Introduction

Skeletal muscle wasting or atrophy is characterized by a weakness generated by the loss of muscle mass concomitantly with a decrease of the ability to produce strength. One of the main mechanisms involved in skeletal muscle atrophy is an increase in protein degradation dependent on the activation of the ubiquitin–proteasome system (UPS) (Rezk et al. 2012; Sanders et al. 2005). The muscle-specific E3 ubiquitin ligases atrogin-1 and MuRF-1 belong to the UPS and are involved from the initial stages of skeletal muscle atrophy (Foletta et al. 2011; Glass 2003; Gomes et al. 2001; Gumucio and Mendias 2013); thus, they are upregulated in muscle atrophy caused by different situations (Bodine et al. 2001; Gomes et al. 2001; Semprun-Prieto et al. 2011). The causes of skeletal muscle atrophy are diverse and include sepsis, immobilization, and chronic diseases. A key factor involved in some chronic diseases, such as cardiac, renal, and pulmonary failure, is the participation of the renin–angiotensin system (RAS) by a high level of circulating angiotensin II (AngII) (Agarwal 2003; Agarwal et al. 2004; Antoniu 2008; Brecher 1996; Mancini and Khalil 2005).

The RAS is known for its systemic effects as a regulator of blood pressure. In addition, the RAS also affects other local functions that include the normal and pathological processes of several target tissues. The RAS is composed of two axes with contrary functions: the classical RAS includes AngII, its AT1 and AT2 receptors, and the angiotensin-converting enzyme (ACE), the enzyme forming AngII; whereas the non-classical RAS comprises Ang (1–7), its receptor Mas, and ACE-2, the enzyme that produces Ang (1–7). There is evidence that a balance between these axes is normally maintained, but is disturbed in pathological conditions (Ferreira et al. 2012a; Fyhrquist and Saijonmaa 2008; Iwai and Horiuchi 2009; Lubel et al. 2008).

Ang (1–7) promotes several biological actions, including the inhibition of cell proliferation, vasodilation, and antihypertensive effects (Benter et al. 1995; Ferrario et al. 2005; Iwata et al. 2005; Marangoni et al. 2006; Tallant et al. 2005). To date, one of the main effects of Ang (1–7) is the prevention or decrease of fibrosis (Grobe et al. 2006; Iwata et al. 2005). The effects of Ang (1–7) are mediated by the G-protein-coupled transmembrane receptor Mas (Chappell 2007; Ferreira et al. 2012b; Santos et al. 2003). In mammals, the gene is expressed predominantly in testis and in distinct areas of the forebrain, such as the hippocampus and amygdala, and less strongly but detectable in kidney, heart, lung, liver, spleen, tongue, and skeletal muscle (Metzger et al. 1995; Munoz et al. 2010; Villar and Pedersen 1994). Interestingly, Mas expression has been described to be modulated by pathological conditions in cardiac muscle and depending on the stimulus can be observed as an increase or a diminution (Chen et al. 2013; Dias-Peixoto et al. 2012).

Several studies have indirectly demonstrated the expression of Mas in skeletal muscle (Acuna et al. 2014; Morales et al. 2014; Prasannarong et al. 2012), although recently its level in rat soleus muscle was reported (Echeverria-Rodriguez et al. 2013). Other studies have demonstrated the expression of Mas in skeletal muscle through its participation in the beneficial effects of its ligand Ang (1–7) (Munoz et al. 2010, 2012). In this context, the participation of Mas was reported in the Ang (1–7)-dependent regulation of the insulin-signalling pathway and in glucose transport in skeletal muscle (Munoz et al. 2010, 2012; Prasannarong et al. 2012). Our group recently demonstrated that Ang (1–7), through Mas, counteracts the effects and signalling dependent on AngII in skeletal muscle tissue and cells (Morales et al. 2014), which is according to the report by Acuna et al. (2014) showing the critical role of Mas in controlling skeletal muscle damage and fibrosis by Ang (1–7) in Duchenne muscular dystrophy.

