Elevated sensitivity to diet-induced obesity and insulin resistance in mice lacking 4E-BP1 and 4E-BP2 (original) (raw)
Increased adiposity in 4E-BP1 and 4E-BP2 DKO mice. Both 4E-BP1 and 4E-BP2 are highly expressed in tissues involved in glucose and lipid homeostasis, including adipose tissue, pancreas, liver, and muscle (10). Because both 4E-BP1 and 4E-BP2 are expressed in these tissues, we chose to investigate their roles in metabolism by generating 4E-BP1 and 4E-BP2 DKO mice. The deletion of 4E-BP1 and 4E-BP2 was confirmed by PCR and Western blotting (Supplemental Figure 1; supplemental material available online with this article; doi:10.1172/JCI29528DS1). Eight-week-old DKO and WT mice were fed a control normal chow diet and monitored over a period of 16 weeks. The weight gain was 29% higher in DKO than in WT mice (7.2 ± 0.2 g and 5.6 ± 0.2 g, respectively; P < 0.05). This difference was not a result of hyperphagia, as food intake was identical for WT and DKO mice (9.9 ± 0.6 kcal/d/mouse and 10.6 ± 0.6 kcal/d/mouse, respectively). The observed increase in body weight in DKO mice can be explained at least in part by an increase in fat accumulation (Figure 1A). Histological examination of epididymal adipocytes by hematoxylin and eosin staining showed a 40% increase in cell size in DKO mice (P < 0.01; Figure 1, B and C). The change in fat accumulation was associated with a rise in serum insulin and cholesterol levels (Table 1). Taken together, these data demonstrate that disruption of 4E-BP1 and 4E-BP2 results in the development of an obese phenotype.
Increased obesity and insulin resistance in 4E-BP1 and 4E-BP2 DKO mice. (A) Weight of the heart, gonadal (GWAT), retroperitoneal (RWAT), and inguinal (IWAT) white adipose tissue (n = 7–10) of WT and DKO mice fed HFD or normal chow diet (control). (B) Histological analysis of WT and DKO gonadal white adipose tissue from mice fed normal chow or HFD. Sections obtained from 4 different animals were stained with hematoxylin and eosin. Original magnification, ×400. (C) Adipocyte cell size in WT and DKO gonadal adipose tissue. (D) Mean VO2 in WT and DKO mice fed normal chow or HFD (n = 5). (E) Respiratory quotient, calculated as the ratio of VO2 to carbon dioxide production, in WT and DKO mice fed normal chow or HFD during light and dark phases. (F) Insulin resistance test. Fed mice given normal chow or HFD received an intraperitoneal injection of 0.75 U/kg insulin, and blood samples were taken at the indicated times (n = 7–14). (G) Glucose tolerance test. Mice fed normal chow or HFD were fasted overnight before receiving an intraperitoneal injection of 2 g/kg glucose, and blood samples were taken at the indicated times (n = 7–14). Data are mean ± SEM. *P < 0.05 versus WT (2-tailed, unpaired Student’s t test).
4E-BP1 and 4E-BP2 DKO mice display altered metabolic parameters
To further characterize the DKO phenotype, 8-week-old mice were subjected to a HFD for 16 weeks. While on the HFD, DKO mice gained 22% more weight than did WT mice (10.8 ± 0.4 g and 8.8 ± 0.4 g, respectively; P < 0.01). The increased body weight observed in the DKO mice was not due to increased food intake (WT, 12.7 ± 0.9 kcal/d/mouse; DKO, 12.6 ± 0.6 kcal/d/mouse), but can be explained by a large accumulation of white adipose tissue, mainly due to increased adipocyte size (Figure 1, A–C). The increased body weight of the DKO mice was also associated with increased liver weight (Figure 2B). Histological analysis of WT and DKO mouse livers clearly demonstrated the development of steatosis (Figure 2A), with a more pronounced accumulation of triglycerides (TGs) in DKO liver (Figure 2C), reinforcing the finding of increased systemic adiposity in DKO mice.
