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Departments of Medicine and Pharmacology (L.W., J.H.), University of Kansas Medical Center, Kansas City, Kansas 66160; Departments of Molecular and Cellular Biology (P.S., L.C., D.D.M.) and Medicine (M.H., L.C., A.S.R.), Baylor College of Medicine, Houston, Texas 77030; Joslin Diabetes Center, Harvard Medical School (R.N.K.), Boston, Massachusetts 02215; and Department of Internal Medicine (Y.K., K.P., I.L.), Keimyung University School of Medicine, Daegu 700-712, Korea
Address all correspondence and requests for reprints to: Li Wang, Departments of Medicine and Pharmacology, University of Kansas Medical Center, Kansas City, Kansas 66160. E-mail: lwang2{at}kumc.edu.
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(PPAR) coactivator-1
-stimulated glucose transporter 4 expression and glucose uptake. SHP–/– hepatocytes showed markedly decreased basal glucose production in cultures, and SHP–/– livers had increased glycogen stores and were more sensitive to insulin inhibition of glucose output, which were concomitant with decreased expression for PPAR1, fatty acid translocase, glucose-6-phosphatase, and phosphoenol/pyruvate carboxykinase, and increased mRNAs for glucokinase and pyruvate kinase. In white fat, SHP deficiency resulted in up-regulation of genes involved in insulin sensitizing, including PPAR2 and adiponectin. We show that, at the transcriptional level, SHP directly represses adiponectin promoter activity by PPAR/liver receptor homolog-1. The results suggest that the increases in insulin sensitivity through multiple signaling pathways in muscle, liver, and fat, with an increase in islet secretory function, represent the complex mechanism whereby SHP deficiency leads to improvement in insulin sensitivity, secretion, and diabetes. |
INTRODUCTION |
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TOPABSTRACTINTRODUCTIONRESULTSDISCUSSIONMATERIALS AND METHODSREFERENCES |
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(3, 4), retinoid X receptor (RXR), and hepatocyte nuclear factor-4 (HNF4) (5), and especially liver receptor homolog-1 (LRH-1) (6) and estrogen receptor-related receptor (7), and inhibits their transcriptional activity. Nuclear receptors are key regulators of metabolic pathways, and previous studies with SHP knockout mice have confirmed that SHP functions in bile acid, cholesterol, and triglyceride homeostasis (8, 9, 10, 11). In humans, loss of SHP function has been associated with mild obesity in Japanese subjects (12), but genetic variation in SHP is not commonly associated with obesity in United Kingdom Caucasians (13, 14). In contrast, we did not observe weight differences between wild-type and SHP–/– mice on a regular chow diet, but found that SHP–/– mice are resistant to high-fat diet-induced obesity, apparently as a consequence of increased peroxisomal proliferator-activated receptor coactivator-1 (PGC-1) expression in brown adipose tissue (15). Recently, the potential role of SHP in glucose-stimulated insulin secretion of pancreatic ß-cell was investigated in an in vitro system (16), in which overexpression of SHP in islets by adenovirus enhanced secretion (16). However, the molecular basis for this observation is not clear. A potential role of SHP in regulating pancreatic function was further supported by the observation that SHP represses transactivation by neurogenic differentiation factor (NeuroD) (2) and forkhead box a 2 (Foxa2) (17), which both play important roles in regulating insulin secretion by pancreatic islets (18, 19, 20). This suggests that SHP may function as a novel negative regulator in controlling islet insulin secretion. Here, for the first time, we provide direct in vivo evidence to support this notion. We report that the loss of SHP function in SHP–/– mice results in a progressive decrease in circulating insulin levels that is associated with both an increased secretory response of islets to glucose and increased peripheral insulin sensitivity. We show that the increased insulin sensitivity is associated with increased expression of glucose-responsive genes in major insulin action tissues including muscle, fat, and liver. These results suggest that SHP is involved in diverse signaling pathways controlling islet insulin secretion and peripheral insulin sensitivity and that it may function as a negative modulator of type 2 diabetes.