Despite the participation of the classical RAS axis in skeletal muscle atrophy, the participation and expression of Mas in muscle wasting has not been described. The purpose of this study was to assess the expression of Mas in skeletal muscle atrophy caused by several stimuli, such as AngII, lipopolysaccharide (LPS), and immobilization. Our results demonstrate that AngII and LPS increase mRNA and protein levels of Mas in myotubes in vitro and gastrocnemius and tibialis anterior muscles. In addition, in a model of unilateral immobilization by casting, Mas expression is upregulated. This is the first report that demonstrates a regulation of Mas expression under atrophic conditions.

Materials and methods

Animals

We used the C57BL/10J (12-week old) strain of mice (male). Animals were kept at room temperature with a 24 h night–day cycle, water available ad libitum, and paired feeding with pellets. The animals were randomized and separated into experimental groups (four to six animals/group), and three independent experiments were performed. At the end of each experiment, the animals were euthanized under anaesthesia and the gastrocnemius and tibialis anterior muscles were dissected, removed, and rapidly frozen and stored at −80 °C until processing (Morales et al. 2013a). All protocols were conducted in strict accordance and with the formal approval of the Animal Ethics Committee at the Universidad Andrés Bello.

Treatments

AngII (1 μg/kg/min) was osmotically infused through micropumps (Alzet-Durect, USA) implanted under ketamine/xylazine anaesthesia in the dorsal area of the animal for 1 or 14 days (Acuna et al. 2014). LPS from E. coli (Sigma, USA; 1 mg/kg) was i.p injected for 18 h or 14 days. Unilateral immobilization was performed in the lower hindlimb for 14 days using a 3M™ Scotchcast™ Soft Cast Casting Tape (Madaro et al. 2008).

Cell cultures

The skeletal muscle cell line C2C12 (American Type Culture Collection) was grown and differentiated until day 5, as described previously (Cabello-Verrugio et al. 2011; Painemal et al. 2013). The myotubes were incubated with 500 nM AngII (Sigma, USA) during 1 or 3 h for quantitative real-time PCR (RT-qPCR) or 24 h for Western blot. Myotubes were treated with 500 ng/ml of LPS (Sigma, USA) during 1–5 h for RT-qPCR or 48 h for Western blot. For treatment with the AngII receptor type 1 (AT-1) and type 2 (AT-2) blockers, the myotubes were pre-incubated for 1 h with losartan (10 μM) or PD-123319 (10 μM) (both from Tocris Bioscience, USA), respectively, and subsequently incubated with AngII. A similar procedure was followed for the inhibitor of the TLR-4 receptor CLI-095 (5 μM) (Invivogen, USA) prior to the incubation with LPS.

RNA isolation, reverse transcription and quantitative real-time PCR (RT-qPCR)

Total RNA was isolated from the gastrocnemius muscles, using TRIzol (Invitrogen, USA) according to the manufacturer’s instructions. The total RNA (1 μg) was reverse transcribed to cDNA using random hexamers and Superscript II reverse transcriptase (Invitrogen). TaqMan RT-qPCRs were performed in triplicate using an Eco Real-Time PCR System (Illumina, USA) with pre-designed primer sets for mouse Mas, atrogin-1, MuRF-1, and the housekeeping gene beta actin (TaqMan Assays-on-Demand; Applied Biosystems, USA). mRNA expression was quantified using the comparative ΔCt method (2−ΔΔCT) with beta actin as the reference gene. The mRNA levels were expressed relative to the mean expression in the control condition (Morales et al. 2012).

Immunoblot analysis

For the skeletal muscle extracts, the gastrocnemius muscles were homogenized in Tris–EDTA buffer with a cocktail of protease inhibitors and 1 mM PMSF. Proteins were subjected to SDS-PAGE, transferred onto PDVF membranes (Millipore, USA), and probed with goat anti-Mas (1:500) (sc-54848, Santa Cruz Biotechnology, USA) and mouse anti-tubulin (1:5,000) (sc-5286, Santa Cruz Biotechnology, USA). All immunoreactions were visualized by enhanced chemiluminescence (Thermo Scientific, USA).

