Increased Insulin Sensitivity and Hypoinsulinemia in APS Knockout Mice

The generation of APS-knockout (APS^−/−) mice has been described elsewhere (M.I., C.K., S.-M.K., A.Y., Y.K., N.Y., K.T., S.T., unpublished observations). Briefly, we disrupted by homologous recombination coding exons 1–6, which contained the translation initiation codon. The non-APS-knockout littermates (APS^+/+) served as age-matched control subjects for the APS^−/− mice. The mouse strain used for crossbreeding was C57BL/6. All the mice used in this study were F2 and F3 siblings. Body weight was measured at weekly intervals through the experimental period.

Mouse APS wild-type cDNA was generated as described (5). The tyrosine 618 site at the COOH-terminus of mouse APS was mutated to phenylalanine (Y618F mutant; YF) as described (5). Recombinant mouse APS wild-type and Y618F mutant adenoviral constructs were generated using adenovirus expression systems (21).

Polyclonal anti-APS NH₂-terminus, anti-APS COOH-terminus, and anti-APS antibodies were obtained by immunizing rabbits with a glutathione S-transferase-fused NH₂-terminal (1–102 amino acids) (10), COOH-terminal (534–621 amino acids) (10), and COOH-terminal (508–591 amino acids) (5) mouse APS fragments. A monoclonal anti-phosphotyrosine antibody 4G10 was purchased from Upstate Biotechnology (Lake Placid, NY). Polyclonal antibodies against insulin receptor-β (sc-711) and Cbl (sc-170) were purchased from Santa Cruz Biotechnology (Santa Cruz, CA). Polyclonal antibodies against phosphospecific Akt (Ser473) and phosphospecific mitogen-activated protein kinase (Thr180/Tyr182) were obtained from Cell Signaling (Beverly, MA). Anti-insulin receptor C40 and anti-IRS-1 antibodies were prepared as described (22). Secondary horseradish peroxidase-conjugated goat anti-mouse IgG and anti-rabbit IgG were obtained from BioSource (Camarillo, CA). All other reagents used were of analytical grade and were purchased from Wako (Osaka, Japan) or Nacalai (Kyoto, Japan).

Various tissues of the mice were stored in liquid nitrogen, and total RNA was extracted from these tissues using a guanidine-isothiocyatate preparation (Trizol Reagent) (Invitrogen, Carlsbad, CA). Poly (A)⁺ mRNA was purified using Oligotex-dT30 Super (Takara, Shiga, Japan) and separated on 1.0% agarose gel and transferred to nitrocellulose membrane (Schleicher & Schuell, Dassel, Germany). The membranes were baked and hybridized with ³²P-labeled APS cDNA probes using BcaBest labeling kits (Takara).

Various organs preserved in liquid nitrogen were homogenized in a motor-driven polytron PT10–35 (Kinematica, Switzerland) in ice-cold lysis buffer containing 25 mmol/l Tris-HCl (pH 7.4), 10 mmol/l Na₄P₂O₇, 100 mmol/l NaF, 10 mmol/l EDTA, 10 mmol/l EGTA, 1% Nonidet P-40, 1 mmol/l Na₃Vo₄ and 1 mmol/l phenylmethanesulfonyl fluoride. Culture cells were incubated in Krebs-Ringer-Hepes buffer (KRH) (23) for 20 min at 37°C, stimulated with or without 100 nmol/l insulin for 5 min at 37°C, and lysed in ice-cold lysis buffer containing 20 mmol/l Tris-HCl (pH 8.0), 140 mmol/l NaCl, 1 mmol/l MgCl₂, 1 mmol/l CaCl₂, 1 mmol/l dithiothreitol, 10% glycerol, 0.02 mmol/l p-amidinophenylmethanesulfonyl fluoride, 1% Nonidet P-40, 0.5 mmol/l Na₃VO₄ and 20 mmol/l Na₄P₂O₇. After centrifugation at 15,000 rpm at 4°C for 20 min, the supernatant was measured using Bio-Rad protein assay kits (Bio-Rad Laboratories, Richmond, CA). The total lysate was subjected to immunoprecipitation and immunoblotting using the indicated antibodies.

