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. Author manuscript; available in PMC: 2014 Feb 5.
Published in final edited form as: Cell Metab. 2013 Feb 5;17(2):197–209. doi: 10.1016/j.cmet.2013.01.009

LRP6 Enhances Glucose Metabolism by Promoting TCF7L2-dependent Insulin Receptor Expression and IGF Receptor stabilization in Humans

Rajvir Singh 1, Renata Belfort De Aguiar 1, Sarita Naik 1, Sheida Mani 2, Kamal Ostadsharif 3, Detlef Wencker 4, Masoud Sotoudeh 5, Reza Malekzadeh 6, Robert S Sherwin 1, Arya Mani 1,7
PMCID: PMC3589523  NIHMSID: NIHMS438420  PMID: 23395167

Abstract

Common genetic variations in Wnt signaling genes have been associated with metabolic syndrome and diabetes by mechanisms that are not well understood. A rare nonconservative mutation in Wnt-coreceptor LRP6 (LRP6R611C) has shown to underlie autosomal dominant early onset coronary artery disease, type 2 diabetes and metabolic syndrome. We examined the interplay between Wnt and insulin signaling pathways in skeletal muscles and skin fibroblasts of healthy non-diabetic LRP6R611C mutation carriers. LRP6 mutation carriers exhibited hyperinsulinemia and reduced insulin sensitivity compared to the non-carrier relatives in response to oral glucose ingestion, which correlated with a significant decline in tissue expression of the insulin receptor (IR) and insulin signaling activity. Further investigations showed that LRP6R611C mutation diminishes TCF7L2-dependent transcription of IR while it increases the stability of IGFR and enhances mTORC1 activity. These findings identify Wnt/LRP6/TCF7L2 axis as a regulator of glucose metabolism and a potential therapeutic target for insulin resistance.

Introduction

Despite widespread recognition of metabolic syndrome as a major risk factor for coronary artery disease and diabetes (Mallika et al., 2007; Mottillo et al., 2010; Vaidya et al., 2010), its underlying causes have remained poorly understood (Gade et al., 2010). Genome-wide association studies have linked polymorphisms within the gene encoding Wnt transcription cofactor TCF7L2 to risk for type 2 diabetes in independent large cohorts (Saxena et al., 2006; Wang et al., 2007). Genetic variations of Wnt5B gene have been similarly associated with susceptibility to type 2 diabetes (Kanazawa et al., 2004). These findings have implicated altered Wnt signaling in impaired glucose metabolism and diabetes.

While the associations between these genetic variants and type 2 diabetes appear as extremely robust, their effects on cellular insulin response are not understood (Liu et al., 2009; Rasmussen-Torvik et al., 2009). In principal, the biological effects imparted by the common polymorphisms are too small to be measurable in artificial cell systems and animal models. Disruptions of the genes that encode Wnt signaling proteins in animal models have similarly indicated the critical role of this pathway in glucose homeostasis (Leng et al., 2010; Pagel-Langenickel et al., 2008; Summers et al., 1999). However, the demonstrated effects of Wnt/β-catenin on insulin signaling (Abiola et al., 2009; Liu et al., 2011; Liu et al., 2012) and hepatic gluconeogenesis have been often inconsistent (Liu et al., 2011). These discrepancies once again reveal a recognized problem with animal models for study of glucose homeostasis, indicating urgent need for human studies.

The canonical Wnt signaling pathway consists of cascades of events that initiate after Wnt proteins bind to the cell-surface receptor frizzled and its coreceptors LRP5/6, triggering inactivation of GSK3β and stabilization of β-catenin (Bilic et al., 2007; Go and Mani, 2012; Schweizer and Varmus, 2003). The final steps include translocation of β-catenin from cytoplasm to the nucleus where it interacts with TCF/LEF family of transcription activators to promote gene expression. We have recently reported a large outlier kindred in whom a rare non-conservative loss of function mutation in LRP6 (LRP6R611C) underlies a monogenic form of early onset CAD, hypertension, dyslipidemia and diabetes, suggesting insulin resistance (Mani et al., 2007). Genetic studies of rare families segregating single gene mutations that impart very large effects on disease pathogenesis have shown greatest potentials to identify genes and pathways that have paved the way for discovery of novel and potent therapeutics (Goldstein and Brown, 1973; Hobbs et al., 1992; Hobbs et al., 1989). Accordingly, the identification of a mutation with major effect in the Wnt-coreceptor LRP6 provided us with an exceptional opportunity to study the role of Wnts and their downstream signaling proteins on glucose metabolism.

The current study was undertaken to examine the interplay between Wnt and insulin signaling pathways and its potential role in development of insulin resistance and type 2 diabetes. For this purpose, we examined young, healthy non-diabetic LRP6R611C mutation carriers and their non-mutation carrier relatives matched for gender, weight and physical activity levels for the presence of impaired insulin sensitivity. Insulin signaling was investigated in the skeletal muscle, the major organ for glucose disposal, and in the skin fibroblast cultures of the mutation carriers and non-carrier relatives after they underwent extensive clinical evaluations and oral glucose tolerance tests (OGTT). These studies were complemented by comprehensive dissection of metabolic pathways in the cultured skin fibroblasts of LRP6R611C mutation carriers and their unaffected relatives as well as in vitro-expression systems.

Results

LRP6R611C mutation impairs glucose tolerance and insulin response

Eight LRP6R611C mutation carriers (n=8) and non-carriers (n=7) who denied history of diabetes participated in the study. The clinical characteristics are summarized in the Supplement Table. Overall, the mutation carriers had significantly higher LDL, and triglyceride levels and similar BMI and serum HDL as compared to non-carriers relatives (Mani et al., 2007).

All 15 family members recruited for the study underwent an OGTT prior to skeletal muscle and skin fibroblast biopsies. Skeletal muscle and skin biopsies were also obtained from mutation carriers with diabetes for comparison. Both serum glucose and insulin levels during 2h OGTT were significantly higher in mutation carriers compared to non-carriers (Fig. 1A, B). Analysis of the plasma glucose levels at 2h post glucose ingestion in mutation carriers revealed that one subject has developed diabetes (DM, blood glucose greater than 200 mg/dl at two h).

