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Gestational Protein Restriction Impairs Glucose Disposal in the Gastrocnemius Muscles of Female Rats Chellakkan S. Blesson,1 Vijayakumar Chinnathambi,3 Sathish Kumar,3 and Chandrasekhar Yallampalli2 1

Division for Reproductive Endocrinology and Infertility and 2Basic Sciences Perinatology Research Laboratories, Department of Obstetrics and Gynecology, Baylor College of Medicine, Houston, Texas 77030; and 3Department of Obstetrics & Gynecology, University of Texas Medical Branch, Galveston, Texas 77555

Gestational low-protein (LP) diet causes hyperglycemia and insulin resistance in adult offspring, but the mechanism is not clearly understood. In this study, we explored the role of insulin signaling in gastrocnemius muscles of gestational LP-exposed female offspring. Pregnant rats were fed a control (20% protein) or an isocaloric LP (6%) diet from gestational day 4 until delivery. Normal diet was given to mothers after delivery and to pups after weaning until necropsy. Offspring were euthanized at 4 months, and gastrocnemius muscles were treated with insulin ex vivo for 30 minutes. Messenger RNA and protein levels of molecules involved in insulin signaling were assessed at 4 months. LP females were smaller at birth but showed rapid catchup growth by 4 weeks. Glucose tolerance test in LP offspring at 3 months showed elevated serum glucose levels (P , 0.01; glycemia D area under the curve 342 6 28 in LP vs 155 6 23 in controls, mmol/L * 120 minutes) without any change in insulin levels. In gastrocnemius muscles, LP rats showed reduced tyrosine phosphorylation of insulin receptor substrate 1 upon insulin stimulation due to the overexpression of tyrosine phosphatase SHP-2, but serine phosphorylation was unaffected. Furthermore, insulin-induced phosphorylation of Akt, glycogen synthase kinase (GSK)–3a, and GSK-3b was diminished in LP rats, and they displayed an increased basal phosphorylation (inactive form) of glycogen synthase. Our study shows that gestational protein restriction causes peripheral insulin resistance by a series of phosphorylation defects in skeletal muscle in a mechanism involving insulin receptor substrate 1, SHP-2, Akt, GSK-3, and glycogen synthase causing dysfunctional GSK-3 signaling and increased stored glycogen, leading to distorted glucose homeostasis. (Endocrinology 158: 756–767, 2017)

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ype 2 diabetes (T2D) affects 29 million Americans (1) and 347 million worldwide (2), but its etiology and pathophysiology are not clearly understood. T2D originates from various genetic and environmental factors. Gestational exposure to a low-protein (LP) diet programs the offspring to become susceptible to metabolic diseases, including diabetes, during their adult life. There is a strong association between in utero growth retardation and the development of T2D (3, 4). We have recently developed a rat model using 6% protein during gestational days 4 to 21. This model of maternal LP-induced glucose intolerance and insulin resistance in offspring is notable for the absence of obesity (5).

Protein restriction during gestation and lactation causes insulin resistance in both male and female offspring (6–8), and the onset and severity of the disease vary between the sexes (9). Data from this model show that at 20 weeks, male offspring of dams fed an LP diet were relatively insulin resistant and hyperinsulinemic compared with the control animals, but the female offspring of the same age were not affected (10). At 15 months, male offspring showed impaired glucose tolerance and higher plasma insulin levels (6), but females of the same age showed neither glucose intolerance nor elevated insulin levels (11). Furthermore, at 17 months, males developed frank diabetes (7) but females developed

ISSN Print 0013-7227 ISSN Online 1945-7170 Printed in USA Copyright © 2017 Endocrine Society Received 13 September 2016. Accepted 23 January 2017. First Published Online 30 January 2017

Abbreviations: Glut4, glucose transporter type 4; GS, glycogen synthase; GSK, glycogen synthase kinase; HOMA-IR, homeostatic model assessment–insulin resistance; HOMA-IS, homeostatic model assessment–insulin sensitivity; IR, insulin receptor; IRS-1, insulin receptor substrate 1; LP, low protein; mRNA, messenger RNA; OGTT, oral glucose tolerance test; pGS, phospho-GS; pGSK, phospho-GSK; T2D, type 2 diabetes.

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hyperinsulinemia only when they reached 21 months old (8). However, this rat model was obese, and their dietary manipulations involved both gestational and lactational intervention. Importantly, this model and ours exhibited sex differences in the disease presentation. Our lean diabetes rat model showed that females develop glucose intolerance and insulin resistance earlier with faster disease progression compared with males (5). In our previous study, we showed that the insulin resistance in LP-programmed males is due to a series of phosphorylation defects leading to impaired glucose transporter 4 (Glut4) translocation (12). Our current aim was to investigate the role of insulin signaling molecules in LP-programmed female offspring to understand their role in LP-induced glucose intolerance and if the mechanisms are different from that of males. We hypothesized that the insulin signaling cascade pertaining to glucose transport and storage will be impaired in LPprogrammed female offspring in gastrocnemius muscle.