Liver histology in WT and DKO mice. (A) Histological analysis of WT and DKO liver from mice fed normal chow or HFD. Sections obtained from 4 different animals were stained with hematoxylin and eosin. Original magnification, ×400. (B) Liver weight of WT and DKO mice fed HFD or normal chow. (C) Liver total TG content. Data are mean ± SEM. *P < 0.05 versus WT (2-tailed, unpaired Student’s t test).
In DKO mice on the HFD, serum glucose, insulin, cholesterol, and HDL-cholesterol levels were all significantly increased (Table 1). Leptin, the product of the ob gene, is a satiety hormone that is mainly synthesized and secreted by white adipose tissue (24). Its production is regulated by nutritional signals, and it is an indicator of long-term energy surplus. Consistent with the increased adipose tissue mass, circulating leptin levels were increased in HFD-fed DKO mice by 75% compared with HFD-fed WT mice (P < 0.05; Table 1). The levels of circulating TGs and nonesterified fatty acids (NEFAs) were not significantly different (Table 1). The growth curves and HFD-induced fat accumulation and insulin resistance test results for single 4E-BP1 KO and 4E-BP2 KO mice (Supplemental Table 1, Supplemental Figures 2 and 3, and Supplemental Results) suggest a synergetic effect of 4E-BP1 and 4E-BP2 in the obesity phenotype of the DKO mice.
Importantly, the metabolic rate was reduced in the DKO mice, as indicated by a 10% decrease in oxygen consumption (VO2) compared with WT mice (Figure 1D). The normal chow diet–fed DKO mice demonstrated a significantly higher respiratory quotient (calculated as the ratio of VO2 to carbon dioxide production) than did normal chow diet–fed WT mice (0.868 ± 0.007 versus 0.817 ± 0.008, respectively; P < 0.0001, 2-way ANOVA; Figure 1E) during the light phase. This indicates that the DKO mice preferentially maintained carbohydrate utilization as their fuel source, possibly to preserve their fat reserves. In contrast, WT mice shifted toward a greater utilization of fat. In both WT and DKO mice fed the HFD, the respiratory quotient was reduced to a similar level, demonstrating a shift to fat utilization (Figure 1E).
Increased insulin resistance in DKO mice. It is thought that obesity and lipid accumulation are responsible for reduced insulin action, leading to the development of insulin resistance and metabolic syndrome (25, 26). Therefore, WT and DKO mice were treated with insulin to monitor insulin resistance. The glucose clearance level was similar in DKO and WT mice fed the normal chow diet (Figure 1F). HFD feeding caused the development of insulin resistance in both DKO and WT mice, as demonstrated by the reduced ability of insulin to decrease blood glucose levels (Figure 1F). The higher level of insulin found in DKO mice (Table 1) indicates that the 4E-BPs are required for a normal response to insulin. Indeed, we observed a greater impairment of insulin action in the DKO mice (Figure 1F). Furthermore, both WT and DKO mice fed the HFD displayed impaired glucose tolerance (Figure 1G), with a more pronounced effect in the DKO mice.
Deletion of 4E-BP1 and 4E-BP2 promotes S6K activity. A negative feedback loop by which S6K1 inhibits the PI3K/Akt pathway has been previously described (11, 12, 18, 27). S6K1 activation inhibits insulin signaling by phosphorylating IRS-1, which leads to its degradation and subsequent inhibition of signaling to Akt. Because the DKO mice display an obese phenotype opposite to the lean S6K1 KO phenotype (11), and because 4E-BPs and S6Ks are both targets of mTOR, we suspected that S6K1 could be deregulated in DKO mice. In control mice, Ser473Akt phosphorylation was dramatically increased in adipose tissue, muscle, and livers 10 minutes after insulin injection (Figure 3A and Supplemental Figure 4). However, Akt phosphorylation was increased to a much lesser extent in DKO mouse tissues. The decrease in Akt phosphorylation correlated with an increase in S6K activity, as indicated by increased S6K1 and S6 phosphorylation (Figure 3A and Supplemental Figure 4). HFD and obesity are associated with overactivation of the mTOR pathway, particularly with increased S6K activity in muscle and liver (12). S6K activity is also increased in 2 models of genetic obesity, ob/ob and K/KAy mice (11). HFD feeding caused an increase in S6K1, S6, and 4E-BP1 phosphorylation in WT mice (reflected by an increase in upper slower migrating bands; Figure 3A, compare lanes 1 and 5) indicative of elevated mTOR activity. Akt phosphorylation in HFD-fed DKO mice was drastically impaired, which is consistent with increased obesity, insulin resistance, and glucose intolerance (Figure 3A and Supplemental Figure 4). Overactivation of the mTOR pathway is also associated with a reduction in IRS-1 protein level (18, 27). Because S6K activity was increased in our model and is involved in the control of IRS-1 activity, we examined IRS-1 expression in adipose tissue. IRS-1 protein levels were reduced by approximately 35% in normal chow diet–fed DKO mice and approximately 60% in HFD-fed DKO mice as compared with WT mice (P < 0.05; Figure 3, B and C). Under such conditions, the insulin receptor–β expression in adipose tissue was reduced by approximately 35% in both WT and DKO mice fed the HFD (Supplemental Figure 5). These data suggest that reduced insulin signaling is a consequence of reduced IRS-1 protein levels.
