Induction of ATF4-Regulated Atrogenes Is Uncoupled from Muscle Atrophy during Disuse in Halofuginone-Treated Mice and in Hibernating Brown Bears

Abstract

Activating transcription factor 4 (ATF4) is involved in muscle atrophy through the overexpression of some atrogenes. However, it also controls the transcription of genes involved in muscle homeostasis maintenance. Here, we explored the effect of ATF4 activation by the pharmacological molecule halofuginone during hindlimb suspension (HS)-induced muscle atrophy. Firstly, we reported that periodic activation of ATF4-regulated atrogenes (Gadd45a, Cdkn1a, and Eif4ebp1) by halofuginone was not associated with muscle atrophy in healthy mice. Secondly, halofuginone-treated mice even showed reduced atrophy during HS, although the induction of the ATF4 pathway was identical to that in untreated HS mice. We further showed that halofuginone inhibited transforming growth factor-β (TGF-β) signalling, while promoting bone morphogenetic protein (BMP) signalling in healthy mice and slightly preserved protein synthesis during HS. Finally, ATF4-regulated atrogenes were also induced in the atrophy-resistant muscles of hibernating brown bears, in which we previously also reported concurrent TGF-β inhibition and BMP activation. Overall, we show that ATF4-induced atrogenes can be uncoupled from muscle atrophy. In addition, our data also indicate that halofuginone can control the TGF-β/BMP balance towards muscle mass maintenance. Whether halofuginone-induced BMP signalling can counteract the effect of ATF4-induced atrogenes needs to be further investigated and may open a new avenue to fight muscle atrophy. Finally, our study opens the way for further studies to identify well-tolerated chemical compounds in humans that are able to fine-tune the TGF-β/BMP balance and could be used to preserve muscle mass during catabolic situations.

1. Introduction

Many unloading conditions (e.g., microgravity, bed rest, or physical inactivity) lead to a loss of muscle mass and strength. This muscle atrophy is associated with adverse health effects such as autonomy decline and increased morbidity and mortality [1,2,3]. Considering the lack of proven, easy-to-use therapeutic or preventive treatment, muscle atrophy remains a major public health issue (World Health Organisation data) [4].

The underlying molecular mechanisms of muscle atrophy involve the dysregulation of a complex network of intracellular pathways leading to an imbalance in protein turnover [5,6,7,8,9]. Activating transcription factor 4 (ATF4) is overexpressed in many conditions of muscle atrophy [10,11,12,13,14] and is, therefore, considered as an atrogene, i.e., one of the genes with expression at the mRNA level that is commonly altered during atrophy [14]. ATF4 belongs to the integrated stress response (ISR) pathway, a conserved intracellular network activated in response to various intrinsic and extrinsic stresses (e.g., amino acid (AA) depletion and endoplasmic reticulum (ER) stress) to restore cellular homeostasis [15]. Activation of the ISR involves phosphorylation of eukaryotic translation initiation factor 2 (eIF2α) by several kinases (i.e., general control nonderepressible 2 (GCN2), protein kinase RNA-like ER kinase (PERK), protein kinase R (PKR), heme-regulated inhibitor (HRI), and microtubule affinity-regulating kinase 2 (MARK2)), resulting in the global inhibition of protein synthesis but the increased translation of certain mRNAs, including ATF4 [15,16]. Inhibition of ATF4 in skeletal muscle limits starvation-, immobilisation-, and ageing-induced atrophy, whereas ATF4 induction results in muscle wasting [11,12,13,17]. ATF4 target genes include some atrogenes, such as GADD45A and CDKN1A, that are required for ATF4-mediated muscle atrophy [11,12,13,18], and TRIB3, which is involved in fasting- and ageing-induced muscle atrophy [19,20]. However, ATF4 targets also include genes that may be involved in the maintenance of muscle homeostasis. Indeed, ATF4 contributes to the transcription of autophagy-related genes [21,22,23,24] and is activated during mitochondrial perturbations (e.g., oxidative stress) to restore mitochondrial homeostasis [25,26]. In fact, activation of the eIF2α-ATF4 pathway by the pharmacological molecule halofuginone (HF), prior to stressful events (i.e., ischemia-reperfusion injury models) has shown positive effects on the preservation of kidney and liver function [27]. Moreover, the maintenance of autophagy and mitochondria homeostasis are essential for maintaining muscle mass [28,29,30]. Altogether, this led to the hypothesis that the ATF4 pathway may have a dual role in skeletal muscle.

