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How does glucose delivered intraperitoneal (i.p.) get into the peripheral circulation?

How does glucose delivered intraperitoneal (i.p.) get into the peripheral circulation?



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Intraperitoneal delivery of drugs or fluids is something that occurs much more frequently in veterinary medicine than clinical medicine. In veterinary medicine or scientific studies using animals, compounds are frequently given via this route that it is typically referred to as "i.p. injection" or "i.p. delivery".

When glucose is delivered via this route (by injection or catheter directly into the peritoneal space), how does the glucose actually get into the systemic circulation? That is, where does it actually go anatomically?

I can't find much data on this or an explanation that is collectively agreed upon. In particular for drug studies, some people say that the drugs or metabolites get into the portal circulation of the liver, while other people say that the drugs are absorbed by the peritoneum and enter the systemic circulation via the lymphatic system? Some individuals would argue that substances diffuse across the peritoneal membrane and then make their way into the interstitial fluid first, and then passively diffuse into the body's fluid compartments. Could it be another route? Could it be an active process or is it completely passive? Or, alternatively, could it be a combination of several of these processes?


Intestinal Fructose and Glucose Metabolism in Health and Disease

55%–71%) and, to a lesser extent, by kidneys (<20%) [16]. Dietary fructose moves from the intestinal lumen to the circulation through a facilitated passive transport [3] across enterocyte membranes by members of the facilitative glucose transport (GLUT Slc2a) family [14]. Upon its intestinal absorption, fructose reaches the liver through the hepatic portal vein and undergoes metabolization in hepatocytes [16]. Fructose transport and metabolism has been extensively reviewed (see refs. [3,14,17]). Exhaustive description of fructose hepatic metabolism is out of the scope of this review. Here, we briefly describe the regulation of intestinal fructose transport and transporters and its intracellular metabolism. We focus on revisiting the role of the liver and small intestine in fructose clearance, the relevance of endogenous fructose production in human diseases, and plant extract inhibitors of fructose transporters.

2.1. Intestinal Fructose Transport

6 mM in Xenopus oocytes expressing the mammalian GLUT5 [19]. In contrast, Kane et al. reported a Km of 11–15 mM using the same expression system for human GLUT5 [25]. Similar values ( Km of 11–13 mM) were found for mouse and rabbit GLUT5 transporter expressed in oocytes [26,27]. Finally, Mate et al. reported a Km of

8–11 mM in ileal brush border membrane vesicles of normotensive Wistar-Kyoto rats and their spontaneously hypertensive rats [28]. Assuming a Km value ranging from 11–15 mM for GLUT5, this Km is similar to that reported for intestinal luminal fructose concentrations (26 mM) in rats fed dietary fructose [29]. On the other hand, the Km of GLUT2 for fructose is

2.2. Dietary Fructose Metabolism

0.2–0.5 mM) [30,31,32,33], which is still very low compared to fasting blood glucose levels (5.5 mM). Finally, type 1 and type 2 diabetic patients exhibited 0.016 mM and 0.009–0.013 mM fasting fructose concentrations, respectively [34,35]. The low fructose concentrations in peripheral blood support the notion that the liver and kidneys are much more sensitive to small changes in circulating fructose levels than the small intestine. Nonetheless, it is unclear how hepatocytes or nephrons reabsorb fructose from the sinusoidal capillaries or glomerular filtrates, respectively, containing very low fructose levels.

2.3. Regulation of GLUT5

2.4. Fructose Metabolism in Human Diseases

2.5. Revisiting the Role of Liver and Small Intestine in Fructose Clearance

90% of fructose phosphorylation occurs in the jejunum, duodenum, or ileum. Most of this fructose is metabolized in the small intestine, appearing in the portal circulation as glucose and lactate (

60%), and the remaining as fructose (<20%). In contrast, high-doses of fructose (≥1 g kg −1 ) saturate the absorption and catabolism of fructose in the small intestine, leading to fructose spill-over into the liver (>30%) and the colonic microbiota in mice [92] (Figure 2). This work challenges our current knowledge about the role of the small intestine in dietary fructose metabolism and spurs the notion that the small intestine shields the liver from toxic fructose exposure. However, several questions arise from this work and remain to be fully addressed: (1) A limitation of the study is regarding the dose-response to fructose, which may vary between mice and humans. Humans may saturate the capacity for fructose metabolism in the small intestine at relatively lower doses than mice. It is necessary to understand the associated dose-response pattern in humans. (2) The role of the small intestine in fructose metabolism in mice and humans may have diverged across evolution. In fact, humans have a relative shorter gut and smaller intestinal area than rodents [93]. (3) The long standing view is that the liver and kidneys are the only gluconeogenic organs in humans, but not the small intestine because it does not express glucose-6-phosphatase (G-6-Pase) [16]. This critical issue is important to translate experimental evidences from mice to humans. In this line, one study have shown the expression of G-6-Pase in the small intestine of humans [94], and another one showed some evidence of the existence of a conversion of fructose to glucose in human jejunum [95].

2.6. Relevance of Endogenous Fructose Production in Human Diseases

30 g) in healthy individuals. Tracer dilution analysis estimated endogenous fructose production

55 mug kg −1 ·min −1 . This work evidenced, for the first time, the capacity for endogenous fructose production in humans [97]. Further research demonstrated the presence of an active polyol pathway in tissues other than those involved in metabolizing dietary fructose, such as the human brain [98,99,100]. Numerous studies using animal models have linked the polyol pathway to metabolic alterations such as obesity, insulin resistance, diabetes, diabetic nephropathy, chronic kidney disease, acute kidney injury, blood pressure, and MetS [101,102,103,104]. Nonetheless, although the presence of an active polyol pathway has been described in humans, and mounting evidences obtained in animal models of the importance of this pathway in diseases, its significance in human metabolic diseases awaits further confirmation.