At age 8–9 weeks, the mice underwent an intraperitoneal insulin tolerance test (IPITT) or intraperitoneal glucose tolerance test (IPGTT) after a 14-h (1900–0900) fast. Insulin (Novolin R; Novo Nordisk, Bagsvared, Denmark), 0.75 units/kg body wt, or glucose, 2 g/kg body wt in normal saline (0.9% NaCl), was administered intraperitoneally. Blood samples were taken from the tail vein at 0, 15, 30, 60, and 120 min after loading, and plasma glucose levels were measured. Blood samples were also taken at 0, 15, and 30 min for measurement of plasma insulin levels during IPGTT.

Plasma glucose levels were determined using the glucose oxidation method (Glucocard; Aventis Pharma, Frankfurt, Germany). Insulin levels were determined using enzyme-linked immunosorbent assay kits (Morinaga Institute of Biological Science, Tokyo, Japan) with mouse insulin as standard.

Insulin-mediated whole-body glucose uptake was determined in anesthetized mice aged 9–11 weeks using euglycemic glucose clamp. After a 6-h fast (0700–1300), the mice were anesthetized by giving an intraperitoneal administration of sodium pentobarbital (50 mg/kg body wt; Nembutal injection; Dainippon Pharmaceutical, Osaka, Japan), and catheters (Sp8 polyethylene tube; Natsume, Tokyo, Japan) were inserted in jugular veins. The mice were given a constant 2-h infusion of insulin (60 pmol · kg body wt⁻¹ · min⁻¹). A glucose solution (100 g/l) was initiated at the time when blood glucose level was <100 mg/dl, and the rate of infusion was adjusted to maintain the plasma concentration of glucose at ∼110 mg/dl. The whole-body glucose uptake represents the glucose infusion rate (GIR) during the last 20 min.

After a 14-h fast (1900–0900), blood samples were collected from the inferior vena cava for biochemical analysis. Plasma levels of total cholesterol, triglycerides, and free fatty acids were determined using conventional enzymatic methods (Wako). Leptin levels were determined using enzyme-linked immunosorbent assay kits (Morinaga) with mouse leptin as standard. Adiponectin and glucagon levels were determined using radioimmunoassay kits (Linco, Charles, MO) with recombinant adiponectin or glucagon as standard.

The 3T3L1-G4myc-CARΔ1 were 3T3L1 cells stably expressing myc-tagged GLUT4 (GLUT4myc) (23) and CARΔ1 (deleted cytoplasmic domain of CAR; cellular receptor for adenovirus and coxsackievirus [24]), which increases the efficiency of adenovirus infection to 3T3L1 adipocytes. Differentiation to adipocytes was induced as described elsewhere (23). 3T3L1-G4myc-CARΔ1 adipocytes were exposed to an adenovirus encoding a green fluorescent protein (GFP), APS wild-type, or Y618F mutant (multiplicity of infection [MOI] 10–20 pfu/cell) and used for experiments after 36 h.

3T3L1-G4myc-CARΔ1 adipocytes overexpressing APS wild-type or Y618F mutant were incubated in KRH for 20 min or in serum-free medium containing 10 mmol/l 2-deoxyglucose for 4 h at 37°C and then treated with or without 100 nmol/l insulin for 10 min at 37°C. 2-Deoxyglucose uptake was measured by incubating cells with 2-[³H]deoxy-d-glucose for 10 min, as described (23).

3T3L1-G4myc-CARΔ1 adipocytes overexpressing APS wild-type or Y618F mutant were incubated in KRH for 20 min at 37°C and and then stimulated with or without 100 nmol/l insulin for 10 min at 37°C. GLUT4myc translocation was measured after fixation with 2% paraformaldehyde, as described elsewhere (23,25).

Adipocytes were isolated from epididymal fat pads by collagenase digestion, as described with some modification (26). The number of isolated adipocytes was adjusted to 10⁶ cells/ml suspension and 2-deoxyglucose uptake was measured, as described (27).