Interestingly, halofuginone also improved muscle performance during dystrophies, mainly through its antifibrotic properties [31,32,33,34]. Whether these beneficial effects involve a regulation of the ATF4 pathway has never been investigated. However, they mainly involved the inhibition of the transforming growth factor-β (TGF-β) pathway [35,36,37]. The TGF-β signalling pathway acts as a negative regulator of muscle mass, notably through the transcriptional induction of the atrogenes TRIM63/MuRF1 and FBXO32/Atrogin-1 [38,39,40]. When inhibited, it promotes a profound muscle hypertrophy phenotype in various conditions and species [41,42]. Members of TGF-β signalling belong to the TGF-β superfamily [38,39], as do the bone morphogenetic protein (BMP) signalling members. The BMP signalling pathway [39,43] instead acts as a positive regulator of muscle mass through the transcriptional repression of the atrogene FBXO30/Musa1, which is required for denervation-induced muscle loss [44,45]. When inhibited, it profoundly exacerbates denervation-induced muscle atrophy [44,45].

This study aimed to explore the impacts of HF-induced ATF4 signalling on skeletal muscle under basal conditions and hindlimb suspension-induced atrophy in mice. We further deciphered the molecular mechanisms of HF by focusing on protein metabolism and TGF-β/BMP signalling. We also took advantage of a model of muscle atrophy resistance that we previously explored [46], and studied the regulation of the ATF4-induced atrogenes in hibernating brown bear muscle.

2. Results

2.1. Induction of ATF4-Regulated Atrogenes Does Not Affect Muscle Mass in Mice

We used halofuginone (HF) to induce ATF4 transcriptional activity in mouse muscles, and we investigated the effect on skeletal muscle mass. For that purpose, mice were treated with HF three times a week for up to 4 weeks. Six hours after the last HF administration at the end of each week, we measured the mRNA levels for some ATF4 target genes, involved in muscle atrophy, i.e., Trib3, Cdkn1a, Gadd45a, and Eif4ebp1 (Figure 1A–F). Except for Atf4, for which mRNA levels were elevated during the first 2 weeks of HF treatment, Trib3, Cdkn1a, and Gadd45a were all overexpressed in muscles after 2 weeks of HF treatment compared to H₂O-treated mice. Of note, Eif4ebp1 mRNA levels increased in mouse muscles after 4 weeks of HF-treatment compared to H₂O, with a noticeable trend after 3 weeks of treatment (Figure 1F). We also investigated the regulation of other ATF4 target genes and showed the overexpression of Asns over the 4 weeks of HF treatment as well as a trend for Ddit3 and Ppp1r15a (Supplementary Figure S1).

These data underline an activation of the ATF4 transcriptional activity. However, despite the overexpression of the ATF4-regulated atrogenes, the mass of the gastrocnemius, soleus, tibialis anterior, and extensor digitorum longus (EDL) muscles remained unchanged during the 4 weeks of HF treatment (Figure 1G, Supplementary Figure S1). Altogether, these data suggest that a long-term halofuginone administration induced ATF4-regulated atrogenes without leading to muscle atrophy.

2.2. Overexpression of ATF4-Regulated Atrogenes during Hindlimb Suspension Is Uncoupled from Muscle Atrophy in HF-Treated Mice