2.7. Plant Extracts Inhibitors of Fructose Transporters

117 μM for (−)-epigallocatechin-gallate and (−)-epicatechin-gallate, respectively] than by catechins lacking this group [apparent Ki values >500 μM for (−)-epicatechin and (−)-epigallocatechin] [105]. In this line of evidence, it has been shown that chamomile tea and green tea [containing (−)-epigallocatechin gallate (240 mg/g extract), (−)-epigallocatechin (70 mg/g extract), (−)-epicatechin (40 mg/g extract), and (+)-catechin (17 mg/g extract)] effectively inhibited fructose transport through GLUT2 in differentiated Caco-2 cells [106]. In addition, chamomile also inhibits D-fructose transport via GLUT5 in Caco-2 cells and in Xenopus oocytes expressing the mammalian GLUT5 [106]. Likewise, Satsu et al. demonstrated that epicatechin gallate inhibited fructose uptake in Caco-2 cells. Interestingly, this reduction in fructose uptake was not related to changes in the affinity ( Km ) of GLUT5 for fructose, but with a decrease in the maximal velocity ( Vmax ) [107]. Furthermore, authors demonstrated that epicatechin gallate suppressed fructose permeation in Caco-2 cells, suggesting that this compound suppressed the transepithelial transport of fructose across epithelial cell monolayers, in addition to its effect on fructose uptake. Lastly, authors reported that similar effects on fructose uptake and permeation were observed with nobiletin, another phytochemical tested in this study [107].


Hormones of the Hypothalamus and Pituitary Gland

The hypothalamic–pituitary axis can be considered the coordinating center of the endocrine system. The hypothalamus is the main neural control center, also known as the “master switchboard,” which coordinates nervous and endocrine system functions. The hypothalamus consolidates inputs derived from higher brain centers, various environmental cues, and endocrine feedback. Neurons within the hypothalamus produce and secrete releasing hormones, such as corticotropin-releasing factor (CRF), luteinizing hormone–releasing hormone (LHRH), thyrotropin-releasing hormone (TRH), and growth hormone–releasing hormone (GRH), as well as inhibiting hormones, such as somatostatin and dopamine, directly into the blood vessel connecting the hypothalamus with the pituitary gland (i.e., the hypothalamic– hypophyseal portal vein). These hormones then control the synthesis and release of hormones in the pituitary gland. The pituitary gland comprises two sections—the adenohypophysis, or anterior lobe, and the neurohypophysis, or posterior lobe. In response to signals from the hypothalamus, the anterior pituitary produces and secretes trophic hormones, which are hormones that have a growth effect on the organs or tissues they are targeting. They include, among others, adrenocorticotropic hormone (ACTH), thyroid-stimulating hormone (TSH), follicle-stimulating hormone (FSH), luteinizing hormone (LH), prolactin, and growth hormone (GH) and modulate the functions of several peripheral endocrine glands (i.e., adrenal glands, thyroid, and gonads) and tissues (e.g., breast, muscle, liver, bone, and skin) (see the table).

Main Function or Target Organ

Luteinizing hormone–releasing hormone

Growth hormone–releasing hormone

Growth hormone/insulin-like growth factor-1

Growth hormone/insulin-like growth factor-1, Hypothalamic–pituitary–thyroid axis

Growth hormone/insulin-like growth factor-1

Growth and repair of all cells

Hypothalamus/ Posterior Pituitary Gland

Uterus, mammary glands, male reproductive organs

Glucocorticoids (cortisol, corticosterone)

Body stress, metabolism, glucose maintenance

Estrogen (estrone, estradiol, estriol)

Female reproductive glands and tissues, bones, heart

Maintenance of pregnancy and preparation of breast tissue

Masculinity, sperm production, bone

Heart rate, temperature, metabolism

The posterior or neurohypophyseal lobe of the pituitary contains the terminals of certain neurons (i.e., magnocellular vasopressin- and oxytocin-producing neurons) originating in two specific sections (i.e., the paraventricular nuclei [PVN] and supraoptic nuclei) of the hypothalamus. These neurons secrete primarily two hormones from the posterior pituitary into the systemic blood: arginine vasopressin (AVP), which controls the renal water handling and cardiovascular functions, and oxytocin, which regulates milk ejection during lactation and uterine contractions during birth. Evidence also indicates that both AVP and oxytocin act not only as hormones but also as neuromodulators and neurotransmitters within the central nervous system (de Wied et al. 1993 Stoop 2014). However, AVP and oxytocin also can be produced in another group of neurons in the PVN and supraoptic nuclei (i.e., in the parvocellular neurons) and released into the hypothalamic–hypophyseal portal vessels to reach the anterior pituitary. There, AVP acts synergistically with CRF to promote secretion of ACTH (Plotsky 1991). In contrast, oxytocin acts on specialized cells in the anterior pituitary to promote prolactin secretion (Sarkar and Gibbs 1984).


