Glucocorticoid Receptor Signaling Impairs Protein Turnover Regulation in Hypoxia-Induced Muscle Atrophy in Male Mice

Glucocorticoid Receptor Signaling Impairs Protein Turnover Regulation in Hypoxia-Induced Muscle... Abstract Hypoxemia may contribute to muscle wasting in conditions such as chronic obstructive pulmonary disease. Muscle wasting develops when muscle proteolysis exceeds protein synthesis. Hypoxia induces skeletal muscle atrophy in mice, which can in part be attributed to reduced food intake. We hypothesized that hypoxia elevates circulating corticosterone concentrations by reduced food intake and enhances glucocorticoid receptor (GR) signaling in muscle, which causes elevated protein degradation signaling and dysregulates protein synthesis signaling during hypoxia-induced muscle atrophy. Muscle-specific GR knockout and control mice were subjected to normoxia, normobaric hypoxia (8% oxygen), or pair-feeding to the hypoxia group for 4 days. Plasma corticosterone and muscle GR signaling increased after hypoxia and pair-feeding. GR deficiency prevented muscle atrophy by pair-feeding but not by hypoxia. GR deficiency differentially affected activation of ubiquitin 26S-proteasome and autophagy proteolytic systems by pair-feeding and hypoxia. Reduced food intake suppressed mammalian target of rapamycin complex 1 (mTORC1) activity under normoxic but not hypoxic conditions, and this retained mTORC1 activity was mediated by GR. We conclude that GR signaling is required for muscle atrophy and increased expression of proteolysis-associated genes induced by decreased food intake under normoxic conditions. Under hypoxic conditions, muscle atrophy and elevated gene expression of the ubiquitin proteasomal system–associated E3 ligases Murf1 and Atrogin-1 are mostly independent of GR signaling. Furthermore, impaired inhibition of mTORC1 activity is GR-dependent in hypoxia-induced muscle atrophy. Chronic obstructive pulmonary disease (COPD) is globally a leading cause of morbidity and disability (1). Weight loss and muscle atrophy are common features of advanced COPD. Muscle atrophy significantly increases disease burden and is a strong predictor of mortality (1–5). In addition to nutritional imbalance, glucocorticoid use, systemic inflammation, and progressive inactivity, hypoxemia has been implicated as a trigger for muscle atrophy in COPD patients (6). Previously, it was shown that hypoxia causes muscle atrophy in rat models (7, 8). Our group confirmed these findings in a mouse model of normobaric hypoxia (9) and revealed that muscle atrophy was attributable to a hypoxia-induced reduction of food intake and hypoxia-specific effects. The mechanisms of fasting-induced muscle atrophy involve elevated glucocorticoid concentrations and glucocorticoid receptor (GR) signaling (10), which affect protein synthesis and proteolysis in skeletal muscle. Adrenalectomized rats display reduced muscle protein breakdown, which is restored by glucocorticoid administration (11). Moreover, in muscle-specific (MLC-cre) GR knockout mice, fasting-induced reductions in fiber cross-sectional area (FCSA) were absent and the activation of the proteolytic machinery was attenuated (12). GR affects muscle protein turnover by genomic and nongenomic actions (13). Genomic effects include activation of protein degradation by the ubiquitin proteasomal system (UPS). Forkhead box protein O1 (FOXO1) is upregulated in a GR-specific manner and contributes to the subsequent induction of expression of atrogenes, such as tripartite motif containing 63 (Trim63/Murf1), F-box only protein 32 (Fbxo32/Mafbx/Atrogin-1), muscle ubiquitin ligase of SCF complex in atrophy-1 (Musa1, Fbxo30), and specific of muscle atrophy and regulated by transcription (Smart/Fbxo21) (14–17). GR is also involved in upregulation of Klf15 messenger RNA (mRNA) expression. In addition to its involvement in the control of proteolysis via regulation of MuRF1 and Atrogin-1, Krüppel-like factor 15 (KLF15) also contributes to the regulation of protein synthesis (18). Glutamate-ammonia ligase (GLUL) is an enzyme involved in muscle amino acid metabolism during fasting (19), the expression of which is also stimulated by glucocorticoids through GR-responsive elements (GRE) in the Glul promoter (20). Myostatin (MSTN) is associated with dexamethasone- (21) and hypoxia-induced muscle atrophy (8) and expression is increased by activation of the GR (21, 22). Furthermore, GR-mediated transcription of DNA damage–inducible transcript 4 protein (Ddit4/Redd1) followed by inhibition of mammalian target of rapamycin (mTOR) results in suppressed protein synthesis signaling (23, 24) and activation of autophagy (25). Nongenomic GR actions involve suppression of protein synthesis through the inhibition of the phosphatidylinositol-4,5-bisphosphate 3-kinase (PI3K)/protein kinase B (PKB/AKT)/TSC1/2/mTOR signaling pathway, in which GR binds to PI3K subunits p110 and p85 and prevents association with insulin receptor substrate-1 (IRS-1) (12). As we previously reported (26), hypoxia-exposed mice show altered activation of signaling pathways of proteolysis and protein synthesis in skeletal muscle. Furthermore, it has been shown that hypoxia can elevate blood glucocorticoid concentrations (27–30). We therefore hypothesized that hypoxia elevates circulating corticosterone concentrations resulting in enhanced GR signaling in muscle, which causes elevated protein degradation signaling and also dysregulates protein synthesis signaling during hypoxia-induced muscle atrophy. Methods Animals and tissue collection All mouse studies were carried out according to a protocol approved by the Institutional Animal Care Committee of Maastricht University. Electroporation of the muscles was done in 12-week-old C57BL/6J male mice (Charles River Laboratories). Twelve-week-old male, muscle‐specific GR knockout (mGRKO) mice were generated by crossing transgenic Mlc1f‐Cre+/‐ mice (Cre recombinase under control of the fast myosin light chain 1 promoter) (31) with mice bearing floxed Gr alleles (Grfl/fl) (32) on a C57BL/6 background, resulting in the ablation of GR in MyHC type II (fast) muscle cells, which accounts for 95% of the muscle fibers (33). Twenty-four control (littermates) and 24 mGRKO mice were randomly divided into three groups: normoxia, normoxia pair-fed to hypoxic animals, and normobaric hypoxia. The pair-fed group was included to assess the effects of reduced food intake induced by hypoxia. All mice were housed in sets of four animals per cage in experimental chambers at 21°C with a 12-hour dark/light cycle. Mice received standard chow (V1534-000 Ssniff R/M-H; Ssniff Spezialdiäten) and water ad libitum. After 5 days of acclimatization, normoxic and pair-fed groups were kept at ambient air (21% O2) and the hypoxic group to normobaric hypoxia. Using the proOX system P110 (BioSpherix), oxygen levels were reduced in a stepwise manner to 12% (day 0), 10% (day 1), and finally 8% (61 mm Hg) on day 2, which was then maintained for the remainder of the experiment. To assess the effects of reduced food intake during hypoxia, pair-fed animals received the amount of food consumed by the hypoxic mice daily at a standardized time (5:00 pm). Tissue and plasma were isolated on day 4 and frozen immediately in liquid nitrogen for further analysis. Timing of the tissue collection was standardized in the early dark phase (9:00 to 10:00 am) to reduce hormonal variation. Tissue weights were corrected for body weight at the start of the experiment. Corticosterone measurement Plasma corticosterone concentrations were determined by high-performance liquid chromatography. A carboxylic acid derived from cortisol (4-androsten-11β,17α-diol-3-one-17β-carboxylic acid) was synthetized and purified as described previously (34) and was added as internal control to plasma and corticosterone standards. Samples and standards were acidified with phosphoric acid, and steroid hormones were subsequently extracted from plasma with diethyl ether and dried with nitrogen gas. The residues were dissolved in acetone and incubated with sulfuric acid for 20 minutes to convert the steroids into fluorescent products (35). After the addition of 100 volumes of diethylether and 5 volumes of water, the fluorescent products were extracted from the aqueous sulfuric acid phase. After evaporating the organic phase under a stream of nitrogen, the residues were dissolved and analyzed by high-performance liquid chromatography as described (34). Ratios of corticosterone and internal standard peak areas were calculated and converted into concentrations using calibration curves. In vivo electroporation of gastrocnemius muscle To assess the activity of the GR in response to hypoxia and reduced food intake, a reporter plasmid was electroporated into the gastrocnemius muscle. Plasmids were prepared using an endotoxin-free maxi prep kit (Qiagen) and dissolved in 0.9% saline. The plasmid mix consisted of 1:1 GRE-luciferase construct (36) and pVAX-lacZ plasmid (Invitrogen) encoding β-galactosidase. Mice were anesthetized with isoflurane and injected subcutaneously with buprenorphine (Temgesic) as analgesic. Skin covering the gastrocnemius muscle was shaved and disinfected. Using a bent 0.3-mL syringe with a 30-gauge needle (Becton Dickinson), five times 10 µL of plasmid (50 µg total) was injected at different sites into the gastrocnemius muscle. Custom-built mouse electrodes with electrolyte crème (Parker Laboratories) were placed on both sides of the gastrocnemius muscle on top of the skin. Eight pulses of 150 V/cm, with 20-ms pulse length and a 1-second interval were applied using the BTX square wave electroporator (BTX/Harvard Apparatus). The direction of the pulse was inverted after four pulses to increase efficiency. After 21 days of recovery, mice were exposed to experimental conditions. Muscles were snap-frozen in liquid nitrogen and ground to a powder using an N2-cooled steel mortar. Muscle powder was lysed in passive lysis buffer (Promega) and assessed for luciferase (Promega) and β‐galactosidase (Tropix) activity. Real-time quantitative polymerase chain reaction Total RNA was isolated using TRI Reagent (Sigma-Aldrich) and further purified by precipitation with 2 M LiCl. Complementary DNA synthesis was performed with random hexamer primers on denatured RNA using the transcriptor first-strand complementary DNA synthesis kit (Roche). Real-time quantitative polymerase chain reaction (qPCR) was performed in the iQ5 thermal cycler (Bio-Rad) using the qPCR SYBR Green fluorescein mix (Abgene). qPCR primers were designed using Primer Express 2.0 software (Applied Biosystems) and ordered from Sigma Genosys (Table 1). All primers were intron spanning to avoid contamination of the amplification products with genomic DNA. Expression of genes was normalized to 18S ribosomal RNA. mRNA expression data in the figures are shown as fold change of the wild-type normoxic group. Table 1. Genes Used in This Study Gene  NCBI Access No.  Forward Primer (5′ to 3′)  Reverse Primer (5′ to 3′)  18S  NR_003278.1  AGTTAGCATGCCAGAGTCTCG  TGCATGGCCGTTCTTAGTTG  Gr  NM_008173.3  CGCCAAGTGATTGCCGC  TGTAGAAGGGTCATTTGGTCATCCA  Glul  NM_008131.3  GGCCATGCGGGAGGAGA  GGTGCCTCTTGCTCAGTTTGTCA  Foxo1  NM_019739.3  AAGAGCGTGCCCTACTTCAAG  CCATGGACGCAGCTCTTCTC  Klf15  NM_023184.3  TGCAGCAAGATGTACACCAAGAG  ATCGCCGGTGCCTTGAC  Redd1  NM_029083.2  CGGGCCGGAGGAAGACT  CTGCATCAGGTTGGCACACA  Murf1  NM_001039048.2  TGTCTGGAGGTCGTTTCCG  CTCGTCTTCGTGTTCCTTGC  Atrogin-1  NM_026346.2  ACCGGCTACTGTGGAAGAGA  CCTTCCAGGAGAGAATGTGG  Map1lc3B  NM_026160.4  GAGCAGCACCCCACCAAGAT  CGTGGTCAGGCACCAGGAA  Bnip3  NM_009760.4  CCATGTCGCAGAGCGGG  GACGGAGGCTGGAACGC  Musa1  NM_027968.3  GGACGTTTGTGGCAGTTTACTTC  GCAGTACTGAATCGCCATACCTTC  Smart  NM_145564.3  CACAGGGATGTCTGCTACTC  ACACAGTTGTAGCCGTACCTC  Mstn  NM_010834.2  GGCCATGATCTTGCTGTAACCT  CGGCAGCACCGGGATT  Smad3  NM_016769  CCCGAGAACACTAACTTCCCTG  AACCTGCGTCCATGCTGTG  Gene  NCBI Access No.  Forward Primer (5′ to 3′)  Reverse Primer (5′ to 3′)  18S  NR_003278.1  AGTTAGCATGCCAGAGTCTCG  TGCATGGCCGTTCTTAGTTG  Gr  NM_008173.3  CGCCAAGTGATTGCCGC  TGTAGAAGGGTCATTTGGTCATCCA  Glul  NM_008131.3  GGCCATGCGGGAGGAGA  GGTGCCTCTTGCTCAGTTTGTCA  Foxo1  NM_019739.3  AAGAGCGTGCCCTACTTCAAG  CCATGGACGCAGCTCTTCTC  Klf15  NM_023184.3  TGCAGCAAGATGTACACCAAGAG  ATCGCCGGTGCCTTGAC  Redd1  NM_029083.2  CGGGCCGGAGGAAGACT  CTGCATCAGGTTGGCACACA  Murf1  NM_001039048.2  TGTCTGGAGGTCGTTTCCG  CTCGTCTTCGTGTTCCTTGC  Atrogin-1  NM_026346.2  ACCGGCTACTGTGGAAGAGA  CCTTCCAGGAGAGAATGTGG  Map1lc3B  NM_026160.4  GAGCAGCACCCCACCAAGAT  CGTGGTCAGGCACCAGGAA  Bnip3  NM_009760.4  CCATGTCGCAGAGCGGG  GACGGAGGCTGGAACGC  Musa1  NM_027968.3  GGACGTTTGTGGCAGTTTACTTC  GCAGTACTGAATCGCCATACCTTC  Smart  NM_145564.3  CACAGGGATGTCTGCTACTC  ACACAGTTGTAGCCGTACCTC  Mstn  NM_010834.2  GGCCATGATCTTGCTGTAACCT  CGGCAGCACCGGGATT  Smad3  NM_016769  CCCGAGAACACTAACTTCCCTG  AACCTGCGTCCATGCTGTG  Abbreviation: NCBI, National Center for Biotechnology Information. View Large Western blotting Western blotting was performed according to procedures previously described (26). In short, gastrocnemius muscle was ground to a powder and lysed. Total protein concentration was determined and 12.5 μg per lane was used for Western blotting. The membrane was incubated overnight at 4°C with primary antibodies (Table 2). Blots were probed with a horseradish peroxidase–conjugated secondary antibody and visualized by chemiluminescence in an LAS-3000 luminescent image analyzer (Fujifilm). Bands were quantified using AIDA software (Fujifilm). Figures display ratios of phosphorylated over total protein of interest; in case total levels of the protein of interest are altered between groups, total levels are also shown separately. PonceauS staining was used to control and correct for protein loading. Protein data in the figures are shown as fold change of the wildtype normoxic group. Protein ratios are calculated by dividing the phosphorylated protein by the total amount of protein. Table 2. Antibodies Used in This Study Target  Product No.  