# Muscle mTOR controls iron homeostasis and ferritinophagy via NRF2, HIFs and AKT/PKB signaling pathways

Agnès Conjard-Duplany, Alexis Osseni, Aline Lamboux, Sandrine Mouradian, Flavien Picard, Vincent Moncollin, Céline Angleraux, Tiphaine Dorel-Dubois, Hélène Puccio, Pascal Leblanc, et al.

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## To cite this version:

Agnès Conjard-Duplany, Alexis Osseni, Aline Lamboux, Sandrine Mouradian, Flavien Picard, et al. Muscle mTOR controls iron homeostasis and ferritinophagy via NRF2, HIFs and AKT/PKB signaling pathways. Cellular and Molecular Life Sciences, 2025, 82, pp.178. 10.1007/s00018-025-05695-9. hal-05347328

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# Muscle mTOR controls iron homeostasis and ferritinophagy via NRF2, HIFs and AKT/PKB signaling pathways

Agnès Conjard-Duplany¹ Ⓓ · Alexis Osseni¹ Ⓓ · Aline Lamboux² Ⓓ · Sandrine Mouradian¹ Ⓓ · Flavien Picard¹ Ⓓ · Vincent Moncollin¹ Ⓓ · Céline Angleraux³ · Tiphaine Dorel-Dubois³ · Hélène Puccio¹ Ⓓ · Pascal Leblanc¹ Ⓓ · Bruno Galy⁴,⁵ Ⓓ · Vincent Balter² Ⓓ · Laurent Schaeffer¹,⁶ Ⓓ · Yann-Gaël Gangloff¹ Ⓓ

Received: 2 September 2024 / Revised: 28 March 2025 / Accepted: 3 April 2025
© The Author(s) 2025

## Abstract

Balanced mTOR activity and iron levels are crucial for muscle integrity, with evidence suggesting mTOR regulates cellular iron homeostasis. In this study, we investigated iron metabolism in muscle-specific mTOR knockout mice (mTORmKO) and its relation to their myopathy. The mTORmKO mice exhibited distinct iron content patterns across muscle types and ages. Slow-twitch *soleus* muscles initially showed reduced iron levels in young mice, which increased with the dystrophy progression but remained within control ranges. In contrast, the less affected fast-twitch muscles maintained near-normal iron levels from a young age. Interestingly, both mTORmKO muscle types exhibited iron metabolism markers indicative of iron excess, including decreased transferrin receptor 1 (TFR1) and increased levels of ferritin (FTL) and ferroportin (FPN) proteins. Paradoxically, these changes were accompanied by downregulated *Ftl* and *Fpn* mRNA levels, indicating post-transcriptional regulation. This discordant regulation resulted from disruption of key iron metabolism pathways, including NRF2/NFE2L2, HIFs, and AKT/PKB signaling. Mechanistically, mTOR deficiency impaired transcriptional regulation of iron-related genes mediated by NRF2 and HIFs. Furthermore, it triggered ferritin accumulation through two NRF2 mechanisms: (1) derepression of ferritin translation via suppression of the FBXL5-IRP axis, and (2) autophagosomal sequestration driven by NCOA4-dependent ferritin targeting to autophagosomes, coupled with age-related impairments of autophagy linked to chronic AKT/PKB activation. Three-week spermidine supplementation in older mTORmKO mice was associated with normalized AKT/PKB-FOXO signaling, increased endolysosomal FTL and reduced total FTL levels in the dystrophic *soleus* muscle. These findings underscore mTOR's crucial role in skeletal muscle iron metabolism and suggest spermidine as a potential strategy to address impaired ferritinophagy due to autophagy blockade in dystrophic muscle.

**Keywords** Dystrophy · Autophagy · Iron-sulfur cluster · Oxidative stress · Myoglobin · Glycogen

Agnès Conjard-Duplany and Alexis Osseni contributed equally to this paper.

☒ Agnès Conjard-Duplany
agnes.duplany@univ-lyon1.fr

☒ Yann-Gaël Gangloff
yann-gael.gangloff@univ-lyon1.fr

¹ Laboratoire Physiopathologie et Génétique du Neurone et du Muscle (PGNM), Institut NeuroMyoGène, Université Claude Bernard Lyon 1, INSERM U1315, CNRS UMR 5261, 8 avenue Rockefeller, Lyon 69008, France

² Laboratoire de Géologie de Lyon: Terre, Planètes, Environnement, UMR 5276, Ecole Normale Supérieure de Lyon, 46, allée d'Italie, Lyon, Cedex 07 69364, France