RESULTS

The apoE −/− mice had markedly increased fasting plasma cholesterol levels compared with apoE +/+ mice when fed either control or diabetogenic diets (Table 1). Fasting plasma TG levels were markedly higher in apoE −/− mice compared with apoE +/+ mice, regardless of the diet (Table 1). Diabetogenic diet increased fasting TG levels significantly in both groups of mice but to a larger extent in apoE −/− mice. Although we found similar TG levels in nonfasted apoE +/+ and apoE −/− mice under control dietary conditions, the nonfasting TG levels were higher in apoE −/− mice compared with apoE +/+ mice when they were fed diabetogenic diet. Fractionation of plasma lipoproteins by FPLC showed markedly increased cholesterol levels in the VLDL and IDL/LDL fractions of apoE −/− mice compared with apoE +/+ mice (Fig. 1A and B) as described previously (24). Feeding the diabetogenic diet resulted in a marked increase in HDL cholesterol in both apoE −/− and apoE +/+ mice compared with control diet–fed groups (Fig. 1A and B). In addition, significant increase in cholesterol level was observed in fractions 20–26 of diabetogenic diet–fed apoE +/+ mice (Fig. 1A). Because of the expected overlaps in particle sizes between IDL/LDL and HDL1 lipoproteins, immunoblot analysis was performed with antibodies against apoAI and apoE, two proteins present in HDL1 but not in LDL. Results confirmed that the cholesterol increase in these fractions was due primarily to increase levels of HDL1 (Fig. 1C). The diabetogenic diet–induced elevation of HDL1 level was not apparent in apoE −/− mice nor in mice fed control diet (Fig. 1B and C). The increased VLDL and decreased HDL1 concentrations in apoE −/− mice are consistent with the well-established defects in receptor-mediated lipid transport pathways associated with apoE deficiency. These results also established that apoE −/− mice are appropriate models for investigating whether suppression of receptor-mediated lipid transport to high–energy metabolism tissues affects insulin responsiveness and diet-induced obesity.

The control diet–fed apoE −/− mice exhibited lower fasting glucose but not insulin levels compared with apoE +/+ mice (Table 1). Importantly, both glucose and insulin levels were significantly lower in apoE −/− mice than in apoE +/+ mice during the fed state when the animals were maintained on the control diet (Table 1), suggesting that apoE −/− mice on control diet were more insulin sensitive than apoE +/+ mice. Diabetogenic diet feeding increased insulin levels in both groups of mice under both fasting and fed states, but the increases were significantly less in apoE −/− mice compared with apoE +/+ mice (6.86- vs. 9.5-fold in fasted state and 5.36- vs. 7-fold in fed state, respectively). Although glucose levels were similar in both groups of diabetogenic diet–fed mice, the markedly lower fasting and fed insulin levels in apoE −/− mice are indicative of their reduced sensitivity to diet-induced insulin resistance. These latter findings were confirmed by ipGTT after a 6-h fasting period. In these experiments, the response to glucose was found to be significantly improved in apoE −/− mice compared with apoE +/+ mice regardless of the diet (Fig. 2A).

The highly efficient glucose tolerance mechanism in the lipid transport–defective apoE −/− mice suggested that the delayed uptake of lipids into tissues during the postprandial state may reduce intracellular lipid accumulation and thereby lead to increased insulin sensitivity. This hypothesis was examined directly by performing ipGTT with or without a simultaneously administered oral fat tolerance test (oFTT) in mice on control diet after an overnight fast. This experimental design allowed us to compare glucose disposal in a truly fasted state with or without an acute influx of lipids. We found that whereas plasma TG clearance after an oral lipid load was significantly impaired in apoE −/− mice (Fig. 2B), glucose tolerance was notably improved in apoE −/− mice compared with apoE +/+ mice when an oFTT was conducted simultaneously (Fig. 2C). In contrast, no difference in glucose response between apoE −/− and apoE +/+ mice was observed when ipGTT was performed after an overnight fast without concomitant influx of dietary lipids (Fig. 2D). The latter results contrasted with results obtained after a 6-h fasting period (Fig. 2A). The difference is likely due to ongoing lipid uptake and metabolism in apoE +/+ mice 6 h after feeding, which was not apparent in apoE −/− mice with defective receptor-mediated lipid uptake.

The impact of defective lipid uptake on glucose disposal by various tissues was investigated by intravenous GTT using deoxy[ 3 H]glucose as a tracer after its infusion with lipids into apoE +/+ and apoE −/− mice. Based on our previous observations that circulating glucose levels peaked at 2 min and were eliminated thereafter by first-order kinetics after intravenous administration of glucose with plasma glucose levels reducing to 50% levels after 15 min (25), we examined the disposition of the deoxy[ 3 H]glucose in various tissues 15 min after intravenous injection as an indication of the rate of glucose uptake by each tissue. We observed a significant increase in glucose uptake by BATs and skeletal muscle but not the heart of apoE −/− mice compared with apoE +/+ mice on diabetogenic diet (Fig. 3A).

Additional experiments were also conducted to assess the impact of apoE-mediated lipid uptake on glucose metabolism in various tissues by analyzing tissue distribution of deoxy[ 3 H]glucose with and without a concomitant oral lipid load in apoE +/+ and apoE −/− mice maintained on control low-fat diet. When deoxy[ 3 H]glucose was administered in the presence of an oral fat load, a significant increase in deoxy[ 3 H]glucose uptake by the subcutaneous white adipose tissue (SAT) was observed in control diet–fed apoE −/− mice compared with that in apoE +/+ mice (Fig. 3B). The uptake of deoxy[ 3 H]glucose by other tissues was similar between both groups under these conditions. The adipose-specific difference in deoxyglucose uptake between apoE +/+ and apoE −/− mice with a fatty meal may explain the slight but significant improvement of glucose tolerance in the apoE −/− mice when glucose was administered together with a fat load. Interestingly, whereas the difference in SAT uptake of deoxy[ 3 H]glucose between control diet–fed apoE +/+ and apoE −/− mice was sustained in the absence of an oral lipid load, significantly less deoxyglucose uptake was observed in the skeletal muscle of apoE −/− mice compared with that in apoE +/+ mice in the absence of an oral fat load (Fig. 3C). The latter observation is consistent with data showing no difference in glucose tolerance between chow-fed apoE +/+ and apoE −/− mice in the absence of an oral lipid load.