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RESULTS |
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TOPABSTRACTINTRODUCTIONRESULTSDISCUSSIONMATERIALS AND METHODSREFERENCES |
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). However, fasting serum insulin levels were decreased by 50% in 6-month-old SHP–/– mice, and by 60% in 12-month-old mice lacking SHP (Fig. 1B). The adipokine leptin inhibits insulin biosynthesis and secretion in pancreatic ß-cells (21) and insulin stimulates leptin secretion from adipose tissue. This hormonal regulatory feedback loop, the "adipo-insular axis" (22), predicts that the hypoinsulinemia observed in SHP–/– mice should also result in reduced leptin secretion. As expected, serum leptin was decreased by 50% under nonfasted and fasted SHP–/– mice at 12 month of age, consistent with lower leptin mRNA in white fat (Fig. 1C). The hypoinsulinemia in SHP–/– mice could be due to increased insulin clearance, primarily by the liver, and/or decreased synthesis or secretion of insulin. To assess insulin clearance, we measured the ratio of the serum levels of insulin and C-peptide, which is produced in a 1:1 molar ratio with insulin but is cleared by a different mechanism than insulin (23, 24). SHP–/– mice showed a 65% higher C-peptide:insulin ratio than the SHP+/+ controls (Fig. 1D), indicating an increased rate of insulin clearance. We next assessed insulin synthesis by determining pancreatic insulin content and islet preproinsulin mRNA levels. SHP–/– mice had 30% less insulin than controls in acid-ethanol extracts of pancreas (Fig. 1E). In addition, the insulin mRNA in isolated islets from SHP–/– mice was much lower than in those from control mice under nonstimulatory conditions (3 mM glucose) (Fig. 1F). Although the decrease in basal insulin mRNA correlated with the decreased pancreatic insulin in SHP–/– mice, stimulation of insulin expression by incubating the islets in the presence of 25 mM glucose increased insulin mRNA in SHP–/– islets to a level similar to that of SHP+/+ mice, resulting in a greater glucose induced fold increase of insulin mRNA (2-fold in controls vs. 8-fold in SHP–/– mice). Thus, the islets of SHP–/– mice maintain a high capacity to synthesize insulin in response to increased glucose. This suggests that the reduction in pancreatic insulin and insulin mRNA in SHP–/– mice may be secondary to increased insulin sensitivity and a decreased demand for insulin to maintain glucose balance, rather than a decreased effect of SHP on insulin gene transcription.
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), demonstrating SHP expression in islets. Consistently, earlier reports also demonstrated SHP expression in pancreas by Northern blotting (7) and SHP expression in islet cells (2). In addition, analysis of pancreas sections using a cocktail of antibodies to non-ß-cell hormones showed no apparent differences in the size of islets at 2 and 6 months of age (data not shown). Assessment of ß-cell morphology was further carried out by immunofluorescence with insulin (red) and glucagon (green) antibodies specific in 12-month-old mice. Well-preserved architecture was observed with no apparent differences in the average size of most islets (Fig. 2, B and C). However, detailed quantitative analyses revealed that there were significant decreases in the number of islets per pancreatic section and islet area per pancreatic section (Fig. 2, D and E), but not in islet size (Fig. 2F). Because the pancreas weights were similar between the SHP–/– and SHP+/+ mice, these results suggest that the decrease in total islet number occurs due to long-term SHP deficiency, which may be associated with a decrease in total pancreatic insulin content (Fig. 1E
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, we predicted that the pancreatic islets of SHP–/– mice would show an increased insulin secretory response to glucose. Thus, we examined insulin secretion in cultured islets. Size-matched pancreatic islets were isolated from 12-month-old control and SHP–/– mice, calibrated overnight, and were perfused with varying concentrations of glucose the next day. No significant differences in basal insulin secretion were observed between the two genotypes. Insulin secretion revealed significant differences in response to high glucose concentrations or potassium chloride (KCl). In the first-phase perfusion, islet insulin secretion was 3-fold higher upon 16.7 mM glucose stimulation in SHP–/– islets than in SHP+/+ islets (Fig. 2G). A similar elevated response was observed upon two successive restimulations with 16.7 mM glucose and KCl (Fig. 2H). The results suggest that there is no impaired coupling of stimulus and secretion in SHP–/– ß-cells. In contrast, the increase in glucose-induced insulin secretion in SHP–/– islets suggests that the islets of SHP–/– mice maintain a high capacity to secrete insulin, consistent with the marked up-regulation of insulin mRNA by glucose (Fig. 1F). On the other hand, the insulin content appeared to be similar in size-matched SHP–/– and SHP+/+ islets (data not shown), suggesting normal insulin synthesis and stores. Thus, the decreased basal insulin mRNA (Fig. 1F) is suggestive of a secondary effect of a decreased need for insulin that may result from increased insulin sensitivity.
