The Role of α₁-Adrenergic Receptors in Regulating Metabolism: Increased Glucose Tolerance, Leptin Secretion, and Lipid Oxidation

Abstract

The role of α₁-adrenergic receptors (α₁-ARs) and their subtypes in metabolism is not well known. Most previous studies were performed before the advent of transgenic mouse models and utilized transformed cell lines and poorly selective antagonists. We have now studied the metabolic regulation of the α_1A- and α_1B-AR subtypes in vivo using knock-out (KO) and transgenic mice that express a constitutively active mutant (CAM) form of the receptor, assessing subtype-selective functions. CAM mice increased glucose tolerance while KO mice display impaired glucose tolerance. CAM mice increased while KO decreased glucose uptake into white fat tissue and skeletal muscle with the CAM α_1A-AR showing selective glucose uptake into the heart. Using indirect calorimetry, both CAM mice demonstrated increased whole body fatty acid oxidation, while KO mice preferentially oxidized carbohydrate. CAM α_1A-AR mice displayed significantly decreased fasting plasma triglycerides and glucose levels while α_1A-AR KO displayed increased levels of triglycerides and glucose. Both CAM mice displayed increased plasma levels of leptin while KO mice decreased leptin levels. Most metabolic effects were more efficacious with the α_1A-AR subtype. Our results suggest that stimulation of α₁-ARs results in a favorable metabolic profile of increased glucose tolerance, cardiac glucose uptake, leptin secretion, and increased whole body lipid metabolism that may contribute to its previously recognized cardioprotective and neuroprotective benefits.

Introduction

α₁-adrenergic receptors (ARs) are G-protein coupled receptors that mediate the sympathetic nervous system. They are well documented for their role in regulating neurotransmission, heart function, cardiac hypertrophy and blood pressure (reviewed in 1). There are nine AR subtypes that all bind epinephrine and norepinephrine (β₁, β₂, β₃, α_2A, α_2B, α_2C, α_1A, α_1B, α_1D), but each with both common, yet distinctive, and sometimes opposite, functional roles. Previous studies on AR regulation of metabolism concentrated on β-ARs or α₂-ARs, not α₁-ARs. α₁-AR-mediated metabolism was explored mostly in the 1980s and used poorly selective antagonists and utilized mostly transformed cell lines to assess functions before transgenic approaches were made available.

The autonomic nervous system is well known to regulate both glucose and fatty acid metabolism through ARs via both neurotransmission as well as hormonal effects. In general, catecholamines direct metabolic effects towards substrate mobilization and utilization in order to meet the increased energy requirements of stress or the “fight or flight” response.

It is well known that norepinephrine mediates multi-faceted aspects of metabolism. Early studies indicated that α₁-ARs stimulate gluconeogenesis and ketogenesis in the liver (2) and suppress triglyceride secretion (3). α₁-ARs have also been shown to regulate glucose uptake into various cell lines (4–6), but never shown to regulate glycolysis. α_1B-AR KO mice were previously reported to be insulin resistant and impaired glucose homeostasis (7). The β-ARs are well known mediators of adipocyte metabolism through their regulation of cAMP levels in both white and brown fat (8) and are lipolytic (9). However, the β₁/β₂/β₃-AR triple KO mice still display norepinephrine-induced lipolysis, suggesting involvement of another AR subtype (10). α₂-AR and in particular α_2A-AR stimulation inhibits insulin secretion (11) and are anti-lipolytic (12). Therefore, there was significant previous data to suggest that α₁-ARs can regulate whole body and tissue-specific metabolism that was different from either β-or α₂-ARs.

Early studies also indicted that α₁-ARs stimulate fatty acid oxidation in hepatocytes but there are no reports outside of the liver or in whole body metabolism (3, 13–16). In the present study, we made use of novel transgenic mice with systemic overexpression of the α_1A- and α_1B-ARs using large fragments of the isogenic promoters. These receptors also contain mutations that render them constitutively active and are chronically activated even when an agonist is not present. These mice provide tools for assessing α₁-AR subtype-selective in vivo signaling. Using these and the corresponding KO mice, we now describe the in vivo metabolic functions regulated by this receptor family.