The soleus muscle was separated from the hindlimb, and glucose transport activity was assessed by measuring of 2-deoxyglucose uptake, as described (28,29).

Tissue glycogen was measured by the methods of Abraham et al. (30) and Hassid (31).

Data are expressed as means ± SD or SE. Data were analyzed by ANOVA plus Bonferroni multiple-comparison tests for determination of the significance of the differences. A P value <0.05 was considered statistically significant.

APS^−/− mice were generated (M.I., C.K., S.-M.K., A.Y., N.Y., K.T., S.T., unpublished data). First, we examined APS mRNA expression in various tissues. APS mRNA in normal mice was strongly expressed in adipose tissue, an insulin target organ, and detected in skeletal muscle, spleen, brain, lung, liver, and kidney (Fig. 1A). These data are almost consistent with some previous reports (5,7,10). We verified complete deletion of APS mRNA in both skeletal muscle and adipose tissue of APS^−/− mice, although there was a smaller size of transcript (1.4 kb) only in adipose tissue of APS^−/− mice (Fig. 1B). The small band of the transcript did not express as a protein (see discussion). The expression of APS protein in liver, skeletal muscle, and adipose tissue was identified by Western blot analysis only in APS^+/+ mice (Fig. 1C). The expression levels of APS protein were high in adipose tissue and low in liver and skeletal muscle. In APS^−/− mice, the expression of APS was not detected in these tissues. Thus, complete deletion of intact APS mRNA and protein was verified by Northern analysis and by the immunoblotting of tissue extracts (Fig. 1B and C).

To examine effects of APS deficiency on insulin actions in APS^−/− mice, we did insulin and glucose tolerance tests in vivo.

In the IPITT, blood glucose levels after fasting did not differ between APS^+/+ and APS^−/− mice. APS^−/− mice had significantly lower blood glucose levels at 30, 60, and 120 min after insulin injection compared with findings in APS^+/+ mice (Fig. 2A). The blood glucose-lowering effect of insulin was greater in the APS^−/− mice. Mice underwent an IPGTT to determine the plasma insulin and blood glucose levels after glucose loading. The blood glucose levels were slightly lower in APS^−/− mice than in APS^+/+ mice during IPGTT and at 15 min after glucose injection; the difference in blood glucose levels was statistically significant (Fig. 2B). Blood glucose levels in IPITT and IPGTT were not statistically different between APS^+/+ and APS^+/− mice (heterozygous) (data not shown). Fasted plasma insulin levels and the plasma insulin levels during IPGTT were significantly lower in APS^−/− mice than in APS^+/+ mice (Fig. 2C). These data indicated that a lower insulin level results in a normal concentration of blood glucose in the APS^−/− mice. To confirm the increased insulin sensitivity in APS^−/− mice, we did hyperinsulinemic-euglycemic clamp studies. The glucose infusion rate was significantly elevated in APS^−/− mice compared with findings in APS^+/+ mice (Fig. 2D). The clamp studies supported the phenotype of APS^−/− mice. All these data showed that APS^−/− mice have an increased insulin sensitivity and hypoinsulinemia.

To determine the physiological effects of APS knockout, we examined body weight, food intake, epididymal fat weight, and metabolic parameters in the blood. APS^+/+ and APS^−/− mice had similar transition curves regarding body weight and identical food consumption and epididymal fat weight (Table 1). APS^−/− mice had a significantly lower fasted insulin concentration compared with data on APS^+/+ mice (Fig. 2C and Table 1). Glucagon levels showed slightly higher levels, but there were no statistically significant differences (Table 1). Triglyceride and total cholesterol levels were somewhat lower in APS^−/− mice, and free fatty acid levels were slightly higher, albeit with no significant difference between the two groups. Therefore, the increased insulin sensitivity is not caused by low glucagon levels or improvements in lipid metabolism.