Fig. 1. Skeletal muscle insulin resistance in LRP6R611C mutation carriers.

Fig. 1

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LRP6R611C mutation carriers (R611C), with or without impaired glucose tolerance (IGT and NGT), had higher circulating glucose levels in response to oral glucose load compared to non-carrier relatives (WT) (A). Serum insulin levels were higher in LRP6R611C mutation carriers compared to non-carrier relatives during the last stages of the OGTT (B). The insulin sensitivity index (ISI) was significantly lower in LRP6R611C mutation carriers with NGT compared to non-carriers (C). IR mRNA expressions (D), IR protein levels and AKT phosphorylation (E) were significantly lower in mutation carriers with NGT (lanes3–4) and DM2 (lanes 5–6) compared to non-carriers (lanes1–2). Fig. 1F and G show quantification of IR and pAKT by densitometry (* specifies p<0.05, ** p<0.005). Error bars represent standard deviation (SD).

Circulating glucose levels in the mutation carriers were similar to non-carrier relatives in the first 45 minutes following glucose ingestion, but subsequently increased to higher levels between 60–120 minutes as compared to non-carrier relatives (Fig. 1A). The majority of patients (4 male, 1 female) had still normal glucose tolerance test by definition (NGT) (Supplement Table). Only two (1 male and 1female) had impaired glucose tolerance (IGT, blood glucose between 140 mg/dl and 200 mg/dl at two h). Fasting plasma glucose and insulin levels prior to the 2h oral glucose tolerance in the NGT LRP6R611C mutation carriers were normal and comparable to the seven non-carrier relatives (5 male, 2 female) who were matched for age, weight and physical activity levels (see supplement table). These findings are consistent with normal baseline hepatic glucose output. However, mean glucose levels in the NGT probands at 2h (118 ± 2 mg/dl) were significantly higher than those in non-carrier relatives (76 ± 12mg/dl (P<0.001). Plasma insulin levels were also initially similar but rose to significantly higher levels in the mutation carriers between 90–120 minutes (Fig. 1B). The delayed decay of the plasma glucose in LRP6 mutation carriers, despite elevated insulin levels, is consistent with a subclinical skeletal muscle insulin resistance as described by prior studies (Abdul-Ghani et al., 2007). Accordingly, the insulin sensitivity index (ISI) was reduced in the mutation carriers with NGT (5.17 ± 1.18) as compared to non-carriers (7.7 ± 1.2) (p<0.005) (Fig. 1C). The remaining three mutation carriers (2 IGT and 1 DM) had also diminished ISI as expected (4.9 ± 0.77, p<0.005). Taken together, these findings suggested significantly greater predisposition to skeletal muscle insulin resistance in mutation carriers vs. their non-carrier relatives and prompted study of insulin signaling in the skeletal muscles of the mutation carriers and non-carrier relatives.

Three male mutation carriers (subjects III-1, III-2, III-7 in the supplement table) and 3 matched non-carrier controls also agreed to participate in a hyperinsulinemic-euglycemic clamp study. This study was carried out to confirm OGTT findings. The glucose infusion rate required to maintain euglycemia during the higher insulin dose clamp was diminished in each of the 3 mutation carriers compared to non-carriers (233 ± 25 mg/m2/min vs. 396 ± 56, respectively, p=0.01). Accordingly, the glucose disposal rate (GDR) during high dose insulin in mutation carriers (with or without IGT/diabetes) was significantly lower compared to non-carriers (256 ± 23 vs. 443 ± 79 mg/m2/min, p<0.05) (Suppl. fig. S1). While the number of subjects studied was small, the differences between mutation carriers and non-carriers were drastic. Accordingly, a power calculation estimated the number to based on 30% difference in GDR to be less than 3, see supplement material). The sample size was too small to exclude hepatic insulin resistance based on hepatic glucose output (Suppl. fig. S1B) or OGTT. Taken together, these finding further supported the presence of skeletal muscle insulin resistance in LRP6R611C mutation carriers.

LRP6R611C impairs skeletal muscle IR expression and insulin signaling activity

To examine the effect of LRP6R611C mutation on insulin signaling prior to development of impaired glucose tolerance, tissues (skeletal muscle and fibroblasts) from individuals with NGT (n=5) and five matched non-carriers relatives were compared for expression of genes involved in insulin signaling. The Q-PCR demonstrated a 60% reduction in skeletal muscle IR mRNA in LRP6R611C mutation carriers as compared to non-carriers (Fig. 1D, p<0.005), suggesting impaired transcription of the IR. Accordingly, Western blot analysis showed a greater than 50% reduction of protein abundances for IR-β in the skeletal muscles of the mutation carriers compared to non-carriers (Fig. 1E and F). As anticipated, IR expression was also reduced in the skeletal muscles of LRP6R611C mutation carriers with diabetes. The analysis also showed reduced AKT phosphorylation in the mutation carriers as compared to non-carriers (Fig. 1E and G). Taken together, these results suggest major impairment in the very initial step of the skeletal muscle insulin signaling in LRP6R611C mutation carriers compared to non-carriers relatives.

Insulin signaling is impaired in cultured fibroblasts of the LRP6R611C mutation carriers

LRP6 phosphorylation in response to Wnt stimulation was significantly lower in the skin fibroblasts of LRP6R611C mutation carriers compared to non-carriers, without having significant effects on adipogenic differentiation (Suppl. fig. S2A &B). Analogous to the skeletal muscle, the IR mRNA (Fig. 2A) and protein (Fig. 2B and 2D) expression levels were reduced in the fibroblasts of LRP6R611C mutation carriers compared to their non-carrier relatives. To assess the impact of these changes on downstream signaling, skin fibroblast cultures obtained from NGT mutation carriers and non-carrier relatives were stimulated with insulin. Insulin stimulated activations of IRS-1, AKT and GSK3β were significantly reduced in fibroblasts of LRP6R611C mutation carriers compared to non-carriers (Fig. 2C and 2E–G).

Fig. 2. Reduced IR expression and insulin response in the skin fibroblasts of LRP6R611C mutation carriers.