Materials and Methods

Animal use.

Transgenic mouse models (mixed B6CBA background) that systemically over express the α_1A-AR or α_1B-AR subtypes were previously described and characterized in our laboratory (17–18). These mice express constitutively active mutants (CAM) of the receptors under the control of their native mouse promoter to increase subtype-selective signaling in tissues that naturally express that subtype. The α₁-AR KO mice were obtained from Paul Simpson, MD (19–20), maintained on a C57BL6 background, and backcrossed every 10 generations. B6CBA WT mice were obtained from Jackson Laboratory, then bred internally and used as controls for the CAM mice. C57BL/6 WT were also obtained from Jackson Laboratory, bred internally, the used as controls for the KO mice. Equal numbers of both male and female mice (unless otherwise indicated) from 2–4 mo of age were used in a randomized fashion in each experiment. Mice were given a normal standard laboratory chow diet (Harlan, #2918), housed on a 12-hour light-dark cycle, temperature-controlled facility at 70°F with free access to food and water unless otherwise indicated. Mice were provided veterinary care in an AAALAC-accredited animal care facility. The experimental protocols employed in this study conform to the Guide for the Care and Use of Laboratory Animals published by the US National Institutes of Health and was approved by the Animal Care and Use Committee at our institutions (Protocol 0844).

Blood Profiles.

Mice were transferred to wood chip bedding and fasted for 6 hours. Mice were then anesthetized with Nembutal (ip, 50mg/ml) and blood collected from the vena cava into 1.5ml polypropylene microfuge tubes. The blood samples were allowed to sit at room temperature for 30 min to coagulate, then centrifuged at 2000 x g for 10 min at 4°C. The sera was transferred to fresh tubes and re-centrifuged to remove blood cells. Samples are stored at −80°C. Measurement of cholesterol, triglycerides, urea and electrolytes were analyzed by the Veterinary Diagnostic Services of Marshfield Laboratories (Cleveland, Ohio) using standard clinical procedures.

Glucose Tolerance.

Mice were fasted for 6 hours on hardwood bedding and weighed. Sterile 20% D-Glucose (Sigma) solution was injected intraperitoneal at a concentration of 2g/kg body weight. Whole blood samples were taken from a tail cut immediately before the injection of glucose (designated as time 0 min) and at 30, 60, 30, 90 and 120 min after the injection of glucose. Glucose levels were measured using a Nova Max Plus glucometer according to manufacture’s instructions (Nova Biomedical Corporation).

Indirect Calorimetry.

Case Western Reserve University Mouse Metabolic Phenotyping Center (MMPC) performed the indirect calorimetry using established procedures (21–22). Metabolic rates were measured by indirect calorimetry in mice using an 8-chamber open-circuit Oxymax system (CLAMS, Columbus Instruments, Columbus, OH). Briefly, mice were acclimated to the experimental room for 1 week prior to the experiment. The mice were individually housed in acrylic calorimeter chambers through which air with a known O₂ concentration is passed at a constant flow rate. The system automatically withdraws gas samples from each chamber hourly for 24 h. The system then calculated the volumes of O₂ consumed (VO₂) and CO₂ generated (VCO₂) by each mouse in 1 h. The respiratory quotient (RQ), which is the ratio of VCO₂ to VO₂, is calculated. Heat or energy expenditures were measured throughout the study and measurements are carried out in both light and dark cycles and both fed and fasting conditions. Mice were maintained at 25°C and had free access to water in all conditions.

Tissue 2-Deoxyglucose Uptake.