Deletion of 4E-BP1 and 4E-BP2 led to impairment of insulin signaling. (A) Increased S6K activity and reduced Ser473 phosphorylation of Akt in muscle, liver, and adipose tissue from WT and DKO animals. Mice were fasted for 6 hours before receiving a 0.75 U/kg insulin injection in the tail vein. Animals were sacrificed 10 minutes later, and tissues were collected for Western blotting. An immunoblot of WT and DKO mouse tissue is shown. S473 pAkt, phosphorylated Ser473 of Akt; T389 pS6K1, phosphorylated Thr389 of S6K1; S240/244 pS6, phosphorylated Ser240/244 of S6. (B) Reduced IRS-1 expression in DKO adipose tissue. (C) Quantification of IRS-1 protein levels in WT and DKO adipose tissue. Levels were normalized to actin (n = 6–7). Data are mean ± SEM. *P < 0.05 versus WT (2-tailed, unpaired Student’s t test). (D) Immunoblot analysis showed increased inhibitory serine phosphorylation of IRS-1 (S636/639 pIRS-1 and S1101 pIRS-1) and sustained Thr389 phosphorylation of S6K1 in DKO MEFs following insulin treatment.
To examine the effect of 4E-BP1 and 4E-BP2 disruption on insulin signaling in a cell-autonomous system and to study the long-term effects of insulin, mouse embryonic fibroblasts (MEFs) were used. In DKO and WT MEFs, insulin treatment led to an increase in S6K1 phosphorylation (Figure 3D). However, S6K1 phosphorylation was already elevated in serum-starved DKO MEFs (Figure 3D, compare lanes 1 and 6), and the kinetics of the insulin-stimulated S6K1 phosphorylation were accelerated — with maximum phosphorylation at 20 minutes after insulin administration — and maintained for 8 hours (Figure 3D, compare lanes 2 and 7). In contrast, in WT MEFs, S6K1 phosphorylation was sustained up to 1 hour after insulin treatment and then returned to basal levels by 4 hours after treatment (Figure 3D). Constitutive activation of S6K1 in DKO MEFs was also associated with reduced phosphorylation of Akt, indicating that the DKO MEFs were less sensitive to insulin treatment (Figure 3D, compare lanes 2 and 7). Because S6K1 activation is associated with obesity and increased serine phosphorylation of IRS-1 (11, 12), IRS-1 phosphorylation was also examined. In DKO MEFs the accelerated S6K1 phosphorylation correlated with a faster phosphorylation of IRS-1 on Ser636/639 and Ser1101 residues (Figure 3D, compare lanes 2 and 7), explaining the reduced activation of Akt. Both of these sites are associated with inhibition of insulin signaling (23, 28), and Ser636/639 phosphorylation has been demonstrated to be associated with type 2 diabetes (23).
