification ×200, HE) in randomly selected fields for each group (n = 6). E: The GSI was calculated to evaluate diabetic nephropathy based on three given PAS staining sections for each group (n = 6). F: Urate crystal areas were calculated by the Image-Pro Plus 6.0 system with arbitrary units (n = 3). Scale bars = 50 μm in HE (original magnification ×200), PASM, PAS, and polarized light sections. Scale bars = 100 μm in HE (original magnification ×400). *P < 0.05, ***P < 0.001. Data are expressed as mean ± SEM.

    STZ-KO mice showed dilated Bowman spaces and tubules and collapsed and necrotic nephrons by pathological analysis (Fig. 7C). The tubular damage was significantly severe, with tubular dilation, detachment of tubular epithelial cells, and condensation of tubular nuclei appearance in STZ-KO mice (Fig. 7C). ULT prevented the development of these lesions indicated by decreased percentage of tubular damage in ULT-KO mice (Fig. 7D). PASM staining exhibited obvious glomerular mesangial hyperplasia, increased glomerular matrix, and thickened glomerular basement membrane in diabetic STZ-KO mice compared with their WT controls (Fig. 7C). PAS staining documented glomerular mesangial expansion and glomerular sclerosis, early features of diabetic nephropathy, in STZ-KO mice compared with STZ-WT mice (Fig. 7C). The GSI, calculated by PAS staining sections, was increased in STZ-KO mice compared with STZ-WT mice (P < 0.05) (Fig. 7E). However, ULT did not prevent the glomerular injury in STZ-induced HU mice as quantified by GSI (Fig. 7E). HU STZ-KO mice showed significant renal urate crystal deposits (P > 0.05) (Fig. 7C). Crystals were dissolved by ULT in STZ-KO mice with decreased urate crystal areas (P < 0.001) (Fig. 7F). The histological analysis presented that ULT exhibited a significant therapeutic effect of HU-crystal–associated kidney injury and tubulointerstitial injury manifestation in diabetic nephropathy.

    Differentially Expressed Genes in HU and/or Diabetic Mice

    We then wondered whether differentially expressed genes (DEGs) in islets of the HU and/or diabetic mice would explain the molecular mechanism of urate on impaired glucose metabolism. Microarray data were represented by heat maps (Fig. 8A and B) in subgroup comparisons of Uox-KO mice versus Uox-WT mice and STZ-KO mice versus STZ-WT mice. We selected the genes based on adjusted P < 0.05 and absolute fold change >2. In Uox-KO versus Uox-WT groups, 850 of 2,018 genes were upregulated and 1,168 of 2,018 genes were downregulated (Fig. 8C). Whereas in STZ-KO versus STZ-WT groups, 29 of 171 genes were upregulated and 142 of 171 genes were downregulated (Fig. 8D). When compared between Uox-KO versus Uox-WT and STZ-KO versus STZ-WT groups together, one gene (Stk17β) was shared in the upregulated DEGs and five genes (Fut4-ps1, Erich3, 1700027H10Rik, Kcnh2, and Klhl32) in the downregulated gene set (Fig. 8C and D). These shared genes were the urate primacy functioning genes. It is notable that the shared upregulated gene, Stk17β, plays a key role in a wide variety of cell death signaling pathways.

    Figure 8
    Figure 8

    Gene set enrichment analysis comparing isolated islets from HU and/or diabetic mice. Heat maps from mouse isolated islets are shown. Each colored box represents the normalized expression level of a given gene in a particular experimental condition of WT vs. KO (A) and STZ-WT vs. STZ-KO (B). Red denotes upregulation and blue denotes downregulation according to the color scale. Shared upregulated (C) and downregulated (D) DEG numbers were enriched in WT vs. KO and in STZ-WT vs. STZ-KO groups. One gene (Stk17β) was shared in the upregulated gene set and five genes (Fut4-ps1, Erich3, 1700027H10Rik, Kcnh2, and Klhl32) were shared in the downregulated gene set, with adjusted P < 0.05 and absolute fold change >2 (8-week-old WT and KO mice and 11-week-old STZ-WT and STZ-KO mice, n = 3 in each group).

    Discussion

    Whether HU is a causal or noncausal factor for diabetes remains controversial. Although numerous clinical studies have showed that HU predicts the development of diabetes and that the ULT could reduce the diabetes incidence or fasting glucose levels accordingly, the causal relationship between HU and diabetes and related mechanisms remains elusive. The current study firstly demonstrated that HU augmented the existing glycometabolism abnormality induced by MLD-STZ and induced diabetes by promoting not only the loss of β-cell mass but also pancreatic β-cell apoptosis in STZ-induced Uox-KO male mice. Although HU increased the incidence of diabetes when accompanied with STZ stimuli, ULT can only ameliorate the incidence by a small proportion without significant statistical differences. In addition, our transcriptomic results indicated that Stk17β is a possible target gene in HU-induced β-cell apoptosis.