We then investigated the effect of the induction of the ATF4 pathway by HF treatment during the muscle atrophy induced by hindlimb suspension (HS). Briefly, mice received HF three times a week for 3 weeks and were then, 3 days after the last HF administration, hindlimb-suspended or not for 3 or 7 days (Figure 2A). Measurements were thus performed at least 6 days after the last HF administration. HF induces ATF4 activation through the phosphorylation of eIF2α [47]. We observed that HF treatment resulted in overall higher phosphorylated and total eIF2α protein levels compared to H₂O-treated mice (Figure 2B–D). In addition, HS led to an overall decrease in phosphorylated eIF2α protein levels compared to the controls (Ctrls) (Figure 2B,C). In addition, levels of the mRNA encoding the phosphatase GADD34 (Ppp1r15a) were higher at 3 days of hindlimb suspension (HS3) compared to the Ctrls in both H₂O- and HF-treated mice (Supplementary Figure S2). The expression of ATF4-regulated genes was not different between Ctrl-HF and Ctrl-H₂O groups (Figure 2 and Supplementary Figure S2). The mRNA levels of Atf4 and its target genes involved in muscle atrophy, i.e., Trib3, Cdkn1a, and Eif4ebp1, were higher during HS in both H₂O- and HF-treated mice (Figure 2E–H), while the mRNA levels of the ATF4-regulated atrogene Gadd45a remained unchanged (Figure 2I). Of note, mRNA expression of other ATF4 target genes remained unchanged for Asns or slightly decreased upon HS for Ddit3. (Supplementary Figure S2). Altogether, Figure 1 and Figure 2 shows that (i) HF administration induced ATF4-regulated atrogenes after 6 h, but no more after 6 days, indicating that this effect was rapid and transient, and (ii) HS induced overexpression of these atrogenes.

We next investigated the outcomes on skeletal muscle. Gastrocnemius muscle had atrophied only in H₂O-treated mice after 3 days of HS (H₂O-HS3) and in both H₂O- and HF-treated mice after 7 days of HS. Surprisingly, the average of the fibre cross-sectional area (CSA) did not change in H₂O-HS3 mice compared to the Ctrls but was lower after HS7 regardless of the treatment (Figure 3B). We further analysed the distribution of gastrocnemius fibre CSA (Supplementary Figure S3A,B). We reported (i) a lower proportion of small fibres and (ii) a higher proportion of large fibres in HF-treated mice compared to the H₂O group (Supplementary Figure S3A). This observation was restricted to fast-twitch fibres (i.e., 2X/2B) (Supplementary Figure S3B). Taken together, our data suggest that induction of ATF4-regulated atrogenes is not associated with muscle atrophy after 3 days of hindlimb suspension in HF-treated mice and, thus, that HF slightly preserves muscle mass during HS.

2.3. Halofuginone Treatment Inhibits TGF-β While Promoting BMP Signalling in Gastrocnemius Muscle

HF inhibits the TGF-β pathway [36,48]. We and others have recently reported that inhibition of the TGF-β signalling is associated with the concomitant activation of BMP signalling [45,46]. Therefore, we investigated how HF treatment and subsequent HS treatment affected these pathways in skeletal muscle. The nuclear localisation of SMADs mirrors the upstream activation of the TGF-β or BMP pathway [49]. We, thus, measured protein levels for the transcription factors SMAD2/3 (TGF-β signalling), SMAD1/5 (BMP signalling), and SMAD4 (TGF-β and BMP signalling) in nuclear and cytosolic fractions (Figure 4A–D and Supplementary Figure S4A–F). The ratio of nuclear SMAD2/3 to total SMAD2/3 was very low in HF-treated mice compared to H₂O-treated mice and decreased upon HS only in H₂O-treated mice (Figure 4A,B). Consistently, the overall mRNA levels of several collagens, which are well-known target genes of TGF-β signalling, decreased upon HS (Supplementary Figure S5). Moreover, the ratio of nuclear SMAD1/5 to total SMAD1/5 was higher in HF-Ctrl mice than in H₂O-Ctrl mice. This ratio was reduced at HS7 compared to the Ctrl in HF-treated mice, while it was increased in H₂O-treated mice (Figure 4A,C). Finally, the ratio of nuclear SMAD4 to total SMAD4 was overall lower in HF- vs. H₂O-treated mice (Figure 4A–D), with a decrease in HF-treated mice at HS7 compared to the Ctrl.