Lipid metabolism is altered in DKO adipose tissue. Next, in order to determine whether fat accumulation in DKO mice is a consequence of increased lipogenesis or reduced lipolysis, TG synthesis, glycerol release, and reesterification were examined in isolated gonadal adipose tissue. Basal TG synthesis was similar in DKO and WT mice. Insulin caused a 2.5-fold increase in TG synthesis in WT mice, whereas the same treatment in DKO mice fed the normal chow diet led to only a 1.7-fold increase (Figure 4A). This result is consistent with the impaired insulin signaling observed in DKO mice (Figure 3). Treatment with isoproterenol, a specific activator of lipolysis, reduced TG synthesis to the same extent in WT and DKO mice (Figure 4A). Lipolysis was then examined using glycerol release as an index of lipid breakdown. Basal lipolysis was lower in adipose tissue of DKO mice compared with WT mice (approximately 32% on chow diet and 47% on HFD as compared to WT levels; P < 0.05; Figure 4B). The decreased lipolysis could be a major contributor to the development of obesity. Upon treatment with insulin, lipolysis was inhibited in WT mice, but not in DKO mice. This is also consistent with our finding that DKO mice were insulin resistant (Figure 1F). Isoproterenol treatment increased lipolysis to the same extent in both WT and DKO mice (Figure 4B). Further investigation revealed no significant differences in the levels of 2 proteins involved in the control of lipolysis in adipocytes, hormone-sensitive lipase (HSL), Ser565-phosphorylated HSL (29, 30), and perilipin (31, 32), in DKO adipose tissue (Figure 4C). Thus, alterations in components of the lipolysis pathway are not responsible for decreased basal lipolysis in DKO adipose tissue.
Deletion of 4E-BP1 and 4E-BP2 altered lipid metabolism. Adipose tissue was collected from fasted mice on control or high fat diet and incubated in 5 mM glucose Ca2+-free Krebs-Ringer buffer containing 1% fatty acid-free BSA (basal), with 100 nM insulin or 10 μM isoproterenol as described in Methods (n = 5). (A) TG synthesis was measured in isolated adipose tissue. (B) Lipolysis was reduced in DKO adipose tissue. (C) Western blot analysis of lipolysis-associated proteins. S565 pHSL, phosphorylated Ser565 of HSL. (D) Increased fatty acid reesterification in DKO adipose tissue. Data are mean ± SEM. *P < 0.05 versus WT (2-way ANOVA).
Fatty acid reesterification could also contribute to the difference in fat accumulation between WT and DKO mice. During lipolysis, NEFAs and glycerol are released from adipocyte TGs (at a ratio of 3:1). Although the glycerol cannot be reused by the adipocytes, the NEFAs can be reesterified to produce new TGs. Thus, the ratio of glycerol levels to NEFA levels provides an index of reesterification (33). In mice fed the normal chow diet, approximately 75%–80% of fatty acids were efficiently taken up by adipocytes and stored as TGs, with no difference between WT and DKO mice (Figure 4D). Fatty acid reesterification was decreased by approximately 30%–40% in mice fed the HFD due to an increase in fatty acids made readily available by the diet. Insulin dramatically reduced fatty acid reesterification in WT mice fed the HFD, but reesterification was maintained at 35% in HFD-fed DKO mice (Figure 4D). Taken together, these data demonstrate that reduced lipolysis and increased insulin-stimulated reesterification cause an increase in fat accumulation in DKO mice.
Increased adiposity is associated with enhanced adipocyte differentiation in DKO cells. Because increased adiposity in the DKO mice could also be due to increased adipocyte differentiation, adipogenesis was investigated using primary MEFs. Adipocyte differentiation was induced in MEFs using 3-isobutyl-1-methylxanthine, dexamethasone, and indomethacin (IBMX/DEX/IND), and lipid droplets were detected with oil red O staining (34). Adipocyte differentiation was increased in DKO MEFs compared with WT MEFs (Figure 5A). Remarkably, the difference in differentiation was not caused by increased DKO MEF proliferation, as this was identical to that of WT MEFs under standard culture conditions (data not shown). The extent of differentiation for WT, DKO, 4E-BP1 KO, and 4E-BP2 KO cells was determined by quantifying oil red O staining (Figure 5, A and B, and Supplemental Figure 6) and leptin secretion (Figure 5C and Supplemental Figure 6). TG content was 3-fold higher (P < 0.05; Figure 5B), and leptin secretion was twice as high, in the DKO-differentiated MEFs as in WT MEFs after 10 days of differentiation (P < 0.01; Figure 5C).
