Journal of Steroid Biochemistry & Molecular Biology 143 (2014) 357–364

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Low-intensity resistance training attenuates dexamethasone-induced atrophy in the flexor hallucis longus muscle Anderson G. Macedo b , André L.O. Krug b , Naiara A. Herrera a , Anderson S. Zago a , James W.E. Rush c , Sandra L. Amaral a,b,∗ a

Department of Physical Education – UNESP, Science Faculty, Av. Eng. Luiz Edmundo Carrijo Coube, 14-01 – Vargem Limpa, Bauru, SP, Brazil Joint Graduate Program in Physiological Sciences, PIPGCF UFSCar/UNESP, Rodovia Washington Luiz, km 235 Monjolinho, 676, São Carlos, SP, Brazil c Department of Kinesiology, Faculty of Applied Health Sciences, University of Waterloo, 200 University Avenue West, Waterloo, Ontario, Canada b

a r t i c l e

i n f o

Article history: Received 12 November 2013 Received in revised form 13 May 2014 Accepted 15 May 2014 Available online 23 May 2014 Keywords: Glucocorticoids Ladder-resistance training Skeletal muscle

a b s t r a c t This study investigated the potential protective effect of low-intensity resistance training (RT) against dexamethasone (DEX) treatment induced muscle atrophy. Rats underwent either an 8 week period of ladder climbing RT or remained sedentary. During the last 10 days of the exercise protocol, animals were submitted to a DEX treatment or a control saline injection. Muscle weights were assessed and levels of AKT, mTOR, FOXO3a, Atrogin-1 and MuRF-1 proteins were analyzed in flexor hallucis longus (FHL), tibialis anterior (TA), and soleus muscles. DEX induced blood glucose increase (+46%), body weight reduction (−19%) and atrophy in FHL (−28%) and TA (−21%) muscles, which was associated with a decrease in AKT and an increase in MuRF-1 proteins levels. Low-intensity RT prevented the blood glucose increase, attenuated the FHL atrophy effects of DEX, and was associated with increased mTOR and reductions in Atrogin-1 and MuRF-1 in FHL. In contrast, TA muscle atrophy and signaling proteins were not affected by RT. These are the first data to demonstrate that low-intensity ladder-climbing RT specifically mitigates the FHL atrophy, which is the main muscle recruited during the training activity, while not preventing atrophy in other limb muscle not as heavily recruited. The recruitment-dependent prevention of atrophy by low intensity RT likely occurs by a combination of attenuated muscle protein degradation signals and enhanced muscle protein synthesis signals including mTOR, Atrogin-1 and MuRF-1. © 2014 Elsevier Ltd. All rights reserved.

1. Introduction

Abbreviations: AKT/PKB, protein kinase B; AMPK, adenosine monophosphate kinase; BSA, bovine serum albumin; CaMKK, calcium/calmodulin-dependent protein kinase; CEUA, Committee for Ethical Use of Animals; DEX, dexamethasone; EDTA, ethylenediaminetetraacetic acid; FHL, flexor hallucis longus; FOXO3a, forkhead box 3A; GLUT-4, glucose transporter type 4; IGF-1, insulin growth factor-1; i.p., intraperitoneal injection; MHC, miosin high chain; mTOR, protein kinase mammalian target of rapamycin; MuRF-1, muscle ring finger-1; MVCC, maximum voluntary carrying capacity; NaCl, sodium chloride; PI3k, phosphoinositide 3 kinase; PMSF, phenylmethylsulfonyl fluoride; P70S6K – 70 kDa, ribossonal protein S6 kinase; RIPA, radioimmunoprecipitation assay; RT, resistance training; S, sedentary; SC, sedentary control; SD, sedentary treated with DEX; SE, standard error; SOL, soleus; T, training; TBST, tris-buffered saline with Tween; TC, resistance training; TD, resistance training treated with DEX; Tris–HCl, Tris hydrochloride. ∗ Corresponding author at: Department of Physical Education, Science Faculty, São Paulo State University – UNESP, Bauru, SP, Brazil. Tel.: +55 14 3103 6082; fax: +55 14 3103 6082. E-mail addresses: [email protected] (A.G. Macedo), [email protected] (A.L.O. Krug), [email protected] (N.A. Herrera), [email protected] (J.W.E. Rush), [email protected] (S.L. Amaral). http://dx.doi.org/10.1016/j.jsbmb.2014.05.010 0960-0760/© 2014 Elsevier Ltd. All rights reserved.

Dexamethasone (DEX) is used as an anti-inflammatory and antiallergic drug, however, high doses or chronic use may lead to several side effects including hyperglycemia, hypertension, dyslipidemia and cachexia [1–4]. With respect to skeletal muscle, some studies have shown that chronic treatment with DEX promotes muscle atrophy [2–6] which, in some circumstances, is also associated with body weight loss [6–8]. As reviewed by Egerman and Glass [9], normal maintenance of muscle mass occurs via a tight control of signaling processes involved in both synthesis (including IGF-1, insulin growth factor-1; PI3k, phosphoinositide 3 kinase; AKT/PKB, protein kinase B; mTOR, protein kinase mammalian target of rapamycin and P70S6K, ribosomal protein S6 kinase) and degradation (including FOXO3a, forkhead box 3A; MAFbx, muscle atrophy Fbox (also named Atrogin-1), and MuRF-1, muscle ring finger-1) of muscle proteins. Atrogin-1 and MuRF-1 were the first proteins considered crucial for regulation of muscle atrophy [10]. Although skeletal muscle atrophy induced by DEX has been shown to be associated with both increased catabolism [6,7] and reduced