The decrease in postprandial fat clearance in apoE −/− mice compared with apoE +/+ mice suggested that the apoE −/− mice may also be resistant to diet-induced adiposity. Therefore, body weight and fat mass of apoE −/− and apoE +/+ mice were analyzed after feeding them with either control or diabetogenic diet. Despite similar body weights between the two groups of animals during regular control diet feeding, the apoE −/− mice had significantly lower whole-body fat mass compared with that observed in apoE +/+ mice (Fig. 4A). The apoE −/− mice also gained significantly less whole-body fat mass after diabetogenic diet feeding compared with apoE +/+ mice, resulting in significantly reduced total body weights after 16 weeks (Fig. 4A and B). The differences in fat mass and body weight gain between apoE +/+ and apoE −/− mice were not related to major differences in energy use, because both groups of mice showed similar energy expenditure (base on lean body mass) when fed either the basal or diabetogenic diet (Fig. 5A and B). Caloric intake and respiratory quotient were also similar between apoE +/+ and apoE −/− mice on both diets (Fig. 5C and D Supplemental Figure S1, which is detailed in the online appendix [available at http://dx.doi.org/10.2337/db07-0403]). Interestingly, a slight increase in UCP-1 expression in BAT of apoE −/− mice was consistently observed, but the increase did not reach statistical significance (Fig. 5E).


2 MATERIALS AND METHODS

2.1 Antibodies, primers, and chemical reagents

All reagents were purchased from Sigma-Aldrich, Australia unless stated. Primary antibodies used were anti-TBC1D1 (#4629), anti-S6RP (#2317), anti-pS6RP Ser235/236 (#4856), anti-4E-BP1 (#9452), anti-p4E-BP1 Ser65 (#9451), anti-SMAD2/3 (#8685), anti-insulin (#9016), and anti-glucagon (#2760) all from Cell Signaling Technologies anti-GAPDH (#SC32233) from Santa Cruz Biotechnology anti-pSMAD3 Ser423/Ser425 (#ab52903) from Abcam and anti-FST (#AF669) from R&D Systems. Secondary antibodies linked to HRP for western blotting of tissue lysates were anti-mouse (#1706515), anti-rabbit (#1706516), and anti-goat (#1721034) from Bio-Rad Technologies. Secondary antibodies conjugated to fluorophores for immunofluorescent labeling of tissue sections were anti-rabbit (A-11037) and anti-goat (A-11058) from Thermo Fisher Scientific.

2.2 Animal husbandry

All experiments were conducted in accordance with the relevant codes of practice for the care and use of animals for scientific purposes (National Institutes of Health, 1985 and the National Health and Medical Council of Australia, 2004). All animal work performed in this study was conducted with approval from the Alfred Medical Research and Education Precinct Animal Ethics Committee (AMREP AEC), Melbourne, Australia. Cohorts of BKS.Cg-Dock7 m +/+ Lepr db /J mice (referred to as db/db herein, and sourced from Jackson Laboratories) were used to model hyperphagia-induced obesity and T2D, 24, 25 with BKS.Cg-Dock7 m +/+ Lepr db /+ heterozygous mice (referred to as +/db herein, and also sourced from Jackson Laboratories) used as nondiabetic controls. All mice were housed in groups of at least three on a 12:12 hours dark/light cycle with ad libitum access to standard chow diet (Specialty Feeds) and water, except where indicated. Body weight and composition were determined by quantitative magnetic resonance (EchoMRI) the day prior to AAV vector administration, and then weekly until the study end.

2.3 Recombinant AAV vector production and administration

Recombinant AAV vectors comprising pseudotype-6 capsids (rAAV6) and carrying constructs for the short isoform of human Follistatin under the control of a CMV promoter (rAAV6:Fst317), or a noncoding control vector (rAAV6:CON) were produced as described previously. 20 Administration of rAAV6:FST317 or rAAV6:CON to mice was performed by intravenous injection of a lateral tail vein, which has been shown to transduce the striated musculature of adult mice. 26, 27 Cohorts of mice treated at 5 weeks of age were administered 2 × 10 12 vector genomes of rAAV6:FST317 or rAAV6:CON. Cohorts of mice treated at 14 weeks of age were administered 6 × 10 12 vector genomes of either vector.

2.4 Sample preparation and immunoblot analysis

Harvested skeletal muscles were lysed in RIPA buffer (25 mM Tris-HCl pH 7.6, 150 mM NaCl, 1% NP-40, 1% sodium deoxycholate, 0.1% SDS), reduced with 2.5% β-mercaptoethanol, supplemented with protease, and phosphatase inhibitor cocktail. Protein concentrations were determined using the bicinchoninic acid (BCA) assay (Thermo Fisher Scientific, Victoria, Australia). Proteins were separated on 4%-12% SDS PAGE gels (Life Technologies) and transferred to PVDF membranes (Millipore, Australia). Primary antibody incubation was performed overnight followed by incubation for 1 hour at RT with HRP-conjugated secondary antibodies and detection using ECL detection reagent (GE Healthcare life sciences, Australia). Densitometry was performed using ImageJ software (http://rsb.info.nih.gov/ij/index.html) and the abundance of GAPDH was used for normalization unless indicated.