³ Université Claude Bernard Lyon 1, CNRS UAR3444, Inserm US8, ENS de Lyon, AniRA-PBES, SFR Biosciences, Lyon 69007, France

⁴ German Cancer Research Center (DKFZ), Division of Virus-associated Carcinogenesis (F170), Heidelberg, Germany

⁵ IB-Cancer Research Foundation, Science Park 2, 66123 Saarbrücken, Germany

⁶ Centre de Biotechnologie Cellulaire, Hospices Civils de Lyon, Lyon, France
Introduction

Approximately 10 to 15% of total body iron (Fe) is present in skeletal muscle. The metal is distributed among various ferroproteins, including myoglobin, which is responsible for oxygen transport, and enzymes involved in essential physiological processes, such as the maintenance of mitochondrial function. Notably, iron assumes pivotal roles as iron-sulfur clusters (ISC), contributing to electron transfer, enzymatic catalysis and protein stabilization [1]. Conversely, surplus iron is stored within the protein ferritin, serving as a reserve.

Both iron excess and deficiency have deleterious effects on muscle function. Accumulation of iron in muscles is associated with oxidative stress, mitochondrial dysfunction, insulin resistance, impaired regeneration and muscle wasting [2]. Muscle iron overload has been documented in conditions such as aging [3] and immobilization [4], as well as in rodent models of amyotrophic lateral sclerosis (ALS) [5] and Duchenne Muscular Dystrophy (DMD) [6]. Likewise, muscle iron deficiency can compromise muscle integrity in various chronic diseases [7, 8].

Cellular iron homeostasis is controlled at both the transcriptional and post-transcriptional levels, involving various proteins responsible for iron uptake (transferrin receptor/TFR1), storage (ferritin-L/FTL and ferritin-H/ FTH), and export (ferroportin/FPN) [9]. At the transcriptional level, cellular iron metabolism is predominantly regulated by Nuclear Factor Erythroid 2-Related Factor 2 (NRF2/NFE2L2) [10] and Hypoxia-Inducible Factors (HIF1α/HIF2α) [11]. NRF2 directly increases the transcription of the ferritin subunits FTL and FTH, as well as FPN/SLC40A1 [10]. HIF1α specifically enhances the transcription of TFR1 [12], while HIF2α is involved in regulating the transcription of both Divalent Metal Transporter 1 (DMT1/SLC11A2) [13] and FPN [14]. At the post-transcriptional level, iron homeostasis is tightly regulated by the iron regulatory proteins IRP1 (ACO1) and IRP2 (IREB2), which play a critical role in maintaining cellular iron balance. In iron-depleted cells, IRPs bind to cis-regulatory iron-responsive elements (IREs) located in the untranslated regions (UTRs) of target mRNAs, blocking their translation (e.g., Ftl, Fth and Fpn) or increasing their stability (e.g., Tfr1) [9]. In iron-replete cells, IRP1 incorporates an ISC that converts it to a cytosolic aconitase, whereas IRP2 undergoes degradation mediated by the E3 ubiquitin ligase F-box and leucine rich repeat protein 5 (FBXL5) [15]. FBXL5 also targets apo-IRP1 for degradation. Another level of regulation involves ferritinophagy, a form of selective autophagy that is triggered in response to iron limitation, enabling the release of iron from ferritin for intracellular utilization. Ferritinophagy is mediated by nuclear receptor coactivator 4 (NCOA4),

a cargo receptor recruiting ferritin to autophagosomes [16]. Interestingly, HIF2α indirectly promotes ferritin degradation and iron mobilization by stimulating NCOA4 transcription [17]. Additionally, NRF2 orchestrates both ferritin synthesis and turnover by increasing the transcription of HERC2 (HECT and RLD domain containing E3 ubiquitin protein ligase 2) that controls the FBXL5/ IRP regulatory axis and NCOA4-mediated ferritinophagy [18].