Fig. 2

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IR mRNA (A) and IR-β protein levels (B) were significantly reduced in skin fibroblasts of LRP6R611C mutation carriers with NGT (lanes 3–4) or diabetes (lanes 5–6) compared to non-carriers relatives. Insulin response assessed by phosphorylation of IRS-1, AKT, and GSK3β was significantly impaired in the skin fibroblasts of the LRP6R611C mutation carriers compared to non-carrier relatives(C). Fig. 2D–G show quantification of IR, p-IRS-1, p-AKT and p-GSK3β by densitometry. (* specifies p<0.05, ** p<0.005, *** p<0.001). Error bars represent SD.

LRP6 is required for normal expression of the IR

LRP6R611C phosphorylation in response to Wnt3a is significantly reduced in skin fibroblasts of mutation carriers compared to wildtype LRP6 (Suppl. fig. 4, upper and lower panels). This confirmed our earlier findings that LRP6R611C mutation is a loss of function mutation. To examine whether LRP6 is required for normal expression of IR, LRP6 was knocked down by RNA interference. LRP6 specific shRNA diminished expression of LRP6 by more than 90% (Fig. 3A). IR mRNA and protein levels in LRP6 knockdown 3T3L1 cells were reduced by greater than 50% (p<0.05) (Fig. 3A–C). This corresponded to markedly reduced IRS-1 and AKT activities in response to insulin in the differentiated LRP6 knockdown 3T3L1 cells (Fig. 3D–E). Treatment of 3T3L1 cells with actinomycin D and cylcohexamide did not alter IR protein levels (data not shown), indicating that the regulation of IR by LRP6 is specifically at transcriptional levels. These findings once again underscore the critical role of LRP6 in IR transcription and maintaining the integrity of the insulin signaling.

Fig. 3. RP6 knockdown reduces IR transcription and impairs insulin signaling.

Fig. 3

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mRNA interference using LRP6 specific shRNA in 3T3L1 preadipocytes reduced expression of LRP6 by more than 90% (A) and reduced expressions of IR protein (A) and mRNA (B) by greater than 50%. Figure C shows IR quantification by densitometry. In LRP6 knockdown 3T3L1 adipocytes there was reduced response to insulin stimulation assayed by phosphorylation of IRS-1, AKT and GSK-3β (fig. D). Figures 3E–G shows quantification of IRS-1, p-AKT and p-GSK3β by densitometry (* specifies p<0.05, ** p<0.005,*** p<0.001). Error bars represent SD.

The canonical Wnt signaling enhances IR expression

Given the above findings a broader question was raised as to whether IR transcription is regulated by Wnt stimulation. To address this issue, mRNA and protein expressions were measured at different time points in 3T3L1 cells stimulated with Wnt3a (Fig. 4A, B, C). IR mRNA expression increased continuously until it reached a maximum after 8 h. IR protein began to increase 6 h after stimulation and reached a maximum after 8 h. Wnt stimulation completely failed to increase IR expression when LRP6 was knocked down (Fig. 4D). We then treated LRP6 knockdown 3T3L1 cells with GSK3β antagonist LiCl for 24h to rescue IR expression. Upon Wnt3a stimulation IR expression was significantly increased in cells treated with LiCl vs. those untreated (Fig. 4E). This finding strongly suggests that LRP6 mediates Wnt3a-dependent IR transcription. Earlier studies have shown that LRP6 behaves as physiological inhibitor of the non-canonical Wnt signaling and many of the phenotypes of LRP6 knockout mice are due to gain of function in the non-canonical pathway (Bryja et al., 2009). To examine the effect of non-canonical pathway on IR transcription, 3T3L1 cells were stimulated with Wnt5a. Wnt5a failed to alter IR expression (Fig. 4F). The role of canonical Wnt signaling in inducing IR expression was further explored by knockdown of β-catenin, an integral component of this pathway. Accordingly, the β-catenin knockdown by RNA interference led to significant reduction in expression of the IR protein (Fig. 4G).

Fig. 4. LRP6-dependent IR transcription is mediated by the canonical Wnt signaling.

Fig. 4

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Stimulation of 3T3L1 preadipocytes by Wnt3a (30 ng/ml) resulted in a continuous rise of IR mRNA(A) and protein levels (B), reaching a maximum 8 h. post stimulation; β-catenin and cyclin-D1 were used as positive controls. IR, cyclinD1 and β– catenin quantification by densitometry are shown (C). The stimulatory effect of Wnt3a (30 ng/ml) on IR transcription was completely blunted when LRP6 was knocked down (D). LiCl (20 mM/ml) treatment of the 3T3L1 cells significantly rescued IR expression (E). Non-canonical stimulation of 3T3L1 cells with Wnt5a (50 ng/ml), in contrast to Wnt3a, did not have any effect on IR-β expression (F). The role of canonical Wnt signaling in IR expression was further investigated by lentivirus Sh-RNA knockdown of β-catenin in 3T3L1 cells, which resulted in lower expression of IR-β compared to GFP-shRNA (G). (* specifies p<0.05, ** p<0.005, *** p<0.001). Error bars represent SD.

Transcription of the IR is regulated by TCF7L2

We examined the expression levels of high mobility group protein [HMGI (Y)] in the skeletal muscles and skin fibroblasts of mutation carriers and their unaffected family members. HMGI (Y) is encoded by HMGA and regulates IR transcription (Brunetti et al., 2001). Mice deficient for HMGA1 express low levels of the IR in insulin sensitive tissue like the skeletal muscles and are diabetic (Foti et al., 2005). HMGA has shown to be inducible by Wnt/β-catenin in different malignant cells. There were no differences at mRNA or protein expression levels of HMGI (Y) in the skeletal muscles or skin fibroblasts of mutation carriers and their unaffected relatives (Suppl. fig. 3A–F).