2.5 Blood glucose and insulin measurement and tolerance tests

Blood samples were collected via tail vein bleed and analyzed for glucose concentration by glucometer (AccuVue) and insulin concentration via mouse insulin ELISA (Alpco) performed according to manufacturer's instructions. Intraperitoneal insulin tolerance tests (ipITT) were performed after mice were fasted for 6 hours. Each mouse was injected with 1 U/kg of lean body mass insulin and blood samples were collected via the tail vein at the indicated timepoints to measure the glucose concentration with a glucometer (AccuVue). Oral glucose tolerance test (oGTT) was performed after a 6 hours fast. Mice were administered glucose at 2 g/kg of lean body mass via oral gavage and blood samples were collected via the tail vein at the indicated timepoints to measure glucose concentration with a glucometer (AccuVue).

2.6 Tracer measurements of glucose uptake

Mice were administered 3H-2-deoxyglucose (2-DOG 2 µCi/25 g body weight) with glucose (1g/kg lean body mass) via intraperitoneal injection. Blood samples were obtained at 3, 15, 30, 45, 60, 90, and 120 minutes using 5 µL heparinized hematocrit tubes (Drummond) and immediately added to 100 µL saturated BaOH solution. Radioactivity was measured from blood supernatants after saturated ZnSO4 solution precipitation. Frozen tissues were homogenized in 500 µL H2O, and soluble supernatant was collected. Supernatant (150 µL) was diluted to 1 mL and added to 4 mL scintillation fluid (Ultima Gold, PerkinElmer) for total counts. Free (nonphosphorylated) glucose counts were measured from supernatant samples after elution off anion exchange columns prepared using AG 1-X8 resin (Bio-Rad). Vortexed samples were read on a Beckman LS 6500 β-counter. 2-Deoxyglucose uptake and phosphorylation in tissues was determined by subtracting AG 1-X8 resin eluate readings from total, homogenized tissue readings and normalized for tissue weight, blood glucose concentrations during the GTT, and radioactive AUC determined from blood samples taken during the GTT.

2.7 Assessment of metabolic parameters, albuminuria, and glycated hemoglobin

For estimation of oxygen consumption (VO2) mice were placed in a 12-chamber indirect calorimeter (Oxymax series Columbus Instruments, Columbus, OH) at a constant environmental temperature (22°C) with an airflow of 0.6 L/min and ad libitum access to food and water. After 12 hours of acclimation, VO2 was measured in individual mice at 12-min intervals over a 24-hour period. For analysis of food consumption and urine production, mice were placed individually into metabolic cages (Iffa Credo, L’Arbresele) for 24-hour monitoring and collection. Albumin excretion rates were measured in 24 hours urine collections using a mouse albumin ELISA quantitation kit (Bethyl Laboratories, Montgomery, TX). Glycated hemoglobin was measured from tail vein bleeds, as described previously. 28

2.8 Microscopic assessment of pancreas and kidney attributes

At experimental endpoints, the pancreas was dissected from each mouse immediately after euthanasia and fixed for 1-2 hours in 4% (w/v) paraformaldehyde. Samples were washed and stored in 70% (w/v) ethanol and embedded in paraffin. For immuno-labeling, 5-8 micron-thick sections were rehydrated in PBS. Sections labeled for Glucagon and ALDH1A3 were incubated in Histo-One VT (Nacalai Tesque, Japan) for 20 minutes at 80°C. Sections labeled for Follistatin were incubated in sodium citrate buffer and heated in a pressure cooker for 10 minutes. Following antigen retrieval, sections were incubated in blocking buffer (5% goat or horse serum, PBS, 0.3% v/v Triton X-100) for 1 hour at RT and incubated overnight at 4°C with anti-Glucagon antibody (1:200), anti-ALDH1A3 (1:100), or anti-FST (1:200). For Glucagon and ALDH1A3 labeling, sections were washed in PBS, and incubated overnight at 4°C with Alexa 594-conjugated anti-rabbit IgG (1:400). For Follistatin labeling, sections were incubated for 1 hour at RT with Alexa 594-conjugated anti-goat IgG (1:250). The sections were then washed in PBS and incubated with Alexa 488-labeled anti-insulin antibody (1:200) for 2 hours at RT. Sections were washed, incubated with 0.1uM DAPI (ThermoSciences) for 5 minutes, and mounted with Mowiol mounting medium. Histological images of tissues were acquired on a fluorescent microscope at 20x magnification (AR1, Nikon, and Axio Imager M2, Zeiss). At least 10 islets were acquired from four to five mice from each treatment and genotype for Glucagon and ALDH1A3 immuno-labeling and at least three islets were acquired from 3-4 mice for Follistatin immuno-labeling. Quantification was performed using Fiji imaging software. 29 Analyses were blinded for genotype and treatment. Kidney sections (3µm) were stained with Periodic Acid-Schiff (PAS) to measure the mesangial expansion and glomerulosclerotic injury (GSI). Mesangial area was analyzed (percentage of glomerular area) from digital pictures of glomeruli (20 glomeruli per kidney per animal) using Image-Pro plus 7.0 software (Media Cybernetics, Bethesda, MD), as described previously. 28 GSI was graded based on the severity of glomerular damage, including mesangial matrix expansion, hyalinosis with focal adhesion, capillary dilation, glomerular tuft occlusion, and sclerosis, as previously described. 30 Twenty glomeruli per kidney were assessed in a masked fashion.