Immunohistochemical analysis

For immunohistochemistry, fresh-frozen gastrocnemius and tibialis anterior muscles cryosections (7 μm) were fixed in acetone, incubated for 1 h with anti-Mas (1:50) (sc-54848, Santa Cruz Biotechnology, USA) in 5 % goat serum in PBS, and blocked for 15 min in methanol–H2O2 3 %. After 1 h incubation with rabbit anti-goat-HRP (Thermo Scientific, USA) followed by 30 min incubation with Envision™ Dual Link System-HRP (Dako, USA), enzyme activity was detected by the use of a 3′,3′-diaminobenzidine tetrahydrochloride liquid system (Dako). Nuclei were stained with haematoxylin (Morales et al. 2011).

Skeletal muscle histology

Fresh-frozen gastrocnemius and tibialis anterior muscles were sectioned, and cryosections (7 µm) were placed on glass slides. Haematoxylin and eosin staining was performed according to standard procedures (Cabello-Verrugio et al. 2012b).

Fibre diameter determination and quantification

Fibre diameter was detected by the analysis of the H&E staining. Briefly, the sections stained with H&E were viewed and photographed on the Motic BA310 microscope (Motic, Hong Kong). Fibre sizes were determined using the ImageJ software (NIH, USA) on five randomly captured images of gastrocnemius and tibialis anterior muscles of each experimental condition (in a blind fashion). Fibres were manually selected and the minimal Feret’s diameter of each fibre was quantified by the ImageJ software (Morales et al. 2013b).

Statistics

Statistical analysis was evaluated using the Student’s t test or the one-way analysis of variance (ANOVA) with a post hoc Bonferroni multiple-comparison test (Sigma Stat software). A difference was considered statistically significant at a P value <0.05.

Results

Angiotensin II increases the expression of the Mas receptor in skeletal muscle in vitro and in vivo

We evaluated the effects of AngII on the mRNA levels of Mas receptor in C2C12 myotubes. For this, the cells were incubated with AngII for several times for RNA extraction. Figure 1a shows an increment in Mas expression between 1 and 9 h after AngII incubation, reaching a maximum effect of 3.5-fold at 3 h and a return to basal levels at 14 h. This increase was dependent on the AT-1 receptor (Fig. S1), which is demonstrated by the complete inhibition of Mas upregulation induced by AngII in the presence of losartan (an AT-1 receptor blocker) but not with PD123319 (an AT-2 receptor blocker). At the protein level, AngII induces the expression of Mas receptor in the myotubes twofold (Fig. 1b, c). In addition, an increase in atrogin-1 (Fig. 1d) and MuRF-1 (Fig. 1e) was detected, which verified the atrophic effect of AngII.

Fig. 1
figure 1

AngII increases the expression of the Mas receptor in C2C12 myotubes. a C2C12 myotubes from day 5 were incubated with AngII (500 nM) for the times indicated in the figure. At the end of this treatment, the mRNA levels of Mas were determined by RT-qPCR using β-actin as the reference gene. The expression was expressed as the fold of induction, normalized to the levels in the control cells. The values correspond to the mean ± SD of three independent experiments (*P < 0.05 relative to the control cells). b Myotubes were incubated with AngII (500 nM) for 24 h. At the end of the treatment, the levels of Mas were detected by Western blot analysis. The levels of tubulin are shown as the loading control. Molecular weights are indicated in kilodaltons (kDa). c The values of the quantification are expressed as the fold of induction relative to the control cells. The expressions of atrogin-1 (d) and MuRF-1 (e) were detected to corroborate the atrophic effect of AngII after 1- and 3-h treatment, respectively. The value corresponds to the mean ± SD of three independent experiments (*P < 0.05 relative to the control cells treated with the vehicle)

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To evaluate the effect of AngII on the Mas receptor in vivo, we administrated AngII to mice through osmotic pumps. Figure 2a shows a twofold increase of Mas mRNA levels in gastrocnemius muscles of mice treated with AngII relative to control mice. This result is according to the twofold increase induced by AngII in the protein levels of Mas (Fig. 2b, c). The increase of Mas levels induced by AngII was corroborated in gastrocnemius muscle sections by immunohistochemistry (Fig. 2d) which was also observed in tibialis anterior (Fig. S2a). Atrophy induced by AngII was determined by a decrease in fibre diameter in gastrocnemius (Fig. 2e, f) and in tibialis anterior (Fig. S2b), and also for an increase in the expression of atrogin-1 (Fig. 2g) and MuRF-1 (Fig. 2h). Together, these results suggest that AngII increases Mas expression concomitant with the induction of atrophy in vitro and in vivo.