Materials and Methods Animals Pregnant (day 4) Wistar rats were bought from Harlan Sprague Dawley (Indianapolis, IN). Rats were given unlimited access to food and water and were housed in a temperature-controlled room (23°C) with a 12:12-hour light/dark cycle. Pregnant rats were fed with a control (20% protein, n = 4) or an isocaloric LP (6%, n = 4) diet (Harlan Teklad, Indianapolis, IN) from day 4 of pregnancy until delivery. Normal diet was given to mothers after delivery until weaning, and pups were given a normal diet after weaning. Pups with extreme weights were culled on day 1 after birth to make the litter size to 10 pups per mother, ensuring equal nutrient access for all offspring. Only female pups were used for the current study. All rats were euthanized during diestrus phase at 4 months, and tissues were collected, snap frozen in liquid nitrogen, and stored at 280°C until analysis. All experimental procedures involving rats were approved by the Institutional Animal Care and Use Committee of the University of Texas Medical Branch.

Oral glucose tolerance test Overnight fasted rats were given glucose orally (2 g/kg body weight) at 3 months of age. Blood was collected at 0, 30, 60, and 120 minutes by orbital sinus puncture. Plasma was isolated from the blood by centrifugation and stored at 280°C until analysis. Plasma glucose levels were measured using a glucose colorimetric assay (Cayman Chemical Company, Ann Arbor, MI) following the manufacturer’s protocol as reported earlier (12).

Insulin measurements Insulin was measured using a rat insulin enzyme-linked immunosorbent assay kit (Mercodia, Uppsala, Sweden) following the manufacturer’s instruction as reported earlier (10). Absorbance was read at 450 nm using a BMG CLARIOstar plate reader (BMG Labtech Gmbh, Ortenberg, Germany), and the results were calculated with cubic spline regression fit using omega CLARIOstar data analysis software.

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Homeostatic model assessment Homeostatic model assessment–insulin resistance (HOMAIR) and homeostatic model assessment–insulin sensitivity (HOMA-IS) were calculated to assess insulin resistance and insulin sensitivity of control and LP rats using the following equations (13). HOMA-IR ¼ ½Fasting Glucoseðmg=dLÞ 3 Fasting InsulinðmU=LÞ=405 HOMA-IS ¼ 10; 000=½Fasting Glucoseðmg=dLÞ 3 Fasting InsulinðmU=LÞ

Ex vivo insulin treatment Gastrocnemius muscle strips from control and LP-programmed rats were preincubated for 30 minutes with Krebs-Henseleit bicarbonate buffer (120 mM NaCl, 4.7 mM KCl, 1.25 mM MgSO4, 1.2 mM KH2PO4, 2.5 mM CaCl2, and 25 mM NaHCO3, pH 7.4) containing 5.5 mM glucose, 2 mM sodium pyruvate, and 0.1% bovine serum albumin followed by incubation with and without insulin (10 mU/mL; Sigma, St Louis, MO) for 30 minutes (12, 14). The incubations were gassed continually with 95% O2 and 5% CO2. After 30 minutes of incubation, muscle strips were blotted rapidly on filter paper and frozen in liquid nitrogen and stored at 280°C until analysis.

Protein extraction Total and membrane protein extraction from gastrocnemius muscles was performed as reported earlier (10). Briefly, tissues were weighed and homogenized in 13 RIPA buffer (Cell Signaling Technology, Danvers, MA) containing 1 mM phenylmethylsulfonyl fluoride, protease inhibitor cocktail (Roche, Branford, CT), and phosphatase inhibitor cocktails (Sigma). The lysates were then sonicated and centrifuged (14,000g for 10 minutes), and the supernatants were stored at 280°C until further analysis. Membrane proteins were isolated using a plasma membrane protein extraction kit (Biovision, Milpitas, CA) following the manufacturer’s instruction for plasma membrane proteins. Proteins were quantified using the Pierce BCA kit (Pierce Biotechnology, Rockford, IL).

Western blot Protein extract (10 to 30 mg) from each sample was resolved on 4% to 15% precast gradient polyacrylamide gels (MiniPROTEAN TGX Precast Gels; Bio-Rad, Hercules, CA). Proteins from the gel were transferred to a polyvinylidine fluoride membrane (Millipore, Billerica, MA). Primary antibodies were incubated overnight at 4°C after blocking the membranes in 5% bovine serum albumin or nonfat dried milk in Tris buffered saline containing 0.1% Tween or Odyssey blocking buffer (insulin receptor b and Akt; LI-COR, Lincoln, NE) for 1 hour at room temperature. Summaries of the primary antibodies and their dilutions are as given here. Antibodies for insulin receptor substrate 1 (IRS-1; #3407), phospho–IRS-1 (Ser307, #2381; Ser318, #5610; and Ser612, #3203), Akt (#9272), phospho– AS-160 (Thr642, #4288), phospho-Akt (Ser473, #9271 and Thr308, #9275), SHP-2 (#9793), glycogen synthase kinase (GSK)–3a (#4337), phospho-GSK-3a (pGSK-3a) (#9316), GSK-3b (#12456), phospho-GSK-3b (pGSK-3b) (#5558), glycogen synthase (GS; #3893), and phospho-GS (pGS) (#3891) were diluted 1:1000 and obtained from Cell Signaling Technology.