TGF-β catabolic action involves the inhibition of protein synthesis [40,50,51], whereas the anabolic action of BMP involves its promotion [44]. The overall protein-synthesis rates measured by puromycin incorporation were reduced during hindlimb suspension. However, this decrease was only significant in H₂O-HS7 compared to H₂O-Ctrl mice (Figure 5A,B). TGF-β signalling acts also as a negative regulator of muscle mass through the induction of the atrogenes TRIM63/MurF1 and FBXO32/Atrogin-1 [38,39], while BMP signalling acts as a positive regulator with the transcriptional repression of the atrogene FBXO30/Musa1 [44,45]. Trim63 and Fbxo32 mRNA levels were upregulated only at HS3 in both H₂O and HF-treated mice, while mRNA levels for the atrogene Fbxo30 remained unchanged (Figure 5C–E). Our data suggest that while HF inhibits TGF-β signalling, it also promotes BMP signalling in the control gastrocnemius muscles. We also showed that HF partially attenuates the drop in protein synthesis during hindlimb suspension.

2.4. ATF4-Regulated Atrogenes Are Overexpressed in Atrophy-Resistant Hibernating Brown Bear Muscle

Our data strongly suggest that the induction of ATF4 signalling is not always associated with muscle atrophy, either in basal conditions or in HS-induced muscle atrophy. We took advantage of a natural model, i.e., the hibernating brown bear, which experiences only a moderate loss of muscle protein content while remaining completely inactive for up to 6 months [52,53,54]. As with HF treatment, we recently reported a concomitant TGF-β pathway inhibition and BMP pathway activation in hibernating brown bear muscle [46]. We, thus, explored whether ATF4-regulated atrogenes may also be induced in this model. Interestingly, as shown in Figure 6, Atf4 was upregulated in hibernating brown bear muscle compared to the active counterpart. In addition, the two main ATF4-regulated atrogenes (Gadd45a and Cdkn1a) and Trib3 were also induced in hibernating brown bear muscle. Other ATF4 target genes were either down- (Eif4ebp1, Ppp1r15a, and Asns) or upregulated (Ddit3). These data show that ATF4-regulated atrogenes are induced in hibernating brown bear muscle, even if they are resistant to atrophy.

3. Discussion

Several muscle-wasting conditions, including fasting or physical inactivity, are associated with eIF2α phosphorylation [55,56] and/or ATF4 overexpression, which trigger muscle atrophy [10,11,12,13,18,57]. Moreover, muscle atrophy is hampered during fasting or ageing in mice with reduced ATF4 expression or expressing a phosphorylation-resistant form of eIF2α [11,12,13]. Both CDKN1A and GADD45A are referred to as atrogenes and are required for ATF4-mediated muscle atrophy [11,12,13,18], and TRIB3 is another ATF4 target gene involved in fasting- and ageing-induced muscle atrophy [19,20]. We showed here that overexpression of ATF4-regulated atrogenes was dissociated from muscle wasting (1) in a basal condition, (2) during hindlimb suspension, and (3) in a natural model of muscle-atrophy resistance (Figure 7).

We first observed that the overexpression of the ATF4-regulated atrogenes Trib3, Cdkn1a Gadd45a, and Eif4ebp1, as well as Atf4 itself, induced by halofuginone treatment for up to 4 weeks, did not coincide with atrophy in all hindlimb muscles, including gastrocnemius. Subsequently, we then reported that pre-treatment with halofuginone mitigated the atrophy of the gastrocnemius muscle during hindlimb suspension. These positive effects of halofuginone treatment are consistent with reports showing that this dose (i.e., 0.25 µg/g) and frequency of administration (i) were very well-tolerated in mice for up to 3 months and (ii) improved muscle-cell survival, promoted membrane repair, and improved muscle performances in models of muscular dystrophies [31,33,58,59,60]. However, none of these studies explored whether the potential effect of HF would involve the ATF4 pathway. Here, we showed that induction of ATF4-regulated atrogenes was uncoupled from muscle atrophy during hindlimb suspension. Indeed, although ATF4-regulated atrogenes were overexpressed during hindlimb suspension, halofuginone-treated mice displayed a partial preservation of gastrocnemius muscle mass and CSA. In addition, we took advantage of a natural model of resistance to muscle atrophy to examine the expression of ATF4-regulated atrogenes. The brown bear remains completely inactive during hibernation for up to 6 months but, surprisingly, is not sensitive to muscle atrophy [52,53,54], which provides an interesting model for finding new molecular mechanisms to fight muscle atrophy in humans. We showed that the atrogenes CDKN1A, GADD45A, ATF4 itself, and TRIB3 were upregulated in atrophy-resistant hibernating brown bear muscle compared to active bear muscle. Of note, ATF4 is mainly regulated at the translational level [15]. However, in most of the previous studies on the topic, the authors only measured Atf4 mRNA expression and expression of its target genes as evidence of its activity in skeletal muscle. Indeed, endogenous ATF4 protein cannot be reliably detected in skeletal muscle, presumably due to its low abundance, very short half-life, and lack of a high quality antibody [11,12,13,17,18,61]. Altogether, these data strongly suggest that the induction of the ATF4 pathway can be dissociated from muscle atrophy. ATF4 target genes are highly dependent on the type and duration of stress stimuli [21,62], and the ability to restore homeostasis may be overwhelmed when the stress is too severe or sustained, resulting in cell death through the transcription of pro-apoptotic genes [63,64,65,66]. Therefore, to avoid chronic and acute activation, halofuginone was administrated periodically to activate the eIF2α-ATF4 pathway in mice. In our conditions, the ATF4 transcriptional program may, thus, (i) differ from the transcriptional program induced by a severe and sustained activation and (ii) include genes that might counteract the effect of ATF4-induced atrogenes.