Manufacturer  Made in  Size (kDa)  RRID  GR  12041  Cell Signaling Technology  Rabbit  91–94  AB_2631286  FOXO1  2880  Cell Signaling Technology  Rabbit  78–82  AB_2106495  GLUL  G45020  BD Transduction Laboratory  Mouse  40  AB_2313767  S6  2217  Cell Signaling Technology  Rabbit  32  AB_331355  p-S6 (S235/236)  4856  Cell Signaling Technology  Rabbit  32  AB_2181037  4E-BP1  9452  Cell Signaling Technology  Rabbit  15–20  AB_331692  p-4E-BP1 (S65)  9451  Cell Signaling Technology  Rabbit  15–20  AB_330947  p-4E-BP1 (T37/46)  9459  Cell Signaling Technology  Rabbit  15–20  AB_330985  mTOR  2983  Cell Signaling Technology  Rabbit  289  AB_2105622  p-mTOR (S2448)  2971  Cell Signaling Technology  Rabbit  289  AB_330970  p-mTOR (S2481)  2974  Cell Signaling Technology  Rabbit  289  AB_2231885  AKT  9272  Cell Signaling Technology  Rabbit  60  AB_329827  p-AKT (S473)  9271  Cell Signaling Technology  Rabbit  60  AB_329825  TSC2  4308  Cell Signaling Technology  Rabbit  200  AB_10547134  p-TSC2 (S939)  3615  Cell Signaling Technology  Rabbit  200  AB_2207796  p-TSC2 (T1462)  3617  Cell Signaling Technology  Rabbit  200  AB_490956  p-TSC2 (S1387)  5584  Cell Signaling Technology  Rabbit  200  AB_10698883  Target  Product No.  Manufacturer  Made in  Size (kDa)  RRID  GR  12041  Cell Signaling Technology  Rabbit  91–94  AB_2631286  FOXO1  2880  Cell Signaling Technology  Rabbit  78–82  AB_2106495  GLUL  G45020  BD Transduction Laboratory  Mouse  40  AB_2313767  S6  2217  Cell Signaling Technology  Rabbit  32  AB_331355  p-S6 (S235/236)  4856  Cell Signaling Technology  Rabbit  32  AB_2181037  4E-BP1  9452  Cell Signaling Technology  Rabbit  15–20  AB_331692  p-4E-BP1 (S65)  9451  Cell Signaling Technology  Rabbit  15–20  AB_330947  p-4E-BP1 (T37/46)  9459  Cell Signaling Technology  Rabbit  15–20  AB_330985  mTOR  2983  Cell Signaling Technology  Rabbit  289  AB_2105622  p-mTOR (S2448)  2971  Cell Signaling Technology  Rabbit  289  AB_330970  p-mTOR (S2481)  2974  Cell Signaling Technology  Rabbit  289  AB_2231885  AKT  9272  Cell Signaling Technology  Rabbit  60  AB_329827  p-AKT (S473)  9271  Cell Signaling Technology  Rabbit  60  AB_329825  TSC2  4308  Cell Signaling Technology  Rabbit  200  AB_10547134  p-TSC2 (S939)  3615  Cell Signaling Technology  Rabbit  200  AB_2207796  p-TSC2 (T1462)  3617  Cell Signaling Technology  Rabbit  200  AB_490956  p-TSC2 (S1387)  5584  Cell Signaling Technology  Rabbit  200  AB_10698883  Abbreviations: 4E-BP1, 4E-binding protein 1; RRID, Research Resource Identifier. View Large FCSA determination Gastrocnemius muscle was embedded in Tissue‐Tek (Sakura Finetek) and sectioned on a Leica CM3050 S cryostat at −20°C. Subsequently, serial cross‐sections (5 μm) were stained with anti‐laminin (no. L‐9393; Sigma‐Aldrich) to determine the FCSA The sections were incubated with Alexa Fluor 350 (no. A‐21426; Invitrogen) as secondary antibody. Digital images of the stained sections were taken at ×200 total magnification using an Eclipse E800 microscope (Nikon) connected to a digital camera (DXM, 1200 NF, Nikon). The FCSA in the glycolytic region of the gastrocnemius was measured for >100 individual fibers per animal, using the Lucia software (version 4.81). A distribution curve was composed of all measured fibers within the groups (>800 fibers). Statistical analysis Data are shown as means ± standard error of the mean. Comparisons were computed with SPSS version 20. Statistical significance between groups within a genotype (wild-type or mGRKO) was tested using a one-way analysis of variance (ANOVA) with a post hoc test. The type of post hoc analysis was chosen on the basis of the data variance (Levene’s test), with the Tukey test for data with equal variance and the Games–Howell test for all other data. Statistical significance between genotypes of the normoxic group was assessed using the independent samples t test. A two-way ANOVA was used to determine the effect of GR deficiency on the response of the individual treatments. A χ2 test was used to compare FCSA distributions. A P values < 0.05 was considered to be statistically significant and 0.05 ≤ P ≤ 0.1 as indicating a trend. Results Hypoxia increases plasma corticosterone concentrations and induces muscle GR signaling Four days of hypoxia resulted in elevated corticosterone concentrations (Fig. 1A). This increase was in part attributable to the reduced food intake (pair-fed, 2.0-fold; hypoxia, 3.0-fold). To address whether the elevated circulating corticosterone concentrations activated muscle GR signaling in vivo, a GRE reporter (luciferase) plasmid was electroporated into the gastrocnemius muscle prior to 4 days exposure to normoxic, hypoxic, or pair-fed conditions (Fig. 1B). The transcriptional activity of GR was increased under hypoxic conditions (1.9-fold) and even more so in the pair-fed group (3.1-fold). Figure 1. View largeDownload slide Hypoxia raises corticosterone plasma concentrations and induces muscle GR signaling. (A) Plasma corticosterone concentration at day 4. (B) GR transcriptional activity in gastrocnemius muscle at day 4. GRE reporter plasmid (luciferase) data were corrected for control (β-galactosidase) plasmid data. Significant differences between groups at a given time point are indicated as follows: *P < 0.05; trends (P ≤ 0.1) are indicated by the specific P value (n = 8 per group). Figure 1. View largeDownload slide Hypoxia raises corticosterone plasma concentrations and induces muscle GR signaling. (A) Plasma corticosterone concentration at day 4. (B) GR transcriptional activity in gastrocnemius muscle at day 4. GRE reporter plasmid (luciferase) data were corrected for control (β-galactosidase) plasmid data. Significant differences between groups at a given time point are indicated as follows: *P < 0.05; trends (P ≤ 0.1) are indicated by the specific P value (n = 8 per group). GR target genes are differentially expressed in mGRKO To assess the importance of GR signaling during hypoxia-induced muscle atrophy, mGRKO mice were used. In these mice, GR protein (Fig. 2A) and Gr mRNA (Fig. 2B) were reduced by 60% in the gastrocnemius muscles. Interestingly, both in control and mGRKO mice, Gr mRNA levels were increased by hypoxia, but not by reduced food intake alone. Baseline plasma corticosterone concentrations were comparable in both genotypes and showed similar increases in response to reduced food intake and hypoxia (Fig. 2C). mRNA expression of postulated GR target genes related to muscle protein turnover was evaluated. Both Glul (Fig. 2D) and Foxo1 (Fig. 2E) expression were GR-dependent, as no increased expression was found by hypoxia and reduced food intake in mGRKO mice. Protein levels of FOXO1 and GLUL also show a GR-dependent increase in response to hypoxia (Supplemental Fig. 1). Overall, the expression of Mstn was reduced by GR deficiency, although the responses to reduced food intake and hypoxia remained similar (Fig. 2F). In contrast, expression of Klf15 (Fig. 2G) and Redd1 (Fig. 2H) was similar in response between control and mGRKO mice and therefore not GR-dependent. To conclude, corticosterone concentrations increased equally in control and mGRKO mice in response to hypoxia. Hypoxia-induced expression of Glul and Foxo1 in skeletal muscle was attributable to the reduced food intake and was GR-dependent. Figure 2. View largeDownload slide GR target genes are differentially expressed in mGRKO. (A) GR protein in control and mGRKO gastrocnemius muscle. (B) mRNA expression of Gr. (C) Plasma corticosterone concentration. mRNA expression of (D) Glul, (E) Foxo1, (F) Mstn, (G) Klf15, and (H) Redd1 is shown. All mRNA concentrations were corrected by 18S concentration. Significant differences between groups within control or mGRKO mice are indicated by asterisks on horizontal lines; basal differences in normoxia groups between control and mGRKO mice are indicated by a hash sign: */#P < 0.05, **/##P ≤ 0.01, ***P ≤ 0.001; trends (P ≤ 0.1) are indicated by the specific P value. Significant effects (P ≤ 0.05) of GR deficiency on the response of the treatment analyzed by two-way ANOVA are indicated as follows: &, normoxia vs hypoxia; $, pair-fed vs hypoxia (n = 6–8). Figure 2. View largeDownload slide GR target genes are differentially expressed in mGRKO. (A) GR protein in control and mGRKO gastrocnemius muscle. (B) mRNA expression of Gr. (C) Plasma corticosterone concentration. mRNA expression of (D) Glul, (E) Foxo1, (F) Mstn, (G) Klf15, and (H) Redd1 is shown. All mRNA concentrations were corrected by 18S concentration. Significant differences between groups within control or mGRKO mice are indicated by asterisks on horizontal lines; basal differences in normoxia groups between control and mGRKO mice are indicated by a hash sign: */#P < 0.05, **/##P ≤ 0.01, ***P ≤ 0.001; trends (P ≤ 0.1) are indicated by the specific P value. Significant effects (P ≤ 0.05) of GR deficiency on the response of the treatment analyzed by two-way ANOVA are indicated as follows: &, normoxia vs hypoxia; $, pair-fed vs hypoxia (n = 6–8). Hypoxia-induced muscle atrophy is not prevented by muscle GR deficiency Gastrocnemius muscle weights of mGRKO animals tended to be increased (+4%, P = 0.06) compared with control mice under basal conditions (Fig. 3A). A significant increase under basal conditions was found when combining weights of gastrocnemius, plantaris, tibialis, and extensor digitorum longus (Supplemental Fig. 2). Despite a trend in muscle weights, FCSA under normoxic conditions between control and mGRKO was not significantly different (Fig. 3C; Supplemental Fig. 3A). No alterations in fiber type composition were observed between control and mGRKO mice (Supplemental Fig. 3B). In control animals, the mass of the gastrocnemius muscle decreased under hypoxia, which was in part attributable to reduced food intake (pair-fed, −7.1%; hypoxia, −12.9%). In contrast, in mGRKO mice, muscle mass was preserved under pair-fed conditions. Accordingly, reduced food intake resulted in a lower FCSA in control mice, but not in the mGRKO mice (Fig. 3B–E). Hypoxia-induced muscle atrophy was still observed in mGRKO mice, and muscle loss (hypoxia vs normoxia; control, −12.9% vs mGRKO, −8.5%) was not significantly different from control (Fig. 3A). Hypoxia resulted in a change in fiber size distribution (χ2: P < 0.05, Fig. 3D) and lower average FCSA (Fig. 3C) in control mice, comparable to effects of reduced food intake. In the mGRKO mice, hypoxia also shifted fiber size distribution (χ2: P < 0.05, Fig. 3E) accompanied by an apparent (P = 0.07) decrease in average FCSA (Fig. 3C), despite GR deficiency. Collectively, these data revealed that muscle atrophy attributable to the hypoxia-induced reduction in food intake was GR-dependent, whereas the hypoxia-specific reduction of muscle mass was not. Figure 3. View largeDownload slide Hypoxia-induced muscle atrophy is not prevented by muscle GR deficiency. (A) Gastrocnemius muscle weight after 4 days of hypoxia or pair-fed conditions in control and mGRKO mice as percentage of the muscle mass of normoxic control mice. (B) Representative images of laminin-stained cryosections (magnification: ×200). (C) Average FCSA of glycolytic region in gastrocnemius muscle of control and mGRKO mice. (D) FCSA distribution of the glycolytic region in gastrocnemius muscle of control mice (no significance indicated). (E) FCSA distribution of the glycolytic region in gastrocnemius muscle of mGRKO mice (no significance indicated). Significant differences between groups within control or mGRKO mice are indicated by asterisks on horizontal lines; basal differences in normoxia groups between control and mGRKO mice are indicated by a hash sign: */#P < 0.05, **P ≤ 0.01, ***P ≤ 0.001; trends (P ≤ 0.1) are indicated by the specific P value. Significant effects (P ≤ 0.05) of GR deficiency on the response of the treatment analyzed by two-way ANOVA are indicated as follows: $, pair-fed vs hypoxia (n = 6–8). Figure 3. View largeDownload slide Hypoxia-induced muscle atrophy is not prevented by muscle GR deficiency. (A) Gastrocnemius muscle weight after 4 days of hypoxia or pair-fed conditions in control and mGRKO mice as percentage of the muscle mass of normoxic control mice. (B) Representative images of laminin-stained cryosections (magnification: ×200). (C) Average FCSA of glycolytic region in gastrocnemius muscle of control and mGRKO mice. (D) FCSA distribution of the glycolytic region in gastrocnemius muscle of control mice (no significance indicated). (E) FCSA distribution of the glycolytic region in gastrocnemius muscle of mGRKO mice (no significance indicated). Significant differences between groups within control or mGRKO mice are indicated by asterisks on horizontal lines; basal differences in normoxia groups between control and mGRKO mice are indicated by a hash sign: */#P < 0.05, **P ≤ 0.01, ***P ≤ 0.001; trends (P ≤ 0.1) are indicated by the specific P value. Significant effects (P ≤ 0.05) of GR deficiency on the response of the treatment analyzed by two-way ANOVA are indicated as follows: $, pair-fed vs hypoxia (n = 6–8). GR deficiency differentially suppresses UPS and autophagy lysosomal pathway proteolytic signaling Muscle atrophy may involve elevated protein degradation through the UPS and the autophagy lysosomal pathway (ALP). Expression of E3-ubiquitin ligases Murf1, Atrogin-1, Musa1, and Smart was investigated to assess UPS activation. Basal Murf1, Atrogin-1, Musa1, and Smart expression was similar in control and mGRKO mice (Fig. 4A–4D). In control mice, reduced food intake and hypoxia equally elevated the expression of the investigated ubiquitin ligases. GR deletion diminished the response of Murf1 and Atrogin-1 expression to reduced food intake, whereas under hypoxic conditions the response was hardly diminished. Increased Musa1 expression in response to reduced food intake and hypoxia was absent in the mGRKO mice. In contrast, increased Smart expression by reduced food intake and hypoxia was not affected by GR deletion. Basal expression levels of the ALP-related genes Bnip3 and Map1lc3B were similar in control and mGRKO animals (Fig. 4E and 4F). Reduced food intake and hypoxia increased expression of both ALP-related genes to a similar extent in both controls and mGRKO mice, although the response was blunted in the latter. In both control and mGRKO mice, MAP1LC3B-I was reduced and MAP1LC3B-II levels were unaltered in response to reduced food intake, resulting in an increased II/I ratio (Fig. 4G–4I). Under hypoxic conditions, MAP1LC3B-I and -II were reduced similarly, resulting in an unaltered II/I ratio compared with normoxia in control and GRKO mice. To summarize, whereas reduced food intake under normoxia induced mRNA expression of Murf1 and Atrogin-1 GR-dependently, their increased transcript levels observed after hypoxia were minimally affected by GR deficiency. This differential response to reduced food intake under normoxic vs hypoxic conditions mediated by muscle GR expression was not observed for the other E3-ubiquitin ligases Musa1 and Smart. Furthermore, GR deficiency similarly blunted ALP-related gene expression in response to hypoxia or reduced food intake, but did not affect changes in MAP1LC3B-I and -II protein turnover. Figure 4. View largeDownload slide GR deficiency differentially suppresses UPS and ALP proteolytic signaling. (A) Murf1 expression. (B) Atrogin-1 expression. (C) Musa1 expression. (D) Smart expression (E) Bnip3 expression. (F) Map1lc3B expression. (G) MAP1LC3B-I protein. (H) MAP1LC3B-II protein. (I) MAP1LC3B-II/I ratio. All mRNA concentrations were corrected by 18S concentration. Significant differences between groups within control or mGRKO mice are indicated by asterisks on horizontal lines; basal differences in normoxia groups between control and mGRKO mice are indicated by a hash sign: *P < 0.05, **/##P ≤ 0.01, ***P ≤ 0.001; trends (P ≤ 0.1) are indicated by the specific P value. Significant effects (P ≤ 0.05) of GR deficiency on the response of the treatment analyzed by two-way ANOVA are indicated as follows: €, pair-fed vs normoxia; &, normoxia vs hypoxia; $, pair-fed vs hypoxia (n = 6–8). Figure 4. View largeDownload slide GR deficiency differentially suppresses UPS and ALP proteolytic signaling. (A) Murf1 expression. (B) Atrogin-1 expression. (C) Musa1 expression. (D) Smart expression (E) Bnip3 expression. (F) Map1lc3B expression. (G) MAP1LC3B-I protein. (H) MAP1LC3B-II protein. (I) MAP1LC3B-II/I ratio. All mRNA concentrations were corrected by 18S concentration. Significant differences between groups within control or mGRKO mice are indicated by asterisks on horizontal lines; basal differences in normoxia groups between control and mGRKO mice are indicated by a hash sign: *P < 0.05, **/##P ≤ 0.01, ***P ≤ 0.001; trends (P ≤ 0.1) are indicated by the specific P value. Significant effects (P ≤ 0.05) of GR deficiency on the response of the treatment analyzed by two-way ANOVA are indicated as follows: €, pair-fed vs normoxia; &, normoxia vs hypoxia; $, pair-fed vs hypoxia (n = 6–8). Hypoxia-induced impairment of protein synthesis signaling is GR-dependent The mTOR complex 1 (mTORC1) pathway is central in the regulation of protein synthesis signaling (37, 38). Ribosomal protein S6 kinase, 70 kDa, polypeptide 1 (P70S6K1) is directly phosphorylated by mTOR. P70S6K1 phosphorylates ribosomal protein S6 (RPS6), thereby controlling protein synthesis. P70S6K1 and RPS6 concentrations were unaffected by both experimental conditions and mouse model. As expected, reduced food intake lowered the P70S6K1 and RPS6 phosphorylation ratio, suggesting suppressed protein synthesis (Fig. 5A–5D). In contrast, the food-dependent reduction of the P70S6K1 and RPS6 phosphorylation ratios was impaired under hypoxic conditions. GR deficiency reduced baseline P70S6K1 and RPS6 phosphorylation ratios of normoxic controls. P70S6K1 and RPS6 phosphorylation ratios were further lowered by reduced food intake in mGRKO muscle, and, importantly, in contrast to control mice, P70S6K1 and RPS6 phosphorylation ratios were similarly suppressed by hypoxia. Alterations in the eukaryotic translation initiation factor 4E-binding protein 1 (4E-BP1) Ser65 phosphorylation ratio (Fig. 5E, 5H) mirrored the changes observed for P70S6K1 and RPS6 phosphorylation. 4E-BP1 threonine Thr37 and Thr46 phosphorylation ratios marginally differed between conditions and mouse models (Fig. 5F and 5H). 4E-BP1 protein concentrations were equally increased by hypoxia and reduced food intake in control but not in GR-deficient mice (Fig. 5G and 5H). Collectively, these data indicate that inhibition of mTORC1 by reduced food intake under normoxia does not require muscle GR signaling. In contrast, retained mTORC1 activity under hypoxic conditions, despite reduced food intake, is GR-dependent. Figure 5. View largeDownload slide Hypoxia-associated dysregulation of protein synthesis regulation is GR-dependent. (A) Phosphorylation ratio (phosphorylated/total protein) of P70S6K1 at Thr389. (B) Phosphorylation ratio (phosphorylated/total protein) of RPS6 at Ser235/236. (C) Representative images of P70S6K Western blots. (D) Representative images of S6 Western blots. (E) Phosphorylation ratio (phosphorylated/total protein) of 4E-BP1 at Ser65. (F) Phosphorylation ratio (phosphorylated/total protein) of 4E-BP1 at Thr37/46. (G) Total 4E-BP1. (H) Representative images of 4EBP1 Western blots. Significant differences between groups within control or mGRKO mice are indicated by asterisks on horizontal lines; basal differences in normoxia groups between control and mGRKO mice are indicated by a hash sign: */#P < 0.05, **/##P ≤ 0.01, ***P ≤ 0.001; trends (P ≤ 0.1) are indicated by the specific P value. Significant effects (P ≤ 0.05) of GR deficiency on the response of the treatment analyzed by two-way ANOVA are indicated as follows: €, pair-fed vs normoxia; &, normoxia vs hypoxia; $, pair-fed vs hypoxia (n = 6–8). Figure 5. View largeDownload slide Hypoxia-associated dysregulation of protein synthesis regulation is GR-dependent. (A) Phosphorylation ratio (phosphorylated/total protein) of P70S6K1 at Thr389. (B) Phosphorylation ratio (phosphorylated/total protein) of RPS6 at Ser235/236. (C) Representative images of P70S6K Western blots. (D) Representative images of S6 Western blots. (E) Phosphorylation ratio (phosphorylated/total protein) of 4E-BP1 at Ser65. (F) Phosphorylation ratio (phosphorylated/total protein) of 4E-BP1 at Thr37/46. (G) Total 4E-BP1. (H) Representative images of 4EBP1 Western blots. Significant differences between groups within control or mGRKO mice are indicated by asterisks on horizontal lines; basal differences in normoxia groups between control and mGRKO mice are indicated by a hash sign: */#P < 0.05, **/##P ≤ 0.01, ***P ≤ 0.001; trends (P ≤ 0.1) are indicated by the specific P value. Significant effects (P ≤ 0.05) of GR deficiency on the response of the treatment analyzed by two-way ANOVA are indicated as follows: €, pair-fed vs normoxia; &, normoxia vs hypoxia; $, pair-fed vs hypoxia (n = 6–8). Perturbations in signaling upstream of mTOR correspond to hypoxia-induced decreases in food intake and are independent of GR mTOR activity is controlled through direct phosphorylation of Ser2448 by AKT, and AKT-mediated inhibition of TSC2 through phosphorylation of sites Ser939 and Thr1462 in TSC2. The basal AKT S473 phosphorylation ratio in mGRKO mice was slightly reduced compared with control (P = 0.06) (Fig. 6A and 6G). In control mice, hypoxia and reduced food intake equally lowered the AKT phosphorylation ratio, whereas in mGRKO mice no further reduction was seen. GR deficiency significantly lowered the mTOR S2448 phosphorylation ratio (Fig. 6B and 6H). Reductions in mTOR S2448 phosphorylation ratio in response to hypoxia and decreased food intake were similar and did not differ between control and mGRKO mice. mTOR Ser2481 autophosphorylation is postulated to reflect mTOR activity (39). Except for a decrease in the phosphorylation ratio in response to hypoxia in mGRKO mice, no changes were observed (Fig. 6C and 6H). TSC2 phosphorylation (S939 and T1462) ratios were not affected by any condition (Fig. 6D, 6E, and 6I). Total TSC2 was lowered by reduced food intake in control mice alone (Fig. 6F and 6I). AKT-independent activation of TSC2 by phosphorylation on Ser1387 was unaffected (Supplemental Fig. 4). Collectively, these data indicate that TSC2 signaling is not involved in suppression of mTOR by reduced food intake, whereas the reduction of the AKT/mTOR signaling by reduced food intake is intact under hypoxic conditions and is independent of GR signaling. Figure 6. View largeDownload slide Perturbations in signaling upstream of mTOR correspond to hypoxia-induced decreases in food intake and are GR-independent. (A) Phosphorylation ratio (phosphorylated/total protein) of AKT at Ser473. (B) Phosphorylation ratio (phosphorylated/total protein) of mTOR at Ser2448. (C) Phosphorylation ratio (phosphorylated/total protein) of mTOR at Ser2481. (D) Phosphorylation ratio (phosphorylated/total protein) of TSC2 at Ser939. (E) Phosphorylation ratio (phosphorylated/total protein) of TSC2 at Ser1462. (F) Protein levels of TSC2. (G) Representative images of AKT Western blots. (H) Representative images of mTOR Western blots. (G) Representative images of TSC2 Western blots. Significant differences between groups within control or mGRKO mice are indicated by asterisks on horizontal lines; basal differences in normoxia groups between control and mGRKO mice are indicated by a hash sign: */#P < 0.05, **/##P ≤ 0.01, ***P ≤ 0.001; trends (P ≤ 0.1) are indicated by the specific P value. Significant effects (P ≤ 0.05) of GR deficiency on the response of the treatment analyzed by two-way ANOVA are indicated as follows: €, pair-fed vs normoxia (n = 6–8). Figure 6. View largeDownload slide Perturbations in signaling upstream of mTOR correspond to hypoxia-induced decreases in food intake and are GR-independent. (A) Phosphorylation ratio (phosphorylated/total protein) of AKT at Ser473. (B) Phosphorylation ratio (phosphorylated/total protein) of mTOR at Ser2448. (C) Phosphorylation ratio (phosphorylated/total protein) of mTOR at Ser2481. (D) Phosphorylation ratio (phosphorylated/total protein) of TSC2 at Ser939. (E) Phosphorylation ratio (phosphorylated/total protein) of TSC2 at Ser1462. (F) Protein levels of TSC2. (G) Representative images of AKT Western blots. (H) Representative images of mTOR Western blots. (G) Representative images of TSC2 Western blots. Significant differences between groups within control or mGRKO mice are indicated by asterisks on horizontal lines; basal differences in normoxia groups between control and mGRKO mice are indicated by a hash sign: */#P < 0.05, **/##P ≤ 0.01, ***P ≤ 0.001; trends (P ≤ 0.1) are indicated by the specific P value. Significant effects (P ≤ 0.05) of GR deficiency on the response of the treatment analyzed by two-way ANOVA are indicated as follows: €, pair-fed vs normoxia (n = 6–8). Discussion In this study, the contribution of muscle GR signaling to hypoxia-induced muscle atrophy was addressed. As food intake is reduced in mice exposed to hypoxia (26), a pair-fed group was included to assess the GR dependency of responses induced by hypoxia and by semistarvation under normoxic conditions. Muscle GR signaling was responsible for muscle atrophy in the pair-fed group but not in the hypoxic group. The increased expression of UPS-related genes was mainly independent of GR in muscle of hypoxic mice. Suppression of mTORC1 activity under semistarvation conditions was impaired by hypoxia in a GR-dependent manner and independent of AKT signaling. Fasting-induced muscle atrophy is associated with elevated corticosterone concentrations (40). In line with this, hypoxia-induced reduction of food intake increases corticosterone concentrations. Mice subjected to hypoxia