2.9 Immunoassays for protein measurements

Activin A was measured in serum and tissue lysates using a specific Activin A ELISA employing antibodies supplied by Oxford Brookes University, as described previously. 31 The limit of detection was 11.3 pg/mL. Activin B was measured in serum and tissue lysates using a specific Activin B ELISA, as reported by Ludlow and coworkers, using antibodies from Oxford Brookes University. 32 The assay was validated for measurement of Activin B in mouse tissues. The limit of detection was 28.4 pg/mL.

2.10 Statistical analyses

Allocation of rAAV6:FST317 or rAAV6:CON treatments within genotype was random. Studies were performed unblinded except for the microscopic analyses of pancreas and kidney sections. To compare repeated measures for statistical differences across treatment and genotype conditions, two-way ANOVAs were performed. Pre-planned comparisons were made across conditions with Fisher's Least Square Difference. To assess the statistical differences across conditions, one-way ANOVAs were performed, with Tukey's post hoc test for comparisons between the specific group means, using GraphPad Prism v.7 (GraphPad). Comparisons between two conditions used the Student's t test. Differences between groups where P < .05 are indicated by character labels. Data are presented as the means ± SEM.


Discussion

The aim of this study was to determine potential sex differences in metabolic effects of chronic Ang-(1-7) treatment in HFD-induced obese mice. The main findings are that (1) male and female mice develop a similar obese metabolic phenotype in response to HFD, with the exception of a milder hyperinsulinemia in females (2) chronic Ang-(1-7) treatment reduces body mass and adiposity and improves lean mass in obese mice of both sexes, with no effect on body composition in control diet-fed mice and (3) Ang-(1-7) reverses HFD-induced insulin resistance in both sexes but only improves glucose tolerance in females. These collective data provide new evidence for sexual dimorphism in effects of chronic Ang-(1-7) treatment in obese mice, with females potentially being more responsive in terms of glucose tolerance. These findings advance our limited understanding of sex differences in RAS mechanisms involved in glucose homeostasis and provide new insight for the potential for targeting Ang-(1-7) as a novel therapeutic strategy for metabolic complications in obesity.

The HFD-induced obese mouse has been used extensively as a model for obesity, given its similarity in terms of pathophysiology to the human condition [29]. C57BL/6 mice, in particular, are susceptible to increased adiposity, hyperglycemia, hyperinsulinemia, insulin resistance, and glucose intolerance when chronically exposed to a HFD. Historically, most studies in this model have been performed in males as they develop a more severe degree of obesity and related metabolic complications and to avoid potential estrous-associated physiological alterations [4, 29, 30]. Recent studies, however, have explored sex differences in body composition and glucose homeostasis in this model. For example, one study showed that while HFD-induced obese female mice accumulate more subcutaneous and epididymal fat compared with males, they have reduced circulating insulin levels and develop milder glucose intolerance than their male counterparts [30]. Similarly, HFD-fed female mice are reported to exhibit greater weight gain and adiposity compared with male mice and are protected from obesity hypertension [23]. These findings appear to support clinical literature showing that despite having higher adiposity, females may be protected from obesity-related metabolic and cardiovascular complications.

In the present study, we observed that HFD increases body mass in both sexes but to a greater extent in male mice. Despite lower weight gain, HFD-fed female mice exhibited similar adiposity when compared with males. A limitation of our study is that we did not systematically assess for differences in visceral versus subcutaneous adipose depot distribution between sexes, or in response to diet or drug treatment. Interestingly, we found that female mice develop obesity-induced hyperinsulinemia to a lesser extent compared with male mice, despite having similar mild hyperglycemia. This may suggest obese female mice are more insulin responsive than obese males, as they appear to require less insulin to maintain blood glucose levels however, we found that HFD produced similar insulin resistance in both sexes when measured by ipITT. The finding that obese female mice were insulin resistant despite lack of marked hyperinsulinemia contrasts what is typically seen in the human population where hyperinsulinemia is an early indicator of prediabetes and T2DM and is closely linked with concurrent insulin resistance [31, 32]. Conversely, genetically altered mice in which insulin secretion is limited are resistant to HFD-induced obesity [33]. Unlike these mice, however, we found that female mice develop obesity and increases in adiposity, suggesting an alternative mechanism of action for their maintenance of normoinsulinemic levels.

Previous studies have shown that Ang-(1-7) reduces body mass and adiposity [13, 20,21,22] and has protective effects on skeletal muscle composition and function [34], in male rodents. Similar to these findings, we found that Ang-(1-7) improves overall body composition in obese male and female mice by reducing percentage of fat and fluid masses and increasing percentage of lean mass. It is important to note, however, that these mice still remained obese, which may reflect the short 3-week duration of Ang-(1-7) treatment in our study. Since energy balance is tightly regulated, it may take more extended time frames to manifest changes in body mass. In support of this, one study found that male fructose-fed rats supplemented with Ang-(1-7) for 4 weeks had similar weight gain as the corresponding saline group [35]. When the length of treatment was extended to 6 months, however, fructose-receiving rats had similar body mass and adiposity compared with controls. Therefore, extending the length of treatment may result in more profound improvements in body composition in both sexes.

There are conflicting reports involving Ang-(1-7) effects on fasting glucose and insulin levels. One group found that Ang-(1-7) significantly reduces baseline blood glucose, with no effect on basal insulin levels, in male fructose-fed rats [35]. Other studies, however, have shown Ang-(1-7) has no effect on fasting glucose levels with a trend to decrease baseline insulin concentrations [17, 22]. The discrepancy may correlate with differences in species (rats versus mice), obesity models (HFD versus fructose), and length of treatment. Our results showed that Ang-(1-7) has no effect on fasting plasma glucose or insulin levels, regardless of sex or diet received. This is consistent with a recent study from our laboratory showing that a similar duration of Ang-(1-7) treatment did not produce significant effects on fasting glucose or insulin levels, although a trend for a reduction in insulin was observed [22]. The reason for this outcome is unclear but again may reflect Ang-(1-7) therapy duration. Since improvements in insulin sensitivity often occur prior to correction of hyperglycemia, it is possible that longer durations of treatment are needed to manifest changes in glucose and insulin levels. In support of this, a recent study showed changes in plasma insulin at 4 weeks, followed by a reduction in glucose at 9 weeks, after chronic Ang-(1-7) therapy in the db/db diabetic mouse model [36].