Equal numbers of male and female mice were fasted for 6 hours then injected with [³H]-2-deoxyglucose (2DG)(20uCi/mouse; ip). Blood samples were taken from the tail vein at 30, 60, 90 mins post-injection to determine blood glucose levels using a Nova Max Plus glucometer according to manufacturer’s instructions (Nova Biomedical Corporation, Waltham, MA). Plasma samples were also prepared at the same time to determine [³H]-2DG specific activity. After the final collection of blood, mice were euthanized with pentobarbitol (i.p. 60mg/kg body weight) and various tissues removed, rinsed in PBS, diced and frozen in dry ice. For skeletal muscles, hindlimb muscle was obtained from the gatrocnemius while forelimb muscle was from the tricep. Tissues were processed with perchloric acid, barium hydroxide/zinc sulfate and glucose uptake rate was calculated according to the method of Ferre’, et al (23), using the following equation, Rate= [2-deoxyglucose 6-phosphate]τ/ LC 0 ∫ ^τ (C_B*/C_B)dt where τ is the sampling time, C_B* is the blood 2-deoxyglucose expressed in terms of radioactivity and C_B is the blood glucose concentration. The lump constant (LC) is a correction factor for the discrimination against 2-deoxyglucose in glucose transport and phosphorylation pathways determined in vitro by comparing glucose and 2-deoxyglucose fractional extraction by the different tissues.

Leptin.

Mice were put into cages with wood-chip bedding and were fasted for 6 hours prior to drawing blood. Blood was drawn from mice by making a small cut in the tail after anesthesia with ketamine (87.5mg/kg) /xylazine (12.5mg/kg) cocktail. The blood was centrifuged for 10 minutes at 2000x g, the sera collected, and stored at −80° until processed. The sera were analyzed for leptin using the mouse/rat leptin enzyme immunoassay kit from SPI BIO (Montigny le Bretonneux, France) (cat#A05176) according to manufacturer’s procedures. Briefly, samples and standards were pipetted onto the 96-well plate pre-coated with a polyclonal anti-mouse leptin antibody. The plate was covered and incubated for 1 hour with shaking. The wells were rinsed and biotin-labelled anti-leptin antibody was dispensed in each well and incubated as before for another hour. The wells were washed followed by incubation with Streptavidin-HRP conjugate for 30 minutes. The wells were washed and substrate solution was added to each well and the plate was incubated in the dark for 10 minutes at room temperature. The color development was stopped and the absorbance at 450 nm was read within 5 minutes. The concentration of the samples was read comparing the absorbance to the standards plotted on a log-log graph.

Statistical Analysis.

One Way Analysis of Variance and Newman-Keuls post-test were used to compare functional and signaling parameters in the different mouse models and experimental conditions. A repeated-measures One Way Analysis of Variance was assessed on the glucose tolerance and respiration quotient data. A probability value p< 0.05 was considered statistically significant. Prism software (GraphPad, San Diego, CA) was used for all data analyses.

Results

α₁-AR activation increased glucose clearance from the blood.

We recently found that α₁-ARs protect against low glucose damage during ischemia by increasing the uptake of glucose into myocytes through activating GLUT transporters GLUT1 and GLUT4 (24). Therefore, we hypothesized that α₁-AR activation may also affect blood glucose levels. Utilizing a glucose tolerance test, we found that both CAM α₁-AR mice (p<0.01) selectively enhanced glucose clearance from the blood using the Repeated Measures ANOVA followed by the Newman-Keuls Multiple Comparison Test. (F(2,4)=8.78, p<0.01, N=8 or 10 mice) (Fig 1A). While both α_1B-AR KO or α_1A/B double KO mice displayed elevated blood glucose levels, only the α_1A-AR KO (p<0.01) mice displayed significantly reduced glucose uptake compared to the control (F(3,4)=44.39, p<0.001).

Figure 1. α₁-AR regulation of glucose tolerance.

Mice are fasted for 6 hours and weighed. 20% glucose is injected intra-peritoneal at a concentration of 2g/kg body weight. Whole blood samples are taken from a tail cut immediately before the injection of glucose (designated as time 0 min) and at 30, 60, 30, 90 and 120 min after the injection of glucose. Glucose levels were measured using a Nova Max Plus glucometer according to manufacture’s instructions. A. Glucose tolerance levels in CAM α_1A-AR (squares), CAM α_1B-AR (triangles) mice and WT control (circles). B. Glucose tolerance levels in α_1A-AR KO (squares), α_1B-AR KO (triangle), α_1A/B-AR double KO (diamonds) and WT controls (circles). * p≤ 0.01, statistically significant from WT control using the Repeated Measures ANOVA followed by the Newman-Keuls Multiple Comparison Test. N=8 or 10 individual mice.