SHP–/– Mice Have Increased Insulin Sensitivity
The results described above suggest that the progressive hypoinsulinemia in aged SHP–/– mice is not due to decreased islet function, but rather the result of increased insulin sensitivity. Glucose and insulin tolerance tests were first performed and we observed increased glucose tolerance and insulin sensitivity in 12-month-old SHP–/– mice (data not shown). Next, the hyperinsulinemic euglycemic clamp was used to more critically test this. Basal glucose production was comparable between the SHP+/+ and SHP–/– mice. Low-dosage insulin infusion decreased glucose production more profoundly in SHP–/– mice than in SHP+/+ mice (6.48 ± 2.65 vs. 11.54 ± 2.82 mg·kg–1·min–1, respectively; P = 0.02). The rate of glucose infusion necessary to maintain euglycemia during low dose insulin infusion was nearly twice as high in SHP–/– as in SHP+/+ mice (Fig. 3A), and slightly more than 2-fold higher in the high-dose clamp (Fig. 3B). Under basal conditions, glucose disposal rate was similar in SHP+/+ and SHP–/– vehicle-infused mice. Hyperinsulinemia increased glucose disposal in both groups, but the disposal rate was elevated in SHP–/– animals compared with SHP+/+ controls under both low-dose (Fig. 3C) and high-dosage insulin infusion (Fig. 3D). In addition, insulin-mediated whole-body glucose uptake was greater in both low- and high-dosage insulin-infused SHP–/– animals than controls (Fig. 3, E and F), which reached statistical significance (P = 0.01). These results demonstrate that the loss of SHP function results in an increase in peripheral insulin sensitivity.
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). Consistent with the up-regulation of Glut4 mRNA, the glucose uptake in soleus muscles was also markedly increased in the same aged SHP–/– mice (Fig. 4B). To further determine the cell-autonomous effect of SHP on Glut4, an in vitro cell system was used with murine muscle-derived C2C12 cells, because C2C12 cells have been widely used to study glucose uptake (26), even though basal Glut4 level is relatively low in this cell line. Overexpressing PGC-1 markedly induced Glut4 mRNA, whereas overexpressing SHP decreased it by 90% (Fig. 4C). When PGC-1 and SHP were coexpressed, SHP mRNA was up-regulated by approximately 50%, whereas PGC-1 mRNA was down-regulated by approximately 50% (Fig. 4C). This suggests a dual regulatory loop, in which PGC-1 activates SHP, and SHP in turn inhibits PGC-1 expression. The more sensitive semiquantitative RT-PCR analysis further revealed direct repression of Glut4 mRNA by SHP (Fig. 4C). Consistently, the basal and insulin-stimulated glucose uptake in C2C12 cells was significantly increased by PGC-1, which was further inhibited by SHP (Fig. 4D). These results demonstrate that SHP represses basal and PGC-1-stimulated glucose uptake in muscle by inhibiting Glut4. However, it does not rule out the possibility for SHP repression of basal glucose uptake through other glucose transporter.