Fig. 2
figure 2

Systemic administration of AngII in mice increases the expression of the Mas receptor in skeletal muscle. C57BL10 mice were systemically treated with the vehicle (control; Ctrl) or AngII for 1 or 14 days, as described in “Materials and methods”. At the end of the treatment, the muscle gastrocnemius was removed and further processed. a mRNA levels of Mas were determined after 1 day of AngII administration by RT-qPCR using β-actin as the reference gene. The expression was expressed as the fold of induction, normalized to the levels in the control cells. Values correspond to the mean ± SD of three independent experiments (*P < 0.05 relative to the control cells). b Protein levels of Mas receptor determined after 14 days of AngII administration by Western blot analysis. The levels of tubulin are shown as the loading control. Molecular weights are indicated in kilodaltons (kDa). c Quantitative analysis of the experiments shown in (b). Values are expressed as the fold of induction relative to the control cells. d Cryosections of the gastrocnemius obtained after 14 days of AngII administration were used to immunodetect Mas through immunohistochemical analysis. Nuclei were labelled with haematoxylin. The bar corresponds to 50 μm. The images are representative of three independent experiments, using four mice for each experimental condition. Upper ×10 magnification; lower ×40 magnification. e Histological analysis of gastrocnemius from control and AngII-treated mice for 14 days through haematoxylin and eosin stain in cryosections. The bar corresponds to 50 μm. The images are representative of three independent experiments, using three mice for each experimental condition. Upper ×10 magnification; lower ×40 magnification. f Quantitative analysis of the fibre diameter from three independent experiments using three mice for each experimental condition. The values are expressed as the percentage of the total fibres quantified (*P < 0.05 relative to the control). g Atrogin-1 and h MuRF-1 mRNA levels were determined by RT-qPCR at 1 day after treatment with AngII, using β-actin as the reference gene. Levels were expressed as the fold of induction, normalized to the levels in the control cells. Values correspond to the mean ± SD of three independent experiments (*P < 0.05 relative to the control)

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Expression of the Mas receptor is increased by lipopolysaccharide in myotubes and skeletal muscle

The effect of LPS on the mRNA levels of the Mas receptor was evaluated by RT-qPCR in C2C12 myotubes. Figure 3a shows a 2.9- and 2.2-fold transient increase in Mas expression at 3 and 4 h, respectively, after LPS incubation, reaching basal levels at 5 h. This increase was dependent on the TLR-4 receptor for LPS since complete inhibition of LPS-induced Mas expression was observed in the presence of CLI-095 (a blocker of TLR-4) (Fig. S3). In addition, LPS also increased Mas protein levels (Fig. 3b). This augmented level was twice that of the control myotubes (Fig. 3c) and was concomitant with the increase observed for atrogin-1 (Fig. 3d) and MuRF-1 (Fig. 3e). We then evaluated the effect of LPS injected i.p. in mice on the expression of Mas in gastrocnemius muscles. Mas mRNA levels (Fig. 4a) and protein levels (Fig. 4b, c) in gastrocnemius muscles of mice treated with LPS increased twofold relative to control mice. In addition, we observed a strong increase in Mas levels in the plasma membrane of fibres evaluated in muscle sections by immunohistochemical analysis (Fig. 4d). Similar results were obtained for Mas levels in muscle TA (Fig. S4a). The atrophy induced by LPS was corroborated by a decrease in fibre diameter in gastrocnemius (Fig. 4e, f) and tibialis anterior (Fig. S4b) and also for an increase in the expression of atrogin-1 (Fig. 4g) and MuRF-1 (Fig. 4h). Thus, our results obtained in vitro and in vivo suggest that LPS induces an increase of the expression of Mas receptor together with typical parameters of skeletal muscle atrophy.