α₁-AR activation increased 2-deoxyglucose uptake into heart, white fat, and faster twitch skeletal muscle.

We next used deoxyglucose to determine which tissues were utilizing glucose through α₁-AR regulation. Deoxyglucose is used for tissue uptake studies because it is not metabolized and becomes trapped inside the cell after phosphorylation by hexokinase. We could not analyze liver due to the presence of an enzyme that interferes with the assay. We found that CAM α_1A-AR mice selectively enhanced glucose uptake into the heart while both α₁-AR subtypes increased glucose uptake in white fat and tricep or gastrocnemius skeletal muscle (p<0.05) (Fig 2A). Utilizing the KO mice, we found that only α_1A-AR KO selectively reduced glucose uptake in the heart, but either KO was sufficient in reducing glucose uptake in either fat tissue or skeletal muscle (p<0.05)(Fig 2B).

Figure 2. α₁-AR regulation of ³H-deoxyglucose uptake.

Mice were fasted for 6 hours then injected with [³H]-2DG (20uCi/mouse; ip). Blood samples were taken from the tail vein at 30, 60, 90 mins post-injection using a Nova Max Plus glucometer. After the final collection of blood, mice were euthanized and tissues removed and frozen in dry ice. Tissues were processed and the glucose uptake rate was calculated according to the method of Ferre’, et al (1985). A. ³H-deoxyglucose uptake in tissues isolated from CAM α_1A-AR (grey bars), CAM α_1B-AR (black bars) mice and WT control (white bars). B. ³H-deoxyglucose uptake in tissues isolated from α_1A-AR KO (grey bars), α_1B-AR KO (black bars), and WT controls (white bars). * p≤ 0.05, statistically significant from WT control. N=6–14 mice (in parentheses). Gastroc (gastrocnemius muscle).

α₁-AR activation increased whole body fatty acid oxidation.

To investigate α₁-AR effects on whole body energy metabolism, we performed indirect calorimetry on conscious mice through collaboration with the NIH-sponsored Mouse Metabolic Phenotyping Core (MMPC) at nearby Case Western Reserve University. The Respiratory Quotient (RQ) is the ratio of CO₂ eliminated/O₂ consumed and can indicate which type of fuel is being preferentially metabolized. RQ range from 1.0–0.7 where 1.0 is expected for pure glucose oxidation and 0.7 for pure fatty acid oxidation (25). Proteins are oxidized in the range of 0.8–0.9 RQ. We found that both CAM α_1A-AR and α_1B-AR mice showed a lower RQ than WT control mice (Fig 3A) when feeding suggesting increased lipid metabolism (F(2,75)=36.76, p≤ 0.001) using the Repeated Measures ANOVA. Both α_1A-AR KO and α_1B-AR KO mice had higher RQs than WT control (F(2,130)=48.65, p<.001)(Fig 3B) suggesting that they burn more carbohydrate than WT control mice. CAM α_1A-AR displayed lower RQ than CAM α_1B-AR mice while α_1A-AR KO mice displayed higher RQ than α_1B-AR KO mice, suggesting there were subtype-specific differences in the rate of metabolism.

Figure 3. α₁-AR regulation of whole body metabolism.

Metabolic rates are measured by indirect calorimetry in mice using an 8-chamber open-circuit Oxymax system. Briefly, mice are acclimated to the experimental room for 1 week prior and individually housed in acrylic calorimeter chambers through which air with a known O₂ concentration is passed at a constant flow rate. The system automatically withdraws gas samples from each chamber hourly for 24 h and calculates the volumes of O₂ consumed (VO₂) and CO₂ generated (VCO₂). The respiratory quotient (RQ), which is the ratio of VCO₂ to VO₂, is calculated. A. RQ profile for CAM α_1A-AR (squares), CAM α_1B-AR (triangles) mice and WT control (circles). F(2,75)=36.76, p≤ 0.001. Both CAM statistically significant (P<0.001) from WT control using Repeated Measures ANOVA. B. RQ profile for α_1A-AR KO (squares), α_1B-AR KO (triangles), and WT controls (circles). N=6 or 8 mice. (F(2,130)=48.65, p<.001), Both KO statistically significant (p<0.001) from WT control using Repeated measures ANOVA.