Research conducted on cell lines has uncovered the critical role of mechanistic target of rapamycin (mTOR), a Ser/Thr protein kinase forming the mTORC1 and mTORC2 complexes, in cellular iron homeostasis. It has been shown that mTORC1 inhibition with rapamycin decreases TFR1 levels by increasing the expression of tristetraprolin (TTP/ZFP36), which binds to and enhances degradation of Tfr1 mRNA. The downregulation of TFR1 initially causes iron deficiency, thereby activating the IRP system, which in turn represses FPN synthesis, ultimately resulting in intracellular iron accumulation [19]. Additionally, mTORC2 has been implicated as an epigenetic regulator of iron metabolism genes [20]. Moreover, mTORC1 may promote ISC biogenesis by phosphorylating and stabilizing the mitochondrial iron-sulfur cluster assembly enzyme ISCU [21], whose muscle deficiency leads to mitochondrial myopathy [22]. mTOR signaling has also been linked to transcriptional regulators of iron homeostasis. For instance, mTORC1 was shown to promote NRF2 activation during autophagy [23], whereas mTORC2 could increase NRF2 abundance and enhance its activity [24]. Furthermore, both mTORC1 and mTORC2 are involved in controlling HIF1α and/or HIF2α synthesis and turnover [25]. However, despite indications of mTOR's involvement in iron homeostasis, no obvious alterations in iron balance and iron gene expression were observed in the hearts of mice with selective ablation of mTOR in cardiomyocytes [26]. Further studies are thus needed to elucidate the complex interplay between mTOR signaling and iron homeostasis in various physiological and pathological contexts, notably in skeletal muscle.

Our previous research has shown that targeted mTOR deletion in mouse skeletal muscle leads to progressive myopathy, characterized by reduced myofiber size and muscle weakness, resulting in premature death between 22 and 38 weeks of age due to respiratory failure [27]. Slow-oxidative muscles exhibit pronounced decreases in mitochondrial biogenesis, impaired oxidative metabolism, myoglobin downregulation, all linked to PGC-1α defects, and a marked progression of degenerative changes. Fast-glycolytic fibers display prominent glycogen accumulation, resulting from enhanced glycogen synthesis coupled with reduced glycogen breakdown, and mild dystrophic features. Furthermore,
muscle mTORC1-deficient mice exhibit sustained autoph-
agy induction, yet concurrently experience an age-related
decline in autophagy flux [28, 29].

In this study, we sought to investigate the impact of mTOR deficiency on iron homeostasis in mouse skeletal muscle and explore its potential contribution to the progression of mTOR-associated myopathy.

**Materials and methods**

Animals

The generation of animals with muscle-specific mTOR inactivation (HSA-Cre⁺; *mTOR* flox/flox herein called mTORmKO) on F5; C57BL/6J background has been previously described in [27]. For this study, the animals were backcrossed on a C57BL/6J background for more than 10 generations. *mTOR* flox/flox mice were used as controls. The animals were provided with mouse chow and water ad libitum under a light–dark cycle (12 h), in a restricted-access, specific pathogen–free animal care facility at AniRA PBES, Lyon, France.

Spermidine administration

*Per os* treatment was performed by adding spermidine (Sigma-Aldrich, S0266) to the drinking water at a concentration of 3 mM as described in [30]. Mice had free access to water for 21 days and were then sacrificed. Water with spermidine was replaced every 2 days, in order to avoid excessive deamination of the active compound. Non-treated mice drank water without addition of spermidine.

Immunoblotting

Muscles were dissected on euthanized mice, snap-frozen in
liquid nitrogen and stored at -80 °C until use. Frozen tissues
were crushed with beads in a homogenizer system (Precel-
lys Evolution, Bertin) in RIPA buffer containing 50 mM
tris-HCl (pH 7.5), 150 mM NaCl, 1% triton X-100, 1mM
EDTA, 1 mM dithiothreitol, 0.5% deoxycholate, 0.1%
SDS, with protease and phosphatase inhibitors from Roche.
Lysates were then centrifuged at 13,000 g for 15 min. Pro-
tein concentration was determined using the Pierce BCA
Protein assay. Equal amounts of proteins were subjected
to Western blot analysis. After electrophoresis, transfer to
PVDF membranes and incubation with specific antibodies,
the proteins were visualized using the ChemiDoc Touch
Imaging System (Bio-Rad). Densitometry was performed
using Image Lab v.6.1.0 software (Bio-Rad). Antibodies are
listed in Table S1.

**Transcript expression analyses**

Total RNA from frozen tissues was prepared using TRIzol (TRI-Reagent, Sigma) or a miRNeasy Mini Kit according to the manufacturer's instructions (Qiagen). Complementary DNA was generated using RevertAid H minus Reverse transcriptase (Fermentas) and random hexamer primers. Quantitative PCR was carried out using ONE Green FAST qPCR premix (Ozyme) and was run using a CFX96 Real-Time PCR Detection System (Bio-Rad, Hercules, CA, USA). Relative RNA expression was normalized to *cyclophilin B* and *β-Actin* expression. Delta Delta Ct (threshold cycle) analysis was used to calculate relative gene expression. The results were plotted in arbitrary units as mean±SEM. The primer sequences are listed in Table S2.