TCF7L2 is a HMG box-containing transcription factor, which is downstream of Wnt/β-catenin. Common genetic variations of TCF7L2 have been associated with risk for type 2 diabetes in numerous independent populations across the world (Scott et al., 2007; Sladek et al., 2007). These variations cause reduced protein expression (Shu et al., 2009), but its effect on insulin signaling is not understood. The IR promoter has number of putative binding sites for the TCFs. TCF7L2 knockdown by RNA interference in both 3T3L1 preadipocytes and human skin fibroblasts abolished the stimulatory effect of Wnt3a on IR transcription (Fig. 5A, B). In contrast, overexpression of TCF7L2 in 3T3L1 cells caused greater than 3-fold increase in IR expression (Fig. 5C, p<0.01).

Fig. 5. TCF7L2 regulates IR expression.

Fig. 5

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Lentivirus Sh-RNA knockdown of TCF7L2 reduced expression of IR-β in 3T3L1 preadipocytes (A) and human skin fibroblasts (B) compared to GFP-shRNA. Accordingly, TCF7L2 overexpression in 3T3 L1 cells resulted in enhanced IR expression, which was further augmented with Wnt3a stimulation (C). A luciferase-based reporter gene assay in HEK cells using IR promoter showed 1.5 to 2 fold increase in luciferase activity starting at 8 h after Wnt stimulation compared to empty vector (D). ChIP revealed binding of TCF to a specific promoter region of IR; IgG was used as negative control (E). Top figure shows relative binding of IR promoter precipitated with either TCF7L2 or IgG antibody, before and after treatment with Wnt3a; lower panel shows agarose gel pictures of the PCR products. H3 stands for histone3. EMSA demonstrating TCF7L2 binding to a TCF consensus motif in IR promoter using nuclear extracts from 3T3L1 cells, 30 minutes after Wnt3a stimulation (F), Lane 1: probe with no extract, Lane 2: probe with nuclear extract, Lane 3: probe with nuclear extract and excess of non-biotinylated probe, Lane 4: probe with nuclear extract and IgG as control, Lane 5: probe with nuclear extract and TCF7L2 antibody showing the supershift. Error bars represent SD.

The effect of Wnt3a on IR transcription was further assayed in HEK cells by IR-promoter dependent expression of luciferase (Fig. 5D). There was significant increase in IR-promoter dependent expression of luciferase after 8 h stimulation with Wnt 3a. The role of TCF7L2 in IR transcription was examined in 3T3L1 cells treated with PBS or Wnt-3a by chromatin immunoprecipitation (ChIP) using a specific antibody. Binding of TCF7L2 to IR promoter was verified by Real time PCR, using IR promoter-specific primers. The ChIP analysis showed enhanced binding of TCF7L2 to IR promoter upon stimulation with Wnt3a compared to vehicle alone (Fig. 5E, upper and lower panels). The EMSA analysis using nuclear extracts of the 3T3L1 cells treated with Wnt3a showed binding of TCF7L2 to the TCF consensus motif in IR promoter and a super-shift after using a ChIP-grade antibody (Fig. 5F). Taken together, these findings establish the role of TCF7L2 as a transcription regulator of the IR with major implications for therapy in diabetes particularly in patients with TCF7L2 variants.

TCF7L2 overexpression in skin fibroblasts of LRP6R611C mutation carriers fully restored IR expression (Supplement fig. S4A) and insulin signaling (Supplement fig. S4B –C). This finding once again underscores the biological importance of this transcription factor in regulation of IR transcription and insulin signaling in humans.

mTORC1-dependent IRS-1 serine phosphorylation contributes to Insulin resistance

Mechanisms that regulate glucose metabolism are linked through numerous intricate molecular pathways, raising the possibility that the R611C mutation may have other effects on insulin signaling and glucose homeostasis. This assumption was further supported by increased IRS-1 serine phosphorylations in response to insulin in LRP6R611C vs. wildtype fibroblasts (Fig. 6A). Increased phosphorylation of IRS-1 on multiple serine residues (Ser307, 612, 636/639 and 1101) in LRP6R611C fibroblasts suggested alteration in the function of mTORC1, a major regulator of insulin signaling and metabolism (Mothe and Van Obberghen, 1996). LRP6R611C fibroblasts exhibited significantly higher S6K and S6 phosphorylations compared to wildtype fibroblasts, suggesting enhanced mTORC1 activity. Increased mTORC1 activity correlated with increased ERK1/2 thr202/tyr204 phosphorylation in LRP6R611C fibroblasts compared to wildtype fibroblasts.

Fig. 6. Enhanced ERK1/2 dependent activation of mTOR results in IRS-1 serine phosphorylation at several residues.

Fig. 6

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Increased expression of IGFR results in enhanced ERK1/2-dependent activation of mTORC1 and enhanced phosphorylation of IRS-1 at several serine residues in skin fibroblasts of LRP6R611C mutation carriers compared to non-carriers (A). LRP6 knockdown increases IGFR expression and results in mTOR-dependent IRS-1 phosphorylation at several serine residues (B). Rapamycin treatment of the skin fibroblasts of LRP6R611C mutation carriers significantly reduces IRS-1 serine phosphorylation (C). IGFR protein (B) but not mRNA levels (D) are higher in LRP6 knock down compared to vector alone. IGFR protein expression rose to similar levels after treatment with proteosomal inhibitor MG-132 (5μm/ml) for 8 h. (E). SUMO-1 was immunoprecipitated and IGFR was immunoblotted with specific antibody and vice versa before and after treatment of cells with IGF-1 (100 nM/ml) for 8 h. IGFR was coimmunoprecipitated with SUMO-1 to significantly greater degree in LRP6 knockdown cells compared to vector alone (F). Arrows show bands that correspond to IGFR. (*** in the figure 6D specifies p<0.001). Error bars represent SD.

Insulin increases the activity of ERK1/2 via IR and IGFR. While IR expression was significantly reduced in LRP6R611C fibroblasts as previously shown, IGFR expression was significantly increased in these cells compared to wildtype fibroblasts (Fig. 6A). Consistent with these findings, LRP6 Knock down by RNA interference resulted in increased expression of IGFR and activity of mTORC1 pathway (Fig. 6B). The significance of mTOR pathway in IRS-1 serine phosphorylation was investigated by treating human skin fibroblasts with mTORC1 antagonist rapamycin for 48h. IRS-1 phosphorylation at these residues significantly decreased in response to insulin in LRP6R611C fibroblasts treated with rapamycin (Fig. 6C).