α₁-AR activation lowers fasted triglyceride and glucose levels.

We next determined if there were differences in the blood lipid and glucose profiles that might reflect and confirm the results from indirect calorimetry. Indeed, we found that CAM α_1A-AR mice significantly lowered triglyceride levels (p<0.05)(Fig 4A) than WT control while the α_1A-AR KO mice displayed elevated triglycerides (p<0.05)(Fig 4B). Likewise, we found that CAM α_1A-AR mice significantly lowered fasting glucose levels (p<0.05)(Fig 5A) than WT control while the α_1A-AR KO mice displayed elevated glucose levels (p<0.05)(Fig 5B). Similarly, but not significantly, both CAM α_1B-AR and α_1B-AR KO mice displayed intermediate values from WT controls.

Figure 4. α₁-AR regulation of plasma fasting triglyceride levels in CAM (A) or KO (B) mice.

Mice were first fasted for 6 hours, anesthetized, then blood collected from the vena cava into 1.5ml polypropylene microfuge tubes. The blood samples were allowed to sit at room temperature for 30 min to coagulate, then centrifuged at 2000x g for 10 min at 4°C. The sera was transferred to fresh tubes and re-centrifuged to remove blood cells. Blood samples were analyzed by the Veterinary Diagnostic Services of Marshfield Laboratories (Cleveland, Ohio) using standard clinical procedures. * p≤ 0.05, statistically significant from WT control. N=6–8 mice.

Figure 5. α₁-AR regulation of fasting glucose levels in CAM (A) or KO (B) mice.

Mice were first fasted for 6 hours, anesthetized, then blood collected from the vena cava into 1.5ml polypropylene microfuge tubes. The blood samples were allowed to sit at room temperature for 30 min to coagulate, then centrifuged at 2000x g for 10 min at 4°C. The sera was transferred to fresh tubes and re-centrifuged to remove blood cells. Blood samples were analyzed by the Veterinary Diagnostic Services of Marshfield Laboratories (Cleveland, Ohio) using standard clinical procedures. * p≤ 0.05, statistically significant from WT control. N=6–10 mice.

α₁-AR stimulation results in higher plasma leptin levels.

As leptin is a major regulator of metabolism and energy balance, we next determine plasma levels of leptin in both CAM and KO mice. Both CAM mice had significantly higher levels of plasma leptin with the CAM α_1A-AR being more efficacious (p<0.01 vs p<0.05) (Fig 6A). Either KO mice had significantly lower levels of leptin (p<0.05)(Fig 6B) with the α_1A/B-AR double KO mice (p<0.01) displaying an additive effect. However, we did not find any correlation in body weight. While CAM α_1A-AR mice had higher levels of leptin, it did not have any change in body weight from normal controls (Fig 7A). While CAM α_1B-AR mice did have significantly lower body weight, they also have significant neurodegeneration (17), and autonomic dysfunction (26), which leads to weight loss (27). KO mice only displayed significantly weight gain in the α_1A/B-AR double KO mice (Fig 7B), which are significantly heavier than single KO or WT controls (P<0.05).

Figure 6. α₁-AR regulation of leptin release in CAM (A) or KO (B) mice.

Mice were fasted for 6 hours. Blood was drawn from mice by making a small cut in the tail after anesthesia and centrifuged for 10 minutes at 2000x g. The sera was collected and stored at −80° until processed. The sera were analyzed for leptin using the mouse/rat leptin enzyme immunoassay kit from SPI BIO (Montigny le Bretonneux, France) according to manufacturer’s procedures. * p ≤ 0.05, **p ≤ 0.01, statistically significant from WT control. N=8–10 mice.

Figure 7. Body weight of male CAM (A) or KO (B) mice.

Mice were weighed at 5 months of age. * p ≤ 0.05, statistically significant from WT control. N= 10–12 mice.