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1 (PPAR1), which is elevated in fatty liver (27) and in livers of obese ob/ob and db/db mice (28), was completely diminished in SHP–/– mice (Fig. 5A). This was evident with no basal expression in 2-month-old livers, and was more striking in 6- and 12-month-old mice and high-sucrose (HS)-fed mice (15). Consistent with this, the fatty acid translocase CD36, a PPAR target, was also decreased in SHP–/– livers (Fig. 5A). These results are consistent with recent studies in which overexpressing the human SHP gene markedly increased PPAR and CD36 mRNAs in mouse liver (29). Consistent with the role of insulin in inhibiting hepatic gluconeogenesis and stimulating glycogen synthesis, mRNAs for the gluconeogenic enzymes glucose-6-phosphatase (G6P) and phosphoenolpyruvate carboxykinase (PEPCK) were decreased, whereas mRNAs of glycolytic enzymes glucokinase (GK) and pyruvate kinase (PK) were increased (Fig. 5B), concomitant with increased total glycogen stores in SHP–/– livers of mice fed chow diet, as assayed by both hepatic periodic acid-Schiff (PAS) staining and quantitative analysis of liver glycogen content (Fig. 5C).
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). At low dose of insulin (0.5 ng ml–1), hepatic glucose production was significantly increased in SHP+/+ mice, but not in SHP–/– mice. The effect of SHP on gluconeogenesis was further examined in cultured hepatocytes. Basal glucose production was markedly lower in SHP-deficient cells (Fig. 5E). The addition of insulin significantly decreased glucose output in SHP+/+ cells to a level comparable to SHP–/– cells but did not further reduce glucose production in SHP–/– cells, suggesting a maximal effect of the loss of SHP. Taken together, the results suggest increased sensitivity of SHP–/– livers to insulin inhibition of gluconeogenesis.
SHP–/– Mice Show Increased Expression of Insulin-Sensitizing Genes in White Adipocyte
PPAR2 plays a crucial role in adipogenesis and insulin sensitization in adipose tissue (31). In striking contrast to the elimination of PPAR1 expression in liver (Fig. 5A), a robust increase in PPAR2 expression was observed in SHP–/– white fat (Fig. 6A), as were lipoprotein lipase (LPL) (32), long-chain acyl-CoA dehydrogenase (LCAD) (33), hormone-sensitive lipase (HSL), perilipin (34), and stearoyl-CoA desaturase-1 (SCD-1) (35). Surprisingly, CD36 was not altered in SHP–/– white fat (Fig. 6A), in contrast to its decreased expression in liver (Fig. 5A). No changes for resistin and aP2 (data not shown) mRNA were observed.
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target (36). The potential interaction of SHP with PPAR and LRH-1 on ADP promoter activity was tested by transient transfection (Fig. 6B). Coexpression of PPAR/RXR increased ADP promoter activity (lane 2), which was significantly augmented by cotransfection of LRH-1 (lane 3). Coexpression of SHP dose-dependently repressed the activation of the ADP promoter by PPAR/RXR/LRH-1 (lanes 4–6). Moreover, the inhibitory effect of SHP on PPAR/RXR transactivation of the ADP promoter by rosiglitazone (Rosi) was more profound in the presence of LRH-1 (lanes 4–6 vs. 7). These results demonstrate that SHP directly represses the ADP promoter activity by PPAR/RXR/LRH-1. We further performed chromatin immunoprecipitation (ChIP) assay with specific SHP antibody in 3T3-L1 cells and SHP was recruited to the ADP promoter (Fig. 6B). To determine the in vivo physiological relevance for SHP regulation, ADP mRNA in white fat was examined and we observed only marginal elevation of ADP in SHP–/– 6-month-old mice, but a more clear elevation at 12 months of age (Fig. 6C). Similar results were observed when serum levels of ADP were measured by Western blots. This result was somewhat unexpected based on the marked increase of PPAR in SHP–/– fat of 6-month-old mice (Fig. 6A) and suggests that factors other than PPAR may play a more dominant role on ADP expression in young SHP–/– mice. Nevertheless, the increased ADP mRNA in 12-month-old SHP–/– mice may at least in part account for the increased insulin sensitivity. |
DISCUSSION |
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The results with isolated SHP–/– islets is in contrast with a recent report in which overexpression of SHP by adenovirus transduction enhanced glucose-stimulated insulin secretion in INS-1 cells, especially those overexpressing the mitochondrial uncoupling protein UCP-2 (16). It should be noted that the acute overexpression of SHP in a cultured cell line is very different from the lifelong loss of function in the whole animal, and it is not clear that these contrasting results are in conflict. Several possible mechanisms may explain why increased insulin secretion was observed after loss of SHP function in mice. SHP was shown to repress the transactivation of several important regulators in islet insulin secretion, including HNF4 (5), NeuroD (2), and Foxa2 (17). The loss of SHP function should lead to their increased activity, resulting in increased insulin secretion. In addition, our recent study demonstrates that SHP acts as a corepressor for BETA2 by competing with coactivator p300 for binding to BETA2 on insulin promoter (Park, K. G., K. M. Lee, D. D. Moore, L. Wang, K. C. Won, J. Y. Park, K. U. Lee, H. S. Choi, I. K. Lee, submitted for publication). Collectively, the results reported indicate that multiple regulatory pathways exist for controlling pancreatic islet insulin secretion, and that SHP can play a major role in regulating these pathways by altering transcription of critical genes. It would be of great interest to explore these possible regulatory mechanisms in future in vivo studies.