in our model are hypoxemic (26), and hypoxemia has also been associated with elevated glucocorticoid concentrations and GR signaling independent of malnutrition (41–45). However, as no further increase in glucocorticoid concentration is observed in hypoxic compared with normoxic pair-fed mice, this suggests that the elevated corticosterone concentration is attributable to the reduced food intake alone. Muscle GR signaling as demonstrated by GRE reporter activation indicates that circulating corticosterone affects skeletal muscle. Interestingly, GRE activation induced by restricted feeding and hypoxia does not reflect plasma corticosterone levels. This may be a consequence of differential prereceptor modulation of glucocorticoid availability in skeletal muscle by 11β-HSD1, as its expression and activity are critical regulatory steps in glucocorticoid responses (46). Considering their postulated transcriptional regulation by GR (18, 47), the increases in Glul and Foxo1 likely reflect genomic actions of activated GR in response to raised corticosterone concentrations. Interestingly, in response to hypoxia, muscle Gr mRNA transcript levels increase. However, as increased Glul and Foxo-1 expression and GR reporter activity do not display consistent additive effects of hypoxia compared with reduced food intake, the genomic actions of GR in hypoxic muscle are probably attributable to the reduced food intake. Although increased by semistarvation, mRNA expression of the putative GR target genes Klf15 and Redd1 (18, 47) is minimally affected by GR deficiency. Furthermore, under hypoxic conditions, increases in Klf15 and Redd1 expression are attenuated (Klf15) or absent (Redd1), indicating that hypoxia blocks transcriptional activation of these genes by semistarvation. Testosterone concentrations were not measured in this study, but it has been shown previously that induction of Redd1 expression by glucocorticoids is abolished in the presence of an elevated testosterone concentration (48). This may explain the lack of Redd1 transcript accumulation in muscle of hypoxic mice, as hypoxia is known to increase serum testosterone concentrations within 1 day (49). Protein degradation in skeletal muscle involves the UPS (50) and the ALP (51). MAP1LC3B-I protein levels are reduced despite increased Map1lc3b mRNA concentrations, suggestive of increased conversion to MAP1LC3B-II. Although MAP1LC3B-II protein levels are reduced, this could be the result of its elevated degradation due to an increased autophagic flux (52). The expression of the ALP-related genes Map1lc3b and Bnip3 is mainly upregulated by the hypoxia-mediated reduction in food intake. GR deficiency blunts this response slightly, but significantly. Interestingly, this corresponds with the GR-dependent induction of Foxo-1, which has been implicated in the transcriptional regulation of ALP-related genes (51), and suggests that genomic actions of GR may indirectly contribute to control of autophagy in muscle (12, 18, 53, 54). FOXO1 is a transcriptional regulator of Murf1 and Atrogin-1 (14), and its expression corresponds to the lowered transcript levels of these atrogenes under conditions of reduced food intake. However, in response to hypoxia our data imply a GR-independent mechanism that does not require increased Foxo1 expression in the regulation of these “classical” atrogenes. KLF15 is a key regulator of Murf1 and Atrogin-1 expression and is reported to be a direct target gene of GR (18). Its GR-independent regulation in response to reduced food intake, as well as its decreased expression in response to hypoxia, however, excludes a key regulatory function of de novo synthesized KLF15 on Murf1 and Atrogin-1 expression under hypoxic conditions. As KLF15 and FOXO1 are not responsible for the increased expression of Murf1 and Atrogin-1 during hypoxia, the transcriptional regulators in play remain to be identified. A novel finding is that increased Musa1 expression requires GR signaling in response to reduced food intake and hypoxia, whereas the newly identified E3 ligase Smart (17) is insensitive to GR deletion (22, 55, 56). Mothers against decapentaplegic homolog 3 (SMAD3) is a transcription factor involved in the transforming growth factor-β signaling (57). Increased Smad3 expression and transcriptional activity has been associated with increased Atrogin-1 expression (58). Smad3 expression is elevated by hypoxia independently of reduced food intake or GR signaling (Supplemental Fig. 5). Interestingly, of all protein turnover signaling constituents measured, only Atrogin-1 mRNA levels were increased in hypoxic compared with pair-fed GRKO muscle. This may point at involvement of transforming growth factor-β signaling in GR-independent hypoxia-induced muscle atrophy. Collectively, our data reveal GR-dependent and -independent constituents of proteolysis regulation. GR deficiency attenuates the overall expression of ALP-associated genes. Furthermore, Musa1 is identified as a GR-regulated E3-ligase, and genomic actions of GR are involved in upregulation of Murf1 and Atrogin-1 expression when food intake is reduced, whereas Atrogin-1 and Murf1 induction under hypoxic conditions occurs independently of GR. Hypoxia reduces food intake (26), and fasting results in the suppression of muscle protein synthesis (59, 60). mTORC1 activity (based on 4E-BP1, P70S6K1, and RPS6 phosphorylation ratio) is decreased accordingly in muscle of normoxic control mice in response to reduced food intake. Interestingly, in mGRKO muscle mTORC1 activity is reduced to a similar extent as in pair-fed control animals in response to reduced food intake, indicating that this suppression of mTORC1 activity occurs independently of muscle GR signaling. In addition to increases in endogenous glucocorticoids, nutritional restriction is accompanied by decreased circulating insulin and insulin-like growth factor-1 levels (61). Although plasma insulin was not measured, the low AKT phosphorylation in muscle of pair-fed control mice suggests that reduced anabolic signaling cues, rather than the increased circulating corticosterone concentration, are responsible for the suppression of mTORC1 activity caused by reduced food intake. A remarkable finding concerns the sustained mTORC1 activity in hypoxia despite reduced food intake. Although this confirmed our previous observations (26), we set out to further verify impairment of mTORC1 regulation by feeding cues under hypoxic conditions. Subjecting wild-type mice to 24 hours of complete food deprivation under hypoxic and normoxic conditions revealed that fasting-induced inhibition of RPS6 and 4E-BP1 phosphorylation is only observed in skeletal muscle of normoxic but not hypoxic mice (Supplemental Fig. 6). This illustrates that inhibitory cues toward mTORC1 activity by partial or complete food deprivation are not appropriately sensed in hypoxic conditions. Alternatively, it may reflect a potential (futile) protective response to counter food restriction–induced suppression of protein synthesis. We now show that GR is involved in this dysregulation, because under hypoxic conditions in mGRKO mice mTORC1 activity is suppressed to levels comparable to pair-fed mice. REDD1 inhibits mTORC1 activity through liberating TSC2 by competing for its binding to 14-3-3 (62). Redd1 expression has been postulated to be GR-dependent (63). However, the lack of Redd1 induction in response to hypoxia in both control as well as in mGRKO muscle strongly suggests that Redd1 expression does not control mTORC1 activity in response to hypoxia. KLF15 has also been postulated to operate GR dependently and to inhibit mTORC1 activity through control of BCAT2 expression (18). In contrast to in vitro studies in which dexamethasone was used to induce GR/KLF15 signaling (18), Klf15 expression under hypoxia and reduced food intake in vivo was not GR-dependent and did not correlate inversely with mTORC1 activity. Collectively, this indicates that aberrant regulation of Redd1 and Klf15 by hypoxia is not responsible for the sustained mTORC1 activity in conditions of semistarvation. Nongenomic actions of GR have also been implicated in mTORC1 signaling by inhibiting AKT phosphorylation through an interaction with PI3K (64, 65), resulting in reduced Akt-mediated phosphorylation of downstream TSC2 (66) and mTOR at S2448 (37). However, TSC2 phosphorylation is not affected, whereas suppressed AKT and S2448 mTOR phosphorylation by either reduced food intake alone or hypoxia is not reversed in GRKO muscle, although basal AKT phosphorylation is already lowered by GR deficiency. Therefore, nongenomic actions of GR on AKT signaling are not involved in the lack of mTORC1 inhibition under hypoxic conditions. Autophosphorylation of mTOR at site S2481 has been described and contributes to mTORC1 activity, but its regulation is only partly understood (37). Interestingly, only in mGRKO muscle a reduction of mTOR S2481 phosphorylation under hypoxia was observed, which corresponds to the restoration of an appropriate response to semistarvation even in the presence of hypoxia. Although the exact mechanism remains to be elucidated, this suggests that GR-dependent preservation of mTOR S2481 phosphorylation may contribute to impaired mTORC1 regulation, that is, retained mTORC1 activity during hypoxia. Overall, the GR-dependent impairment of mTORC1 activity regulation in response to hypoxia is neither explained by aberrant upstream TSC2 or AKT/mTOR-S2448 signaling, nor by alterations in Redd1 or Klf15 expression. GR-mediated control of mTOR S2481 phosphorylation therefore appears to contribute to the regulation of protein synthesis independently of the established nongenomic actions of GR. Protein synthesis and degradation signaling change in a coordinated fashion in conditions altering muscle mass (67–69). In line, the decreased protein synthesis signaling in response to semistarvation is accompanied by an increased expression of genes mediating proteolysis in atrophying muscle. Still, increased proteolysis signaling rather than reduced synthesis signaling likely provides the main GR-dependent contribution to muscle atrophy in response to normoxic food restriction. Conversely, most of these attenuating effects of GR deficiency on proteolysis signaling are absent under hypoxia, and muscle atrophy is not prevented in GRKO mice. This implies differential (i.e., GR-independent) regulation of proteolysis under hypoxia compared with normoxic food restriction. Importantly, the coordination of protein synthesis and proteolysis signaling cues appears impaired under hypoxic conditions in a GR-dependent manner. However, considering that muscle atrophy under hypoxia is equal in wild-type and GRKO mice despite suppressed mTOR signaling in GRKO muscle, the physiological relevance of sustained stimulatory signals for protein synthesis signaling under hypoxia remain to be clarified, as our data reveal it does not function as an adaptive response to protect from further aggravation of muscle atrophy induced by reduced food intake. In COPD patients with muscle atrophy, molecular signatures reflecting parallel increases in protein synthesis and degradation signaling in skeletal muscle have been reported (70–73) and are associated with a hypermetabolic state. This impaired coordination of protein synthetic and proteolytic cues may contribute to the loss of muscle mass observed in pathological conditions. In conclusion, GR signaling is required for muscle atrophy and increased expression of proteolysis-associated genes induced by decreased food intake under normoxic conditions. Under hypoxic conditions, muscle atrophy and elevated gene expression of the UPS-associated E3 ligases Murf1 and Atrogin-1 are mostly independent of GR signaling. Importantly, impaired inhibition of mTORC1 activity is GR-dependent in hypoxia-induced muscle atrophy. Abbreviations: 4E-BP1 4E-binding protein 1 ALP autophagy lysosomal pathway ANOVA analysis of variance COPD chronic obstructive pulmonary disease FCSA fiber cross-sectional area FOXO1 forkhead box protein O1 GLUL glutamate-ammonia ligase GR glucocorticoid receptor GRE glucocorticoid receptor–responsive element KLF15 Krüppel-like factor 15 mGRKO muscle‐specific glucocorticoid receptor knockout mRNA messenger RNA MSTN myostatin mTOR mammalian target of rapamycin mTORC1 mammalian target of rapamycin complex 1 PI3K phosphatidylinositol-4,5-bisphosphate 3-kinase P70S6K1 protein S6 kinase, 70 kDa, polypeptide 1 qPCR quantitative polymerase chain reaction RPS6 ribosomal protein S6 SMAD3 mothers against decapentaplegic homolog 3 UPS ubiquitin proteasomal system. Acknowledgments The GRE luciferase construct was provided by R. Hoffmann (University Medical Center Groningen, Groningen, Netherlands). 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Google Scholar CrossRef Search ADS PubMed  Copyright © 2018 Endocrine Society http://www.deepdyve.com/assets/images/DeepDyve-Logo-lg.png Endocrinology Oxford University Press

Glucocorticoid Receptor Signaling Impairs Protein Turnover Regulation in Hypoxia-Induced Muscle Atrophy in Male Mice

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Oxford University Press
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Copyright © 2018 Endocrine Society
ISSN
0013-7227
eISSN
1945-7170
D.O.I.