Ang-(1-7) improves insulin sensitivity in lean, obese, and diabetic male rodent models via numerous mechanisms including positive effects on intracellular insulin signaling pathways and increasing glucose uptake in peripheral tissues [11,12,13,14, 22]. A previous study from our laboratory showed that Ang-(1-7) improves whole-body insulin sensitivity in HFD-induced obese male mice by enhancing glucose uptake within skeletal muscle through increased expression of sarcolemmal glucose 4 transporters (GLUT4) [22]. In the current study, we similarly found that Ang-(1-7) reverses insulin resistance in HFD-induced obese male mice. We expand on these previous findings by demonstrating Ang-(1-7) also improves insulin sensitivity to a similar extent in HFD-induced obese females. The mechanism of action for this return of insulin sensitivity in females is currently unknown but is anticipated to reflect skeletal muscle insulin sensitization similar to what has been previously seen in males [22].

Chronic Ang-(1-7) administration or ACE2 activation also improves glucose tolerance in male rodent models of metabolic syndrome and T2DM [13, 14, 21, 35, 36]. In this study, we found that Ang-(1-7) improved the ability to dispose of exogenous glucose from the bloodstream in HFD-fed female mice, but not in males. Since earlier studies demonstrated that Ang-(1-7) improves pancreatic β cell function to increase glucose-mediated insulin secretion [17, 37, 38], we assessed for insulin receptivity in response to dextrose administration. We found that Ang-(1-7)-treated mice had higher glucose-stimulated insulin concentrations independent of sex or diet. In addition to insulin secretion, glucose tolerance tests induce multiple physiological responses including intestinal glucose absorption, insulin sensitivity, and uptake of glucose in peripheral tissues, glucose effectiveness, and counter-regulatory mechanisms, any of which could account for these sex differences [39]. In addition, while not explored in this study, Ang-(1-7)-mediated vasodilation is more pronounced in women versus men [27], which could serve to increase rate of glucose shuttling to peripheral tissues to enhance glucose tolerance.

There are currently limited studies examining sex differences in circulating Ang peptides in rodent models [23, 24, 40, 41]. In the present study, there were no significant main effects of diet or sex on Ang II concentrations. Similar to our findings, one study showed no difference in Ang II in HFD versus control diet-fed male mice. Another study showed, however, that HFD increases Ang II in males, with no effect on levels of this hormone in females. Similar to our findings, a few studies have shown no sex differences in Ang II levels in normotensive rats and healthy humans however, others have shown that males have higher levels of Ang II compared with females in obese mice and in control, hypertensive, and diabetic rats. Ang-(1-7) infusion elevated circulating Ang II levels in this study, which was more conspicuous in chow-fed mice and with no sex interaction. Our results parallel previous findings in chow- and HFD-fed groups [22], with this counterintuitive elevation in Ang II perhaps reflecting a physiological balance response.

In terms of Ang-(1-7), a significant diet effect was not detected, although a trend was apparent for HFD to decrease levels in males and increase levels in females. This is consistent with our previous report showing reduced Ang-(1-7) levels in HFD-induced obese male mice. An additional report showed no effect in male mice, but an increase in Ang-(1-7) in female mice in response to HFD as a potential compensatory mechanism to protect against development of hypertension [23]. In this study, we found a significant main effect for sex, with males exhibiting higher levels compared with females, particularly under control diet conditions. This finding is consistent with a previous report in chow-fed mice [23]. It contrasts, however, with studies showing higher circulating Ang-(1-7) concentrations in healthy women and hypertensive rats, and higher renal Ang-(1-7) in female rats [27, 41,42,43]. Additionally, studies have shown no sex differences in Ang-(1-7) levels in obese mice, normotensive rats, and diabetic rats [23, 40, 41]. Similar to our previous study [22], chronic Ang-(1-7) infusion increased plasma Ang-(1-7) levels, with effects most prominent in males and under chow diet conditions.

Overall, these previous studies have shown inconsistent results for diet and sex effects on circulating Ang II and Ang-(1-7) concentrations. These disparate findings may reflect differences in species (e.g., rats, mice, humans), disease models (e.g., diet-induced obesity, type I diabetes, hypertension, healthy), and assays used (e.g., radioimmunoassay, ELISA). In addition, we observed large variability in Ang peptide levels among individual mice, which may reflect inter-assay variability as well as differences in cohorts.


Guidelines & Statements

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INTRODUCTION

Transdermal drug delivery (TDD) offers a convenient and patient-friendly way for the treatment of both local disorders and diseases of other organs. It allows drugs to bypass the first pass metabolism while providing sustained and controlled delivery (1, 2). While traditional cream or gel formulations passively deliver small and lipophilic drugs through the stratum corneum (SC) via diffusion, macromolecules (>500 Da) with very low permeability cannot diffuse through the compact lipid-rich matrix of SC (3). Chemical and biochemical enhancement techniques have been developed over the past decades to disrupt SC for the delivery of large and hydrophilic drugs. However, these enhancers are potent irritants at high concentrations and thus affect therapeutic compliance (4, 5). Physical enhancement methods amiable for routine and effective delivery have also been developed (e.g., electroporation, laser ablation, and iontophoresis) (1, 6, 7). Nonetheless, these physical means are accomplished through utilization of additional, specialized equipment alongside trained users, to modulate skin permeability, thereby presenting challenges in terms of patient convenience and home use (8). Another technology worth mentioning is the microneedle (MN), which has received vast attention recently (9, 10). They can be fabricated with different needle length and width (150- to 1500-μm long, 50- to 250-μm wide), with a few drug loading options (10). However, because of their small sizes, the amount of drugs that can be delivered is usually within microgram range (11, 12).