Our gene expression studies identified several critical genes in skeletal muscle, liver, and fat that may contribute to the increased insulin sensitivity seen in SHP–/– mice (Fig. 7). Increased skeletal muscle mRNA for Glut4 contributes to the increased muscle glucose uptake in SHP–/– mice, because SHP represses Glut4 and its associated glucose uptake. The activation of Glut4 expression and glucose uptake by PGC-1, and a potential dual regulatory loop between PGC-1 and SHP, represent a new hypothesis regarding mechanisms for SHP regulation of muscle insulin sensitivity. Detailed transcriptional analysis will be required to characterize these regulatory mechanisms. Another striking observation is the diminished hepatic PPAR1 in SHP–/– mice, which provides an explanation for the lower lipid content in SHP–/– livers (8, 15) that will result in improved hepatic insulin sensitivity (28). This result is in agreement with the up-regulation of PPAR1 in SHP-overexpressed liver (29). On the other hand, the strong up-regulation of GK and down-regulation of G6P and PEPCK is consistent with the decreased gluconeogenesis in SHP–/– livers and in cultured hepatocytes. However, this result is in contrast to the reported inhibitory role of SHP on G6P and PEPCK by bile acids (38). Thus, decreased expression of G6P and PEPCK in SHP–/– livers may occur as a secondary effect of other metabolic changes.
Similarly, increased white adipose tissue expression of PPAR2 is consistent with increased insulin sensitivity, because adipose-specific PPAR–/– mice exhibit insulin resistance in fat (39). The induction of both LPL/SCD-1 and HSL/perilipin in adipose tissue of SHP–/– mice is in agreement with the role of PPAR in promoting both lipid uptake (LPL) and triglyceride synthesis (SCD-1), as well as lipolysis (HSL, perilipin), and increases in expression of these genes are associated with increased insulin sensitivity (34, 35, 40). Surprisingly, no increase in CD36 mRNA was observed, indicating fatty acid uptake may not be increased in SHP–/– fat, because we do not know whether fatty acid transporter protein is up-regulated to increase fatty acid uptake (34). No increase in ADP expression in young SHP–/– mice is unexpected and suggest that ADP expression in young SHP–/– mice is regulated by factors other than PPAR. In addition, no changes of ADP receptors (26) are observed in both liver and muscle of SHP–/– mice (data not shown), suggesting that the increased insulin sensitivity is not due to a secondary effect through ADP receptors. The marked improvement in hepatic function and obese phenotype in SHP–/– mice becomes more evident when the mice are older or challenged with an excess fat load (8, 15), which may explain the lack of apparent glucose or insulin metabolism phenotypes in 2-month-old SHP–/– mice. We anticipate that a more pronounced phenotype could be obtained if the mice are subjected to an appropriate stress, such as streptozotocin, to induce diabetes. It is also possible that chronic and progressive alterations in multiple overlapping systems contribute to the emergence of the improved insulin sensitivity in SHP–/– mice. Moreover, age-dependent changes in glucose homeostasis are also seen in other mouse models (41), and in SH-2B–/– mice (42) and Pten–/– mice (43); thus, it is not surprising to see them in SHP–/– mice. Lastly, the striking differential expression pattern for PPAR2 in white fat and PPAR1 in liver of SHP–/– mice is extremely intriguing, which suggests a differential regulation by SHP in cell- and tissue context-dependent manner. Future studies will be required to provide more mechanistic insights into SHP regulation.