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synthesis of muscle proteins [5,9], the possible alterations in molecular signaling mechanisms causing this are not fully understood. Low intensity physical exercise has been used as a nonpharmacological intervention to control several side effects of DEX including diabetes, hypertension and sarcopenia [11,12,13]. Pinheiro et al. [1] have shown that endurance exercise is effective in preventing alterations in lipid metabolism induced by DEX. Similarly, Barel et al. [2] demonstrated that 8 weeks of treadmill endurance training attenuated DEX-induced increases in blood glucose, but was not able to prevent muscle atrophy induced by DEX. In contrast, other studies demonstrate that aerobic continuous exercise has been shown to attenuate [14], or prevent [15], the muscle mass reduction induced by glucocorticoids. It is important to investigate different types of exercise in order to better understand whether exercise can prevent muscle mass reductions induced by DEX, and to identify mechanisms that might be involved. Resistance exercise training has been shown to be efficient in the attenuation of muscle mass reduction in some pathologies [4,14,15]. However, the role of resistance training in preventing muscle atrophy induced by DEX is not completely understood. The purpose of the current study is to examine the hypothesis that low intensity resistance training will protect against DEX-induced muscle atrophy and the underlying molecular mechanisms responsible for this effect, by attenuation of signals associated with muscle protein degradation and elevation of signals associated with muscle protein synthesis. These responses will be specific to the flexor hallucis longus (FHL) muscle that is heavily recruited during the training protocol used in this study, and will not occur in other limb muscles that are not as heavily recruited during this type of training [16]. To our knowledge, this is the first study to investigate the effects of this type of low intensity resistance training on FHL muscle atrophy induced by DEX. 2. Materials and methods 2.1. Animals All methods used were approved by the Committee for Ethical Use of Animals (CEUA) of the UNESP-São Paulo State University, Arac¸atuba (protocol # 2012-02253). Forty-four male rats (250–300 g) were housed in group cages and maintained under controlled environmental conditions (12 h dark–light cycle, 22 ◦ C of temperature) with ad libitum access to standard diet (Biobase, Brazil) and water. Body weight was measured weekly during the exercise protocol and daily during DEX treatment. 2.2. Maximum voluntary carrying capacity (MVCC) Initially, the animals were adapted to the climbing apparatus which included a vertical ladder (110 cm, 80◦ incline) and a tail weight attachment as previously described by Hornberger and Farrar [16], Prestes et al. [17] and Sanches et al. [18]. Briefly, after a 10 day familiarization period, each rat performed a test in order to evaluate its maximum voluntary carrying capacity (MVCC). In this test, each rat started to climb the ladder carrying 75% of its own body weight and was allowed to rest at the top of the ladder for 120 s. After each completed climb 30 more grams were added to the total carried mass. This procedure was successively repeated until the rat failed to climb the entire length of the ladder on three consecutive attempts; MVCC was defined as the highest load successfully carried in this protocol. 2.3. Blood glucose determination After overnight fasting (12 h), animals were gently restrained, tail blood samples were obtained from tail nicks, and blood glucose

levels were determined using a digital glucometer system (One Touch Ultra – Johnsons & Johnsons). This procedure was performed for each animal on three occasions during the protocol: (1) at the beginning of the sedentary/exercise protocol; (2) before DEX/saline administration, and (3) just prior to euthanasia. For occasion 2, the blood sample was obtained 24 h after the last training session. 2.4. Experimental treatment protocol After preliminary assessments were completed, rats were allocated into four experimental groups balanced to ensure equal initial body weight, fasting blood glucose and MVCC across groups: (1) sedentary control (SC); (2) sedentary treated with DEX (SD); (3) resistance trained control (TC) and (4) resistance trained treated with DEX (TD). DEX-treated rats received daily injections (at 9 a.m.) of DEX (Decadron® , 0.5 mg/kg of body weight, i.p., dissolved in saline) during the last 10 days of the entire experimental protocol (8 weeks + 10 days). Control animals received daily injections of saline during the last 10 days of the experimental protocol (same volume as DEX treated rats). In the trained group, resistance training continued throughout the DEX or saline administration period. Trained groups performed 8 weeks of ladder resistance training at 60% of MVCC, 5 days a week [19]. Each training session consisted of 14–20 ladder climbs. The maximal voluntary carrying test was repeated after 4 weeks (MVCC-2) and after 8 weeks (MVCC-3) of training. An additional MVCC (MVCC-4) was carried out after the 10 day DEX (or saline) treatment period. Sedentary rats also performed MVCC tests at the same time intervals as indicated for the trained animals, but otherwise remained in their cages and did not undergo the training protocol. 2.5. Tissue harvesting After euthanasia by excess of anesthesia (Ketamine 160 mg/kg and Xylazine 20 mg/kg), entire FHL, tibialis anterior (TA), and soleus muscles were removed, weighed, immediately frozen in dry ice and then stored at −80 ◦ C until used for Western blotting analysis. Muscle weights were normalized to tibia bone length since DEX treatment is known to affect body weight. The tibia bone length was measured from the proximal extremity of the lateral condyle up to the posterior process, in the distal extremity. The three muscles selected for study have different fiber type composition. While FHL is composed predominantly of myosin heavy chain 2a/x (MHC IIa/x) with a small percentage of myosin heavy chain 2b (MHC IIb) [16,20], TA muscle is composed mainly of MHCIIb [16,20]. Soleus muscle is composed mainly by slow-fiber type I. It has been previously demonstrated that, of these three limb muscles, FHL muscle is the heavily recruited muscle during the type of ladder-climbing resistance exercise training protocol used in this study [16,20]. 2.6. Western blotting procedures Samples of TA, FHL and SOL were homogenized in RIPA solution (10X, Millipore) containing 0.5 M Tris–HCl, pH 7.4, 1.5 M NaCl, 2.5% deoxycolic acid, 10% NP-40, 10 mM EDTA using a polytron homogenizer. Just prior to homogenization, a protease inhibitor cocktail (Pic, Sigma) and 1% PMSF were added to the samples. Samples were centrifuged at 10,000 × g for 5 min and the supernatant was collected and stored in −20 ◦ C freezer for future analysis. Bradford assays were used to determine the protein concentration of the samples (Bio-Rad Kit, Protein Assay Standard II, Hercules, CA) as previously published [21]. Absorbance values were determined using a spectrophotometric plate reader (BMG Labtech, Spectro Star Nano). Western blotting was performed according to previously reported procedures [22]. In summary, samples containing 50–80 ␮g of protein (depending on the specific target in