Amidst the popular technologies mentioned above, rubbing serves as a potential method to deliver drugs transdermally. While the mechanism of this method is still unclear (13, 14), the limited literature review on rubbing suggests that the locally applied pressure can benefit TDD. One example is the jet injector. High-pressure liquid jet of

14 MPa and a diameter of 340 μm can pierce the skin, with a depth of

2 mm, for delivery of medication such as corticosteroids (15), local anesthesia (16), bleomycin (17), etc. However, the piercing of the skin is a concern to many researchers and clinicians due to risk of infection, pain, or injury of operator’s finger (18). Another technology using similar concept is the ultrasound-mediated TDD that administers an acoustic oscillating pressure wave. The oscillation increases the size of cavitation bubbles in the fluids of the skin. When imploded, these bubbles create an intense local shockwave, disrupting the SC for improved TDD. The range of frequencies is in the range of 20 kHz to 16 MHz to deliver insulin, mannitol, glucose, morphine, and lidocaine (19). While ultrasound-mediated TDD has proven to be effective, it is limited by the complex machinery and associated skin tissue heating, which can damage the deeper tissue (20). Therefore, there still remains a need for simple, cost-effective, and minimally invasive TDD technology to enable convenient transdermal drug administration by the patients.

This article introduces a pressure-based TDD methodology to address this unmet need (Fig. 1A and fig. S1). Tapping on established studies where magnets are used to mimic pressure ulcer injury in vivo due to its well-defined magnetic field and force (21), this study chose neodymium magnets as the proof-of-concept tool to produce the required pressure force to induce temporal skin barrier changes for drug delivery. Specifically, a local pressure (0.14, 0.28, and 0.4 MPa) is generated by applying two neodymium magnets to pinch skin. After a given time, the magnets are removed, and therapeutics in the moisturizer are topically applied to the pressure-processed area. Here, we report the successful delivery of nanoparticles (NPs up to 500 nm), dextran molecules (up to 20 kDa), and insulin across the skin of mice. The improved penetration is observed through inter- and transcellular routes. We optimize this process and propose 0.28 MPa and 1 min of application as the ideal procedure to achieve an effective penetration without compromising the skin barrier. Furthermore, we demonstrate the effective delivery of insulin into the systemic circulation of both normal and diabetic mice and achieve the 65 and 80% reduction in blood glucose levels, respectively.

(A) Schematic diagram demonstrating the effect of temporal pressure application leading to the occurrence of microphysiological changes, allowing the delivery of drugs across the skin barrier. (B) Representative images of mice with topically applied fluorescent nanoparticles (NPs) after the pressure and MN treatment. (C) Quantification of the fluorescent signal in (A). (D) Quantification of NPs in the dermis. (E) Fluorescence imaging [blue, 4′,6-diamidino-2-phenylindole (DAPI) red, NPs] of histological skin samples in (A). (F) Left: H&E staining of skin samples. Right: The appearance of mouse skin with or without the pressure treatment and MN. Scale bars, 100 μm. n = 3, all data are means ± SD, *P < 0.05. Photo credit: Daniel Chin Shiuan Lio, School of Chemical and Biomedical Engineering, Nanyang Technological University.


The studies involving human participants were reviewed and approved by Northern Sydney Local Health District Human Research Ethics Committee. The patients/participants provided their written informed consent to participate in this study. The animal study was reviewed and approved by Sydney Local Health District Animal Ethics committee.

KB, DR, SC, MF and ZC performed all the animal experiments. KB, MT, MF and OT performed all the cell experiments and tissue analysis. KB, MK, SP and GF conceived and designed the experiments. KB and GF wrote the manuscript. All authors drafted and approved the final manuscript.


ACKNOWLEDGMENTS

This study was supported by an American Diabetes Association Junior Faculty Award (1-08-JF-58) and a National Institutes of Health (NIH) grant (DK085129) to B.W., an NIH grant (DK074966) to M.W.R., NIH grants to L.H.P. (DK48494 and DK063493), and by University of Chicago Diabetes Research and Training Center funding from the NIH (DK020595) to B.W. and L.H.P. K.K. was supported by a T32 National Institute of Diabetes and Digestive and Kidney Diseases training grant (DK087703) and by an NIH F31 grant (AG035620).

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

K.A.K. and B.W. designed this study. K.A.K., L.M.D., D.A.J., and B.W. conducted the experimental research and analyzed the data. K.A.K., L.M.D., D.A.J., M.W.R., L.H.P., and B.W. contributed to the preparation of the manuscript. N.T., M.W.R., and L.H.P. provided the MIP-CreERT mice, an essential reagent for this study. B.W. is the guarantor of this work and, as such, had full access to all the data in the study and takes responsibility for the integrity of the data and the accuracy of the data analysis.

The authors are grateful to Dr. Stanley McKnight (University of Washington, Seattle, Washington) for providing the PKA-CαR mice.


Watch the video: Glucose Transporter GLUT: How Does it Work? (August 2022).