10.1210/en.2017-00603
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Abstract

Abstract Hypoxemia may contribute to muscle wasting in conditions such as chronic obstructive pulmonary disease. Muscle wasting develops when muscle proteolysis exceeds protein synthesis. Hypoxia induces skeletal muscle atrophy in mice, which can in part be attributed to reduced food intake. We hypothesized that hypoxia elevates circulating corticosterone concentrations by reduced food intake and enhances glucocorticoid receptor (GR) signaling in muscle, which causes elevated protein degradation signaling and dysregulates protein synthesis signaling during hypoxia-induced muscle atrophy. Muscle-specific GR knockout and control mice were subjected to normoxia, normobaric hypoxia (8% oxygen), or pair-feeding to the hypoxia group for 4 days. Plasma corticosterone and muscle GR signaling increased after hypoxia and pair-feeding. GR deficiency prevented muscle atrophy by pair-feeding but not by hypoxia. GR deficiency differentially affected activation of ubiquitin 26S-proteasome and autophagy proteolytic systems by pair-feeding and hypoxia. Reduced food intake suppressed mammalian target of rapamycin complex 1 (mTORC1) activity under normoxic but not hypoxic conditions, and this retained mTORC1 activity was mediated by GR. We conclude that GR signaling is required for muscle atrophy and increased expression of proteolysis-associated genes induced by decreased food intake under normoxic conditions. Under hypoxic conditions, muscle atrophy and elevated gene expression of the ubiquitin proteasomal system–associated E3 ligases Murf1 and Atrogin-1 are mostly independent of GR signaling. Furthermore, impaired inhibition of mTORC1 activity is GR-dependent in hypoxia-induced muscle atrophy. Chronic obstructive pulmonary disease (COPD) is globally a leading cause of morbidity and disability (1). Weight loss and muscle atrophy are common features of advanced COPD. Muscle atrophy significantly increases disease burden and is a strong predictor of mortality (1–5). In addition to nutritional imbalance, glucocorticoid use, systemic inflammation, and progressive inactivity, hypoxemia has been implicated as a trigger for muscle atrophy in COPD patients (6). Previously, it was shown that hypoxia causes muscle atrophy in rat models (7, 8). Our group confirmed these findings in a mouse model of normobaric hypoxia (9) and revealed that muscle atrophy was attributable to a hypoxia-induced reduction of food intake and hypoxia-specific effects. The mechanisms of fasting-induced muscle atrophy involve elevated glucocorticoid concentrations and glucocorticoid receptor (GR) signaling (10), which affect protein synthesis and proteolysis in skeletal muscle. Adrenalectomized rats display reduced muscle protein breakdown, which is restored by glucocorticoid administration (11). Moreover, in muscle-specific (MLC-cre) GR knockout mice, fasting-induced reductions in fiber cross-sectional area (FCSA) were absent and the activation of the proteolytic machinery was attenuated (12). GR affects muscle protein turnover by genomic and nongenomic actions (13). Genomic effects include activation of protein degradation by the ubiquitin proteasomal system (UPS). Forkhead box protein O1 (FOXO1) is upregulated in a GR-specific manner and contributes to the subsequent induction of expression of atrogenes, such as tripartite motif containing 63 (Trim63/Murf1), F-box only protein 32 (Fbxo32/Mafbx/Atrogin-1), muscle ubiquitin ligase of SCF complex in atrophy-1 (Musa1, Fbxo30), and specific of muscle atrophy and regulated by transcription (Smart/Fbxo21) (14–17). GR is also involved in upregulation of Klf15 messenger RNA (mRNA) expression. In addition to its involvement in the control of proteolysis via regulation of MuRF1 and Atrogin-1, Krüppel-like factor 15 (KLF15) also contributes to the regulation of protein synthesis (18). Glutamate-ammonia ligase (GLUL) is an enzyme involved in muscle amino acid metabolism during fasting (19), the expression of which is also stimulated by glucocorticoids through GR-responsive elements (GRE) in the Glul promoter (20). Myostatin (MSTN) is associated with dexamethasone- (21) and hypoxia-induced muscle atrophy (8) and expression is increased by activation of the GR (21, 22). Furthermore, GR-mediated transcription of DNA damage–inducible transcript 4 protein (Ddit4/Redd1) followed by inhibition of mammalian target of rapamycin (mTOR) results in suppressed protein synthesis signaling (23, 24) and activation of autophagy (25). Nongenomic GR actions involve suppression of protein synthesis through the inhibition of the phosphatidylinositol-4,5-bisphosphate 3-kinase (PI3K)/protein kinase B (PKB/AKT)/TSC1/2/mTOR signaling pathway, in which GR binds to PI3K subunits p110 and p85 and prevents association with insulin receptor substrate-1 (IRS-1) (12). As we previously reported (26), hypoxia-exposed mice show altered activation of signaling pathways of proteolysis and protein synthesis in skeletal muscle. Furthermore, it has been shown that hypoxia can elevate blood glucocorticoid concentrations (27–30). We therefore hypothesized that hypoxia elevates circulating corticosterone concentrations resulting in enhanced GR signaling in muscle, which causes elevated protein degradation signaling and also dysregulates protein synthesis signaling during hypoxia-induced muscle atrophy. Methods Animals and tissue collection All mouse studies were carried out according to a protocol approved by the Institutional Animal Care Committee of Maastricht University. Electroporation of the muscles was done in 12-week-old C57BL/6J male mice (Charles River Laboratories). Twelve-week-old male, muscle‐specific GR knockout (mGRKO) mice were generated by crossing transgenic Mlc1f‐Cre+/‐ mice (Cre recombinase under control of the fast myosin light chain 1 promoter) (31) with mice bearing floxed Gr alleles (Grfl/fl) (32) on a C57BL/6 background, resulting in the ablation of GR in MyHC type II (fast) muscle cells, which accounts for 95% of the muscle fibers (33). Twenty-four control (littermates) and 24 mGRKO mice were randomly divided into three groups: normoxia, normoxia pair-fed to hypoxic animals, and normobaric hypoxia. The pair-fed group was included to assess the effects of reduced food intake induced by hypoxia. All mice were housed in sets of four animals per cage in experimental chambers at 21°C with a 12-hour dark/light cycle. Mice received standard chow (V1534-000 Ssniff R/M-H; Ssniff Spezialdiäten) and water ad libitum. After 5 days of acclimatization, normoxic and pair-fed groups were kept at ambient air (21% O2) and the hypoxic group to normobaric hypoxia. Using the proOX system P110 (BioSpherix), oxygen levels were reduced in a stepwise manner to 12% (day 0), 10% (day 1), and finally 8% (61 mm Hg) on day 2, which was then maintained for the remainder of the experiment. To assess the effects of reduced food intake during hypoxia, pair-fed animals received the amount of food consumed by the hypoxic mice daily at a standardized time (5:00 pm). Tissue and plasma were isolated on day 4 and frozen immediately in liquid nitrogen for further analysis. Timing of the tissue collection was standardized in the early dark phase (9:00 to 10:00 am) to reduce hormonal variation. Tissue weights were corrected for body weight at the start of the experiment. Corticosterone measurement Plasma corticosterone concentrations were determined by high-performance liquid chromatography. A carboxylic acid derived from cortisol (4-androsten-11β,17α-diol-3-one-17β-carboxylic acid) was synthetized and purified as described previously (34) and was added as internal control to plasma and corticosterone standards. Samples and standards were acidified with phosphoric acid, and steroid hormones were subsequently extracted from plasma with diethyl ether and dried with nitrogen gas. The residues were dissolved in acetone and incubated with sulfuric acid for 20 minutes to convert the steroids into fluorescent products (35). After the addition of 100 volumes of diethylether and 5 volumes of water, the fluorescent products were extracted from the aqueous sulfuric acid phase. After evaporating the organic phase under a stream of nitrogen, the residues were dissolved and analyzed by high-performance liquid chromatography as described (34). Ratios of corticosterone and internal standard peak areas were calculated and converted into concentrations using calibration curves. In vivo electroporation of gastrocnemius muscle To assess the activity of the GR in response to hypoxia and reduced food intake, a reporter plasmid was electroporated into the gastrocnemius muscle. Plasmids were prepared using an endotoxin-free maxi prep kit (Qiagen) and dissolved in 0.9% saline. The plasmid mix consisted of 1:1 GRE-luciferase construct (36) and pVAX-lacZ plasmid (Invitrogen) encoding β-galactosidase. Mice were anesthetized with isoflurane and injected subcutaneously with buprenorphine (Temgesic) as analgesic. Skin covering the gastrocnemius muscle was shaved and disinfected. Using a bent 0.3-mL syringe with a 30-gauge needle (Becton Dickinson), five times 10 µL of plasmid (50 µg total) was injected at different sites into the gastrocnemius muscle. Custom-built mouse electrodes with electrolyte crème (Parker Laboratories) were placed on both sides of the gastrocnemius muscle on top of the skin. Eight pulses of 150 V/cm, with 20-ms pulse length and a 1-second interval were applied using the BTX square wave electroporator (BTX/Harvard Apparatus). The direction of the pulse was inverted after four pulses to increase efficiency. After 21 days of recovery, mice were exposed to experimental conditions. Muscles were snap-frozen in liquid nitrogen and ground to a powder using an N2-cooled steel mortar. Muscle powder was lysed in passive lysis buffer (Promega) and assessed for luciferase (Promega) and β‐galactosidase (Tropix) activity. Real-time quantitative polymerase chain reaction Total RNA was isolated using TRI Reagent (Sigma-Aldrich) and further purified by precipitation with 2 M LiCl. Complementary DNA synthesis was performed with random hexamer primers on denatured RNA using the transcriptor first-strand complementary DNA synthesis kit (Roche). Real-time quantitative polymerase chain reaction (qPCR) was performed in the iQ5 thermal cycler (Bio-Rad) using the qPCR SYBR Green fluorescein mix (Abgene). qPCR primers were designed using Primer Express 2.0 software (Applied Biosystems) and ordered from Sigma Genosys (Table 1). All primers were intron spanning to avoid contamination of the amplification products with genomic DNA. Expression of genes was normalized to 18S ribosomal RNA. mRNA expression data in the figures are shown as fold change of the wild-type normoxic group. Table 1. Genes Used in This Study Gene  NCBI Access No.  Forward Primer (5′ to 3′)  Reverse Primer (5′ to 3′)  18S  NR_003278.1  AGTTAGCATGCCAGAGTCTCG  TGCATGGCCGTTCTTAGTTG  Gr  NM_008173.3  CGCCAAGTGATTGCCGC  TGTAGAAGGGTCATTTGGTCATCCA  Glul  NM_008131.3  GGCCATGCGGGAGGAGA  GGTGCCTCTTGCTCAGTTTGTCA  Foxo1  NM_019739.3  AAGAGCGTGCCCTACTTCAAG  CCATGGACGCAGCTCTTCTC  Klf15  NM_023184.3  TGCAGCAAGATGTACACCAAGAG  ATCGCCGGTGCCTTGAC  Redd1  NM_029083.2  CGGGCCGGAGGAAGACT  CTGCATCAGGTTGGCACACA  Murf1  NM_001039048.2  TGTCTGGAGGTCGTTTCCG  CTCGTCTTCGTGTTCCTTGC  Atrogin-1  NM_026346.2  ACCGGCTACTGTGGAAGAGA  CCTTCCAGGAGAGAATGTGG  Map1lc3B  NM_026160.4  GAGCAGCACCCCACCAAGAT  CGTGGTCAGGCACCAGGAA  Bnip3  NM_009760.4  CCATGTCGCAGAGCGGG  GACGGAGGCTGGAACGC  Musa1  NM_027968.3  GGACGTTTGTGGCAGTTTACTTC  GCAGTACTGAATCGCCATACCTTC  Smart  NM_145564.3  CACAGGGATGTCTGCTACTC  ACACAGTTGTAGCCGTACCTC  Mstn  NM_010834.2  GGCCATGATCTTGCTGTAACCT  CGGCAGCACCGGGATT  Smad3  NM_016769  CCCGAGAACACTAACTTCCCTG  AACCTGCGTCCATGCTGTG  Gene  NCBI Access No.  Forward Primer (5′ to 3′)  Reverse Primer (5′ to 3′)  18S  NR_003278.1  AGTTAGCATGCCAGAGTCTCG  TGCATGGCCGTTCTTAGTTG  Gr  NM_008173.3  CGCCAAGTGATTGCCGC  TGTAGAAGGGTCATTTGGTCATCCA  Glul  NM_008131.3  GGCCATGCGGGAGGAGA  GGTGCCTCTTGCTCAGTTTGTCA  Foxo1  NM_019739.3  AAGAGCGTGCCCTACTTCAAG  CCATGGACGCAGCTCTTCTC  Klf15  NM_023184.3  TGCAGCAAGATGTACACCAAGAG  ATCGCCGGTGCCTTGAC  Redd1  NM_029083.2  CGGGCCGGAGGAAGACT  CTGCATCAGGTTGGCACACA  Murf1  NM_001039048.2  TGTCTGGAGGTCGTTTCCG  CTCGTCTTCGTGTTCCTTGC  Atrogin-1  NM_026346.2  ACCGGCTACTGTGGAAGAGA  CCTTCCAGGAGAGAATGTGG  Map1lc3B  NM_026160.4  GAGCAGCACCCCACCAAGAT  CGTGGTCAGGCACCAGGAA  Bnip3  NM_009760.4  CCATGTCGCAGAGCGGG  GACGGAGGCTGGAACGC  Musa1  NM_027968.3  GGACGTTTGTGGCAGTTTACTTC  GCAGTACTGAATCGCCATACCTTC  Smart  NM_145564.3  CACAGGGATGTCTGCTACTC  ACACAGTTGTAGCCGTACCTC  Mstn  NM_010834.2  GGCCATGATCTTGCTGTAACCT  CGGCAGCACCGGGATT  Smad3  NM_016769  CCCGAGAACACTAACTTCCCTG  AACCTGCGTCCATGCTGTG  Abbreviation: NCBI, National Center for Biotechnology Information. View Large Western blotting Western blotting was performed according to procedures previously described (26). In short, gastrocnemius muscle was ground to a powder and lysed. Total protein concentration was determined and 12.5 μg per lane was used for Western blotting. The membrane was incubated overnight at 4°C with primary antibodies (Table 2). Blots were probed with a horseradish peroxidase–conjugated secondary antibody and visualized by chemiluminescence in an LAS-3000 luminescent image analyzer (Fujifilm). Bands were quantified using AIDA software (Fujifilm). Figures display ratios of phosphorylated over total protein of interest; in case total levels of the protein of interest are altered between groups, total levels are also shown separately. PonceauS staining was used to control and correct for protein loading. Protein data in the figures are shown as fold change of the wildtype normoxic group. Protein ratios are calculated by dividing the phosphorylated protein by the total amount of protein. Table 2. Antibodies Used in This Study Target  Product No.  