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SC SD TC TD

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3.1. Body weight Fig. 1A represents the body weight values during exercise training and DEX treatment periods. All rats had a similar and significant increase of body weight during the first 8 weeks (exercise training). Statistical analysis showed an effect of time on body weight gain, independent of training status. On the other hand, 10 days of DEX treatment significantly decreased body weight of both sedentary (−19%) and trained (−16%) groups. A drug effect, independent of training status, was observed during DEX treatment. The inset shows the body weight of each group at the end of the protocol. Fig. 1B shows the daily food intake in all groups. As observed, food intake was similar among groups before DEX treatment. DEX treatment provoked a reduction in food intake of sedentary and treated animals, however, exercised group exhibited normal food intake after 10 days of treatment.

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Days Fig. 1. (A) Body weight (BW) during the entire protocol (8 weeks of training and 10 days of dexamethasone treatment plus training). Insert represents values of BW on euthanasia day. (B) Daily food intake before treatment (days −2, −1 and 0) and during 10 days of DEX treatment in all groups: sedentary control (SC, n = 11), sedentary treated with DEX (SD, n = 11), resistance trained control (TC, n = 11) and resistance trained treated with DEX (TD, n = 13). Significance: * vs respective control (drug vs saline), # vs first day for all groups analyzed, p < 0.05.

Fig. 2B illustrates the 8 week training effectiveness in improving MVCC, expressed by the difference between MVCC-3 and MVCC-1 grouped across all trained and all sedentary animals. Fig. 2C illustrates the effects of DEX treatment on MVCC, expressed by the difference between MVCC-4 and MVCC-3 in all groups analyzed. This analysis reveals that treatment with DEX did not affect MVCC in sedentary or trained animals compared to their respective controls. 3.3. Blood glucose All groups presented similar values of fasting blood glucose at the beginning of the experimental protocol, as shown in Table 1. Eight weeks of exercise training did not change blood glucose Table 1 Fasting blood glucose during experimental protocol in all groups. Initial (mg/dL)

3.2. MVCC tests As shown in Fig. 2A, the MVCC was similar among all groups at the beginning of the training protocol (MVCC-1). Resistance training significantly increased the MVCC of animals after 8 weeks. The training effect was independent of drug treatment group assignment, as expected since this measurement was performed prior to initiation of the DEX or saline (control) treatment period. Sedentary animals did not improve their MVCC as demonstrated in Fig. 2A.

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2.7. Statistical analysis All values were presented as mean ± SE. Significant differences in the values measured among all groups were detected using a Two way analysis of variance (ANOVA), considering training and treatment as factors. Some variables (body weight, MVCC and fasting glucose) were analyzed using two way Repeated Measures ANOVA. Significant differences were further investigated using a post hoc test (Tukey). For comparisons between two groups we used nonpaired t-test. The level of significance considered was ˛ < 0.05.

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each immunoblot) were separated using 10% denaturing polyacrylamide gels. These gels were then transferred to nitrocellulose membranes (190 mM glycine, 25 mM Tris, 20% methanol, pH 8.3). Equal protein loading across samples for a given immunoblot was confirmed using 0.5% Ponceau staining. Membranes were blocked with 1.5% bovine serum albumin (BSA) in Tris-buffered saline with Tween (TBS-T) for 40 s. The SNAP i.d.® 2.0 Protein Detection system (Millipore) was used which required a 10 min incubation of the membranes with appropriate dilution of primary antibody: Akt (cell signaling #9271s, 1:1000 in 3% BSA), mTOR (cell signaling #2972s, 1:1000 in 3% BSA), FOXO-3a (cell signaling #9464, 1:1000 in 3% BSA), Atrogin-1 (Abcam #a74023, 1:1000 in 3% BSA), MuRF-1 (Santa Cruz (C-20) sc-27642, 1:1000 in 3% BSA) and GAPDH (R&D SYSTEMS #AF5718, 1:1000 in 3% BSA) for 10 min. The membranes were then washed and incubated in the appropriate horse radish peroxidase-conjugated secondary antibody: Akt (IgG anti-rabbit 1:1000), mTOR (IgG anti-rabbit 1:1000), FOXO3a (IgG anti-rabbit 1:1000), Atrogin-1 (IgG anti-rabbit 1:10,000), MuRF-1 (anti-donkey 1:10,000) and GAPDH (IgG anti-rabbit 1:1000)for 10 min. Signals were detected by enhanced chemiluminescence (Super signal® West Pico, Pierce) captured on radiographic film, and bands of interest were analyzed using scanning densitometry (Scion Image Computer, 4.0.2). Raw densitometry data for each target protein signal were normalized to the GAPDH protein signal from the same sample lane. Group data are expressed as a percentage relative to the control group.