In conclusion, our results identify a multifunctional and apparent inhibitory role of SHP in pancreas, muscle, liver, and fat in the regulation of glucose homeostasis and whole-body insulin sensitivity. These are complex processes, but the transcriptional regulatory consequences of the loss of SHP provides a firm ground for detailed studies regarding the mechanisms for changes in insulin sensitivity. By using tissue-specific loss or gain of SHP mouse models, we should be able to elucidate tissue-specific functions of SHP in glucose homeostasis and their impact on the dysregulation of these processes in type 2 diabetes.
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MATERIALS AND METHODS |
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Serum Chemistry
Blood glucose concentration was determined with a Hemocue blood glucose analyzer. Serum glucose was determined using enzymatic assay (Sigma-Aldrich, St. Louis, MO). Serum glucagon, leptin, and C-peptide were determined by RIA using each RIA kit from Linco Research (St. Charles, MO). Serum insulin was measured using ELISA kit (Crystal Chem, Chicago, IL) or RIA (Linco). Pancreas insulin content was measured by RIA after acid/ethanol extraction from homogenized whole pancreas that had been dissected, removed from the animal, and weighed.
In Vivo Glucose Kinetics
Hyperinsulinemic euglycemic clamp study was performed using chronically cannulated, conscious mice as described (44). All groups of mice were fasted 6–8 h before the experimentation. Body weight, hematocrit, and general appearance were used as indices of health. A variable infusion of 50% glucose was used to raise blood glucose levels to approximately 300 mg/dl. A 4-µCi bolus of tracer ([3-3H]glucose; NEN Life Science Products, Boston, MA) was given at –100 min, followed by a constant 0.04 µCi/min for the duration of the study. The glucose turnover rate (in milligrams/kilogram·minute) was calculated as the rate of tracer infusion (in disintegrations per minute/minute) divided by the blood glucose specific activity (in disintegrations per minute/milligram) corrected to the body weight of the mouse. Glycogen synthesis via the direct pathway was determined by measuring incorporation of 3H into glycogen (in disintegrations per minute/gram of liver)/3H specific activity in plasma glucose (in disintegrations per minute/milligram).
Islet Isolation, Culture, and Insulin Secretion
The isolation of islets was described elsewhere (45). Briefly, pancreatic islets were isolated by collagenase P digestion (Roche, Basel, Switzerland), hand-picked, and transferred to DMEM supplemented with 10% FBS, glucose, glutamine, sodium pyruvate, and antibiotics. Freshly isolated islets were cultured overnight at 37 C, size-matched, divided into groups, and used for perfusion experiments (200 islets). In perfusion experiments, Krebs-Ringer bicarbonate buffer was pumped at a rate of 1 ml/min around islets that were loaded into temperature- and CO2-controlled 300-µl plastic chambers. Islets were washed under basal conditions for 1 h before the experiment. The amount of insulin released into the medium was determined by ELISA. For islet insulin content assay, isolated islets were suspended in 100 µl of acid ethanol, and cellular insulin was extracted and assayed by RIA.
Islet Staining
Generation of the SHP–/– mice was described elsewhere (8). In brief, the first exon of the SHP gene was disrupted by homologous recombination with a LacZ-neo cassette. This insertion provides an endogenous marker of SHP expression by quantification of LacZ expression. Because LacZ is not expressed in tissues of SHP+/+ mice, the background signal is low and therefore is very sensitive to defining SHP expression. Pancreas harvested from animals were infused with paraformaldehyde, isolated, and frozen. The tissue was cryosectioned (5-µm sections were acquired). Pancreatic sections were stained for ß-galactosidase activity using standard techniques. For islet insulin and glucagon staining, immunohistochemistry was performed following standard procedures as described (46). Briefly, pancreas was immersion-fixed in 4.0% (wt/vol) paraformaldehyde, 0.1 M sodium phosphate buffer at 4 C overnight. Diluted guinea pig antiporcine insulin (A564; DakoCytomation, Carpinteria, CA) (1:200), or rabbit antiporcine glucagon (A565; DakoCytomation) (1:200), or rabbit antihuman somatostatin (A566; DakoCytomation) (1:200) was applied to the sections for 45 min at room temperature. The sections were then rinsed with Tris-buffered saline, and then treated with a second antibody followed by visualization using a confocal microscope.