Discussion

This is the first comprehensive in vivo report of the metabolic functions regulated through α₁-ARs. Stimulation of α₁-ARs regulates glucose tolerance, whole body metabolism, leptin release, and is selective in regulating glucose uptake into particular tissues. While both the α_1A- and α_1B-AR subtypes appear to regulate metabolism in a similar fashion, the α_1A-AR is clearly more efficacious and confirmed in the α_1A-AR KO mice. Stimulation of α_1A-ARs results in a favorable metabolic profile of increased glucose tolerance, cardiac glucose uptake, leptin release, and enhanced whole body lipid metabolism, resulting in a favorable lipid and glucose blood profile that may contribute to CAM α_1A-AR’s increased longevity and cardioprotection (1, 18, 28–29). These results suggest that α₁-ARs, and in particular the α_1A-AR, is a significant, but previously unrecognized, regulator of whole body and organ-specific metabolism.

It is well known that norepinephrine mediates multi-faceted aspects of metabolism. Early studies indicated that α₁-ARs stimulate gluconeogenesis and ketogenesis in the liver (2) and suppress triglyceride secretion (3). α₁-ARs have also been shown to regulate glucose uptake into various cell lines (4–6), but never shown to regulate glycolysis. α_1B-AR KO mice were previously reported to be insulin resistant and impaired glucose homeostasis (7). The β-ARs are well known mediators of adipocyte metabolism through their regulation of cAMP levels in both white and brown fat (8) and are lipolytic (9). However, the β₁/β₂/β₃-AR triple KO mice still display norepinephrine-induced lipolysis, suggesting involvement of another AR subtype (10). α₂-AR and in particular α_2A-AR stimulation inhibits insulin secretion (11) and are anti-lipolytic (12). Therefore, there was significant previous data to suggest that α₁-ARs can regulate whole body and tissue-specific metabolism that was different from either β-or α₂-ARs.

Early studies also indicted that α₁-ARs stimulate fatty acid oxidation in hepatocytes but there are no reports outside of the liver or in whole body metabolism (3, 13–16). Whole body metabolism is usually regulated through skeletal muscle or adipose tissue because of their mass volume in the body. Both α₁-AR subtypes appear to stimulate whole body fatty acid oxidation (Fig 3A). If α₁-AR stimulation increases whole body fatty acid oxidation, we might expect that triglyceride levels to be decreased as they are being preferentially oxidized. We found that CAM α_1A-AR had decreased plasma triglycerides (Fig 4A) while the α_1A-AR KO had increased levels (Fig 4B). Very early multi-centered clinical studies using non-selective prazosin showed little (30) or no changes in triglyceride levels (31). However, selective blockage of the α_1A-AR has not been tried. However, a more recent cross-over study (i.e. each subject is compared to themselves) using doxazosin showed an 40% increased in plasma non-esterified fatty acid concentrations (32), similar to increased triglyceride levels in α_1A-AR KO mice. Our results suggest that chronic stimulation of the α_1A-AR may be an important regulator of lipid metabolism than previously recognized.

The stimulatory effect of sympathetic activity on glucose uptake in human adipocytes (33) and into rat white adipose in vitro (34) can be mediated by the α₁-AR and confirmed in mouse fat tissue in vivo in our studies (Fig 2). Norepinephrine enhances glucose entry into human adipocytes independently of insulin action that can be blocked with urapidil, an α₁-AR blocker (33). Glucose uptake into adipose tissue plays an important role in glucose homeostasis. In obese subjects with insulin resistance, α₁-AR stimulation may provide an important alternative pathway for glucose uptake into adipose tissue.