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SC SD TC TD

79 78 79 76

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After training/before DEXA treatment (mg/dL) 80 82 78 78

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± ± ± ±

3 9* 2 3+

SC = sedentary control; SD = sedentary treated with DEX; TC = resistance training control; TD = resistance training treated with DEX; DEX: dexamethasone. * vs respective control group, p < 0.05. + vs respective sendentary group, p < 0.05.

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associated with a reduction of AKT (−47% for SD vs SC) and an increase in MuRF-1 (+41%, for SD vs SC). Protein levels of mTOR, FOXO3a, Atrogin-1 in TA were not affected by DEX treatment. In contrast to the response in FHL, TA muscle mass reduction was not attenuated by low intensity RT (−18%, for TD vs −21% for SD). The patterns of treatment effects on AKT and MuRF-1 in TA muscle were similar to those observed in the FHL. However, the training effects observed in FHL with respect to mTOR and Atrogin-1 were not observed in TA. Fig. 5 reveals that DEX treatment did not affect SOL muscle weight, nor the levels of any of the proteins studied. Likewise the low intensity resistance training did not affect the levels of proteins.

4. Discussion

Fig. 2. (A) Maximal voluntary carrying capacity (MVCC) tests performed on the ladder during the experimental protocol: MVCC-1 (beginning), MVCC-2 (after 4 weeks), MVCC-3 (after 8 weeks, just prior to beginning the DEX or saline treatment) and MVCC-4 (after DEX or saline treatment) in all four groups. Gray bar means training period and hatched bar means training + DEX period; (B) Effect of resistance training in the trained (n = 24) and sedentary (n = 24) animals expressed as the difference between MVCC-3 and MVCC-1 (8 weeks vs beginning). Student’s t-test showed that T group was significantly different from S group (p = 0.034); (C) Effect of drug treatment on the maximal voluntary carrying capacity expressed as the difference between MVCC-4 (after DEX or saline treatment) and MVCC-3 (before DEX or saline treatment) in all four groups: sedentary control (SC, n = 11), sedentary treated with DEX (SD, n = 13), resistance trained control (TC, n = 11) and resistance trained treated with DEX (TD, n = 13) + vs respective sedentary; # vs first day for all groups, p < 0.05.

values. However, 10 days of DEX treatment significantly increased blood glucose of sedentary animals (+46%), whereas low intensity resistance training completely prevented this hyperglycemic effect of DEX. 3.4. Muscle atrophy and protein expression Figs. 3–5 illustrate skeletal muscle weight, and the content of some proteins of interest involved in muscle hypertrophy (AKT, mTOR) and muscle atrophy (FOXO3a, Atrogin-1 and MuRF1). All muscle weights were normalized by tibia bone length which was similar among groups (4.16 ± 0.1, 4.23 ± 0.2, 4.21 ± 0.1, 4.26 ± 0.2 cm, for SC, SD, TC and TD, respectively, p > 0.05). Fig. 3 illustrates the responses in FHL muscle which is the most recruited muscle of those studied in the training protocol employed. In the sedentary group, DEX treatment provoked a reduction of ∼28% in FHL muscle mass compared with control. Dexamethasone significantly reduced AKT (−37%) protein levels and increased MuRF-1 protein levels (+67%) in FHL muscle of sedentary animals, compared with control. FOXO3a, mTOR and Atrogin-1 protein levels remained unchanged in FHL after DEX treatment. Although low-intensity resistance training did not increase FHL muscle mass of the TC group (compared to SC), it was effective in attenuating the DEX-induced FHL muscle mass loss compared with sedentary treated rats (−18% for TD vs TC and −28% for SD vs SC). AKT levels were decreased by DEX (SD) and by training (TC and TD) compared to SC, while DEX did not have a significant effect within the trained groups. However, low intensity resistance training significantly attenuated the DEX-induced increase in MuRF-1 protein levels. These responses were associated with reduced Atrogin1 and increased mTOR protein levels as main training effects vs sedentary across control and DEX groups. Fig. 4 illustrates that DEX treatment induced a 21% reduction of TA muscle mass in sedentary animals and this response was