IGFR mRNA was significantly reduced in LRP6 knocked down cells compared to cells infected with vector alone (Fig. 6D) suggesting posttranscriptional regulation of IGFR by LRP6. Accordingly, protein levels rose to similar levels in both cell types after treatment with proteosomal inhibitor MG-132 (Fig. 6E), indicating IGFR ubiquitination by LRP6. Earlier studies have shown that IGFR is target of sumoylation (Sehat et al., 2010), a process that may render stability to the protein. Immunocoprecipitation studies in 3T3L1 cells treated with IGF-1 revealed that IGFR and SUMO-1 (small ubiquitin like-modifier 1) form a complex in LRP6 knocked down cells (Fig. 6F). These findings provide strong evidence that LRP6 regulates nutrient sensing mTOR pathway through a tight network and impairment of this regulation by LRP6R611C mutation results in excess ERK1/2-dependent mTORC1 activity and IRS-1 serine phosphorylation at multiple residues.

Discussion

We have previously reported of a nonconservative mutation in Wnt coreceptor LRP6 (R611C) that underlies autosomal dominant early onset atherosclerosis, diabetes and metabolic syndrome (Keramati et al., 2011; Mani et al., 2007; Ye et al., 2012). In the current study we demonstrate that healthy LRP6R611C mutation carriers suffer from impaired skeletal muscle insulin signaling prior to development of impaired glucose tolerance or diabetes.

Dissection of the insulin-signaling pathway in the skeletal muscle of LRP6R611C mutation carriers with NGT revealed markedly reduced expression of the skeletal muscle IR as one of the underlying causes of insulin resistance. The IR and its downstream effectors constitute an evolutionarily highly conserved pathway and perturbation of their expression and function has been linked to insulin resistance. Reduced transcription of IR has been previously described in type A insulin resistance subjects with mutations in IR gene (Kadowaki et al., 1988; Kahn et al., 1976; Kahn and White, 1988; Kriauciunas et al., 1988; Ojamaa et al., 1988). The affected individuals exhibit extremely high circulating insulin levels in early years of life and require extreme doses of insulin when they develop diabetes. In contrast, reduced IR expression in ob/ob and db/db mice is due to defects in posttranscriptional regulations (Ludwig et al., 1988).

There is relatively little known about the transcriptional regulation of the IR (McKeon, 1993). High mobility group protein HMGI (Y) and FOXO1 have been recently shown to be transcriptional regulators of IR (Brunetti et al., 2001; Puig and Tjian, 2005). HMGI (Y) is a Wnt/β-catenin inducible protein (Akaboshi et al., 2009), which together with TCF7L2 belongs to HMG domain family of DNA-bending proteins (van Houte et al., 1993) and FOXO1 is a pluripotent transcription factor activated by Wnt/β-catenin (Essers et al., 2005). Our studies identified TCF7L2 as a potent transcriptional regulator of the insulin receptor. Genome-wide association studies have associated a minor TCF7L2 (formerly known as TCF4) allele with the risk for type 2 diabetes. The minor allele has been linked to reduced TCF7L2 protein expression and impaired insulin signaling (Yoon et al., 2010). The effect imparted by this variant on insulin signaling is inherently too small to be measurable in biological systems (Liu et al., 2009; Lyssenko et al., 2007). In contrast, the effect of R611C mutation on blood glucose is substantial, allowing careful investigation of Wnt and insulin signaling and the interplay between them. This led to the key finding that Wnt- β-catenin activation triggers binding of TCF7L2 to a TCF/LEF binding motif within IR promoter and enhances transcription of IR. The TCF binding motif is abundantly present in 5′ and 3′ regulatory regions of the IR and is highly conserved from drosophila, to mammals, indicating the important role of TCFs in transcriptional regulation of IR. The restoration of IR expression and insulin signaling in the skin fibroblasts of LRP6R611C mutation carriers by TCF7L2 overexpression further confirmed the biological importance of this transcription factor in regulation of IR transcription and insulin signaling in humans.

LRP6R611C mutation has likely multifaceted effects on glucose homeostasis (fig. 7). Dissection of insulin signaling in the skin fibroblasts of LRP6R611C mutation carriers showed that this mutation impairs insulin signaling also by phosphorylation of IRS-1 at multiple serine residues. IRS-1 Serine phosphorylations are regulated by different kinases. Raptor-mTOR and S6K1, however, are required for phosphorylation of IRS-1 at a large subset of serine residues (Shah and Hunter, 2006). mTORC1 is activated by insulin as well as growth factors like IGF-1. Our findings suggest that impaired ubiquitination of IGFR by the mutant LRP6 results in enhanced expression and activities of IGF-mTOR pathways. High insulin levels in presence of marked reduction of IR likely exacerbate this effect.

Fig. 7. Schematic of Wnt/LRP6 signaling, its role in regulation of IR and IGFR ubiquitination and the effect of LRP6 mutation on these function.

Fig. 7

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LRP6 mediates Wnt3a activation and binding of TCF7L2 to TCF/LEF binding motif within IR promoter and enhances transcription of IR and promotes insulin signaling. Furthermore, it regulates IGFR expression by promoting its ubiquitination (A). The mutant LRP6 protein sumoylates IGFR and increases its stability. IGFR enhances ERK1/2-dependent mTOR activity. mTOR phosphorylates IRS-1 at several serine residues and promotes its ubiquitination (B). In addition, mutant protein impairs Wnt signaling and diminishes TCF7L2-dependent IR transcription.

Taken together our investigations provide mechanisms by which Wnt regulates insulin signaling in preadipocytes and in primary cells of mutation carriers. Association between Wnt and insulin signaling in preadipocytes had been reported without elaborating the mechanisms (Palsgaard et al., 2012). By dissecting insulin signaling pathways in mutant and wildtype primary human cells we discovered that Wnt/LRP6 regulate IR and IGFR expressions via two independent mechanisms. Through rigorous investigations we demonstrated that Wnt signaling regulates transcription of IR, while its coreceptor LRP6 regulates IGFR by ubiquitination.