Fig. 3
figure 3

Expression of Mas is upregulated by LPS in C2C12 myotubes. a C2C12 myotubes from day 5 were incubated with LPS (500 ng/ml) for the times indicated in the figure. At the end of this treatment, the mRNA levels of Mas were determined by RT-qPCR using β-actin as the reference gene. The expression was expressed as the fold of induction, normalized to the levels in the control cells. Values correspond to the mean ± SD of three independent experiments (*P < 0.05 relative to the control cells). b Myotubes were incubated with LPS (500 ng/ml) for 48 h. At the end of the treatment, the levels of Mas were detected by Western blot analysis. The levels of tubulin are shown as the loading control. c Values of the quantification are expressed as the fold of induction relative to the control cells. d Atrogin-1 and e MuRF-1 were detected to corroborate the atrophic effect of LPS. The value corresponds to the mean ± SD of three independent experiments (*P < 0.05 relative to the control cells treated with the vehicle)

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Fig. 4
figure 4

Mas receptor levels are augmented in LPS-induced skeletal muscle atrophy. C57BL10 mice were intraperitoneally injected with LPS (1 mg/kg) for 18 h or 14 days, as described in “Materials and methods”. At the end of the treatment, muscle gastrocnemius was removed and processed. a mRNA levels of Mas were determined after 18 h of LPS treatment by RT-qPCR using β-actin as the reference gene. The expression was expressed as the fold of induction, normalized to the levels in the control cells. Values correspond to the mean ± SD of three independent experiments (*P < 0.05 relative to the control cells). b Mas receptor protein levels were evaluated by Western blot in gastrocnemius from control and LPS-treated mice for 14 days. Tubulin is shown as the loading control. Molecular weights are indicated in kilodaltons (kDa). c Quantitative analysis of the experiments shown in (b). The values are expressed as the fold of induction relative to the control cells. d Cryosections of the gastrocnemius obtained after 14 day of AngII administration were used to immunodetect Mas through immunohistochemical analysis. Nuclei were labelled with haematoxylin. The bar corresponds to 50 μm. The images are representative of three independent experiments, using four mice for each experimental condition. Upper ×10 magnification; lower ×40 magnification. e Histological analysis of gastrocnemius from control and AngII-treated mice for 14 days through haematoxylin and eosin stain in cryosections. The bar corresponds to 50 μm. The images are representative of three independent experiments, using three mice for each experimental condition. Upper ×10 magnification; lower ×40 magnification. f Quantitative analysis of the fibre diameter from three independent experiments using three mice for each experimental condition. The values are expressed as the percentage of the total fibres quantified (*P < 0.05 relative to the control). g Atrogin-1 and h MuRF-1 mRNA levels were determined by RT-qPCR at 18 h after treatment with LPS, using β-actin as the reference gene. The levels were expressed as the fold of induction, normalized to the levels in the control cells. Values correspond to the mean ± SD of three independent experiments (*P < 0.05 relative to the control)

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Expression of the Mas receptor is increased during immobilization-induced skeletal muscle atrophy

To evaluate the effect of immobilization on the expression of Mas, we detected its mRNA levels by RT-qPCR technique in gastrocnemius muscles. Mas mRNA levels in immobilized gastrocnemius muscles increased sixfold relative to the contralateral non-immobilized muscle (Fig. 5a) and protein levels increased 2.3-fold (Fig. 5b, c). In addition, we observed a strong increase in immunodetection of Mas levels in the plasma membrane of muscle fibres (Fig. 5d). Similar results were observed for Mas levels in immobilized tibialis anterior muscle (Fig. S5a). The atrophy induced by immobilization was corroborated by a decrease in fibre diameter in gastrocnemius (Fig. 5e, f) and tibialis anterior (Fig. S5b), and for an increase in the expression of atrogin-1 (Fig. 5g) and MuRF-1 (Fig. 5h). Together, our results suggest that Mas expression is increased under conditions of skeletal muscle atrophy by immobilization concomitant with typical parameters of skeletal muscle atrophy.