Halofuginone is well-described to also target the TGF-β pathway [35,36]. The nuclear translocation of the TGF-β transcription factors SMAD2/3 requires the formation of a complex with SMAD4 [43]. Halofuginone-induced eIF2α phosphorylation has been reported to inhibit the nuclear translocation of this complex in intestinal porcine enterocyte cells in vitro [67]. Consistently, we reported here a concomitant overall (i) increase in phosphorylated eIF2α protein levels and (ii) reduction in SMAD2/3 and SMAD4 nuclear protein levels in HF-treated mice. Thus, this highlights in skeletal muscle, for the first time, the possible role of HF-induced eIF2α phosphorylation in TGF-β inhibition. Although, the concomitant collagen downregulation and decrease in SMAD2/3 nuclear protein levels during hindlimb suspension in H₂O-treated mice suggest a decrease in TGF-β signalling, we cannot exclude that these events are disconnected. Indeed, the SMAD2/3 nuclear protein levels are consistently low in HF-treated mice and, thus, cannot explain the decreased collagen expression during hindlimb suspension. Much remains to be clarified about the mechanisms of action of HF. Indeed, HF is used for its antifibrotic properties mediated by TGF-β inhibition in situations already characterised by fibrosis [48]. This is, however, not the case in our study. In addition, the inhibition of TGF-β signalling in muscles of HF-treated mice could have led to transcriptional changes that remain to be explored. It is possible that TGF-β signalling is induced later during hindlimb suspension. In fact, the TGF-β signalling pathway has previously been reported to be either unchanged in skeletal muscle after 1–3 days or induced after 7–10 days of unloading [46,68,69]. We also reported an upregulation of Trim63 and Fbxo32 during hindlimb suspension. Although these atrogenes are targets of the TGF-β signalling activation, they are also regulated by other signalling pathways [5].