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Insulin receptor (IR) b antibody (cat. no. 610109, 1:500) was obtained from BD Biosciences (Franklin Lakes, NJ). AS-160 antibody (#07-741, 1:1000) was obtained from Millipore. Glut4 (ab654, 1:2000), phospho-IR Tyr972 (ab5678, 1:1000), phospho–IRS-1 Tyr612 (ab66153, 1:500), phospho–IRS-1 Tyr896 (ab46800, 1:500), a1 sodium potassium ATPase (ab7671, 1:5000), and glyceraldehyde 3-phosphate dehydrogenase (ab9485, 1:2500) antibodies were obtained from Abcam (Cambridge, MA). After primary antibody incubations, membranes were washed and incubated for 60 minutes at room temperature with horseradish peroxidase or IRDye 800CWconjugated (LI-COR; for IRb and Akt) secondary antibodies. Membranes were washed and incubated in ECL Western blotting detection reagents (Pierce Biotechnology) for 1 minute and imaged using the Odyssey Fc imaging system (LI-COR). Membranes were then reprobed for glyceraldehyde 3-phosphate dehydrogenase. Densitometric analyses were performed using Image Studio software from LI-COR.

Glycogen estimation Stored glycogen from gastrocnemius muscles was measured using a glycogen assay kit (Cayman Chemical Company) according to the manufacturer’s protocol. Briefly, flash-frozen tissue was thawed and weighed. Tissue (;150 mg) was minced and homogenized in 1 mL assay buffer containing protease inhibitors. The lysate was spun at 800g for 10 minutes at 4°C. The supernatant was removed and diluted (1:2) with assay buffer, and 10 mL of standards/samples was added to designated wells. Each sample well had its own sample background well. Hydrolysis enzyme solution (50 mL) was added to all sample and standard wells, and glycogen hydrolysis buffer (50 mL) was added to background wells. All assays were performed in duplicates. The plate was then incubated at 37°C for 30 minutes. After the incubation, developer solution was added to all the wells and further incubated for 15 minutes at 37°C. The plate was then read using an excitation wavelength of 530 nm and an emission wavelength of 585 nm using a BMG CLARIOstar plate reader (BMG Labtech Gmbh), and the results were calculated with linear regression fit using omega CLARIOstar data analysis software.

Real-time quantitative polymerase chain reaction Total RNA was isolated from gastrocnemius muscles using TRIzol reagent (Life Technologies, Carlsbad, CA) and further purified with the RNeasy clean-up kit (Qiagen, Valencia, CA). All RNA samples were treated with DNase. RNA concentration and purity were determined using an ND-1000 model Nanodrop spectrophotometer (Thermo Fisher Scientific, Newark, DE). In total, 2 mg total RNA was reverse-transcribed using a modified Maloney murine leukemia virus-derived reverse transcription (New England Biolabs, Ipswich, MA) and random hexamer primers (Life Technologies) as reported earlier (12). Complementary DNA was amplified by real-time polymerase chain reaction using SYBR Green (Bio-Rad) in a CFX96 model real-time thermal cycler (Bio-Rad). Specific pairs of primers (IDT, Coralville, IA) were used for each gene amplification. Polymerase chain reaction conditions used were 10 minutes at 95°C for 1 cycle, 15 seconds at 95°C, 30 seconds at 60°C, and 15 seconds at 72°C for 40 cycles, followed by a melt curve analysis (0.5°C/5 seconds from 65°C to 95°C). Results were calculated using the 2–DDCT method and expressed as fold changes of expression of genes of interest. All reactions were

Figure 1. Rat weights and plasma glucose and insulin levels during a glucose tolerance test in control and LP offspring. (a) Weight gain by female pups (n = 9 to 23) born to control and LP diet-fed mothers. (b, c) Glucose (D AUC, mmol/L * 120 minutes) and (d, e) insulin levels (D AUC, pmol/L * 120 minutes) for control and gestational LP diet-fed offspring. Blood samples were collected at 0, 30, 60, and 120 minutes following oral glucose (2 g/kg body weight) administration. The overall plasma glucose levels are expressed as D AUC. D AUC is calculated as a difference between the total AUC and the baseline values calculated after the oral glucose tolerance test at 3 months. Results are presented as mean 6 SEM, n = 4 to 5 per group. *P , 0.05, **P , 0.01, and ***P , 0.001. AUC, area under the curve.

performed in duplicate, and cyclophilin A was used as internal control.