The major finding of this study is that low intensity resistance training attenuated FHL muscle atrophy induced by chronic treatment with DEX, and that this response is associated with training-induced effects on key signaling proteins that regulate muscle protein synthesis and degradation including mTOR, MuRF1 and Atrogin-1. To our knowledge, this is the first study that investigates the effects of low-intensity ladder-climbing resistance training on FHL muscle atrophy induced by DEX. Despite the utility of DEX in treatment of inflammatory and autoimmune disorders, when used in high doses or chronically for longer periods it provokes several side effects such as: peripheral insulin resistance; hypercholesterolemia; hyperinsulinemia; body weight loss and, muscle atrophy [1,2,4,8,23–25]. It has been suggested that DEX treatment may cause changes in liver, pancreas and muscle metabolism, inducing hyperglycemia [2,26,27]. We found in the present study that 10 days of DEX treatment induced a 46% increase in blood glucose concentration. On the other hand, our results demonstrate that low intensity resistance training completely abolished the increased blood glucose observed in sedentary DEX treated rats. In accordance with our observation, Nicastro et al. [4] have shown that some sessions of squat type exercise, performed together with DEX treatment, can attenuate DEX-dependent blood glucose increases. The preventive response resulting from exercise likely involves improvements in both insulin-dependent and independent glucose uptake signaling [3]. It is reasonable to suggest that this beneficial response is dependent on the type and intensity of exercise, because previous studies illustrate that aerobic continuous exercise can attenuate, but not abolish, the increased blood glucose induced by DEX [2], while our own results with low-intensity resistance training demonstrate a more complete protection against DEX-induced hyperglycemia. In the current study, both DEX-treated groups had decreased body weight. This response may be partially explained by reductions in food intake, as observed in our treated animals. Previous studies indicate that this response may be due to DEX effects on leptin and ghrelin levels since DEX increases plasma levels of leptin [29] and inhibits the production of ghrelin [30], both leading to an increased satiety signaling in the central nervous system. In the present study, low intensity resistance training was not able to attenuate body weight loss in either DEX group and these results agree with those of Nicastro et al. [28]. Although body weight was maintained lower throughout the protocol in DEX groups, the food intake returned to control levels after 8 days of exercise, suggesting that decreased food intake, although an important trigger, does not fully explain the body weight reduction. Muscle atrophy is a serious side effect of synthetic glucocorticoids [2,7,9]. There is strong evidence that DEX acts on muscles containing mainly type IIb fast-twitch fibers compared with type I slow-twitch fibers [5,15,28]. In agreement, in the present study we observed atrophy in TA and FHL but not in soleus. Studies

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Fig. 3. FHL muscle weight and densitometry analysis of AKT, mTOR, FOXO3a, Atrogin-1 e MuRF-1 protein level normalized by GAPDH in all groups: sedentary control (SC), sedentary treated with DEX (SD), resistance trained control (TC) and resistance trained treated with DEX (TD). Results are expressed as % of control group (SC). Number of rats used for protein levels is 5–6 in each group. Significance: *vs respective control group, + vs respective sedentary group, ‡ vs sedentary control group (SC), p < 0.05.

examining FHL responses to DEX are lacking, but this muscle was chosen for examination in the current study since it is the most recruited in the ladder climbing type of resistance exercise utilized herein [16]. Skeletal muscle mass is regulated by multiple interconnecting pathways that control both synthesis and degradation of proteins [9]. The cellular and molecular pathways responsible for degradation induced by DEX are beginning to be elucidated. Indeed, two muscle specific E3 ligases, involved in the muscle ubiquitin/proteolysis pathway, MuRF-1 and Atrogin-1, are thought to be key regulators of this process and are excellent markers of muscle atrophy [10,12,31]. Our results indicate that DEX treatment provoked a reduction of ∼28% in FHL muscle mass, and this response is associated with decreases in AKT and increases in MuRF-1. These findings suggest that the IGF1/PI3K/Akt pathway is depressed by DEX, as was first demonstrated by Stitt et al. [32] and Sandri et al. [33]. These authors have shown that, in the presence of atrophy, the activity of the PI3K/AKT pathway decreases, leads to activation of FOXO transcription factors and Atrogin-1 and MuRF-1 induction. In the present study, Atrogin-1 protein levels were not altered

after DEX treatment, even in the presence of muscle atrophy, consistent with the argument established by Baehr et al. [6] that MuRF-1 and Atrogin-1 do not function similarly under all atrophy models. These investigators observed that fiber cross-sectional area of gastrocnemius muscle is preserved in MuRF-1 knockout mice treated with DEX when compared with MAFbx (Atrogin-1) knockout mice treated with DEX, suggesting that MuRF-1 could be more important in DEX muscle atrophy models than Atrogin-1. In accordance, Nicastro et al. [4] also demonstrated that 7 days of DEX treatment causes plantaris muscle atrophy associated with decreases in phospho-AKT and increases in MuRF-1. We did not assess phospho-AKT protein levels in this study which is a limitation, but we have observed in another group of animals that phospho-AKT/total AKT ratio in TA muscle is reduced after DEX treatment [3]. Sandri et al. [33] have demonstrated that forkhead factors (FOXO family) play a crucial role in the muscle atrophy. The present study was limited by only examining total FOXO3a, which was not affected by DEX, and thus we missed the opportunity to probe whether phosphorylation of FOXO3a is related to the atrophy responses we studied in this model. In agreement, Nicastro et al. [4] have observed that FOXO3a was unchanged (similarly

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Fig. 4. TA muscle weight and densitometry analysis of AKT, mTOR, FOXO3a, Atrogin-1 e MuRF-1 protein level normalized by GAPDH in all study groups: sedentary control (SC), sedentary treated with DEX (SD), resistance trained control (TC) and resistance trained treated with DEX (TD). Results are expressed as % of control group (SC). Number of rats used for protein levels is 5–6 in each group. Significance: * vs respective control group, + vs respective sedentary group, ‡ vs sedentary control group (SC), p < 0.05.