As shown in Fig. 2, APS^−/− mice have an increased insulin sensitivity to blood glucose-lowering effects. Since Ahmed and colleagues (15,17) reported that APS with Cbl facilitates insulin-induced insulin receptor ubiquitination, lack of APS may lead to increased insulin receptor numbers on the cell surface, and consequently there is an increase insulin sensitivity. Insulin receptor contents in liver, skeletal muscle, heart, brain, and adipose tissue in APS^−/− and APS^+/+ mice were examined by immunoblotting, but no differences in quantities were found (Fig. 3A). Although insulin receptor contents were not altered, the turnover of insulin receptor might possibly be augmented by APS. To examine the effect of APS on turnover of insulin receptor, we prepared cells overexpressing APS wild-type or Y618F mutant in which tyrosine 618 (insulin-induced tyrosine phosphorylation site) was mutated to phenylalanine. APS Y618F mutant acted as a dominant-negative form, which prevented tyrosine phosphorylation of normal APS by insulin stimulation and prevented the interaction with insulin receptor (5).

We prepared 3T3L1-G4myc-CARΔ1 adipocytes overexpressing APS wild-type and Y618F mutant. We detected the rapid tyrosine phosphorylation of APS after insulin stimulation in 3T3L1-G4myc-CARΔ1 adipocytes overexpressing APS wild-type (Fig. 3B). In 3T3L1-G4myc-CARΔ1 adipocytes expressing GFP by adenovirus as a control, tyrosine phosphorylation of endogenous APS was detected (Fig. 3B). Overexpression of APS wild-type but not Y618F mutant resulted in interaction between APS with Cbl by insulin stimulation (Fig. 3B). Tyrosine phosphorylation of c-Cbl by insulin stimulation was increased in cells overexpressed APS wild-type, but decreased by overexpression of APS Y618F mutant. Using these cells, we examined insulin receptor levels in immunoblotting and cell-surface insulin receptor binding assays. Overexpression of APS wild-type or Y618F mutant did not affect insulin receptor β contents (Fig. 3B) and cell-surface insulin receptor binding (data not shown). Therefore, APS did not affect insulin receptor contents in the APS^−/− mice and cultured cells (see discussion).

We examined the effects of APS on the internalization and turnover of insulin receptor in CHO cells overexpressing insulin receptor (32). The internalization and degradation rate of insulin receptor were not changed in CHO cells overexpressing insulin receptors and APS wild-type or Y618F mutant (data not shown).

Another possibility is that APS might affect cellular insulin signaling and glucose transport. We examined the effects of exogenously overexpressed APS wild-type or Y618F mutant on insulin-stimulated glucose uptake and GLUT4 translocation. Transient expression of APS wild-type or Y618F mutant in 3T3L1-G4myc-CARΔ1 adipocytes using an adenovirus expression system did not affect insulin-stimulated glucose uptake (Fig. 4A). We previously developed a sensitive and quantitative method to measure GLUT4 translocation by detecting C-Myc epitope-tagged GLUT4 (GLUT4myc), which is exogenously expressed on the cell surface (23,33–36). Using this system, we found that insulin-stimulated GLUT4 translocation was not altered by transient expression of APS wild-type or Y618F mutant in 3T3L1-G4myc-CARΔ1 adipocytes (Fig. 4B). APS also did not affect insulin-stimulated GLUT4 translocation and glucose uptake in CHO-insulin receptor-GLUT4myc cells (data not shown).

We studied the effects on insulin signaling of isolated adipocyte of APS^−/− mice. Tyrosine phosphorylations of insulin receptor, phosphorylations of Akt and mitogen-activated protein kinase (Fig. 5), and activation of phosphatidylinositol 3-kinase (data not shown) did not detect any difference between adipocytes of APS^−/− mice and that of APS^+/+ mice. The interaction of APS and Cbl was identified by Western blot analysis only in adipocytes of APS^+/+ mice after insulin stimulation (Fig. 5).