Statistical analyses Statistical analyses were performed using GraphPad Prism software (GraphPad Software, La Jolla, CA). Data are

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presented as mean 6 SEM. Comparison between two groups was performed using the unpaired Student t test. When comparisons between multiple groups with two factors were done, statistics was performed with two-way analysis of variance followed by the Bonferroni test. Differences were considered significant when P , 0.05.

Results LP females were small at birth but exhibited rapid catch-up growth Dams fed with the LP diet gave birth to significantly (P , 0.001) smaller pups (5.1 6 0.1 g) compared with mothers fed with the control diet (6.2 6 0.1 g) when weighed on day 1. LP offspring continued to be smaller until 3 weeks, but they showed rapid catchup growth and their weights were similar to that of controls (control, 80.5 6 1.4 g; LP, 77.6 6 0.7 g) by 4 weeks. Both controls and LP female offspring maintained similar body weights until the termination of the experiment at 4 months (control, 293 6 13 g; LP, 296 6 8 g) [Fig. 1(a)]. LP-programmed females showed impaired glucose tolerance Fasting glucose levels did not change between controls and LP-programmed female offspring (controls, 6.2 6 0.2 mmol/L; LP, 5.9 6 0.2 mmol/L). Oral glucose tolerance test (OGTT) showed that plasma glucose levels

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in LP offspring displayed an increasing tendency at 30 minutes (LP, 10.0 6 0.6 mmol/L; control, 8.5 6 0.5 mmol/L), followed by a significant increase at 60 minutes (LP, 9.8 6 0.7 mmol/L; control, 7.3 6 0.3 mmol/L; P , 0.01) and no change at 120 minutes (LP, 7.9 6 0.6 mmol/L; control, 8.0 6 0.3 mmol/L) compared with controls [Fig. 1(b)]. The overall differences in the plasma glucose concentrations after administering OGTT were analyzed by calculating the D glycemia area under the curve (mmol/L * 120 minutes), which showed that overall glucose levels were significantly (P , 0.01) higher (glycemia: 342 6 28) in LP offspring compared with controls (glycemia: 155 6 23) [Fig. 1(c)]. Insulin levels during fasting (control, 8 6 1 pmol/L; LP, 9 6 3 pmol/L) and OGTT varied between time points but did not show any differences between control and LP rats at 30 minutes (control, 200 6 36 pmol/L; LP, 204 6 39 pmol/L), 60 minutes (control, 125 6 30 pmol/L; LP, 140 6 20 pmol/L), and 120 minutes (control, 135 6 27 pmol/L; LP, 78 6 28 pmol/L) [Fig. 1(d)]. The overall glucose-induced insulin response expressed as D insulin area under the curve (pmol/L * 120 minutes) after OGTT also did not show any differences between the LP offspring (14,842 6 1274) and controls (14,149 6 3054) [Fig. 1(e)]. HOMAIR (control, 0.28 6 0.05; LP, 0.26 6 0.12) and HOMA-IS (control, 82.7 6 27; LP, 73.8 6 28) did not show any difference between the control and LP offspring.

Figure 2. Basal mRNA and protein expression levels of key molecules involved in insulin signaling cascade pertaining to glucose transport as quantified by quantitative polymerase chain reaction (qPCR) and Western blot. (a) mRNA expression of IRb, IRS-1, Akt, AS-160, and Glut4. qPCR data were normalized with the housekeeping gene cyclophilin A. (b) Representative blots and their densitometric analyses showing the protein levels of IRb, IRS-1, Akt, AS-160, and Glut4. Glyceraldehyde 3-phosphate dehydrogenase was used as a loading control for Western blots, and its values were used for normalization. Results are presented as mean 6 SEM, n = 4 to 5 per group. *P , 0.05; **P , 0.01.

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LP female pups had high basal AS-160 messenger RNA and protein levels The basal expression of key molecules involved in insulin signaling pertaining to glucose transport in skeletal muscles, such as IR (b subunit), IRS-1, Akt, AS160, and Glut4, was evaluated in gastrocnemius muscles. Results show that AS-160 messenger RNA (mRNA) and protein levels were significantly upregulated (P , 0.05) in LP-programmed females compared with controls (Fig. 2). All other genes that were probed did not show any changes except the mRNA expression of IRS-1, which was downregulated, but its protein levels did not show any difference compared with controls (Fig. 2). LP programming affected tyrosine and not serine phosphorylation of IRS-1 Glucose transport in skeletal muscle is initiated by insulin through the phosphorylation of IR and IRS-1. We explored if LP programming in female offspring affected the insulin-induced phosphorylation of IRs and IRS-1 (Fig. 3). Our results show that insulin phosphorylated IR at Tyr974 in both controls and LP female offspring