In this study we focus on the earliest effects of the LRP6R611C mutation on skeletal muscle insulin signaling before mutation carriers develop apparent signs of impaired glucose tolerance. Since LRP6 is ubiquitously expressed, parallel changes in IR expression of the liver and pancreas tissues of the mutation carriers may occur. While the abnormalities we demonstrate here are limited to the skeletal muscle, reduced IR expression in the liver and or beta cells may similarly contribute to hyperglycemia later in the disease course. Lack of access to these tissues is a limitation of our study. Our physiological studies may also have been underpowered to show subtle effects of the mutation on hepatic glucose production at the very early stage of the disease. Furthermore, other LRP6 mutations may primarily impair hepatic insulin sensitivity or beta cell function. Larger studies involving carriers of different LRP6 mutations as well as animal models may provide further information on effects of Wnt signaling on glucose homeostasis. In summary, our study presents molecular pathways that links Wnt signaling to insulin action in the skeletal muscle and provides significant insight into pathogenesis of metabolic syndrome. Through molecular dissection of these pathways we have identified a number of potential targets for development of novel therapeutics against inulin resistance ranging from LRP6 and IGFR/mTORC1 to TCF7L2.

Methods

Study Population

The original R611C kindred was extended to 48 mutation carriers and 44 non-carriers. Healthy nondiabetic members of the family were asked to participate in the study. Among these individuals 8 LRP6R611C mutation carriers (6 male, 2 female) and 7 non-diabetic non-mutation carrier (5 male, 2 female) family members were enrolled. Each volunteer was free of major medical problems based upon clinical evaluations and was not taking prescription medications. All probands underwent physical examination, blood testing, and an OGTT followed by skeletal muscle and skin punch biopsies. The institutional review board at the Yale University School of Medicine in the United States and the Tehran University of Medical Sciences and Shahid Motahari Hospital in Iran approved the research protocol; all study participants provided written informed consent for clinical and genetic studies.

OGTT and euglycemic-hyperinsulinemic clamp studies

Glucose tolerance was assessed by administration of a 75 g dextrose drink (Glucola, Curtin Matheson Scientific, Houston, TX). Blood samples for measurement of plasma glucose and insulin concentrations were obtained at 0, 15, 30, 45, 60, 90, 120 minutes and used to estimate insulin sensitivity by the ISI (10−4 dl/min per micro unit/ml) was calculated as ISI=10,000/sqrt (FPGXFPI) X (GX I), where FPG is fasting plasma glucose, FPI is fasting plasma insulin, and G and I are average glucose and insulin concentrations during the OGTT (Matsuda and DeFronzo, 1999).

A selected number of the LRP6R611C mutation carriers (n=3) and non-carriers (n=3) agreed to participate in euglycemic-hyperinsulinemic clamp studies. These subjects were admitted to the Yale-New Haven Hospital Research Unit of the Yale Center for Clinical Investigation (YCCI) on the day of the study, after an overnight fasting. Two indwelling venous catheters were inserted. One into a distal arm or hand vein that was placed in a heated box to allow for the sampling of arterialized venous blood. The second venous catheter was used for the administration of a primed (200 mg/m2) and continuous (2 mg/min−1·m−2) infusion of 6, 6D2-glucose and D5-glycerol (4 μM/m2·min) as well as insulin and glucose. After the basal equilibration period lasting 2 h, a sequential two-step euglycemic-hyperinsulinemic clamp study was performed (Mayerson et al., 2002), during which regular insulin was administered as a primed-continuous infusion at 10 and 80 mU/m2·min, each for 120 min. Plasma glucose levels were measured at 5 minute intervals and a variable glucose infusion initiated to maintain levels at ~90mg/dl. Additional plasma samples were drawn at baseline and throughout the insulin infusion period for measurement of isotope enrichment, insulin, FFA, and glycerol turnover. One of the LRP6R611C mutation carriers only received the higher insulin infusion dose, i.e. 80 mU/m2·min, for 120 min.

Skeletal muscle and skin biopsies

Skeletal muscle biopsies were obtained from M. vastus lateralis of mutation carrier and non-carrier relatives according to the technique of Bergström (Bergström et al., 1967). The first biopsates were quickly frozen in All protect Tissue Reagent (Qiagen) for later protein and mRNA quantification. Skin biopsies were also obtained and immediately placed in culture medium. Subsequently, fibroblasts were harvested by isolating dermis followed by fragmentation and placement 20 mm grid Petri dishes.

Analytical procedures

Plasma glucose concentrations were measured by the glucose oxidase method (Glucose analyzer II; Beckman Instruments Inc., Fullerton, CA) and plasma insulin was determined using a double antibody RIA (Diagnostic Systems Laboratories, Inc., Webster, TX). Glucose isotope data is acquired by a chromatography mass spectrometer at the DERC Stable Isotope Core Facility. Glucose kinetics were determined in the basal state and during euglycemic hyperinsulinemia (Sherwin et al., 1974). Total body glucose disposal during the clamp study was calculated as the sum of endogenous glucose production and the exogenous glucose infusion rate (GIR) in the final 30 minutes of each infusion dose.

Chemicals

Insulin, dexamethasone, 3-isobutyl-1-methylxanthine, puromycin, protease inhibitor cocktail (cat# P8340), phosphatase inhibitor (P2850) cocktail and phenylmethanesulfonyl fluoride (P7626), rapamycin (R8781) and LiCl were purchased from Sigma-Aldrich (St. Louis, MO). Cell lysis buffer (9803) and ChIP kit (9003S) were obtained from Cell Signaling Technology (Boston, MA). The lentiviral LRP6 shRNA(h) (sc-37233-v), LRP6 shRNA(m) (sc-37234-v), β-catenin shRNA (sc-29210-v), TCF-4(TCF7L2) shRNA (sc-43526-v) control shRNA (sc-108080) and polybrene were purchased from Santa Cruz Biotechnology Inc. Wnt-3a (1324-WN) and Wnt-5a (645-WN) were purchased from R & D System, Dulbeco’s Modified Eagle’s medium, fetal bovine serum, penicillin streptomycin cocktail, trypsin-EDTA solution and Trizol from GIBCO/Invitrogen, and PVDF membranes from (Bio-Rad Laboratories, Hercules, CA).