to our results) but the phospho-FOXO3/total FOXO3a ratio was reduced after DEX treatment, suggesting that DEX treatment may decrease the phosphorylation of FOXO3a, and not the total protein level. Although the resistance training applied in this study did not promote increases in muscle mass in the control group, which have been observed after high intensity protocols in previous studies [17,20,31], low-intensity resistance training, performed before and concomitant with DEX treatment, was effective in attenuating FHL muscle atrophy. This effect was associated with a combination of mTOR increases, Atrogin-1 decreases and attenuation of DEX-induced MuRF-1 increase. Our exercise protocol did reduce AKT levels and did not bring back to normal the reduced AKT protein level induced by DEX treatment. Possibly the intensity of the exercise was not enough to fully restore the AKT level and the muscle mass of FHL. Some data provide support for the hypothesis that mTOR signaling, which is downstream to AKT/PKB, is a critical mediator of muscle growth [33,34] but few works have actually demonstrated the effects of resistance exercise on AKT, and the results are still not clear. The majority of the studies that have shown effects of exercise on AKT protein levels have examined

just acute exercise and few have observed chronic effect of exercise. Deldicque et al. [35] showed that the phosphorylated form of AKT was reduced immediately following a high-intensity resistance exercise bout in human quadriceps muscle, while Creer et al. [36] demonstrated increases in phospho-AKT after low-intensity resistance exercise. On the other hand, Zanchi et al. [37] demonstrated that 12 weeks of high intensity resistance training did not change total AKT protein level in plantaris muscle, while Luo et al. [38] observed reductions of phospho-AKT and total AKT in skeletal muscle of aged rats after chronic resistance training, which agrees with the current results for AKT. Even though low intensity exercise training in the current study did not increase FHL muscle mass, it was effective in increasing mTOR levels, which probably contributed to the attenuation of FHL muscle mass loss induced by DEX. Likewise, Zanchi et al. [37] showed increases of mTOR after chronic high intensity resistance training. On the other hand, Nicastro et al. [28] demonstrated that animals that underwent squat type resistance training, showed increases of phospho-mTOR/total mTOR ratio but no change in total mTOR in plantaris muscle of both DEX treated and untreated rats, compared with respective controls. It is possible that the duration

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Fig. 5. SOL muscle weight and densitometry analysis of AKT, mTOR, FOXO3a, Atrogin-1 e MuRF-1 protein level normalized by GAPGH in all groups: sedentary control (SC), sedentary treated with DEX (SD), resistance trained control (TC) and resistance trained treated with DEX (TD). Results are expressed as % of control group (SC). Number of rats used for protein levels is 5–6 in each group.

of exercise protocol is an important determinant of changes in total protein level, since Nicastro et al. [28] observed the effects after only 3 days of training. Our study did not investigate the phospho-mTOR protein levels, which is a limitation of this study, but we have illustrated that 2 months of exercise were effective in increasing total mTOR protein level. The total FOXO3a was not affected by resistance training, in agreement with Nicastro et al. [28], however these authors have previously demonstrated that phosphoFOXO3/total FOXO3a ratio was reduced after resistance training. Our results have revealed that Atrogin-1 was reduced by ∼29% after low intensity resistance training. This result is in agreement with other studies that have shown that both acute and chronic resistance exercise appear to decrease gene expression of Atrogin-1 [36,38]. MuRF-1 did not change significantly with resistance training alone in control rats but our protocol, using a ladder-climbing regimen, attenuated the increase of MuRF-1 observed in FHL muscle of DEX-treated animals. In contrast, Nicastro et al. [28] showed

that MuRF-1 protein level in the plantar muscle did not change after resistance training in animals treated with DEX. In contrast to the responses to ladder-resistance exercise training in FHL, RT did not attenuate TA muscle atrophy induced by DEX. Furthermore, in accordance with maintenance of muscle atrophy in TA after RT, the levels of Atrogin-1 and mTOR were not altered by training, even though MuRF-1 protein levels were normal after training. The most likely explanation for this differential effect of RT on muscles is that FHL is the most recruited in this kind of ladder-regimen resistance exercise [16], consistent with the observation that of several limb muscles analyzed only FHL demonstrated higher cross-sectional area, and total and myofibrillar protein content after 8 weeks of such ladder resistance exercise [16]. It is important to note that the slow fiber-type soleus muscle, unlike the fast fiber-type TA or FHL muscles, did not exhibit any trend toward a decrease in muscle mass in response to DEX, in agreement with the findings of other studies [2,15,23]. Since SOL