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[Fig. 3(a) and 3(b)]. Furthermore, insulin also phosphorylated IRS-1 at both Tyr608 (P , 0.001) and Tyr895 (P , 0.01) positions in the controls [Fig. 3(a), 3(d), and 3(e)]. However, in LP-programmed females, insulin did not increase tyrosine phosphorylation [Fig. 3(a), 3(d), and 3(e)]. Interestingly, insulin induced the phosphorylation of IRS-1 at Ser318 and Ser612 positions in both controls and LP rats [Fig. 3(a), 3(f), and 3(g)], showing that LP programming in female rats specifically affects only tyrosine phosphorylation and not serine phosphorylation of IRS-1. SHP2 was responsible for the dephosphorylation of IRS-1 Tyr phosphorylation To identify the cause for the failure of insulin to induce IRS-1 phosphorylation in LP-programmed female offspring, we screened for possible candidates that could impair IRS phosphorylation. We hypothesized that three possible types of molecules could impede phosphorylation: (1) site access denying competitors, (2) low expression/activation of kinases, or (3) high expression/ activation of phosphatases. We screened for the mRNA

Figure 3. Insulin-induced phosphorylation of insulin receptor and IRS-1. Ex vivo incubation of gastrocnemius muscles was performed with insulin (10 mU/mL) for 30 minutes. (a) Representative blots showing bands for pIRb Tyr974, total IR, pIRS-1 (Tyr608), pIRS-1 (Tyr895), pIRS-1 (Ser318), pIRS-1 (Ser612), and total IRS-1 levels, along with glyceraldehyde 3-phosphate dehydrogenase (GAPDH). Histograms from the densitometric analyses of blots showing (b) pIRb Tyr974, (c) total IR, (d) pIRS-1 (Tyr608), (e) pIRS-1 (Tyr895), (f) pIRS-1 (Ser318), (g) pIRS-1 (Ser612), and (h) total IRS-1 levels in the presence or absence of insulin. Values were normalized with GAPDH. Results are presented as mean 6 SEM, n = 3 to 4 per group. *P , 0.05.

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and protein expression of known adapter molecules that could block and deny access of the IRS-1 phosphorylation site, such as Shc and several SOCS isoforms. Their levels were not altered (data not shown). We also screened for various kinases such as adenosine monophosphate–activated protein kinase (mRNA) and several variants of protein kinase C (mRNA and proteins) and found that they were not differentially regulated (data not shown). We then screened for the key tyrosine phosphatase SHP-2, also known as tyrosine-protein phosphatase nonreceptor type 11. We found that SHP2 mRNA expression [Fig. 4(a) and 4(b)] and protein levels [Fig. 4(c) and 4(d)] were significantly upregulated, suggesting that SHP-2 could play a key role in the selective dephosphorylation of tyrosine residues in IRS-1. IRS-1 Tyr dephosphorylation affected Akt phosphorylation but not Glut4 translocation Akt is downstream to IRS-1 in the insulin signaling cascade following PI3 kinase. Insulin-induced PI3 kinase activation remained unaffected in LP rats, and the phosphorylation was similar in controls and LP female offspring (data not shown). Also, insulin treatment did not affect the gene expression of PI3 kinase (data not shown). However, insulin-induced Akt phosphorylation was affected at both Ser473 and Thr308 in LP females compared with controls [Fig. 5(a–c)]. Total Akt expression, however, was not altered by insulin treatment [Fig. 5(d)]. In skeletal muscles, Glut4 promoted the uptake of glucose by translocating to the plasma membrane. AS-160 is an Akt substrate involved in the regulation of Glut4 translocation. Insulin activated the phosphorylation of AS-160 in both control and LP groups [Fig. 5(a) and 5(e)]. Next, we assessed the insulininduced translocation of Glut4 using the membrane fraction isolated from gastrocnemius muscles after treating with insulin. Interestingly, insulin treatment induced the translocation of Glut4 in LP and control groups, showing that the insulin-induced Glut4-mediated glucose transport mechanism was not affected in LP females [Fig. 5(a) and 5(h)]. GSK-3 and GS phosphorylation levels were affected in LP female rats GSK-3 signaling cascade is downstream to Akt and is involved in the regulation of blood glucose levels by facilitating glycogen synthesis. Our data show that insulin treatment significantly induced the phosphorylation of GSK-3a at Ser21 and GSK-3b at Ser9 in control females compared with vehicle treatment [Fig. 6(a–e)]. However, in LP-programmed females, insulin could not induce phosphorylation in both GSK-3a and GSK-3b isoforms [Fig. 6(a–e)]. Further downstream in the