    Multiple lines of evidence have shown the association between HU and diabetes. A community-based study in the U.S. demonstrated the risk of diabetes incidence increased 18% with every 1 mg/dL SU elevation, and the association remained significant after adjustment for fasting glucose and insulin levels (15). The diabetes incidence was 19% for SU ≤7 mg/dL patients, 23% for SU 7 mg/dL to ≤9 mg/dL, and 27% for SU >9 mg/dL in an 80-month follow-up investigation that included 1,923 U.S. veterans (15). This study also indicated ∼8.7% of all new cases of diabetes were statistically attributed to HU (15). The age- and sex-adjusted hazard ratio for diabetes was 2.83 for the fourth quartile of SU in subjects from the Rotterdam study (16). Although epidemiology studies showed that HU predicts the development of diabetes, no causality was found in the Mendelian randomization study (6).

    Thus, for further phenotype and mechanism explorations, a major obstruction is the lack of an appropriate animal model given that Uox in most mammals, including rodents, is functional. Here, we were able to use Uox-KO mice to study the potential role of urate in glucose metabolism. Traditional establishment strategies, such as potassium oxonate (a chemical inhibitor of uricase) i.p. injection (17), would interfere with the effects of HU per se as the additional exogenous intervention. An advantage here is that this mouse model is a suitable HU model because of its consistent and stable SU elevations to simulate a human-specific biological background of elevated urate (10).

    Next, we further stimulated this spontaneous HU mouse model with the HFD and/or MLD-STZ. A phenotypic heterogeneity in glycometabolism existed between sexes in the Uox-KO model, displaying severe glucose intolerance and more sensitivity to STZ in Uox-KO males than in females in a previous study (10). To avoid confounding factors due to sex or sex hormone differences, only male mice were used in the current study. Further investigations in female mice would help to delineate a full picture of the effect of HU on glucose metabolism.

    Our data show significant elevated blood glucose concentrations at 15 and 30 min of GTT under basal conditions and in response to high-fat feeding (Figs. 2 and 3). Moreover, the plasma insulin drop indicated the compromise of β-cell function is responsible for glucose intolerance. The link of HU and insulin resistance has been extensively reported and discussed (18,19). Spontaneous HU represents one of the metabolic syndromes resulting from the interactions between genetic and environmental factors, including dietary and behavioral factors, which also contribute to insulin resistance in studies (20). Thus, genetically modified HU mouse models lack major complicated genetic or nongenetic risk factors, and HU indeed did not induce insulin resistance, which is normally obesity related. We then fed Uox-KO mice and WT controls the HFD or normal chow diet. The results showed HFD-fed Uox-KO mice displayed a dramatic reduction in plasma insulin levels compared with HFD-fed WT mice, which indicated a failure of compensatory insulin production by β-cells in the Uox-KO mice. Glycol-metabolism evaluations, including GTT, ITT, and GSIS, hint at severe glucose intolerance, with unchanged insulin sensitivity in HFD-induced Uox-KO mice (Fig. 3). To be noted, neither the fasting glucose nor fasting insulin changed in the Uox-KO mice, even with aging (Fig. 1). In this study, the lack of being able to see a urate-to-insulin resistance relationship is based on the use of a high-fat rather than a high-sugar diet. A similar conclusion was drawn by Kelly and colleagues (21) that systemic HU, while clearly a biomarker of the metabolic abnormalities of obesity, does not appear to be causal. Mendelian randomization studies that focused on genetic variants associated with urate, thereby removing confounding factors such as obesity, showed elevated urate could not independently predict diabetes development (6,22), which also supports our study findings. However, in models of sugar-/fructose-induced insulin resistance or in models where liver xanthine oxidase activity is increased, then one does see urate-dependent insulin resistance (23,24), which is likely mediated by gluconeogenesis (25). Further investigations using sugar-/fructose-fed Uox-KO mice are needed to detect the relationship between urate and insulin resistance.