Manufacturer  Made in  Size (kDa)  RRID  GR  12041  Cell Signaling Technology  Rabbit  91–94  AB_2631286  FOXO1  2880  Cell Signaling Technology  Rabbit  78–82  AB_2106495  GLUL  G45020  BD Transduction Laboratory  Mouse  40  AB_2313767  S6  2217  Cell Signaling Technology  Rabbit  32  AB_331355  p-S6 (S235/236)  4856  Cell Signaling Technology  Rabbit  32  AB_2181037  4E-BP1  9452  Cell Signaling Technology  Rabbit  15–20  AB_331692  p-4E-BP1 (S65)  9451  Cell Signaling Technology  Rabbit  15–20  AB_330947  p-4E-BP1 (T37/46)  9459  Cell Signaling Technology  Rabbit  15–20  AB_330985  mTOR  2983  Cell Signaling Technology  Rabbit  289  AB_2105622  p-mTOR (S2448)  2971  Cell Signaling Technology  Rabbit  289  AB_330970  p-mTOR (S2481)  2974  Cell Signaling Technology  Rabbit  289  AB_2231885  AKT  9272  Cell Signaling Technology  Rabbit  60  AB_329827  p-AKT (S473)  9271  Cell Signaling Technology  Rabbit  60  AB_329825  TSC2  4308  Cell Signaling Technology  Rabbit  200  AB_10547134  p-TSC2 (S939)  3615  Cell Signaling Technology  Rabbit  200  AB_2207796  p-TSC2 (T1462)  3617  Cell Signaling Technology  Rabbit  200  AB_490956  p-TSC2 (S1387)  5584  Cell Signaling Technology  Rabbit  200  AB_10698883  Target  Product No.  Manufacturer  Made in  Size (kDa)  RRID  GR  12041  Cell Signaling Technology  Rabbit  91–94  AB_2631286  FOXO1  2880  Cell Signaling Technology  Rabbit  78–82  AB_2106495  GLUL  G45020  BD Transduction Laboratory  Mouse  40  AB_2313767  S6  2217  Cell Signaling Technology  Rabbit  32  AB_331355  p-S6 (S235/236)  4856  Cell Signaling Technology  Rabbit  32  AB_2181037  4E-BP1  9452  Cell Signaling Technology  Rabbit  15–20  AB_331692  p-4E-BP1 (S65)  9451  Cell Signaling Technology  Rabbit  15–20  AB_330947  p-4E-BP1 (T37/46)  9459  Cell Signaling Technology  Rabbit  15–20  AB_330985  mTOR  2983  Cell Signaling Technology  Rabbit  289  AB_2105622  p-mTOR (S2448)  2971  Cell Signaling Technology  Rabbit  289  AB_330970  p-mTOR (S2481)  2974  Cell Signaling Technology  Rabbit  289  AB_2231885  AKT  9272  Cell Signaling Technology  Rabbit  60  AB_329827  p-AKT (S473)  9271  Cell Signaling Technology  Rabbit  60  AB_329825  TSC2  4308  Cell Signaling Technology  Rabbit  200  AB_10547134  p-TSC2 (S939)  3615  Cell Signaling Technology  Rabbit  200  AB_2207796  p-TSC2 (T1462)  3617  Cell Signaling Technology  Rabbit  200  AB_490956  p-TSC2 (S1387)  5584  Cell Signaling Technology  Rabbit  200  AB_10698883  Abbreviations: 4E-BP1, 4E-binding protein 1; RRID, Research Resource Identifier. View Large FCSA determination Gastrocnemius muscle was embedded in Tissue‐Tek (Sakura Finetek) and sectioned on a Leica CM3050 S cryostat at −20°C. Subsequently, serial cross‐sections (5 μm) were stained with anti‐laminin (no. L‐9393; Sigma‐Aldrich) to determine the FCSA The sections were incubated with Alexa Fluor 350 (no. A‐21426; Invitrogen) as secondary antibody. Digital images of the stained sections were taken at ×200 total magnification using an Eclipse E800 microscope (Nikon) connected to a digital camera (DXM, 1200 NF, Nikon). The FCSA in the glycolytic region of the gastrocnemius was measured for >100 individual fibers per animal, using the Lucia software (version 4.81). A distribution curve was composed of all measured fibers within the groups (>800 fibers). Statistical analysis Data are shown as means ± standard error of the mean. Comparisons were computed with SPSS version 20. Statistical significance between groups within a genotype (wild-type or mGRKO) was tested using a one-way analysis of variance (ANOVA) with a post hoc test. The type of post hoc analysis was chosen on the basis of the data variance (Levene’s test), with the Tukey test for data with equal variance and the Games–Howell test for all other data. Statistical significance between genotypes of the normoxic group was assessed using the independent samples t test. A two-way ANOVA was used to determine the effect of GR deficiency on the response of the individual treatments. A χ2 test was used to compare FCSA distributions. A P values < 0.05 was considered to be statistically significant and 0.05 ≤ P ≤ 0.1 as indicating a trend. Results Hypoxia increases plasma corticosterone concentrations and induces muscle GR signaling Four days of hypoxia resulted in elevated corticosterone concentrations (Fig. 1A). This increase was in part attributable to the reduced food intake (pair-fed, 2.0-fold; hypoxia, 3.0-fold). To address whether the elevated circulating corticosterone concentrations activated muscle GR signaling in vivo, a GRE reporter (luciferase) plasmid was electroporated into the gastrocnemius muscle prior to 4 days exposure to normoxic, hypoxic, or pair-fed conditions (Fig. 1B). The transcriptional activity of GR was increased under hypoxic conditions (1.9-fold) and even more so in the pair-fed group (3.1-fold). Figure 1. View largeDownload slide Hypoxia raises corticosterone plasma concentrations and induces muscle GR signaling. (A) Plasma corticosterone concentration at day 4. (B) GR transcriptional activity in gastrocnemius muscle at day 4. GRE reporter plasmid (luciferase) data were corrected for control (β-galactosidase) plasmid data. Significant differences between groups at a given time point are indicated as follows: *P < 0.05; trends (P ≤ 0.1) are indicated by the specific P value (n = 8 per group). Figure 1. View largeDownload slide Hypoxia raises corticosterone plasma concentrations and induces muscle GR signaling. (A) Plasma corticosterone concentration at day 4. (B) GR transcriptional activity in gastrocnemius muscle at day 4. GRE reporter plasmid (luciferase) data were corrected for control (β-galactosidase) plasmid data. Significant differences between groups at a given time point are indicated as follows: *P < 0.05; trends (P ≤ 0.1) are indicated by the specific P value (n = 8 per group). GR target genes are differentially expressed in mGRKO To assess the importance of GR signaling during hypoxia-induced muscle atrophy, mGRKO mice were used. In these mice, GR protein (Fig. 2A) and Gr mRNA (Fig. 2B) were reduced by 60% in the gastrocnemius muscles. Interestingly, both in control and mGRKO mice, Gr mRNA levels were increased by hypoxia, but not by reduced food intake alone. Baseline plasma corticosterone concentrations were comparable in both genotypes and showed similar increases in response to reduced food intake and hypoxia (Fig. 2C). mRNA expression of postulated GR target genes related to muscle protein turnover was evaluated. Both Glul (Fig. 2D) and Foxo1 (Fig. 2E) expression were GR-dependent, as no increased expression was found by hypoxia and reduced food intake in mGRKO mice. Protein levels of FOXO1 and GLUL also show a GR-dependent increase in response to hypoxia (Supplemental Fig. 1). Overall, the expression of Mstn was reduced by GR deficiency, although the responses to reduced food intake and hypoxia remained similar (Fig. 2F). In contrast, expression of Klf15 (Fig. 2G) and Redd1 (Fig. 2H) was similar in response between control and mGRKO mice and therefore not GR-dependent. To conclude, corticosterone concentrations increased equally in control and mGRKO mice in response to hypoxia. Hypoxia-induced expression of Glul and Foxo1 in skeletal muscle was attributable to the reduced food intake and was GR-dependent. Figure 2. View largeDownload slide GR target genes are differentially expressed in mGRKO. (A) GR protein in control and mGRKO gastrocnemius muscle. (B) mRNA expression of Gr. (C) Plasma corticosterone concentration. mRNA expression of (D) Glul, (E) Foxo1, (F) Mstn, (G) Klf15, and (H) Redd1 is shown. All mRNA concentrations were corrected by 18S concentration. Significant differences between groups within control or mGRKO mice are indicated by asterisks on horizontal lines; basal differences in normoxia groups between control and mGRKO mice are indicated by a hash sign: */#P < 0.05, **/##P ≤ 0.01, ***P ≤ 0.001; trends (P ≤ 0.1) are indicated by the specific P value. Significant effects (P ≤ 0.05) of GR deficiency on the response of the treatment analyzed by two-way ANOVA are indicated as follows: &, normoxia vs hypoxia; $, pair-fed vs hypoxia (n = 6–8). Figure 2. View largeDownload slide GR target genes are differentially expressed in mGRKO. (A) GR protein in control and mGRKO gastrocnemius muscle. (B) mRNA expression of Gr. (C) Plasma corticosterone concentration. mRNA expression of (D) Glul, (E) Foxo1, (F) Mstn, (G) Klf15, and (H) Redd1 is shown. All mRNA concentrations were corrected by 18S concentration. Significant differences between groups within control or mGRKO mice are indicated by asterisks on horizontal lines; basal differences in normoxia groups between control and mGRKO mice are indicated by a hash sign: */#P < 0.05, **/##P ≤ 0.01, ***P ≤ 0.001; trends (P ≤ 0.1) are indicated by the specific P value. Significant effects (P ≤ 0.05) of GR deficiency on the response of the treatment analyzed by two-way ANOVA are indicated as follows: &, normoxia vs hypoxia; $, pair-fed vs hypoxia (n = 6–8). Hypoxia-induced muscle atrophy is not prevented by muscle GR deficiency Gastrocnemius muscle weights of mGRKO animals tended to be increased (+4%, P = 0.06) compared with control mice under basal conditions (Fig. 3A). A significant increase under basal conditions was found when combining weights of gastrocnemius, plantaris, tibialis, and extensor digitorum longus (Supplemental Fig. 2). Despite a trend in muscle weights, FCSA under normoxic conditions between control and mGRKO was not significantly different (Fig. 3C; Supplemental Fig. 3A). No alterations in fiber type composition were observed between control and mGRKO mice (Supplemental Fig. 3B). In control animals, the mass of the gastrocnemius muscle decreased under hypoxia, which was in part attributable to reduced food intake (pair-fed, −7.1%; hypoxia, −12.9%). In contrast, in mGRKO mice, muscle mass was preserved under pair-fed conditions. Accordingly, reduced food intake resulted in a lower FCSA in control mice, but not in the mGRKO mice (Fig. 3B–E). Hypoxia-induced muscle atrophy was still observed in mGRKO mice, and muscle loss (hypoxia vs normoxia; control, −12.9% vs mGRKO, −8.5%) was not significantly different from control (Fig. 3A). Hypoxia resulted in a change in fiber size distribution (χ2: P < 0.05, Fig. 3D) and lower average FCSA (Fig. 3C) in control mice, comparable to effects of reduced food intake. In the mGRKO mice, hypoxia also shifted fiber size distribution (χ2: P < 0.05, Fig. 3E) accompanied by an apparent (P = 0.07) decrease in average FCSA (Fig. 3C), despite GR deficiency. Collectively, these data revealed that muscle atrophy attributable to the hypoxia-induced reduction in food intake was GR-dependent, whereas the hypoxia-specific reduction of muscle mass was not. Figure 3. View largeDownload slide Hypoxia-induced muscle atrophy is not prevented by muscle GR deficiency. (A) Gastrocnemius muscle weight after 4 days of hypoxia or pair-fed conditions in control and mGRKO mice as percentage of the muscle mass of normoxic control mice. (B) Representative images of laminin-stained cryosections (magnification: ×200). (C) Average FCSA of glycolytic region in gastrocnemius muscle of control and mGRKO mice. (D) FCSA distribution of the glycolytic region in gastrocnemius muscle of control mice (no significance indicated). (E) FCSA distribution of the glycolytic region in gastrocnemius muscle of mGRKO mice (no significance indicated). Significant differences between groups within control or mGRKO mice are indicated by asterisks on horizontal lines; basal differences in normoxia groups between control and mGRKO mice are indicated by a hash sign: */#P < 0.05, **P ≤ 0.01, ***P ≤ 0.001; trends (P ≤ 0.1) are indicated by the specific P value. Significant effects (P ≤ 0.05) of GR deficiency on the response of the treatment analyzed by two-way ANOVA are indicated as follows: $, pair-fed vs hypoxia (n = 6–8). Figure 3. View largeDownload slide Hypoxia-induced muscle atrophy is not prevented by muscle GR deficiency. (A) Gastrocnemius muscle weight after 4 days of hypoxia or pair-fed conditions in control and mGRKO mice as percentage of the muscle mass of normoxic control mice. (B) Representative images of laminin-stained cryosections (magnification: ×200). (C) Average FCSA of glycolytic region in gastrocnemius muscle of control and mGRKO mice. (D) FCSA distribution