Deletion of 4E-BP1 and 4E-BP2 promotes adipocyte differentiation. MEFs from WT and DKO embryos were grown to confluence and differentiated into adipocytes as described in Methods. (A) Microscopic images of MEFs following induction of adipogenesis. Original magnification, ×400. Lipid droplets were stained with oil red O solution. (B) Quantification of lipid incorporation by measurement of the intensity of oil red O staining in WT and DKO MEFs at day 10 of the adipocyte differentiation process. (C) Leptin secretion in the culture medium was measured throughout induction of adipogenesis and quantified. (D) RT-PCR analysis showing C/EBPβ, C/EBPδ, C/EBPα, and PPARγ expression following differentiation of WT and DKO MEFs. (E) RT-PCR analysis showing C/EBPβ, C/EBPδ, C/EBPα, and PPARγ expression following differentiation of preadipocytes isolated from WT and DKO adipose tissue. (F) Lipid incorporation was quantified by measuring the intensity of oil red O staining in WT and DKO preadipocytes at day 10 of the adipocyte differentiation process. Data are mean ± SEM from 4 different experiments. *P < 0.05 versus WT (2-tailed, unpaired Student’s t test).
Next, we investigated the expression of PPARγ, which is a key regulator of adipocyte differentiation (35). PPARγ is induced prior to the transcriptional activation of most adipocyte-specific genes, and its ectopic expression in nonadipogenic fibroblasts is sufficient to initiate adipocyte differentiation (35). Furthermore, PPARγ is required to promote fat cell differentiation, as PPARγ-deficient cells fail to differentiate into adipocytes (36, 37). PPARγ mRNA was detected 2 days after onset of differentiation in WT MEFs following IBMX/DEX/IND treatment and was further elevated at days 4 and 6 (Figure 5D). Concomitant with the increased lipid accumulation and leptin secretion, 4E-BP1 and 4E-BP2 deletion resulted in higher expression of PPARγ mRNA in MEFs, reflecting the increased number of adipocyte-differentiated cells (Figure 5D). The expression of PPARγ and adipocyte differentiation is controlled by the members of the C/EBP family (35, 38, 39). We therefore examined the expression of C/EBP family mRNAs following induction of adipocyte differentiation by IBMX/DEX/IND in MEFs. Expression of C/EBPα mRNA, which is required for adipocyte differentiation, was increased in DKO MEFs compared with WT MEFs (Figure 5D). Primary preadipocytes freshly isolated from gonadal adipose tissue were also induced to differentiate into mature adipocytes. Differentiation, as well as PPARγ and C/EBPα expression, were markedly increased in the DKO-isolated adipocytes (Figure 5, E and F). The onset and level of C/EBPβ mRNA expression was similar in DKO and WT MEFs (Figure 5D). In contrast, C/EBPδ expression, which is required for the subsequent expression of C/EBPα and PPARγ in the later phases of differentiation (35), was increased in DKO MEFs during the early stages of differentiation (Figure 5D).
To confirm the role of 4E-BP1 and 4E-BP2 in the control of adipocyte differentiation, 4E-BP1 was transfected in 4E-BP1 KO MEFs and 4E-BP2 was transfected in 4E-BP2 KO MEFs prior to differentiation. Single deletion of 4E-BP1 or 4E-BP2 in MEFs led to increased adipocyte differentiation, as demonstrated by increased C/EBPδ, C/EBPα, and PPARγ mRNA expression (Supplemental Figure 6). Reintroduction of 4E-BP1 or 4E-BP2 in the respective KO MEFs caused reduced expression of the former transcription factors to a level similar to that seen in WT mice (Supplemental Figure 6). Taken together, these results implicate 4E-BPs in the control of C/EBPδ expression and suggest that increased expression of C/EBPδ in the DKO cells may be responsible for the subsequent increased expression of PPARγ and C/EBPα and increased adipogenesis.