Glucose uptake in adipocytes is a key regulator of leptin release (35). Leptin (ob) is a mainly adipocyte-secreted hormone that signals through receptors in the hypothalamus that control hunger, energy balance and body fat (36–37). Corroborating with α₁-ARs increasing glucose uptake into adipose tissue, we found that CAM α₁-AR mice increased leptin levels (Fig 6A), while KO mice decreased leptin release (Fig 6B). There are only a few prior studies indicating association of leptin with α₁-ARs. The study of (38) demonstrated that leptin activates hepatic AMPK through α₁-ARs. Blocking α₁-ARs also reduced leptin levels in obese humans (39) without affecting body weight, similar to our results in KO mice. We also could not see correlation of leptin levels with body weight in our mice (Fig 7), although the α_1A/B-AR double KO mice are significantly overweight. α₁-AR agonists also enhance leptin transport across the blood brain barrier (40). Leptin improves insulin sensitivity by reducing fat content in the liver and skeletal muscle (41–42). Leptin can stimulate fatty acid oxidation in skeletal muscle through AMPK signaling (43–46). There is significant data to indicate that leptin is involved in the regulation of peripheral lipid metabolism (47–50). An in vitro study of leptin showed an increase of lipolytic activity in adipocytes (51). An in vivo study demonstrated that leptin accelerated lipolysis in adipose tissues (43). In the liver, peripheral treatment with leptin prevented accumulation of lipids by acceleration of fatty acid oxidation (50). Besides regulating the release of leptin, α₁-ARs and particularly the α_1A-AR are highly expressed in the hypothalamus (52). Thus, we hypothesize that α₁-ARs mediate glucose uptake into adipocytes, which causes the release of leptin that regulates peripheral whole body lipid metabolism through the hypothalamus. Interestingly, leptin is also both cardioprotective (53–55) and neuroprotective with increasing cognition and anti-aging effects (56), all similar phenotypes in our CAM α_1A-AR mice (1, 18, 28).

We also found that the α_1A-AR in vivo appears selective in mediating glucose uptake into the adult heart (Fig 2). Previous studies in the isolated rat heart (57) and neonatal rat myocytes (58) also indicate α₁-AR mediated glucose uptake. In the normal adult heart, fatty acids are the preferred substrate for metabolism and generate 67% of the ATP. This is contrasted with the fetal heart, which relies mostly on glucose (57). However, glucose uptake, the rate-limiting step in glycolysis (59) may play an important alternative energy source in specific pathological conditions, such as ischemia in the adult (60–61) suggesting that an increase of glucose uptake in the heart may result in a favorable metabolic guard against ischemic injury. The selectivity of the α_1A-AR in mediating glucose uptake into the heart may contribute to its cardioprotective phenotype against ischemia as seen in the CAM α_1A-AR mouse (18).

Skeletal muscle as the largest organ in the body with high metabolic rates, utilize a large amount of glucose (62). The increased glucose tolerance (Fig 1) is likely due to the increased glucose uptake into skeletal muscle by α₁-ARs (Fig 2). CAM mice also had decreased fasting glucose levels (Fig 5A) while KO had elevated fasting levels (Fig 5B), suggesting either a diabetic or pre-diabetic state. Both the gastrocnemius and triceps in the mouse are considered fast twitch, high glycolytic muscle (63). Increasing capacity of glucose uptake in the skeletal muscle may be a beneficial therapeutic approach for the prevention and treatment of type 2 diabetes (64).

β₃-ARs agonists, once pursued for metabolic treatment, was cut short in clinical trial due to poor oral availability, pharmacokinetics and poor efficacy in humans compared with initial studies in rodents (65). α_1A-AR partial agonists may overcome pharmacodynamic barriers as novel oral agonists were developed and considered safe for use in treating urinary incontinence (66) with better clinical efficacy in humans. Clinical studies demonstrating effects of α₁-AR blockers on lipid and leptin profiles (32, 39) may also suggest better clinical efficacy in humans than β₃-AR agonists. While α₁-AR agonists have the negative side effect of increasing blood pressure, imidazoline-type partial α_1A-AR agonists allow for the baroreflex response, have high oral availability, biased agonism, and CNS penetration that may temper affects on blood pressure (67–70).

CONCLUSIONS

Our results suggest that stimulation of α₁-ARs regulates glucose tolerance, whole body metabolism, leptin release, and is selective in regulating glucose uptake into particular tissues. While both the α_1A- and α_1B-AR subtypes appear to regulate metabolism in a similar fashion, the α_1A-AR is clearly more efficacious. α_1A-AR regulation of metabolism needs to be furthered explored to determine its potential to treat metabolic syndrome.