Figure 4. Basal expression of tyrosine phosphatase SHP-2 in gastrocnemius muscles in control and LP females. (a) mRNA expression of SHP2-1 isoform and (b) SHP2-2 isoform, along with protein levels of SHP-2 showing (c) representative blots and (d) densitometric analysis. Glyceraldehyde 3-phosphate dehydrogenase was used as a loading control for Western blots. Results are presented as mean 6 SEM, n = 4 to 5 per group. *P , 0.05.

signaling cascade, GS phosphorylation at Ser641 position was also dysregulated [Fig. 6(a) and 6(f)]. Insulin induced the phosphorylation of GS in the controls, whereas in the LP offspring, there was no increase in GS phosphorylation. Interestingly, the LP-programmed offspring had a twofold higher basal GS phosphorylation compared with controls [Fig. 6(a) and 6(f)]. LP female offspring had more stored glycogen in the gastrocnemius muscle Increased basal phosphorylation of GS led us to postulate that glycogen storage and turnover could be affected in LP rats. So, we measured stored glycogen levels in gastrocnemius muscles in fasting rats. Our results show that LP programmed females had significantly more (P , 0.01) glycogen stored in their muscles with a threefold increase compared with controls (Fig. 7).

Discussion T2D has been historically attributed to lifestyle and genetics, but recent studies indicate that an adverse in utero environment is often associated with the development of glucose intolerance and insulin resistance later in life (15). We have characterized a gestational protein restriction

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Figure 5. Insulin-induced phosphorylation of Akt and AS-160 and the translocation of Glut4 in gastrocnemius muscles after incubation with insulin (10 mU/mL) for 30 minutes. (a) Representative blots showing bands for pAkt (Ser473), pAkt (Tyr 308), total Akt, pAS-160 (Thr642), total AS-160, and total Glut4, along with glyceraldehyde 3-phosphate dehydrogenase (GAPDH) (loading control) levels. Blot also showing membrane Glut4 and a subunit of Na+ K+ ATPase (loading control/membrane marker), along with membrane GAPDH (cytoplasmic marker shown to demonstrate the purity of membrane fraction). Densitometric analyses of (b) pAkt (Ser473), (c) pAkt (Tyr308), and (d) total Akt, along with (e) pAS-160 (Thr642) and (f) total AS-160 levels. (g) Total Glut4 and (h) membrane Glut4 showing their levels in control and LP rats in the presence or absence of insulin. Values were normalized with loading control, GAPDH for total Glut4, and a subunit of Na+ K+ ATPase for membrane Glut4. Results are presented as mean 6 SEM, n = 4 to 5 per group. *P , 0.05 and **P , 0.01.

rat model that results in glucose intolerance and insulin resistance during their adult life but is not accompanied by obesity (5). Although various aspects of T2D are well studied, the pathogenesis, progression, and sex differences of the fetal origins of T2D are poorly understood. Using this lean T2D rat model, we had shown earlier that both male and female LP-programmed offspring exhibit progressively worsening glucose intolerance and insulin resistance (5). In the current study, we show the mechanism causing compromised insulin signaling in skeletal muscles leading to insulin resistance in females. LP-programmed female rats weighed less compared with controls but exhibited catchup growth similar to our earlier study on males and other studies in rodents and humans (12, 15–17). Numerous studies in animal models and in humans suggest that catchup growth in itself could be related to causing insulin resistance (15, 18–20). Although the exact mechanism of how catchup growth leads to insulin resistance is unknown, several studies

suggest that it could be due to altered fat metabolism and obesity (16, 21–23). In the current study, rats were not obese but lean, and these programmed rats had a similar fat content to that of controls (4), and hence their insulin resistance cannot be attributed to obesity. The other likely explanation could be due to abnormal mitochondrial function similar to that reported in a calorie restriction rat model (24) along with other mechanisms involving insulin signaling. We used ex vivo insulin treatment studies to demonstrate the insulin signaling mechanisms in skeletal muscles (12, 14). Because in vivo insulin treatment could trigger responses from multiple signaling pathways and various systemic factors such as hormones, cytokines, and growth factors, we preferred ex vivo treatment to exclude the effect of various interactive factors and focus on direct insulin effects on skeletal muscles. Insulin action is transduced via IRS-1, a vital downstream regulator of insulin signaling. It is differentially

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Figure 6. Insulin-induced phosphorylation of GSK-3 and GS. Ex vivo incubation of gastrocnemius muscles was performed with insulin (10 mU/mL) for 30 minutes. (a) Representative blots showing bands for pGSK-3a (Ser21), GSK-3a, pGSK-3b (Ser9), GSK-3b, pGS (Ser641), and GS, along with glyceraldehyde 3-phosphate dehydrogenase (GAPDH) (loading control). Histograms from the densitometric analyses of blots showing (b) pGSK-3a (Ser21), (c) total GSK-3a, (d) pGSK-3b (Ser9), (e) GSK-3b, (f) pGS (Ser641), and (g) GS levels in the presence or absence of insulin. Values were normalized with GAPDH. Results are presented as mean 6 SEM, n = 3 to 4 per group. *P , 0.05 and **P , 0.01.