Point Count for Quantification of Islet
The relative area of islet in the pancreas was counted by the point counting method using a microscope (Carl Zeiss, Oberkochen, Germany) connected to a personal computer monitor with a 90-point transparent overlay. In brief, immunostained pancreatic sections were visualized under x200 magnification and positioned a regular lattice overlay on personal computer monitor. The islets were counted simultaneously in insulin-stained slides. Average of 104 fields and 14,210 points in nonoverlapping fields was counted from each tissue; three sections were counted in each tissue block. The number of islets per each field was recorded and the area of each islet was measured with NIH Image analysis program equipped with WACOM Intuos Tablet System (Wacom, Saitama, Japan). Relative area of islet in pancreatic tissue was represented as follows: number of points corresponding to insulin antibody-stained area/number of points corresponding to remaining pancreatic area (pancreatic area). Average size of islet and islet appearance rate were represented as follows: summation of total area of islet/islet number and total number of islet/pancreatic area, respectively.
Adenoviral Transduction
Recombinant adenoviruses for expression of PGC-1, SHP, and the GFP control were described (15). Cells were plated at 2 x 106 per 10-cm dish, cultured to 70–80% confluence, and infected the next day with viral supernatant at different multiplicities of infection for 2 h. Virus-containing media were removed and cells were continuously cultured for 2 d. Gene expression was analyzed by Northern blotting.
Glucose Uptake Assay
C2C12 mouse muscle myoblasts (American Type Culture Collection, Manassas, VA) were grown in six-well plates to confluence in DMEM with 10% FBS and antibiotics. Cells were differentiated for 1 d in DMEM with 2% horse serum, and then were infected with adenoviruses for GFP, PGC-1, and/or SHP for 2 h. The cells were continuously differentiated for 2 d to myotubes. For the last 5 h on the fourth day, the cells were serum-starved in serum-free DMEM with 25 mM glucose and 0.2% BSA, and insulin was added at 100 nM for the last 20 min. Cells were then rinsed twice in Krebs-Ringer-phosphate (KRPH) buffer, and glucose uptake was assessed with 100 µM 2-deoxy-D-glucose (1 µCi/ml [1,2-3H]-2-deoxy-D-glucose) with or without 10 µM cytochalasin B in KRPH for 10 min at room temperature. Uptake was terminated by adding cold stop medium (50 mM [SCAP];d-glucose in KRPH). Cells were then washed, and radioactivity associated with the cells was determined by cell lysis in NaOH/sodium dodecyl sulfate, followed by scintillation counting. Aliquots of cell lysates were used for protein content determination. 2-Deoxy-D-[1,2-3H]glucose (2-DG) uptake was expressed as picomoles per minute per milligram of protein. For insulin-stimulated glucose uptake in muscles, soleus muscles were isolated and incubated for 30 min in Krebs-Henseleit buffer, with or without insulin (100 nM). The muscles were then rinsed for 10 min at 29 C and incubated for 20 min at 29 C in Krebs-Henseleit buffer containing 8 mM 2-DG (2.25 µCi/ml) and 32 mM [14C]mannitol (0.3 µCi/ml). After incubation, the muscles were rapidly solubilized. Radioactivity in the resultant samples was counted, and 2-DG uptake rates were corrected for extracellular trapping with mannitol counts. The protein concentration was determined by the Bradford method.
Liver Glycogen Staining and Content
Briefly, livers were fixed in 4% formaldehyde, dehydrated, embedded in paraffin, and sectioned (9). Sections were stained with PAS to examine glycogen content in liver. For determination of glycogen, 50-mg fragments of liver were homogenized in KOH and incubated at 97 C for 15 min. The glycogen was precipitated by Na2SO4 and EtOH at –20 C for 16 h. After centrifugation, the pellet was collected and washed with KOH and EtOH and hydrolyzed in H2SO4 at 100 C for 1 h. The solution was neutralized, and glucose was determined by the glucose oxidase method.