of the glycolytic region in gastrocnemius muscle of control mice (no significance indicated). (E) FCSA distribution of the glycolytic region in gastrocnemius muscle of mGRKO mice (no significance indicated). Significant differences between groups within control or mGRKO mice are indicated by asterisks on horizontal lines; basal differences in normoxia groups between control and mGRKO mice are indicated by a hash sign: */#P < 0.05, **P ≤ 0.01, ***P ≤ 0.001; trends (P ≤ 0.1) are indicated by the specific P value. Significant effects (P ≤ 0.05) of GR deficiency on the response of the treatment analyzed by two-way ANOVA are indicated as follows: $, pair-fed vs hypoxia (n = 6–8). GR deficiency differentially suppresses UPS and autophagy lysosomal pathway proteolytic signaling Muscle atrophy may involve elevated protein degradation through the UPS and the autophagy lysosomal pathway (ALP). Expression of E3-ubiquitin ligases Murf1, Atrogin-1, Musa1, and Smart was investigated to assess UPS activation. Basal Murf1, Atrogin-1, Musa1, and Smart expression was similar in control and mGRKO mice (Fig. 4A–4D). In control mice, reduced food intake and hypoxia equally elevated the expression of the investigated ubiquitin ligases. GR deletion diminished the response of Murf1 and Atrogin-1 expression to reduced food intake, whereas under hypoxic conditions the response was hardly diminished. Increased Musa1 expression in response to reduced food intake and hypoxia was absent in the mGRKO mice. In contrast, increased Smart expression by reduced food intake and hypoxia was not affected by GR deletion. Basal expression levels of the ALP-related genes Bnip3 and Map1lc3B were similar in control and mGRKO animals (Fig. 4E and 4F). Reduced food intake and hypoxia increased expression of both ALP-related genes to a similar extent in both controls and mGRKO mice, although the response was blunted in the latter. In both control and mGRKO mice, MAP1LC3B-I was reduced and MAP1LC3B-II levels were unaltered in response to reduced food intake, resulting in an increased II/I ratio (Fig. 4G–4I). Under hypoxic conditions, MAP1LC3B-I and -II were reduced similarly, resulting in an unaltered II/I ratio compared with normoxia in control and GRKO mice. To summarize, whereas reduced food intake under normoxia induced mRNA expression of Murf1 and Atrogin-1 GR-dependently, their increased transcript levels observed after hypoxia were minimally affected by GR deficiency. This differential response to reduced food intake under normoxic vs hypoxic conditions mediated by muscle GR expression was not observed for the other E3-ubiquitin ligases Musa1 and Smart. Furthermore, GR deficiency similarly blunted ALP-related gene expression in response to hypoxia or reduced food intake, but did not affect changes in MAP1LC3B-I and -II protein turnover. Figure 4. View largeDownload slide GR deficiency differentially suppresses UPS and ALP proteolytic signaling. (A) Murf1 expression. (B) Atrogin-1 expression. (C) Musa1 expression. (D) Smart expression (E) Bnip3 expression. (F) Map1lc3B expression. (G) MAP1LC3B-I protein. (H) MAP1LC3B-II protein. (I) MAP1LC3B-II/I ratio. All mRNA concentrations were corrected by 18S concentration. Significant differences between groups within control or mGRKO mice are indicated by asterisks on horizontal lines; basal differences in normoxia groups between control and mGRKO mice are indicated by a hash sign: *P < 0.05, **/##P ≤ 0.01, ***P ≤ 0.001; trends (P ≤ 0.1) are indicated by the specific P value. Significant effects (P ≤ 0.05) of GR deficiency on the response of the treatment analyzed by two-way ANOVA are indicated as follows: €, pair-fed vs normoxia; &, normoxia vs hypoxia; $, pair-fed vs hypoxia (n = 6–8). Figure 4. View largeDownload slide GR deficiency differentially suppresses UPS and ALP proteolytic signaling. (A) Murf1 expression. (B) Atrogin-1 expression. (C) Musa1 expression. (D) Smart expression (E) Bnip3 expression. (F) Map1lc3B expression. (G) MAP1LC3B-I protein. (H) MAP1LC3B-II protein. (I) MAP1LC3B-II/I ratio. All mRNA concentrations were corrected by 18S concentration. Significant differences between groups within control or mGRKO mice are indicated by asterisks on horizontal lines; basal differences in normoxia groups between control and mGRKO mice are indicated by a hash sign: *P < 0.05, **/##P ≤ 0.01, ***P ≤ 0.001; trends (P ≤ 0.1) are indicated by the specific P value. Significant effects (P ≤ 0.05) of GR deficiency on the response of the treatment analyzed by two-way ANOVA are indicated as follows: €, pair-fed vs normoxia; &, normoxia vs hypoxia; $, pair-fed vs hypoxia (n = 6–8). Hypoxia-induced impairment of protein synthesis signaling is GR-dependent The mTOR complex 1 (mTORC1) pathway is central in the regulation of protein synthesis signaling (37, 38). Ribosomal protein S6 kinase, 70 kDa, polypeptide 1 (P70S6K1) is directly phosphorylated by mTOR. P70S6K1 phosphorylates ribosomal protein S6 (RPS6), thereby controlling protein synthesis. P70S6K1 and RPS6 concentrations were unaffected by both experimental conditions and mouse model. As expected, reduced food intake lowered the P70S6K1 and RPS6 phosphorylation ratio, suggesting suppressed protein synthesis (Fig. 5A–5D). In contrast, the food-dependent reduction of the P70S6K1 and RPS6 phosphorylation ratios was impaired under hypoxic conditions. GR deficiency reduced baseline P70S6K1 and RPS6 phosphorylation ratios of normoxic controls. P70S6K1 and RPS6 phosphorylation ratios were further lowered by reduced food intake in mGRKO muscle, and, importantly, in contrast to control mice, P70S6K1 and RPS6 phosphorylation ratios were similarly suppressed by hypoxia. Alterations in the eukaryotic translation initiation factor 4E-binding protein 1 (4E-BP1) Ser65 phosphorylation ratio (Fig. 5E, 5H) mirrored the changes observed for P70S6K1 and RPS6 phosphorylation. 4E-BP1 threonine Thr37 and Thr46 phosphorylation ratios marginally differed between conditions and mouse models (Fig. 5F and 5H). 4E-BP1 protein concentrations were equally increased by hypoxia and reduced food intake in control but not in GR-deficient mice (Fig. 5G and 5H). Collectively, these data indicate that inhibition of mTORC1 by reduced food intake under normoxia does not require muscle GR signaling. In contrast, retained mTORC1 activity under hypoxic conditions, despite reduced food intake, is GR-dependent. Figure 5. View largeDownload slide Hypoxia-associated dysregulation of protein synthesis regulation is GR-dependent. (A) Phosphorylation ratio (phosphorylated/total protein) of P70S6K1 at Thr389. (B) Phosphorylation ratio (phosphorylated/total protein) of RPS6 at Ser235/236. (C) Representative images of P70S6K Western blots. (D) Representative images of S6 Western blots. (E) Phosphorylation ratio (phosphorylated/total protein) of 4E-BP1 at Ser65. (F) Phosphorylation ratio (phosphorylated/total protein) of 4E-BP1 at Thr37/46. (G) Total 4E-BP1. (H) Representative images of 4EBP1 Western blots. Significant differences between groups within control or mGRKO mice are indicated by asterisks on horizontal lines; basal differences in normoxia groups between control and mGRKO mice are indicated by a hash sign: */#P < 0.05, **/##P ≤ 0.01, ***P ≤ 0.001; trends (P ≤ 0.1) are indicated by the specific P value. Significant effects (P ≤ 0.05) of GR deficiency on the response of the treatment analyzed by two-way ANOVA are indicated as follows: €, pair-fed vs normoxia; &, normoxia vs hypoxia; $, pair-fed vs hypoxia (n = 6–8). Figure 5. View largeDownload slide Hypoxia-associated dysregulation of protein synthesis regulation is GR-dependent. (A) Phosphorylation ratio (phosphorylated/total protein) of P70S6K1 at Thr389. (B) Phosphorylation ratio (phosphorylated/total protein) of RPS6 at Ser235/236. (C) Representative images of P70S6K Western blots. (D) Representative images of S6 Western blots. (E) Phosphorylation ratio (phosphorylated/total protein) of 4E-BP1 at Ser65. (F) Phosphorylation ratio (phosphorylated/total protein) of 4E-BP1 at Thr37/46. (G) Total 4E-BP1. (H) Representative images of 4EBP1 Western blots. Significant differences between groups within control or mGRKO mice are indicated by asterisks on horizontal lines; basal differences in normoxia groups between control and mGRKO mice are indicated by a hash sign: */#P < 0.05, **/##P ≤ 0.01, ***P ≤ 0.001; trends (P ≤ 0.1) are indicated by the specific P value. Significant effects (P ≤ 0.05) of GR deficiency on the response of the treatment analyzed by two-way ANOVA are indicated as follows: €, pair-fed vs normoxia; &, normoxia vs hypoxia; $, pair-fed vs hypoxia (n = 6–8). Perturbations in signaling upstream of mTOR correspond to hypoxia-induced decreases in food intake and are independent of GR mTOR activity is controlled through direct phosphorylation of Ser2448 by AKT, and AKT-mediated inhibition of TSC2 through phosphorylation of sites Ser939 and Thr1462 in TSC2. The basal AKT S473 phosphorylation ratio in mGRKO mice was slightly reduced compared with control (P = 0.06) (Fig. 6A and 6G). In control mice, hypoxia and reduced food intake equally lowered the AKT phosphorylation ratio, whereas in mGRKO mice no further reduction was seen. GR deficiency significantly lowered the mTOR S2448 phosphorylation ratio (Fig. 6B and 6H). Reductions in mTOR S2448 phosphorylation ratio in response to hypoxia and decreased food intake were similar and did not differ between control and mGRKO mice. mTOR Ser2481 autophosphorylation is postulated to reflect mTOR activity (39). Except for a decrease in the phosphorylation ratio in response to hypoxia in mGRKO mice, no changes were observed (Fig. 6C and 6H). TSC2 phosphorylation (S939 and T1462) ratios were not affected by any condition (Fig. 6D, 6E, and 6I). Total TSC2 was lowered by reduced food intake in control mice alone (Fig. 6F and 6I). AKT-independent activation of TSC2 by phosphorylation on Ser1387 was unaffected (Supplemental Fig. 4). Collectively, these data indicate that TSC2 signaling is not involved in suppression of mTOR by reduced food intake, whereas the reduction of the AKT/mTOR signaling by reduced food intake is intact under hypoxic conditions and is independent of GR signaling. Figure 6. View largeDownload slide Perturbations in signaling upstream of mTOR correspond to hypoxia-induced decreases in food intake and are GR-independent. (A) Phosphorylation ratio (phosphorylated/total protein) of AKT at Ser473. (B) Phosphorylation ratio (phosphorylated/total protein) of mTOR at Ser2448. (C) Phosphorylation ratio (phosphorylated/total protein) of mTOR at Ser2481. (D) Phosphorylation ratio (phosphorylated/total protein) of TSC2 at Ser939. (E) Phosphorylation ratio (phosphorylated/total protein) of TSC2 at Ser1462. (F) Protein levels of TSC2. (G) Representative images of AKT Western blots. (H) Representative images of mTOR Western blots. (G) Representative images of TSC2 Western blots. Significant differences between groups within control or mGRKO mice are indicated by asterisks on horizontal lines; basal differences in normoxia groups between control and mGRKO mice are indicated by a hash sign: */#P < 0.05, **/##P ≤ 0.01, ***P ≤ 0.001; trends (P ≤ 0.1) are indicated by the specific P value. Significant effects (P ≤ 0.05) of GR deficiency on the response of the treatment analyzed by two-way ANOVA are indicated as follows: €, pair-fed vs normoxia (n = 6–8). Figure 6. View largeDownload slide Perturbations in signaling upstream of mTOR correspond to hypoxia-induced decreases in food intake and are GR-independent. (A) Phosphorylation ratio (phosphorylated/total protein) of AKT at Ser473. (B) Phosphorylation ratio (phosphorylated/total protein) of mTOR at Ser2448. (C) Phosphorylation ratio (phosphorylated/total protein) of mTOR at Ser2481. (D) Phosphorylation ratio (phosphorylated/total protein) of TSC2 at Ser939. (E) Phosphorylation ratio (phosphorylated/total protein) of TSC2 at Ser1462. (F) Protein levels of TSC2. (G) Representative images of AKT Western blots. (H) Representative images of mTOR Western blots. (G) Representative images of TSC2 Western blots. Significant differences between groups within control or mGRKO mice are indicated by asterisks on horizontal lines; basal differences in normoxia groups between control and mGRKO mice are indicated by a hash sign: */#P < 0.05, **/##P ≤ 0.01, ***P ≤ 0.001; trends (P ≤ 0.1) are indicated by the specific P value. Significant effects (P ≤ 0.05) of GR deficiency on the response of the treatment analyzed by two-way ANOVA are indicated as follows: €, pair-fed vs normoxia (n = 6–8). Discussion In this study, the contribution of muscle GR signaling to hypoxia-induced muscle atrophy was addressed. As food intake is reduced in mice exposed to hypoxia (26), a pair-fed group was included to assess the GR dependency of responses induced by hypoxia and by semistarvation under normoxic conditions. Muscle GR signaling was responsible for muscle atrophy in the pair-fed group but not in the hypoxic group. The increased expression of UPS-related genes was mainly independent of GR in muscle of hypoxic mice. Suppression of mTORC1 activity