regulated by the phosphorylation of tyrosine and serine residues by various kinases, leading to diverse functions (25). The phosphorylation of IRS-1 is regulated by various receptor tyrosine kinases, protein tyrosine phosphatases, and serine/threonine kinases (25–27). Tyrosine phosphorylation of IRS-1 is vital for the normal insulindriven action (25). Phosphorylated IRS-1 can interact with protein tyrosine phosphatases such as SHP-2 (28) and can be specifically dephosphorylated by SHP-2 (29, 30). In the current study, we show that insulin-induced IRS-1 tyrosine phosphorylation is significantly decreased in LP-programmed rats. Such reduced insulin-simulated IRS-1 phosphorylation has been shown in other insulinresistant models (31). Furthermore, we also show that SHP-2 expression is upregulated in these rats, indicating that SHP-2 could be attributed to the selective dephosphorylation of tyrosine residues in IRS-1, thereby compromising downstream signaling in these animals. Phosphorylated IRS-1 triggers the phosphorylation of Akt via PI3 kinase and PDK1 (32, 33). Akt then mediates signaling cascades regulating glucose transport involving AS-160 and Glut4 (34–37). Our study shows that Akt phosphorylation is diminished in LP rats at both Ser473 and Thr308. However, AS-160 phosphorylation and Glut4 translocation were intact in these animals. Although phosphorylated Akt is considered the primary activator of AS-160, reports show that AS-160 could also be phosphorylated by adenosine monophosphate– activated protein kinase (38, 39). It is possible that adenosine monophosphate–activated protein kinase phosphorylates AS-160 to promote Glut4 translocation when Akt phosphorylation is compromised. This likely redundant mechanism could offer an explanation for the intact function of AS-160–mediated Glut4 translocation in female LP-programmed rats.

Insulin-induced Akt phosphorylation regulates glycogen synthesis via the GSK-3–GS signaling pathway (40). Our data show that insulin-induced phosphorylation of both GSK-3a and GSK-3b was reduced in LPprogrammed females. Under normal conditions, GSK-3 is constitutively active, and its inactivation by Akt phosphorylation leads to the activation of GS, which in turn facilitates glycogen synthesis (41, 42). Other kinases such as protein kinase A (43) and mitogen-activated protein (44) have also been shown to inactivate GSK-3, but the insulin-induced Akt pathway plays a dominant role in skeletal muscles (40, 41, 45, 46). Reduction in the phosphorylation of GSK-3 in LP rats leads to a hyperactive GSK-3 signaling cascade, and such GSK-3

Figure 7. Histogram showing the stored glycogen levels in the gastrocnemius muscles of female LP-programmed fasting rats. Results are presented as mean 6 SEM, n = 4 per group. **P , 0.01.

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Figure 8. Graphic representation of the impaired insulin signaling cascade in gastrocnemius muscles of 4-month-old maternal protein-restricted female offspring. The asterisk indicates impaired insulin-induced phosphorylations. Phosphorylation of IRS-1 (tyrosine 608 and 895), Akt (threonine 308 and tyrosine 473), GSK-3a (serine 21), and GSKb (serine 9) is severely decreased in LP rats. Increased basal phosphorylation of GS (blue arrow) is seen in LP rats. LP females also have higher fasting glycogen levels in their gastrocnemius muscles.

hyperactivity in skeletal muscles has been shown in several T2D models and humans (47–51). It is apparent from all these studies, including our observations, that impaired insulin-induced GSK-3 phosphorylation causes insulin resistance in skeletal muscles. Further downstream, insulin-induced dephosphorylation of GS leads to its activation, which catalyzes the incorporation of UDP-glucose to glycogen (52, 53). Our data show that in LP rats, the basal inactive form of GS was significantly higher compared with controls, suggesting a flawed baseline activation of GS as reported earlier in other models (54, 55). Furthermore, fasting glycogen levels in the gastrocnemius muscles of LP rats were approximately threefold higher than the controls. Interestingly, few other studies have shown such high glycogen storage in skeletal muscles and have suggested that high glycogen storage could contribute to insulin resistance (56–59). Glycogen levels at the end of overnight fasting should be low as they will be converted to glucose to maintain glucose homeostasis. However, in these T2D female rats, we observed higher levels of stored glycogen, similar to glycogen storage disease

in humans (42). We have also performed stable isotopebased studies and found that the regulation of glycogenolysis is affected in these LP-programmed animals (60). Upon insulin treatment of 30 minutes, inactive phosphorylated GS levels increased in controls and decreased in the LP group contrary to the expectation. This could be due to the ex vivo nature of our study, and the reaction is allosterically regulated by glucose-6-phosphate levels in the tissue (52). It is likely that after 30 minutes, the tissue glucose source is exhausted in the controls and GS is inactivated again due to the lack of substrate, whereas in LP rats, GS is still active, trying to convert glucose to glycogen. Furthermore, euglycemic hyperinsulinemic clamp data (5) and glucose tolerance test in the current study clearly show that the LP-programmed rats are glucose intolerant and insulin resistant. Taken together, our studies show that gestational LP-induced T2D females have an inherent disadvantage for insulin action on GSK-3 signaling due to high basal levels of inactive GS and stored glycogen. Sex differences in disease presentations and developmental programming are a novel concept and not