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did not atrophy in response to DEX, it was not surprising that the regulatory markers assessed also showed no changes. The major conclusions supported by our studies are that low intensity resistance training was effective in mitigating FHL muscle atrophy induced by DEX, while this training effect did not extend to TA muscle, which is less recruited during the training protocol. The atrophic response was associated with increases in MuRF-1 and decreases in AKT protein levels in both muscles. However, low intensity RT attenuated the DEX-induced MuRF-1 increase, decreased Atrogin-1 and increased mTOR protein levels specifically in the highly recruited FHL muscle. Conflict of interest The authors have no conflicts of interest to disclose. Acknowledgments This study was supported by São Paulo Research Foundation (FAPESP # 2011/21522-0 grant to SLA). AGM was a recipient of a scholarship from the National Council for Scientific and Technological Development (CNPq, process # 130232/2011-4). ALOK was a recipient of a scholarship from São Paulo Research Foundation (FAPESP # 2012/21820-3) and Coordination for the Improvement of Higher Education Personnel (CAPES). We gratefully acknowledge the technical assistance of Lidiane M Souza (FAPESP # 2012/03816-9). References [1] C.H. Pinheiro, W.N. Sousa Filho, J. Oliveira Neto, M.J. Marinho, R.N. Motta, M.M. Smith, C.A. Silva, Exercise prevents cardiometabolic alterations induced by chronic use of glucocorticoids, Arq. Bras. Cardiol. 93 (2009) 400–408. [2] M. Barel, O.A. Perez, V.A. Giozzet, A. Rafacho, J.R. Bosqueiro, S.L. do Amaral, Exercise training prevents hyperinsulinemia, muscular glycogen loss and muscle atrophy induced by dexamethasone treatment, Eur. J. Appl. Physiol. 108 (2010) 999–1007. [3] T. Dionisio, J. Louzada, B. Viscelli, A. Martuscelli, M. Barel, O. Perez, J. Bowqueiro, D. Brozoski, C. Santos, S. Amaral, Aerobic training prevents dexamethasoneinduced peripheral insulin resistance, Horm. Metab. Res. 46 (2014) 1–6. [4] H. Nicastro, B. Gualano, W.M. de Moraes, V. de Salles Painelli, C.R. da Luz, A. dos Santos Costa, F. de Salvi, A. Medeiros, P.C. Brum, A.H. Lancha Jr., Effects of creatine supplementation on muscle wasting and glucose homeostasis in rats treated with dexamethasone, Amino Acids 42 (2012) 1695–1701. [5] J.E. Cho, M. Fournier, X. Da, M.I. Lewis, Time course expression of Foxo transcription factors in skeletal muscle following corticosteroid administration, J. Appl. Physiol. 108 (2009) 137–145. [6] L.M. Baehr, J.D. Furlow, S.C. Bodine, Muscle sparing in muscle RING finger 1 null mice: response to synthetic glucocorticoids, J. Physiol. 589 (2011) 4759–4776. [7] K. Ma, C. Mallidis, S. Bhasin, V. Mahabadi, J. Artaza, N. Gonzalez-Cadavid, J. Arias, B. Salehian, Glucocorticoid-induced skeletal muscle atrophy is associated with upregulation of myostatin gene expression, Am. J. Physiol. Endocrinol. Metab. 285 (2003) 363–371. [8] H. Gilson, O. Schakman, L. Combaret, P. Lause, L. Grobet, D. Attaix, J.M. Ketelslegers, J.P. Thissen, Myostatin gene deletion prevents glucocorticoid-induced muscle atrophy, Endocrinology 148 (2007) 452–460. [9] M.A. Egerman, D.J. Glass, Signaling pathways controlling skeletal muscle mass, Crit. Rev. Biochem. Mol. Biol. 49 (2014) 58–59. [10] S.C. Bodine, E. Latres, S. Baumhueter, V.K. Lai, B.A. Clarke, W.T. Poueymirou, Identification of ubiquitin ligases required for skeletal muscle atrophy, Science 23 (294) (2001) 1704–1708. [11] O. Schakman, H. Gilson, J.P. Thissen, Mechanisms of glucocorticoid-induced myopathy, J. Endocrinol. 197 (2008) 1–10. [12] J.P. Little, A. Safdar, G.P. Wilkin, M.A. Tarnopolsky, M.J. Gibala, A practical model of low-volume high-intensity interval training induces mitochondrial biogenesis in human skeletal muscle: potential mechanisms, J. Physiol. 588 (2010) 1011–1022. [13] K. Lenk, G. Schuler, V. Adams, Skeletal muscle wasting in cachexia and sarcopenia: molecular pathophysiology and impact of exercise training, J. Cachexia Sarcopenia Muscle 1 (2010) 9–21.