    In the current study, we report that Uox deficiency predisposes mice to the onset of diabetes, which results from a loss of β-cell mass under conditions of the HFD accompanied by MLD-STZ. The study has provided both functional and mechanistic data showing the proapoptotic effect of HU in β-cells. This is the first in vivo investigation suggesting a critical role of HU in acceleration of the progression from impaired glucose tolerance into diabetes via the action of promoting β-cell death. Increasing incidence of diabetes is also accompanied by decreased islet area and β-cell mass (Figs. 4 and 5), which corroborates that this glucose intolerance induced by HU is primarily caused by β-cell death. Multiple factors and signaling mechanisms have been demonstrated to influence β-cell compensation, of which β-cell apoptosis has emerged as a key event causing decompensation of β-cells and the development of diabetes (26). Insulin deficiency has been demonstrated to play a causative role in the development of diabetes from both animal and human studies (26). In line with this, Uox-KO mice induced by MLD-STZ develop diabetes, accompanied by hypoinsulinemia and increased β-cell apoptosis. The diabetic phenotype of Uox-KO mice not only indicates that loss of functional β-cell mass is a key cause of the disease but also reveals a critical role of HU that is required for maintaining β-cell survival in the STZ-induced condition. However, it is worth noting that no differences were found in β-cell mass and insulin production between Uox-KO mice and WT controls fed the normal chow diet, suggesting that HU is not required for islet development and is dispensable under nonstressed physiological conditions.

    The benefits of ULT are still inconclusive. Randomized trials have reported that ULT by allopurinol improves insulin resistance in asymptomatic HU individuals (27,28), and similar improvement in insulin resistance has been observed with the use of benzbromarone (29). However, in a gout population, the incidence of diabetes was lower in urate-lowering drug (ULD) users than in nonusers (30). In vivo study showed oral administration of ULD dose-dependently reduced the blood glucose level and improved glucose tolerance and insulin resistance in db/db mice (31). Whether asymptomatic HU should be treated remains controversial, as Sluijs et al. (6) concluded that SU is not causally linked to diabetes and that ULTs may therefore not be beneficial in lowering diabetes risk. A long-held notion in diabetes is that macrophages within the islet produce reactive oxygen species and proinflammatory cytokines, creating a β-cell cytotoxic environment (32). Pegloticase, a recombinant uricase, exerts its urate-lowering role by degrading urate with a mild oxidative stress. Lowering SU may also reduce oxidative stress because urate, although an antioxidant in noncellular systems, is a pro-oxidant in cellular systems. The present in vivo ULTs exhibited only a partial reverse function on reducing diabetes incidence (Fig. 4) and could not ameliorate β-cell apoptosis (Fig. 6) or glomerular lesions in diabetes (Fig. 7). Substantial short-term ULT did not have a direct protective effect on β-cell apoptosis or antidiabetic potential, suggesting that, on its own, this might not be an effective strategy for restoring β-cell function in STZ-induced HU mice. Longer-term ULT needs to be done in future work to solidify the conclusion.

    One of the interesting findings in this study is that ULT improved tubulointerstitial injury but not glomerular lesions in STZ-KO mice. Gilbert and Cooper (33) suggested that tubulointerstitial injury is a major feature of diabetic nephropathy and that its development may reflect influences that are common to other forms of renal disease and also those that are unique to diabetes. An in vivo study showed HU plays a pathogenic role in the mild tubulointerstitial injury associated with diabetic nephropathy but not glomerular damage in diabetic mice (34). Other factors, such as high glucose or oxidative stress, could be responsible for the diabetic glomerular lesion because high glucose is well known to be one of the major stimuli to accelerate extracellular matrix deposition in diabetic glomeruli (35).

    Contrary to studies that have previously reported a positive causal association with progression of chronic kidney diseases (CKDs) (36), urate concentration was not causally related to the development of diabetic nephropathy in a Mendelian randomization study by Ahola et al. (37). This Mendelian randomization analysis suggested that the SU concentration does not have any causal effect on diabetic kidney complications but is rather a downstream marker of the kidney damage (37). Moreover, as one Mendelian randomization study suggests that SU is a causal risk factor for CKD in the general population (38), it might be that SU plays a role only in the processes leading to nondiabetic renal disease rather than in pure diabetic nephropathy. Furthermore, the authors of a post hoc analysis in diabetic nephropathy with mean follow-up of 3 years concluded that urate was weakly associated with decline in the glomerular filtration rate (GFR) in patients with type 1 diabetes with overt nephropathy (39). Evidence showed urate may facilitate the development and progression of CKD in people with diabetes (4042). However, the benefits of urate-lowering are still putative and inconclusive. A post hoc analysis of the Febuxostat Open-Label Clinical Trial of Urate-Lowering Efficacy and Safety Study (FOCUS) with 116 HU patients treated with febuxostat for 5 years found an inverse correlation between urate and estimated (e)GFR and projected an improvement in eGFR of 1 mL/min/1.73 m2 for every 1 mg/dL decrease in SU (43). In a randomized clinical trial, the authors found a significant association between higher urate and lower GFR (P = 0.017), while this association was absent in the allopurinol treatment (P = 0.61) (44).