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well understood. Although some aspects are known (9), we do not have deep understanding of the sex differences at the molecular level. Our earlier study in males (12) and the current study in females show clear sex differences in the mechanistic aspects despite showing similar disease outcomes. We had shown earlier that in males, glucose intolerance and insulin resistance are caused by a series of phosphorylation defects in IR, IRS-1, and AS-160, leading to impaired Glut4 translocation affecting glucose transport in gastrocnemius muscles (12). In this study, we explored the mechanism of glucose intolerance and insulin resistance in females. Unlike males, females maintained an intact Glut4 translocation, clearly showing sex-dependent impairment in the molecular signaling involving insulininduced glucose transport. Although the underlying mechanism for the sex differences is not clearly understood, sex steroids could play a vital role in the modulation of insulin signaling (61). Clinical studies have shown that men with decreased testosterone (62–64) and women with decreased estradiol and increased testosterone exhibit varying degrees of insulin resistance (61). In summary, we show that gestational protein restriction in female rats alters the insulin signaling

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cascade that regulates glycogen synthesis, leading to insulin resistance and glucose intolerance (Fig. 8). Our data clearly show that defects in insulin-induced IRS-1 tyrosine phosphorylation start the domino effect in the insulin signaling cascade that contributes to impaired glucose disposal. Interestingly, although LP programming affects both males and females, the mechanisms that contribute to insulin resistance are different, as we have shown now and in our earlier publications. The reason for the sex difference in LP programming is not clearly understood. A better understanding of sex differences in the disease mechanisms may help us to devise appropriate sex-specific treatment strategies.

Acknowledgments Address all correspondence and requests for reprints to: Chandrasekhar Yallampalli, DVM, PhD, Baylor College of Medicine, Department of Obstetrics and Gynecology, One Baylor Plaza, MS: BCM610, Houston, Texas 77030. E-mail: [email protected]. This work was supported by National Institutes of Health Grants HL102866 and HL58144 (to C.Y.) and HD069750 and HL119869 (to K.S.). Disclosure Summary: The authors have nothing to disclose.

Antibodies Used

Peptide/Protein Target Phospho–IRS-1 Ser318 Phospho–IRS-1 Ser612 Akt Phospho–AS-160 Thr642 Phospho-Akt Ser473 Phospho-Akt Thr308 GSK3a Phospho-GSK3a GSK3b Phospho-GSK3b GS Phospho-GS SHP-2 IRb AS-160 Glut4 Phospho–insulin receptor Tyr972 Phospho–IRS-1 Tyr612 Phospho–IRS-1 Tyr896 a1 Sodium potassium ATPase GAPDH

Antigen Sequence (if Known)

Name of Antibody

Manufacturer, Catalog #, and/or Name of Individual Providing the Antibody

Species Raised in; Dilution Monoclonal or Polyclonal Used

Cell Signaling #5610

Rabbit; polyclonal

1:1000

Cell Signaling #3203

Rabbit; polyclonal

1:1000

Cell Signaling #9272 Cell Signaling #4288

Rabbit; polyclonal Rabbit; polyclonal

1:1000 1:1000

Cell Signaling #9271 Cell Signaling #9275 Cell Signaling #4337 Cell Signaling #9316 Cell Signaling #12456 Cell Signaling #5558 Cell Signaling #3893 Cell Signaling #3891 Cell Signaling #9793 BD Biosciences Cat. no. 610109 Millipore #07-741 Abcam ab654 Abcam ab5678

Rabbit; polyclonal Rabbit; polyclonal Rabbit; polyclonal Rabbit; polyclonal Rabbit; polyclonal Rabbit; polyclonal Rabbit; polyclonal Rabbit; polyclonal Rabbit; polyclonal Mouse; monoclonal Rabbit; polyclonal Rabbit; polyclonal Rabbit; polyclonal

1:1000 1:1000 1:1000 1:1000 1:1000 1:1000 1:1000 1:1000 2:1000 1:500 1:1000 1:2000 1:1000

Abcam ab66153

Rabbit; polyclonal

1:500

Abcam ab46800

Rabbit; polyclonal

1:500

Abcam ab7671

Mouse; monoclonal

1:5000

Abcam ab9485

Rabbit; polyclonal

1:2500

766

Blesson et al

Prenatal Protein Deficiency Impairs Insulin Signaling

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