Hepatocyte Culture and Glucose Production
Hepatocytes were isolated (8) with cell viability assessed by the trypan blue exclusion at greater than 80%. Cells were serum-starved overnight and stimulated with insulin (100 nM) for 24 h. Cells were washed with DMEM and incubated with glucose production medium (without glucose) containing 20 mM lactate and 2 mM pyruvate for 3 h. Culture medium was collected and centrifuged, and glucose levels were determined (Sigma-Aldrich), normalized to protein content of the cells.
To measure glucose production in vivo, the liver perfusion was performed on starved SHP+/+ and SHP–/– mice (30). After anesthesia with pentobarbitone sodium (60 mg/kg of body weight, ip), the portal vein and the inferior vena cava were cannulated. The liver was perfused with oxygenated Krebs-Henseleit buffer with varying amounts of insulin at 37 C in a single-pass mode with a total flow rate of 1.5–2 ml min–1. The outflow was collected, and glucose concentrations were measured. Glucose output was calculated by subtracting the amount of glucose contained in the perfusion buffer from that measured in the outflow.
Transient Transfection and ChIP Assay
NIH 3T3 cells and HeLa cells were maintained in DMEM with 10% FBS. For luciferase assays, cells were plated in 24-well plates 1 d before transfection and transfections were carried out using calcium phosphate (15). ADP promoter Luc was cotransfected with PPAR (P; 25 ng), LRH-1 (L; 100 ng), and/or SHP (S; 100, 200, 300 ng) expression vectors, respectively. Forty-eight hours posttransfection, cells were harvested, and luciferase activity was measured and normalized against ß-galactosidase activity as an internal control with a Dual-Luciferase Reporter System (Promega, Madison, WI). The transfection experiments were carried out independently three times with similar efficiency. For ChIP assay, 3T3-L1 cells were used and chromatin cross-linked, immunoprecipitated with anti-SHP antibodies (29), with IgG as negative control.
Northern and Western Blotting
Northern blotting and semiquantitative PCR analysis were performed as previously described (9). The primer sequences of each gene were from the PubMed GenBank database and were designed and used to synthesize various probes for Northern blot analysis. Primer sequences are available upon request. Western blotting was followed by standard procedures.
Statistical Analysis
Data are expressed as mean ± SD. Statistical analyses were carried out using Student’s unpaired t test; P < 0.01 was considered statistically significant.
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ACKNOWLEDGMENTS |
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FOOTNOTES |
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L.W., J.H., P.S., R.N.K., M.H., Y.K., K.P., L.C., A.S.R., I.L., D.D.M. have nothing to declare.
First Published Online June 27, 2006
1 P.S., R.N.K., M.H., and Y.K. contributed equally to this work. 
Abbreviations: ADP, Adiponectin; CD36, fatty acid translocase; ChIP, chromatin immunoprecipitation; 2-DG, 2-deoxy-D-[1,2-3H]glucose; Foxa2, forkhead box a 2; GK, glucokinase; Glut4, glucose transporter 4; G6P, glucose-6-phosphatase; HNF4, hepatocyte nuclear factor-4; HS, high sucrose; HSL, hormone-sensitive lipase; KRPH, Krebs-Ringer-phosphate; LCAD, long-chain acyl-CoA dehydrogenase; LPL, lipoprotein lipase; LRH-1, liver receptor homolog-1; NeuroD, neurogenic differentiation factor; PAS, periodic acid-Schiff; PEPCK, phosphoenol/pyruvate carboxykinase; PGC-1, peroxisomal proliferator-activated receptor coactivator-1; PPAR, peroxisomal proliferator-activated receptor ; RXR, retinoid X receptor ; SCD-1, stearoyl-CoA desaturase-1; SHP, small heterodimer partner.
Received for publication May 24, 2006. Accepted for publication June 23, 2006.
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3-fold) in SHP–/– islets (red lines) than of SHP+/+ islets (black lines). Each time point represented an average of four groups (200 islets/group, each group from 15–20 mice). Values are means ± SEM. *, P < 0.01.