We and others reported that the balance between TGF-β and BMP signalling seems crucial for muscle-mass maintenance during catabolic situations [39,44,45,46,70]. Indeed, using the hibernating bear model, we recently reported that TGF-β signalling, i.e., a negative regulator of muscle mass, was downregulated at the transcriptomic level in muscles that are resistant to atrophy, while BMP signalling, i.e., a positive regulator of muscle mass, was maintained [46]. Previous data suggested that TGF-β inhibition would release SMAD4, i.e., the common actor in TGF-β and BMP signalling, which could, thus, be recruited to BMP signalling and promote hypertrophy and/or counteract atrophy [45]. Here, we reported an increase in the SMAD1/5 nuclear protein levels in the halofuginone-treated control mice, suggesting there was concomitant BMP signalling activation and TGF-β inhibition. In addition, BMP activation was reported to increase during denervation, intensive care disuse, and amyotrophic lateral sclerosis and was described as essential to counteract excessive muscle wasting [44,45]. In agreement, we reported here that BMP transcription factors SMAD1/5 accumulated in the nucleus in H₂O-treated mice but, surprisingly, declined in HF-treated mice after 7 days of hindlimb suspension. Whether the higher basal pools of nuclear SMAD1/5 and their maintenance after 3 days of hindlimb suspension in HF-treated mice contributed to attenuating skeletal muscle atrophy during hindlimb suspension remains to be explored. Of note, BMP signalling has been reported to promote protein synthesis in muscle [44]. Maintenance of the BMP pathway after 3 days of hindlimb suspension may have contributed to the partial preservation of protein synthesis and muscle mass in HF-treated mice. Nuclear translocation of SMAD1/5 represses the transcription of FBXO30/Musa1 [44,45]. However, we did not observe any change in Fbxo30 expression. Mechanisms by which BMP signalling controls muscle mass are still very poorly understood and will require further studies, particularly with a comprehensive characterisation of the BMP target genes in skeletal muscle. We can also speculate that HF-induced BMP activation has helped to limit muscle atrophy induced by ATF4-regulated atrogenes (Figure 7).

In conclusion, halofuginone treatment reproduced the muscle features of hibernating bears in gastrocnemius mice muscles with (i) the activation of ATF4-regulated atrogenes and (ii) the concurrent inhibition of TGF-β signalling and promotion of BMP signalling, without resulting in muscle atrophy (Figure 7). These characteristics were associated with mitigated muscle atrophy during physical inactivity. To date, clinical trials have all attempted to inhibit the TGF-β pathway, mostly with side effects or minimal efficiency [71]. Our study suggests halofuginone, as a well-tolerated chemical compound, already used in human clinical trials [36], was able to tune the TGF-β/BMP balance in vivo and likely sustained muscle mass. Moreover, our data open new ways to further decipher by which precise mechanisms ATF4 induces atrophy and how BMP activation can interfere.

4. Materials and Methods

Ethics, animals housing, and experimental design. All experiments were conducted with the approval of the regional ethics committee (agreement no. D6334515) following the European Directive 2010/63/EU on the protection of vertebrate animals used for experimental and scientific purposes. This study was performed with 12-week-old C57BL6/J male mice (25–30 g), purchased from Janvier Labs (Le Genest-Saint-Isle, France). They were housed individually upon arrival for 10 days of acclimatisation in a controlled room (22 ± 2 °C, 60 ± 5% humidity, 12 h light/dark cycle, and light period starting at 8 h), fed ad libitum a standard rodent diet (pellets A03 from Safe, Augy, France), and given free access to water. Two distinct animal experiments were performed. To evaluate the effects of a periodic halofuginone (HF) (#32481, Sigma, Saint-Quentin-Fallavier, France) administration, we performed a first protocol where mice received either HF (0.25 µg/g) or water (H₂O) by gavage 3 times a week for 1 to 4 weeks (n = 6 animals per group). This dose was reported as well tolerated over longer periods [31,33,60]. Gastrocnemius muscle was sampled 6 h after the last HF/H₂O administration at the end of each week. Subsequently, we performed a second protocol to test whether HF administration before hindlimb unloading had a positive effect on muscle mass and function. For that purpose, we performed two separate animal experiments. In each experiment, mice received either HF (0.25 µg/g) or H₂O by gavage 3 times a week for 3 weeks and were afterwards subjected either to hindlimb unloading through tail suspension (HS) or kept unsuspended (Ctrl) for 3 or 7 days, as previously described [46] (n = 8–19 animals per group). We did not record any difference between Ctrl mice at 3 or 7 days for all the measurements reported in this manuscript. We, therefore, pooled the two groups of Ctrl mice for further analysis and data representation. Food intake and body weight were recorded throughout the different protocols. Unloading in control mice resulted in only a small body weight loss (<10%) that occurred within the first 3 days concomitantly with a decrease in food intake, whereas HF treatment did not modify food intake or body weight ( see Supplementary Figures S1 and S2).

Tissue collection. At the end of the experiments, mice were euthanised by cervical dislocation. The soleus, gastrocnemius, tibialis anterior, and extensor digitorum longus (EDL) muscles were carefully collected and weighed prior to immediate freezing in liquid nitrogen and storage at −80 °C until analyses.