Antibodies, cell lines and adipocyte differentiation

Antibodies for LRP6, p-LRP6 (ser1490), IR β, Akt, p-Akt(ser473), IRS-1, GSK-3β, p-GSK-3β(ser9), β-catenin, cyclin D1, GAPDH, β-actin, IRS-1-P (ser307), IRS-1-P (ser1101), IRS-1-P (ser636/639), IRS-1-P(ser612), S6 ribosomal protein-p (ser235/236), S6 ribosomal protein, p70 S6Kinase-P(thr389), p70 S6 Kinase, mTOR, mTOR-P(ser2448) p44/42 ERK1/2-P(thr202/tyr204), p44/42 ERK1/2, IGFR, p-IGFR (tyr 1316) SUMO-1 and Ubiquitin were purchased from Cell Signaling Technology (Boston, MA). Antibodies for IR β (C-19), TCF-4(TCF7L2) (sc-8631), LRP6 (C-20), HMGI(Y), IRS-1-P (ser265/270) were purchased from Santa Cruz Biotechnology Inc. (Santa Cruz, CA). Antibodies for p-IRS-1 (tyr612) were purchased from Invitrogen Life Technology, (ON, Canada). The specificity of the commercial IR antibodies has been confirmed previously(Cheung et al., 2010). Human skin fibroblasts, murine 3T3L1 cells, and human embryonic kidney (HEK) 293T cells were grown in DMEM supplemented with 10% FBS, and 1% (v/v) pen/strep cocktail. (HEK) 293T cells were used for lentivirus production. 3T3L1 cells (0.5×106/well) were plated on 6-well plate one day before infection. Polybrene (5ug/ml) and lentivirus particles were added into the medium. On day 2 following transduction the medium was replaced with medium supplemented with 5ug/ml of puromycin. Untransduced dead cells were removed on day two and live cells were further expanded and maintained in DMEM supplemented with 10% FBS &1% (v/v) pen/strep cocktail.

The 3T3L1 pre-adipocytes were differentiated into adipocytes by culturing with DMEM supplemented 10% FBS, 1μg/ml insulin, 1μM dexamethasone and 0.5mM isobutylmethylxanthine for 48 h and then for an additional 48h with DMEM supplemented with 10% FBS and 1μg/ml insulin. Subsequently, the cells were re-fed every 48 h with DMED containing 10%FBS. For differentiation of skin fibroblasts 0.2mM indomethacin was added to the cocktail as described earlier (Lysy et al., 2007); cells were treated with this cocktail for 28 days. Medium was changed every 4 days. At completion of differentiation adipogenesis was examined by oil red O staining. The stimulation experiments are carried out after culturing the differentiated cells for 24h in DMEM supplemented 0.2% FBS.

Immunoprecipitation and Immunoblotting

Whole cell lysates from human muscle biopsies and cultured cell lines were prepared using cell lysis buffer (CST# 9803) supplemented with 4 μl/ml protease inhibitor cocktail, 10 μl/ml phosphate inhibitor, and 1mM PMSF. Bio-Rad protein assay regent was used to measure protein concentration. For immunocoprecipitation, 500 μg of total proteins were immunoprecipitated with the appropriate antibodies for overnight and protein A/G agarose beads for 1 h at 4°C. Proteins were solubilized in Laemmli sample loading buffer and resolved by SDS-PAGE. Protein was transferred to a PVDF membrane and immunoblotted using different primary antibodies. Appropriate HRP-conjugated secondary antibodies and enhanced chemiluminescence reagents were used to develop the blots.

Real-time PCR

Total RNA was isolated from human muscle biopsies and cultured cells using Trizol (Invitrogen) and cDNA were generated using High Capacity cDNA Reverse Transcription Kit (Applied Biosystem) according to the manufacturer’s instructions. Real-time-PCR amplification was performed using iQ SYBR Green Supermix (Bio-Rad) in Eppendorf Realplex2 Mastercycler after adjusting the threshold-cycle (Ct). Reactions were performed in quadruple with a β-actin internal control. Relative quantification of mRNA levels was expressed as fold increase relative to the control.

ChIP assay

ChIP was performed according to the manufacturer’s instructions (Cell Signaling Technology, # 9003S). Briefly, the chromatin/DNA proteins complexes were prepared from 3T3L1 cells treated with vehicle (PBS with 0.1% BSA) or Wnt-3a 30ng/ml for 2 h. Chemical cross-linking of DNA-proteins was carried out using 1% formaldehyde for 10 min at room temperature. The Cross-linking was quenched by addition of glycine (0.125 M) for 5 min at room temperature and followed by two washes with ice-cold PBS. Cells were scraped into PBS containing 1mM PMSF. The cell suspension was centrifuged and the pellet was mixed by inverting the tube every 3 min in buffer A + DTT+ PIC+ PMSF, incubated on ice for 10 min. The pellet (nuclei) was dissolved in 1.0 ml Buffer B +DTT + 5 μl of micrococcal nuclease and incubated for 20 min at 37°C with frequent mixing to digest DNA to length of approximately150–900 bp. The lysate was incubated with appropriate chip-grade TCF-4 (TCF7L2) antibody (sc-8631, Santa Cruz) to immunoprecipitate chromatin overnight at 4° C with rotation and followed by ChIP-grade protein G magnetic beads and incubation for 2 h at 4° C with rotation. The magnetic beads were washed using buffers supplied with the kit. The eluted DNA was purified and analyzed by PCR to determine the binding of TCF7L2 to IR promoter. PCR primers were designed by using the criteria described in the kit. The region of mouse IR promoter from −550 to −900 bp was taken to design the primers and was found to bear the putative TCF7L2 binding site (consensus sequence: AGATCAAAGGG) (van de Wetering et al., 1997) identified by online transcription factors binding prediction software (PROMO 3.0) of (http://alggen.lsi.upc.es) and (http://searchlauncher.bcm.tmc.edu/seq-search/alignmenftml). The following primers were used for Mouse IR-P-forward (F) 5′-TAAGACATTGGTAGCATAGGCTGT-3′, M ouse IR-P-reverse (R) 5 ′-CCAAGCACATTTTGTCTTTCTTTA-3′. T he control primers for mouse RPL30 were provided with the kit. Real-time-PCR amplification was performed using iQ SYBR Green Supermix (Bio-Rad) and using Eppendorf Realplex2 Mastercycler.