In isolated epididymal adipocytes from APS^−/− mice, basal glucose uptake was unchanged compared with findings in APS^+/+ mice, but insulin-stimulated glucose transport in adipocytes of APS^−/− mice was significantly increased over that of APS^+/+ mice (Fig. 6A). However, the significant difference of insulin-stimulated glucose transport in isolated skeletal muscle was not detected between the two groups of mice (Fig. 6B). Glycogen content in livers of APS^−/− mice was slightly lower, in either fasted or fed states, but there was no statistically significant difference (Fig. 6C). These results suggest that APS knockout affects insulin-stimulated glucose transport into adipose tissues and that APS deficiency in adipose tissues may induce increased insulin sensitivity directly or indirectly in APS^−/− mice.

Leptin and adiponectin, both major adipocyte secretory proteins, are thought to play important roles in glucose homeostasis and insulin resistance (37–40). Plasma concentrations of leptin and adiponectin in APS^−/− mice were significantly higher than concentrations in APS^+/+ mice (Fig. 7A and B). Leptin content and the expression of leptin mRNA in epididymal adipose tissues from APS^−/− mice were also significantly higher than in APS^+/+ mice (data not shown). Increased leptin and adiponectin levels might contribute to the increased insulin sensitivity in APS^−/− mice (see discussion). Since APS may affect the gene expressions of leptin and adiponectin, we examined these mRNAs by Northern blotting in 3T3-L1 adipocytes. As shown in Fig. 7C, APS did not affect the expressions of these genes. While there is no direct evidence that APS knockout causes the increased leptin and adiponectin levels, the increase in these levels might contribute to the higher insulin sensitivity in APS^−/− mice.

In this study, we found that APS^−/− mice did show the remarkable phenotype of increased insulin sensitivity and hypoinsulinemia, although we were not able to elucidate the molecular mechanisms. Our results suggest that enhancement of insulin sensitivity by lacking APS might be due to increased insulin-response on adipose tissues and increased release of leptin and adiponectin. Our analysis on insulin receptor expression in APS^−/− mice indicated that APS is not likely to be involved in insulin receptor pathway degradation. Insulin receptor expression levels in various tissues of APS^−/− mice were comparable to those of APS^+/+ mice.

It has been reported that large adipocytes increase insulin resistance (41) or increase the number of small adipocytes, leading to improvement in insulin resistance (42). To evaluate the effect of lack of APS on adipose tissue morphology, we carried out histological studies on adipocytes of APS^−/− and APS^+/+ mice but found no distinct differences (data not shown). Therefore, increased insulin sensitivity and glucose transport in adipocytes of APS^−/− mice does not seem to be caused by the cell size or by the morphology.

APS forms an adaptor protein family with SH2-B as insulin receptor activation loop binding proteins that undergo insulin-stimulated tyrosine phosphorylation. APS also interacts with c-Cbl-associated protein (CAP), which is an important molecule that mediates insulin-dependent and phosphatidylinositol 3-kinase-independent glucose transport (43). Therefore, we examined whether the expressions of these molecules might change in APS^−/− mice, but the expressions of SH2-B and CAP were unchanged (data not shown). Protein tyrosine phosphatase 1B (PTP1B) is a major protein tyrosine phosphatase that has been implicated in the regulation of insulin action and is related to insulin resistance. Actually, PTP1B knockout mice showed the phenotype of increased insulin sensitivity in skeletal muscle (44). APS overexpression in CHO cells elevated the expression of PTP1B (45). However, the expression of PTP1B in APS^−/− mice was almost the same as that in APS^+/+ mice (data not shown). Therefore, the changes of expressions of SH2-B, CAP, or PTP1B do not seem to cause the phenotype of APS^−/− mice.

Liu et al. (18) reported that APS facilitates c-Cbl tyrosine phosphorylation in response to insulin and that both APS wild-type and Y618F mutant inhibited insulin-triggered GLUT4 translocation. We found that APS led to c-Cbl tyrosine phosphorylation but had no effects on GLUT4 translocation and glucose uptake (Fig. 3B and 4). The reason for the difference between our data and those of Liu et al. is unclear.