[14] S.M. Czerwinski, T.G. Kurowski, T.M. O’Neill, R.C. Hickson, Initiating regular exercise protects against muscle atrophy from glucocorticoids, J. Appl. Physiol. 63 (1987) 1504–1510. [15] M.T. Falduto, S.M. Czerwinski, R.C. Hickson, Glucocorticoid-induced muscle atrophy prevention by exercise in fast-twitch fibers, J. Appl. Physiol. 69 (1990) 1058–1062. [16] T.A. Hornberger Jr., R.P. Farrar, Physiological hypertrophy of the FHL muscle following 8 weeks of progressive resistance exercise in the rat, Can. J. Appl. Physiol. 29 (2004) 16–31. [17] J. Prestes, M.R. de Cássia, G.E. Shiguemoto, R.D. Leite, G.B. Pereira, H.S. Selistre-de-Araújo, V. Baldissera, S.E.P. de Andrade, Effects of ovariectomy and resistance training on MMP-2 activity in skeletal muscle, Appl. Physiol. Nutr. Metab. 34 (2009) 700–706. [18] I.C. Sanches, F.F. Conti, M. Sartori, M.C. Irigoyen, K. De Angili, Standardization of resistance exercise training: effect in diabetics ovariectomized rats, Int. J. Sports Med. (2013) 1–10. [19] S. Lee, E.R. Barton, H.L. Sweeney, R.P. Farrar, Viral expression of insulin-like growth factor-I enhances muscle hypertrophy in resistance-trained rats, J. Appl. Physiol. 96 (2004) 1097–1104. [20] H. Zhong, R.R. Roy, J. Woo, J.A. Kim, V.R. Edgerton, Differential modulation of myosin heavy chain phenotype in an inactive extensor and flexor muscle of adult rats, J. Anat. 210 (2007) 19–31. [21] S.L. Amaral, P.E. Papanek, A.S. Greene, Angiotensin II and VEGF are involved in angiogenesis induced by short-term exercise training, Am. J. Physiol. Heart Circ. Physiol. 281 (2001) 163–169. [22] S.L. Amaral, L.S. Sanchez, A.J. Chang, L.V. Rossoni, L.C. Michelini, Time course of training-induced microcirculatory changes and of vegf expression in skeletal muscles of spontaneously hypertensive female rats, Braz. J. Med. Biol. Res. 41 (2008) 424–431. [23] A.M. Ahtikoski, E.M. Riso, S.O. Koskinen, J. Risteli, T.E. Takala, Regulation of type IV collagen gene expression and degradation in fast and slow muscles during dexamethasone treatment and exercise, Pflugers Arch 448 (2004) 123–130. [24] R.C. Hickson, S. Czerrwink, L.E. Wegryzyn, Glutamine prevents downregulation of myosin heavy chain synthesis and muscle atrophy from glucocorticoids, Am. J. Physiol. 268 (1998) 730–734. [25] A. Rafacho, V.A. Giozzet, A.C. Boschero, J.R. Bosqueiro, Functional alterations in endocrine pancreas of rats with different degrees of dexamethasone-induced insulin resistance, Pancreas 36 (2008) 284–293. [26] C. Severino, P. Brizzi, A. Solinas, G. Secchi, M. Maioli, G. Tonolo, Low-dose dexamethasone in the rat: a model to study insulin resistance, Am. J. Physiol. Endocrinol. Metab. 283 (2002) 367–373. [27] J. Ruzzin, A.S. Wagman, J. Jensen, Glucocorticoid-induced insulin resistance in skeletal muscles: defects in insulin signalling and the effects of a selective glycogen synthase kinase-3 inhibitor, Diabetologia 48 (2005) 2119–2130. [28] H. Nicastro, N.E. Zanchi, C.R. da Luz, W.M. de Moraes, P. Ramona, M.A.F. de Siqueira, D.F. Chaves, A. Medeiros, P.C. Brum, D. Dardevet, A.H. Lanchar Jr., Effects of leucine supplementation and resistance exercise on dexamethasoneinduced muscle atrophy and insulin resistance in rats, Nutrition 28 (2012) 465–471. [29] J.W. Jhang, N.Y. Kim, V. Ryu, S.B. Yoo, B.T. Kim, D.W. Kang, J.H. Lee, Dexamethasone reduces food intake, weight gain and the hypothalamic 5-HT concentration and increases plasma leptin in rats, Eur. J. Pharmacol. 581 (2008) 64–70. [30] G. Tulipano, J.E. Taylor, H.A. Halem, R. Datta, J.Z. Dong, M.D. Culler, I. Bianchi, D. Cocchi, A. Giustina, Glucocorticoid inhibition of growth in rats: partial reversal with the full-length ghrelin analog BIM-28125, Pituitary 10 (2007) 267–274. [31] K. Baar, K. Esser, Phosphorylation of p70(S6k) correlates with increased skeletal muscle mass following resistance exercise, Am. J. Physiol. 276 (1999) 120–127. [32] T.N. Stitt, D. Duran, B.A. Clarke, F. Panaro, Y. Timofeyva, W.O. Klyne, D.J. Glass, The IGF-1/PI3K/Akt pathway prevents expression of muscle atrophy-induced ubiquitin ligases by inhibiting FOXO transcription factors, Mol. Cell 14 (2004) 395–403. [33] M. Sandri, C. Sandri, A. Gilbert, C. Skurk, E. Calabria, S. Schiaffino, A.L. Goldberg, Foxo transcription factors induce the atrophy-related ubiquitin ligase atrogin-1 cause skeletal muscle atrophy, Cell 117 (3) (2004) 399–412. [34] S.C. Bodine, mTOR signaling and the molecular adaptation to resistance exercise, Med. Sci. Sports Exerc. 38 (2006) 1950–1957. [35] L. Deldicque, P. Atherton, R. Patel, D. Theisen, H. Nielens, M.J. Rennie, M. Francaux, Decrease in Akt/PKB signalling in human skeletal muscle by resistance exercise, Eur. J. Appl. Physiol. 104 (2008) 57–65. [36] A. Creer, P. Gallagher, D. Slivka, B. Jemiolo, W. Fink, S. Trappe, Influence of muscle glycogen availability on ERK1/2 and Akt signaling after resistance exercise in human skeletal muscle, J. Appl. Physiol. 99 (2005) 950–956. [37] N.E. Zanchi, M.A.F. de Siqueira, F.S. Lira, J.C. Rosa, A.S. Yamashita, C.R.C. de Oliveira, M. Seelaender, A.G. Lancha Jr., Chronic resistance training decreases MuRF-1 and Atrogin-1 gene expression but does not modify Akt, GSK-3beta and p70S6K levels in rats, Eur. J. Appl. Physiol. 106 (2009) 415–423. [38] L. Luo, A.M. Lu, Y. Wang, A. Hong, Y. Chen, J. Hu, X. Li, Z.H. Qin, Chronic resistance training activates autophagy and reduces apoptosis of muscle cells by modulating IGF-1 and its receptors, Akt/mTOR and Akt/FOXO3a signaling in aged rats, Exp. Gerontol. 48 (2013) 427–436.