Electrophoretic mobility Shifting Assay

Electrophoretic mobility shift assay (EMSA) was carried out to determine the binding of TCF7L2 to 65 bp IR promoter region 5′ GTAGCATAGGCTGTAAAGAAAAACAGAATCAAAGGTAAAGAATTAAAGTGAATTCCCC ACGGGAG 3′ (corresponding to nt position -694 to -759) bearing one putative TCF7L2 recognition site was synthesized by W.M Keck Oligonucleotide Synthesis Facility of Yale University. The DNA fragment was labeled using the Biotin 3′ End DNA Labeling Kit (Thermo Scientific). Nuclear extracts were prepared using the NE-PER Nuclear Extraction Regent Kit (Thermo Scientific). The binding reactions (20 μl reaction mix) were incubated for 20 minutes at room temperature using the LightShift EMSA Kit (Thermo Scientific) with 20 fM concentration of labeled duplex DNA, 3μg of nuclear protein extract, binding buffer, 50ng/μl Poly (dI-dC), 0.5% NP-40, 2.5% glycerol and 5mM MgCl2 in concentrations based on the manufacturer’s recommendations. For antibody super shift, 1μl of TCF7L2 specific chip grade (C-19, Santa Cruz Biotechnology Inc.) and 1 μl of a nonspecific control IgG antibodies were incubated with nuclear extract proteins for 30 min at room temperature prior to addition of labeled probes. After mixing with 5 μl loading buffer, the reaction mix was immediately loaded on 5% native agarose gel in 0.5× TBE that was pre-electrophoresed in 0.5× TBE followed by electrophoresis and then electrophoretically transferred onto a positively charged nylon membrane (Thermo Scientific) in 0.5× TBE. The DNA-protein complex on membrane was cross-linked by UV and visualized with horseradish peroxidase–conjugated streptavidin kit (Thermo Scientific) according to the manufacturer’s instructions.

Construct and luciferase assay

pIRP-GLuc plasmid for IR promoter was constructed as described previously with some modifications (Dasgupta et al., 2011). Briefly, The IR promoter sequence was amplified by PCR using phINSRP-1 as a template with forward (5′-GGGGGAATTCGGCCATTGCACTCCA-3′) and reverse (5′-AATTGGATCCTGCGGGAGCGCGGGG-3′) primers. The PCR product was cloned into TOPO vector according to the manufacturer’s protocol (Topo TA Cloning kits, Invitrogen, USA). The cloned insert was removed from the TOPO vector by digestion with EcoRI/BamHI and subcloned in the pGLuc vector yielding pIRP-GLuc. The cloned IR promoter construct was confirmed by sequencing. HEK 293 cells were transfected for 48 h with pIRP-GLuc plasmid using X-tremeGENE HP DNA Transfection Reagent (Roche Applied Science, Indianapolis, IN, USA) and luciferase activity was measured after stimulation with recombinant Wnt3a for different time points using BioLux Gaussia Luciferase assay kit (New England BioLabs Inc. MA, USA).

Rescue studies

To further confirm that the transcription regulation of IR is Wnt dependent, LRP6 knockdown 3T3L1 cells were treated with LiCl for 24 h followed by stimulation with Wnt 3a for different time courses. We examined the effect of TCF on IR transcription by permanent expression of TCF7L2 in 3T3L1 cells and human skin fibroblasts. For this purpose a retrovirus vector (LXSN, Clonetech) containing TCF7L2 cDNA using plasmid 16514 (Addgene: (Korinek et al., 1997) as template was generated. Cells were infected with either TCF7L2 containing or empty vectors. Cells were serum starved and were stimulated by wnt3a for different time course and harvested for analysis. To rescue mTORC1-dependent IRS-1 seine phosphorylation, human mutant and wildtype fibroblasts were treated with 100 nM/ml mTORC1inhibitor rapamycin for 48 h and were stimulated with insulin.

Statistical analysis

All laboratory-based experimental data represent results from four independent experiments. Sample size was calculated to provide sufficient power to detect 30% difference in the basal rate of Glucose Disposal rate (GDR) between non-diabetic and diabetic subjects. Sample size justifications were based on ANOVA and an alpha of 0.05 using the following formula:

N=2[Zα+Zβ]2[s]2Δ2

Where N is the required sample size, Za and Zb are the false positive and false negative error rates tolerates (Za =1.96 (95%, 2 tailed); Zb=1.28 (90% 1-tailed), s2 is the variance based on prior studies (square of the standard deviation (SD) and D is the difference in the primary endpoint (GDR). Based on previous studies in normal control subjects (n=7)

N=2(3.24)2(23)2(76.8)2=1.88

Protein expression levels were quantified by densitometry of Western blots. All statistical analyses, including clinical data was carried out with two-factor analysis of variance (ANOVA). Statistical Comparisons were done using GraphPad Prism 5 software. A probability value of <0.05 was considered statistically significant. Error bars represent standard deviation (SD). Power calculation was carried out based on GDR (see supplement material).

Supplementary Material

01

Acknowledgments

We thank members of the kindred studied for their invaluable participation in this project. We thank Karin Allen, Carmen Galarza, Mikhail Smolgovsky, Ralph Jacob, Codruta Todeasa, and Aida Groszmann at the YCCI for their assistance in insulin clamp studies, Dr. Gary Kline and Yale Diabetes Endocrinology Research Center for isotope measurements and Prof. Graeme Bell (University of Chicago, Chicago, Illinois, USA) for providing the plasmid construct of human IR promoter phINSRP-1. This work was supported by National Institutes of Health Grants R01HL094784-01 and R01HL094574-03 (to A.M.) and The National Institute of Diabetes and Digestive and Kidney Diseases Grant T32 DK007058 (to R.S.S).

Footnotes

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