Measurement of protein synthesis in gastrocnemius. At the end of protocol 2, mice received an intraperitoneal injection of 0.040 μmol/g puromycin (#P8833, Sigma, Saint-Quentin-Fallavier, France) dissolved in 100 μL of a saline solution before euthanasia, as described previously [72]. At exactly 30 min post-puromycin injection, gastrocnemius muscle was dissected and frozen in liquid nitrogen for Western blot analysis, as follows.

Histology and morphometric measurements. A part of the gastrocnemius muscle was collected at the end of protocol 2 and frozen in isopentane chilled with liquid nitrogen and stored at −80 °C until use. Serial muscle cross-sections (10 µm thick) were obtained using a cryostat (HM500M Microm International, Fisher Scientific, Illkirch, France) at −20 °C. Cross-sections were labelled with anti-laminin-α1 (L9393 Sigma, Saint-Quentin-Fallavier, France) to outline the fibre cross-sectional area (CSA) and BFF3 antibody (#AB_2266724, DSHB, Iowa City, IA, USA) to determine myosin heavy chain type 2B fibre. Both were subsequently hybridised with a corresponding secondary antibody conjugated to Alexa-Fluor (Invitrogen, Cergy-Pontoise, France). Image acquisitions were performed with a high-resolution ORCA-Flash4.0 LT+ Digital CMOS camera coupled to a IX-73 microscope (Olympus, Münster, Germany) and Cell-Sens dimension software (Olympus Soft Imaging Solutions, Münster, Germany). The CSA was determined for 1000–1500 fibres per animal, using ImageJ software 1.53f51 (http://rsb.info.nih.gov/ij/, accessed on 3 April 2018).

Protein isolation. Gastrocnemius muscles were pulverised in liquid nitrogen. (1) For all targets, ~30 mg of the resulting powders were homogenised using a polytron in 1 mL of an ice-cold buffer (10 mM Tris pH 7.5, 150 mM NaCl, 1 mM EDTA, 1 mM EGTA, 1% Triton X-100, and 0.5% Igepal CA630) containing inhibitors of proteases (Protease Inhibitor Cocktail) and phosphatases (1 mM Na3VO3 and 10 mM NaF) (Sigma, Saint-Quentin-Fallavier, France). The homogenates were stirred for 1h at 4 °C and then centrifuged at 10,000× g for 15 min at 4 °C. The resulting supernatants containing total soluble proteins were then stored at −80 °C until use. (2) For SMADs protein level analysis, subcellular fractionation was performed. For that purpose, ~50 mg of gastrocnemius powder samples were homogenised for 1 min on ice using a polytron in 500 μL of ice-cold extraction buffer (10 mM HEPES, pH 7.5, 10 mM MgCl2, 5 mM KCl, 0.1 mM EDTA, pH 8.0, and 0.1% Triton X-100) [73]. The resulting homogenates were subjected to sequential fractionation steps to separate soluble cytosolic and nuclear proteins as described [74]. Pellets containing nuclear proteins were solubilised in nuclear extraction buffer (20 mM HEPES, pH 7.9, 25% glycerol, 500 mM NaCl, 1.5 mM MgCl2, 0.2 mM EDTA, and pH 8.0) [73]. For all protein extracts, protein concentration was determined using the Bradford Protein Assay Kit (Biorad, Marnes-la-Coquette, France). Proteins were then diluted in Laemmli buffer and stored at −80 °C until use.