    However, the renal status in the above studies was assessed with the eGFR and/or urinary albumin excretion rate, which manifested the glomerular not the tubular function. Despite the current availability of conclusive trial data, Bartáková et al. (45) have recommended 10–15% lower SU cutoff values for people with diabetes to confer protection against kidney disease. In an animal study, Kosugi et al. (34) found that db/db mice developed HU and glomerular mesangial expansion (an early feature of diabetic nephropathy). Allopurinol treatment significantly lowered urate levels and reduced tubulointerstitial injury but without amelioration of glomerular lesions in diabetes (34). These results were consistent with our reported data. However, the mixed conclusion of whether lowering SU benefits diabetic nephropathy from Mendelian randomization and clinical and animal studies needs to be carefully explained.

    The underlying mechanism of how urate interferes with the function of pancreatic β-cells has not been well elucidated. Roncal-Jimenez et al. (46) showed that sugar-induced HU likely played a role in the development of diabetes through an effect that included a direct urate effect on islet insulin secretion. Previous studies demonstrated β-cell toxicity of soluble urate via the NF-κB–iNOS–NO signal axis (7). Our research is the first study to elucidate the molecular changes occurring specifically in HU and/or diabetic islets by microarray. The transcriptomic analysis of isolated islets indicates major alterations in shared DEGs with up- and downregulation that might contribute to the accelerated progression of urate-induced cell death. Death-associated protein kinase–related apoptosis-inducing kinase-2 (Drak2), also known as Stk17β, is a serine/threonine kinase that executes its role by apoptosis-related pathways. Stk17β rapidly induces apoptosis in mouse islet β-cells by inflammatory cytokines (47) and free fatty acids (48). Stk17β overexpression, along with the cytokine assaults, has been demonstrated to lead to aggravated apoptosis of β-cells (47), which was consistent with our transcriptomic results (Fig. 8). Mao et al. (47) suggested the inflammatory cytokine/Stk17β/p70S6 kinase pathway seemed to be critical in islet apoptosis. They also demonstrated Stk17β acted through caspase-9 in apoptosis (47). Together with our transcriptomic profiles, these lines of evidence indicate that Stk17β is a potential gene in urate-mediated islet apoptosis of STZ-induced diabetes.

    Particularly, reduced levels of TG and unchanged TC levels in Uox-KO mice may partially explain the absence of an insulin-resistance phenotype given TG are an independent risk factor for the future development of insulin resistance (49). The decreased TG levels are attributable to sex hormones per se, since it has been reported that testosterone enhances TG in rodents (50) while our HU mice had lower testosterone (data not shown).

    Limitations of this study also need to be pointed out. To meet the animal ethic, the mice were euthanized with CO2, followed by cervical dislocation before islet isolation. CO2, as an additional stress on the islets, may magnify gene expression differences between KO and WT islets. Although Uox-KO mice displayed high SU levels comparable to those observed in adult humans, their germline disruption of Uox produced embryonic or postnatal developmental effects such as nephropathy. We also acknowledge this potential caveat that the mice developed kidney disease and urate crystalluria spontaneously, as we previously described (10). To separate whether the impaired glucose tolerance is due to the impaired renal function or to the urate, mice with a conditional and inducible disruption of Uox (e.g., tamoxifen-responsive cre mice) may be used to obviate the concern.

    In conclusion, the current study demonstrates that urate per se is insufficient to induce diabetes, while it impairs glucose tolerance. For the first time, our research using a constitutive HU model corroborates that high levels of urate predispose mice to diabetes by disrupting β-cell function. Diabetes incidence, β-cell death, or glomerular lesions in diabetes were not reversed by ULT with significance; however, ULT displayed a therapeutic effect on HU-crystal–associated kidney injury and tubulointerstitial damage in diabetic nephropathy. Transcriptomic profiling suggested Stk17β is a urate primacy gene in β-cell apoptosis. We believe that our microarray data specifically from isolated islets of HU mice can serve as a valuable resource for investigators in the field for further exploration of specific genes involved in HU with glycol-metabolic dysfunction.

    Article Information

    Acknowledgments. The authors thank pathologists Shihong Shao, Xiangyan Zhang, and Feng Hou (Department of Pathology, the Affiliated Hospital of Qingdao University) for their professional technical support.

    Funding. This study was supported by research project grants from the National Key Research and Development Program (2016YFC0903400), the National Science Foundation of China (31900413, 81520108007, and 81770869), the Shandong Province Key Research and Development Program (2018CXGC1207), and the Shandong Province Natural Science Foundation (ZR2018ZC1053).

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

    Author Contributions. J.L., Y.H., L.C., Z.L., X.L., H.Z., H.L., W.S., A.J., and Y.W. performed the experiments. J.L., Y.H., H.Y., and C.L. designed the study. J.L., Y.H., X.X., H.Y., and C.L. analyzed and interpreted the data. All authors approved the final version to be published. J.L. and C.L. are the guarantors of this work and, as such, had full access to all the data in the study and take responsibility for the integrity of the data and the accuracy of the data analysis.



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