Fig. 5
figure 5

Mas receptor is upregulated in disuse-induced skeletal muscle atrophy. Skeletal muscle atrophy was induced in C57BL/10J mice by unilateral hindlimb immobilization by casting. a Mas receptor expression from gastrocnemius non-immobilizated (NI) and immobilizated (I) hindlimb for 24 h was determined by RT-qPCR using β-actin as the reference gene. The expression was expressed as the fold of induction, normalized to the levels in the control muscles. Values correspond to the mean ± SD of three independent experiments (*P < 0.05 relative to the control muscles). b Mas receptor protein levels were evaluated by Western blot in gastrocnemius of non-immobilizated and immobilizated hindlimb for 14 days. Tubulin is shown as the loading control. Molecular weights are indicated in kilodaltons (kDa). c Quantification of B. Values correspond to the mean ± SD of three independent experiments and are expressed as fold of induction relative to the expression in non-immobilizated gastrocnemius (NI) (*P < 0.05). d Mas receptor was detected by indirect immunohistochemical analysis in cryosections of gastrocnemius muscles from non-immobilizated and immobilizated hindlimb for 14 days. Nuclei were labelled with haematoxylin. The bar corresponds to 50 μm. The images are representative of three independent experiments, using three mice for each experimental condition. Upper ×10 magnification; lower ×40 magnification. e Histological analysis of non-immobilizated and immobilizated hindlimb for 14 days. Muscle cross sections were stained with haematoxylin and eosin to visualize muscle architecture. The bar corresponds to 50 μm. The images are representative of three independent experiments, using three mice for each experimental condition. Upper ×10 magnification; lower ×40 magnification. f Quantitative analysis of the fibre diameter from three independent experiments using three mice for each experimental condition. The values are expressed as the percentage of the total fibres quantified (*P < 0.05 relative to non-immobilizated muscles). g Atrogin-1 and h MuRF-1 ubiquitin ligase mRNA levels from gastrocnemius of non-immobilizated and immobilizated hindlimb for 18 h were determined by RT-qPCR using β-actin as the reference gene. The levels were expressed as the fold of induction, normalized to the levels in the control cells. Values correspond to the mean ± SD of three independent experiments (*P < 0.05 relative to the non-immobilizated muscles)

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Discussion

In this study, we show for the first time that the levels of Mas receptor for Ang (1–7) are regulated in skeletal muscle under atrophic conditions. We found that Mas expression in mRNA and protein levels increases in response to AngII, LPS, and immobilization together with the upregulation of typical parameters of skeletal muscle atrophy, such as the muscle-specific E3 ubiquitin ligases atrogin-1 and MuRF-1, or the decrease in fibre diameter.

The three different models of skeletal muscle atrophy used for us developed typical features of muscle wasting such as increase of UPS activity (specifically the increase in the expression of atrogin-1 and MuRF-1) and decrease of fibre size. Several signalling pathways dependent on AKT, reactive oxygen species (ROS), and p38 MAPK are involved in the regulation of the atrogin-1 and MuRF-1 expression (Foletta et al. 2011). Thus, it has been reported that AKT phosphorylation is key to prevent the induction of atrogin-1 gene expression by regulating FOXO activity (Foletta et al. 2011; Stitt et al. 2004). ROS is directly involved in the muscle proteolysis and in the increase of UPS in muscle wasting induced by AngII, immobilization, and LPS (Kondo et al. 1993, 1994; Powers et al. 2007, 2012; Sukhanov et al. 2011; Tisdale 2005; Yu et al. 2008). We have previously reported that AngII induces ROS in skeletal muscle cells through a mechanism dependent on NADPH oxidase which is decreased by Ang (1–7) (Morales et al. 2014). Furthermore, p38 MAPK has been demonstrated to be a key signalling in the induction of skeletal muscle atrophy, also in the increase of atrogin-1 and MuRF-1 by oxidative stress (Doyle et al. 2011; Glass 2005; McClung et al. 2010).

Several studies have shown the importance of the balance between the classical and non-classical RAS axes. In this context, our results showed that Mas, the main receptor of the non-classical RAS axis associated with beneficial effects in many tissues including skeletal muscle, is increased in skeletal muscle wasting. It has been reported that LPS induces the Mas upregulation in macrophages (Souza and Costa-Neto 2012) which is in agreement with our findings in skeletal muscle. Regarding to immobilization and AngII treatment, there are not reports showing increase of the Mas receptor levels. In general, the expression of Mas has been reported to be downregulated under some pathological conditions: in the dorsal medulla by exposure to glucocorticoid (Marshall et al. 2013), in cardiomyocytes by endothelin-1 (Chen et al. 2013), in myocardial infarction, cardiac hypertrophy, and damage elicited by isoproterenol treatment (Dias-Peixoto et al. 2012). Despite the most of the evidence suggests a decrease of Ang (1–7)/Mas axis in pathological status in which RAS is involved, there are some exceptions. Mas expression increases in ovary under treatment with chorionic gonadotropin (Pereira et al. 2009), in brain under acute ischaemic stroke (Lu et al. 2013), in heart of DOCA-salt rats (Dias-Peixoto et al. 2012), in hypertensive rats subjected to swimming training (Filho et al. 2008), in gastrointestinal smooth muscles of patients with achalasia (Casselbrant et al. 2014), in jejunal enterocytes from diabetic mellitus type 1 (Wong et al. 2012), and in severe acute pancreatitis (Wang et al. 2012). Thus, our study is not the only evidence of the Mas upregulation under pathological conditions.