under semistarvation conditions was impaired by hypoxia in a GR-dependent manner and independent of AKT signaling. Fasting-induced muscle atrophy is associated with elevated corticosterone concentrations (40). In line with this, hypoxia-induced reduction of food intake increases corticosterone concentrations. Mice subjected to hypoxia in our model are hypoxemic (26), and hypoxemia has also been associated with elevated glucocorticoid concentrations and GR signaling independent of malnutrition (41–45). However, as no further increase in glucocorticoid concentration is observed in hypoxic compared with normoxic pair-fed mice, this suggests that the elevated corticosterone concentration is attributable to the reduced food intake alone. Muscle GR signaling as demonstrated by GRE reporter activation indicates that circulating corticosterone affects skeletal muscle. Interestingly, GRE activation induced by restricted feeding and hypoxia does not reflect plasma corticosterone levels. This may be a consequence of differential prereceptor modulation of glucocorticoid availability in skeletal muscle by 11β-HSD1, as its expression and activity are critical regulatory steps in glucocorticoid responses (46). Considering their postulated transcriptional regulation by GR (18, 47), the increases in Glul and Foxo1 likely reflect genomic actions of activated GR in response to raised corticosterone concentrations. Interestingly, in response to hypoxia, muscle Gr mRNA transcript levels increase. However, as increased Glul and Foxo-1 expression and GR reporter activity do not display consistent additive effects of hypoxia compared with reduced food intake, the genomic actions of GR in hypoxic muscle are probably attributable to the reduced food intake. Although increased by semistarvation, mRNA expression of the putative GR target genes Klf15 and Redd1 (18, 47) is minimally affected by GR deficiency. Furthermore, under hypoxic conditions, increases in Klf15 and Redd1 expression are attenuated (Klf15) or absent (Redd1), indicating that hypoxia blocks transcriptional activation of these genes by semistarvation. Testosterone concentrations were not measured in this study, but it has been shown previously that induction of Redd1 expression by glucocorticoids is abolished in the presence of an elevated testosterone concentration (48). This may explain the lack of Redd1 transcript accumulation in muscle of hypoxic mice, as hypoxia is known to increase serum testosterone concentrations within 1 day (49). Protein degradation in skeletal muscle involves the UPS (50) and the ALP (51). MAP1LC3B-I protein levels are reduced despite increased Map1lc3b mRNA concentrations, suggestive of increased conversion to MAP1LC3B-II. Although MAP1LC3B-II protein levels are reduced, this could be the result of its elevated degradation due to an increased autophagic flux (52). The expression of the ALP-related genes Map1lc3b and Bnip3 is mainly upregulated by the hypoxia-mediated reduction in food intake. GR deficiency blunts this response slightly, but significantly. Interestingly, this corresponds with the GR-dependent induction of Foxo-1, which has been implicated in the transcriptional regulation of ALP-related genes (51), and suggests that genomic actions of GR may indirectly contribute to control of autophagy in muscle (12, 18, 53, 54). FOXO1 is a transcriptional regulator of Murf1 and Atrogin-1 (14), and its expression corresponds to the lowered transcript levels of these atrogenes under conditions of reduced food intake. However, in response to hypoxia our data imply a GR-independent mechanism that does not require increased Foxo1 expression in the regulation of these “classical” atrogenes. KLF15 is a key regulator of Murf1 and Atrogin-1 expression and is reported to be a direct target gene of GR (18). Its GR-independent regulation in response to reduced food intake, as well as its decreased expression in response to hypoxia, however, excludes a key regulatory function of de novo synthesized KLF15 on Murf1 and Atrogin-1 expression under hypoxic conditions. As KLF15 and FOXO1 are not responsible for the increased expression of Murf1 and Atrogin-1 during hypoxia, the transcriptional regulators in play remain to be identified. A novel finding is that increased Musa1 expression requires GR signaling in response to reduced food intake and hypoxia, whereas the newly identified E3 ligase Smart (17) is insensitive to GR deletion (22, 55, 56). Mothers against decapentaplegic homolog 3 (SMAD3) is a transcription factor involved in the transforming growth factor-β signaling (57). Increased Smad3 expression and transcriptional activity has been associated with increased Atrogin-1 expression (58). Smad3 expression is elevated by hypoxia independently of reduced food intake or GR signaling (Supplemental Fig. 5). Interestingly, of all protein turnover signaling constituents measured, only Atrogin-1 mRNA levels were increased in hypoxic compared with pair-fed GRKO muscle. This may point at involvement of transforming growth factor-β signaling in GR-independent hypoxia-induced muscle atrophy. Collectively, our data reveal GR-dependent and -independent constituents of proteolysis regulation. GR deficiency attenuates the overall expression of ALP-associated genes. Furthermore, Musa1 is identified as a GR-regulated E3-ligase, and genomic actions of GR are involved in upregulation of Murf1 and Atrogin-1 expression when food intake is reduced, whereas Atrogin-1 and Murf1 induction under hypoxic conditions occurs independently of GR. Hypoxia reduces food intake (26), and fasting results in the suppression of muscle protein synthesis (59, 60). mTORC1 activity (based on 4E-BP1, P70S6K1, and RPS6 phosphorylation ratio) is decreased accordingly in muscle of normoxic control mice in response to reduced food intake. Interestingly, in mGRKO muscle mTORC1 activity is reduced to a similar extent as in pair-fed control animals in response to reduced food intake, indicating that this suppression of mTORC1 activity occurs independently of muscle GR signaling. In addition to increases in endogenous glucocorticoids, nutritional restriction is accompanied by decreased circulating insulin and insulin-like growth factor-1 levels (61). Although plasma insulin was not measured, the low AKT phosphorylation in muscle of pair-fed control mice suggests that reduced anabolic signaling cues, rather than the increased circulating corticosterone concentration, are responsible for the suppression of mTORC1 activity caused by reduced food intake. A remarkable finding concerns the sustained mTORC1 activity in hypoxia despite reduced food intake. Although this confirmed our previous observations (26), we set out to further verify impairment of mTORC1 regulation by feeding cues under hypoxic conditions. Subjecting wild-type mice to 24 hours of complete food deprivation under hypoxic and normoxic conditions revealed that fasting-induced inhibition of RPS6 and 4E-BP1 phosphorylation is only observed in skeletal muscle of normoxic but not hypoxic mice (Supplemental Fig. 6). This illustrates that inhibitory cues toward mTORC1 activity by partial or complete food deprivation are not appropriately sensed in hypoxic conditions. Alternatively, it may reflect a potential (futile) protective response to counter food restriction–induced suppression of protein synthesis. We now show that GR is involved in this dysregulation, because under hypoxic conditions in mGRKO mice mTORC1 activity is suppressed to levels comparable to pair-fed mice. REDD1 inhibits mTORC1 activity through liberating TSC2 by competing for its binding to 14-3-3 (62). Redd1 expression has been postulated to be GR-dependent (63). However, the lack of Redd1 induction in response to hypoxia in both control as well as in mGRKO muscle strongly suggests that Redd1 expression does not control mTORC1 activity in response to hypoxia. KLF15 has also been postulated to operate GR dependently and to inhibit mTORC1 activity through control of BCAT2 expression (18). In contrast to in vitro studies in which dexamethasone was used to induce GR/KLF15 signaling (18), Klf15 expression under hypoxia and reduced food intake in vivo was not GR-dependent and did not correlate inversely with mTORC1 activity. Collectively, this indicates that aberrant regulation of Redd1 and Klf15 by hypoxia is not responsible for the sustained mTORC1 activity in conditions of semistarvation. Nongenomic actions of GR have also been implicated in mTORC1 signaling by inhibiting AKT phosphorylation through an interaction with PI3K (64, 65), resulting in reduced Akt-mediated phosphorylation of downstream TSC2 (66) and mTOR at S2448 (37). However, TSC2 phosphorylation is not affected, whereas suppressed AKT and S2448 mTOR phosphorylation by either reduced food intake alone or hypoxia is not reversed in GRKO muscle, although basal AKT phosphorylation is already lowered by GR deficiency. Therefore, nongenomic actions of GR on AKT signaling are not involved in the lack of mTORC1 inhibition under hypoxic conditions. Autophosphorylation of mTOR at site S2481 has been described and contributes to mTORC1 activity, but its regulation is only partly understood (37). Interestingly, only in mGRKO muscle a reduction of mTOR S2481 phosphorylation under hypoxia was observed, which corresponds to the restoration of an appropriate response to semistarvation even in the presence of hypoxia. Although the exact mechanism remains to be elucidated, this suggests that GR-dependent preservation of mTOR S2481 phosphorylation may contribute to impaired mTORC1 regulation, that is, retained mTORC1 activity during hypoxia. Overall, the GR-dependent impairment of mTORC1 activity regulation in response to hypoxia is neither explained by aberrant upstream TSC2 or AKT/mTOR-S2448 signaling, nor by alterations in Redd1 or Klf15 expression. GR-mediated control of mTOR S2481 phosphorylation therefore appears to contribute to the regulation of protein synthesis independently of the established nongenomic actions of GR. Protein synthesis and degradation signaling change in a coordinated fashion in conditions altering muscle mass (67–69). In line, the decreased protein synthesis signaling in response to semistarvation is accompanied by an increased expression of genes mediating proteolysis in atrophying muscle. Still, increased proteolysis signaling rather than reduced synthesis signaling likely provides the main GR-dependent contribution to muscle atrophy in response to normoxic food restriction. Conversely, most of these attenuating effects of GR deficiency on proteolysis signaling are absent under hypoxia, and muscle atrophy is not prevented in GRKO mice. This implies differential (i.e., GR-independent) regulation of proteolysis under hypoxia compared with normoxic food restriction. Importantly, the coordination of protein synthesis and proteolysis signaling cues appears impaired under hypoxic conditions in a GR-dependent manner. However, considering that muscle atrophy under hypoxia is equal in wild-type and GRKO mice despite suppressed mTOR signaling in GRKO muscle, the physiological relevance of sustained stimulatory signals for protein synthesis signaling under hypoxia remain to be clarified, as our data reveal it does not function as an adaptive response to protect from further aggravation of muscle atrophy induced by reduced food intake. In COPD patients with muscle atrophy, molecular signatures reflecting parallel increases in protein synthesis and degradation signaling in skeletal muscle have been reported (70–73) and are associated with a hypermetabolic state. This impaired coordination of protein synthetic and proteolytic cues may contribute to the loss of muscle mass observed in pathological conditions. In conclusion, GR signaling is required for muscle atrophy and increased expression of proteolysis-associated genes induced by decreased food intake under normoxic conditions. Under hypoxic conditions, muscle atrophy and elevated gene expression of the UPS-associated E3 ligases Murf1 and Atrogin-1 are mostly independent of GR signaling. Importantly, impaired inhibition of mTORC1 activity is GR-dependent in hypoxia-induced muscle atrophy. Abbreviations: 4E-BP1 4E-binding protein 1 ALP autophagy lysosomal pathway ANOVA analysis of variance COPD chronic obstructive pulmonary disease FCSA fiber cross-sectional area FOXO1 forkhead box protein O1 GLUL glutamate-ammonia ligase GR glucocorticoid receptor GRE glucocorticoid receptor–responsive element KLF15 Krüppel-like factor 15 mGRKO muscle‐specific glucocorticoid receptor knockout mRNA messenger RNA MSTN myostatin mTOR mammalian target of rapamycin mTORC1 mammalian target of rapamycin complex 1 PI3K phosphatidylinositol-4,5-bisphosphate 3-kinase P70S6K1 protein S6 kinase, 70 kDa, polypeptide 1 qPCR quantitative polymerase chain reaction RPS6 ribosomal protein S6 SMAD3 mothers against decapentaplegic homolog 3 UPS ubiquitin proteasomal system. Acknowledgments The GRE luciferase construct was provided by R. Hoffmann (University Medical Center Groningen, Groningen, Netherlands). 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