Western blots. Protein contents for (i) SMAD family members (anti-SMAD1-5, PA5-80036, Thermofisher, Illkirch, France; anti-SMAD2-3, #8685, Cell Signalling Technology, Saint-Cyr-L’Ecole, France; anti-SMAD4, ab230815, Abcam, Cambridge, UK), (ii) total and phosphorylated eukaryotic initiation factor 2 alpha (anti-eIF2α, #9722, Cell Signalling Technology; anti-p-Ser51eIF2α, ab32157, and Abcam), and (iii) incorporation of puromycin (anti-puromycin clone 12D10, MABE343, Millipore, Burlington, MA, USA) were assessed by immunoblotting. Briefly, 20–40 µg of protein extracts were subjected to SDS-PAGE (sodium dodecyl sulfate-polyacrylamide gel electrophoresis) using TGX™ FastCast™ 10% Acrylamide gels (Biorad, Marnes-la-Coquette, France) and transferred onto a PVDF membrane (Hybond P, Amersham, England) using Trans-Blot^® Turbo™ Transfer System standard protocol (Biorad, Marnes-la-Coquette, France). Western blots were blocked for 1 h at room temperature in TBS (Tris-Buffered Saline) buffer with 0.1% Tween-20 (TBS-T, pH = 7.8) with 5% bovine serum albumin (BSA) for all the targets, in accordance with the instructions of the manufacturer. They were then washed thrice in TBS-T and incubated (overnight, stirring, 4 °C) with appropriate primary antibodies diluted at 1:1000, except for anti-puromycin diluted at 1:5000. Western blots were then washed and incubated for 1 h with an appropriate secondary antibody (HRP-conjugated anti-rabbit (#7074) or anti-mouse (#7076) IgGs) (Cell Signalling Technology, Saint-Cyr-L’Ecole, France). For anti-puromycin antibody, an anti-mouse IgG2Ak (115-035-206, Jackson ImmunoResearch Laboratories, West Grove, PA, USA) was used. Signals were detected after incubation with Luminata Crescendo Western HRP substrate (Millipore, Burlington, MA, USA) and visualised using G: BOX ChemiXT4 (XL1) imaging system (Syngene, Frederick, MD, USA). Signals were then quantified using ImageJ 1.53f51 software. Two samples from each group were loaded on each gel. The signal recorded within each lane of one Western blot was normalised to the overall signal of that blot, and then signals were normalised to the total amount of proteins determined by the Biorad’s stain-free system or ponceau S to correct for uneven loading. The normalised values were then averaged by group and expressed as the fold change from the mean of all H₂O-ctrl samples.

RT-qPCR. Total RNA from gastrocnemius muscle samples was extracted with Macherey-Nagel™ NucleoSpin™ 96 RNA Kit and KingFisher™ Duo Prime Purification System, in accordance with the instructions of the manufacturer (Macherey-Nagel, Hoerdt Cedex, France). RNA was quantified by measuring the absorbance at 260 nm on a NanoDrop ND-1000 spectrophotometer (Thermo Scientific, Wilmington, DE, USA). Then, 500 ng of RNA were treated with DNase I (Invitrogen, Cergy-Pontoise, France) prior to reverse transcription using random primers and SuperScript II (Invitrogen, Cergy-Pontoise, France), in accordance with the instructions of the manufacturer. Real-time PCR was carried out using the CFX96 Real-Time PCR detection system (Biorad, Marnes-la-Coquette, France). Primer sequences are provided in Supplementary Table S1. PCR reactions were performed using the IQ SYBR Green Supermix (Biorad, Marnes-la-Coquette, France), in accordance with the instructions of the manufacturer. The comparative threshold cycle (2ΔΔCT) method was used to compare the relative mRNA expression between each group, using TBP (TATA binding protein) as a reference gene for muscle. The relative mRNA abundance was arbitrarily set to 1 for the H₂O-Ctrl group.

Statistics. All data are means ± SEM and were analysed for normality of residuals using the Shapiro-Wilk test. No set of data was transformed for non-normality distribution. For protocol 1 (n = 6/group), we performed a multiple Welch t-test within each week. For protocol 2 (n = 8–19/group), we performed a two-way ANOVA with the factors “Hindlimb suspension” and “Halofuginone” and corrected the data for multiple comparisons using Tukey’s test. These analyses were performed using Prism 9 (GraphPad Prism 9, San Diego, CA, USA).

Transcriptomic Data. We used transcriptomic data from already published studies [46]. The transcriptomic bear data supporting Figure 6 of this study are openly available in the GEO repository database (https://www.ncbi.nlm.nih.gov/geo/query/acc.cgi, reference no. GSE144856, accessed on 1 September 2021). To identify the differentially expressed genes (DEGs) from this list, we selected a winter/summer (FC) > 1.0 with an adjusted p-value < 0.05 as cut-off for the up-regulated genes.