There are few evidences about molecular mechanisms related to the increase of Mas gene expression. A recent study shows a mechanism involved in the epigenetic regulation of the Mas gene through nitration of ZNF274/KAP-1 (KRAB-associated protein-1) protein complex (Prokop et al. 2014). Further experiments could be performed to evaluate this regulatory mechanism of Mas receptor gene expression in skeletal muscle atrophy. In our study, we not have elucidated the signalling pathways involved in the increase of Mas reported in this work under different atrophic stimuli. One possibility is that the increase of Mas levels in vivo with could be explained as an indirect effect dependent on IL-1β production, which is increased by AngII, LPS and immobilization (Borge et al. 2009; Bruells et al. 2013; Cabello-Verrugio et al. 2012a; Wang et al. 2014) and upregulates Mas mRNA and protein expression (Ender et al. 2014). Another possibility is the participation of p38MAPK that is has been recently described to be involved in the transcriptional upregulation of Mas in dorsal root ganglia neurons (Cao et al. 2013). Interestingly, p38 MAPK has been found to be a key player in skeletal muscle atrophy induced by immobilization, AngII treatment and LPS (Doyle et al. 2011; Eley et al. 2008; Kim et al. 2009). Thus, we can speculate that p38MAPK could participate of the Mas upregulation in the atrophic models used for us. Thus, further studies must be performed to evaluate the role of p38 MAPK in this process.

Our results show that Mas is located at least, in the plasma membrane of muscle fibres of the gastrocnemius and tibialis anterior, which indicate that muscle fibres have the ability to respond to Ang (1–7). This fact can have high relevance to evaluate the effect of Ang (1–7) administration on the weakness associated with skeletal muscle atrophy. A pathological condition in which skeletal muscle atrophy is observed is the dystrophic muscles of aged mdx mice (Mouisel et al. 2010). Recent reports have shown a decrease in fibrosis and damage of the skeletal muscles of dystrophic mice treated with Ang (1–7) which are mediated through the Mas receptor (Acuna et al. 2014; Sabharwal et al. 2014). In this context, it is important to determine the signalling pathway activated by Ang (1–7) through Mas that could be involved in a possible effect of Ang (1–7) on skeletal muscle atrophy. Ang (1–7) binds to the Mas receptor which activates intracellular signalling pathways through nitric oxide synthase and AKT (Sampaio et al. 2007), GSK3β (Gomes et al. 2010), SHP-1 (Gava et al. 2009), or by inhibition of NF-κBeta pathway (El-Hashim et al. 2012; Jiang et al. 2012; Santos et al. 2013). In regard to skeletal muscle atrophy, AKT signalling is decreased which activates FOXO proteins and produces the increment of the atrogin-1 and MuRF-1 gene expression (Sandri et al. 2004). In this context, we have data that demonstrate Ang (1–7) has an anti-atrophic effect on AngII-induced muscle wasting through the AKT phosphorylation (data not shown). It has been reported that Ang (1–7) can modulate the NF-κBeta signalling in liver, brain, and lung (El-Hashim et al. 2012; Jiang et al. 2012; Santos et al. 2013). Thus, it would be important to evaluate the effect of Ang (1–7) on the NF-κBeta signalling pathway because it, among others, is directly involved in the modulation of MuRF-1 gene expression.

Our study demonstrates for the first time that the expression of the Mas receptor is modulated under several conditions of muscle wasting and increases the knowledge describing a differential expression of a Mas receptor that might be involved in the pathophysiology of skeletal muscle atrophy.