The mouse APS gene consists of eight coding exons and is colocalized to chromosome 5 (10). We substituted the EcoRI-EcoRI fragment, including exon 1–6, for a neomycin selection cassette and disrupted exon 1–6 of APS. We verified the complete lack of 2.9 kb APS mRNA in APS^−/− mice in both skeletal muscle and adipose tissue. However, we did detect a smaller size band (1.4 kb) of transcription in adipose tissue of APS^−/− mice (Fig. 1B) and in skeletal muscle after long exposure (data not shown). We sought to determine whether a small-sized APS protein existed, including a splicing variant, because there is the possibility that an APS COOH-terminal lesion might remain and that protein of the lesion would be expressed. We did not detect a small sized APS protein using immunoblotting and an anti-APS COOH-terminus antibody as well as an NH₂-terminus antibody (data not shown). In Southern blot analysis using the APS NH₂-terminal region as a probe, no band was seen in case of APS^−/− mice, although the clear band was seen for the APS^+/+ mice and a weaker band in APS heterogeneous mice (data not shown). When the COOH-terminal region of APS was used as a probe, the band was detected in all these mice because the COOH-terminal region of APS gene containing exon 7–8 was not disrupted in APS^−/− mice (M.I., C.K., S.-M.K., A.Y., Y.K., N.Y., K.T., S.T., unpublished data). Therefore, this small band (1.4 kb) of the transcript in APS^−/− mice does not seem to be functional.

APS presumably manifests its negative function in an independent manner from c-Cbl that could bind to COOH-terminal phosphorylated tyrosine residue and from insulin receptor downregulation. These observations present a reasonable synonym with the negative regulation by Lnk in c-Kit signals (14,37). In contrast, positive regulatory roles for SH2-B have been demonstrated in SH2-B^−/− mice (13). Thus, despite the significant structural similarities between Lnk, APS, and SH2-B, their functions appear to be quite different from each other.

Our hyperinsulinemic-euglycemic clamp studies were performed in anesthetized mice; therefore, the clamp data in these studies might not be representative of glucose kinetics in conscious mice. We cannot rule out the possibility of the increased insulin sensitivity in liver in these APS^−/− mice. Adipose tissue is not only a store of excess energy, but also secretes a variety of proteins into the circulating blood, and systemic metabolism is influenced. Leptin and adiponectin, the hormones secreted by adipocytes, regulate energy consumption and glucose metabolism (38,39). Leptin regulates the amount of body fat by reducing food intake and increasing energy consumption, stimulates glucose transport (40), and shows an increased insulin sensitivity in the whole body (46). The serum concentration of leptin was increased in APS^−/− mice compared with APS^+/+ mice (Fig. 7A). Although there was no relation with obesity, the epididymal fat weight of APS^−/− mice increased slightly (Table 1). It might contribute to the increased serum concentration of leptin in APS^−/− mice. Adiponectin exists abundantly in the plasma, enhances insulin action, and expression of adiponectin correlates with insulin sensitivity (47–49). Lack of leptin or adiponectin causes a decrease in insulin sensitivity, glucose intolerance, and development of type 2 diabetes (47,50). The serum concentration adiponectin and leptin was increased in APS^−/− mice compared with APS^+/+ mice (Fig. 7B). The mechanisms of the increases of leptin and adiponectin in APS^−/− mice were not elucidated. Since leptin synthesis and secretion is stimulated by increased glucose flux into adipose cells, it is possible that the increased rate of glucose transport is responsible for the increased levels of leptin. Conversely, increases of leptin and adiponectin might lead to increased insulin sensitivity and insulin-stimulated glucose uptake of adipocytes isolated from APS^−/− mice.

This work was supported by research grants from the Ministry of Education, Science, Technology, Sports, and Culture of Japan, Takeda Science Foundation, Merck, the Naito Foundation, Kowa Life Science Foundation, Kato Memorial Foundation, and the Japan Diabetes Foundation.

We thank Dr. T. Murakami (Kanazawa University) for providing obese (ob) mouse cDNA, Dr. Y. Kawano (Ohtsuka Pharmaceutical, Tokyo, Japan) and Dr. F. Yoshizawa (Utsunomiya University) for technical advice on glucose uptake into isolated skeletal muscle, and Dr. J. DeGregori (University of Colorado) for kindly providing CARΔ1. We also thank M. Ohara for reading the manuscript.