Iron assay

A colorimetric iron assay kit (Mak025, Sigma) was used to measure ferrous [Fe(II)] and ferric [Fe(III)] iron. The assay was performed on snap-frozen muscle tissues according to the manufacturer's instructions. Iron standard was used for calibration.

Carbonylation

Protein carbonyls content in muscle samples was determined after derivatization with the probe NBDH (Carbofax®, Yelen Analytics, France). Concentration of NBDH-reacted carbonyls was determined by fluorescence. Reduced and oxidized bovine serum albumin were used for calibration.

Lipid peroxidation

Malondialdehyde (MDA) content in muscle samples was determined by fluorescence using Lipid Peroxidation Assay Kit (ab118970, Abcam) following the manufacturer's instructions.

RNA electrophoretic mobility shift assay (REMSA)

Frozen muscle samples were homogenized in ice-cold REMSA Binding Buffer (10 mM Hepes pH 7.5, 40 mM KCl, 3 mM MgCl2, 0.2% NP-40, 5% glycerol, Rnasin® 0.4U/µl and 1× protease inhibitors). Lysates were incubated for 30 min and debris were removed by repeated centrifugation at 10,000 g for 10 min at 4 °C. Protein concentration in the supernatant was determined using the Pierce BCA Protein assay. The IRP-IRE interactions were performed for 20 min at room temperature in REMSA Binding Buffer using 100 µg of muscle extracts and 0.05 µM Alexa Fluor® 488 labeled ferritin-H IRE probe (5'-UCCUGCUUCAAC
AGUGCUUGAACGGAAC-3'), and where indicated, 2% 2-mercaptoethanol (2-ME) was added. Afterward, heparin (final concentration 5 mg/ml) was added and incubated for 10 min. The IRP-IRE complexes were visualized by fluorescence on a ChemiDoc Touch Imaging System with Blue Epi Illumination (Bio-Rad).

## Muscle histology, immunohistochemistry, and imaging

Skeletal muscles were collected, embedded in tragacanth gum, and quickly frozen in isopentane cooled in liquid nitrogen. 10-µm-thick sections were obtained from the middle portion of frozen muscles from three animals per group and processed for histological analysis according to standard protocols. Sections were stained with haematoxylin, eosin, and saffron (HES) to assess general muscle architecture; Perls' Prussian blue stain, with a cytoplasmic counterstain, to detect ferric iron deposition; and periodic acid-Schiff (PAS) to identify glycogen deposition. Images were captured on a Zeiss AxioScan Z1 and 7 (Carl Zeiss) equipped with 20X or 40X objectives. The fiber CSA and the number of centrally nucleated fibers were determined after immunofluorescence staining of Laminin and Dapi on sections using FIJI software (ImageJ 2.14./1.54f, National Institutes of Health, USA) and for CSA measurement with the help of Open-CSAM, an ImageJ macro [31].

## Immunostaining on isolated single fibers

Single muscle fibers were isolated from soleus and tibialis anterior (TA) muscles as described previously [32] and fixed 30 min at room temperature (RT) in PBS-4% formaldehyde, permeabilized 30 min in PBS-1% Triton X-100 at 30 °C before saturation in PBS-0.1% Tween 20 supplemented with 0,5% BSA and 2% goat serum at RT for 6 h. Incubations with primary antibodies in PBS-0.1% Tween 20 were performed at 4 °C overnight and washed in PBS-0.1% Tween 20. After incubation in PBS-0.1% Tween 20 for 24 h at 4 °C with fluorescent secondary antibodies, dissociated fibers were washed in PBS-0.1% Tween 20. Nuclear DNA were stained with DAPI for 10 min. Coverslips were mounted on microscope slides with FluorSave™ reagent (Calbiochem). Images were captured at RT on a Zeiss LSM980 microscope (Carl Zeiss) with an AiryScan2 detector equipped with a 63X/1.4-NA objective. All images were processed with either the ZEN blue software, Photoshop CS5 (Adobe Systems, San Jose, CA, USA) or FIJI software (ImageJ 2.0.0-rc-69/1.52n, National Institutes of Health, Bethesda, MD).

Colocalization between p62 or LAMP1 (green channel) and FTL (red channel) was quantified using a binary mask

method as described by [32]. This mask represented the overlapping surface area of p62 or LAMP1 with FTL on a single confocal plane. For each imaged field, the mean gray value of dissociated fibers within this mask was measured. Measurements from the control (CTL) condition were normalized to 1. To quantify FOXO3 nuclear localization, the mean gray value of the green channel (FOXO3) was measured within each nucleus. Measurements from the control (CTL) condition were normalized to 1.

## Minor and major element quantification

Lyophilized muscle were weighed on a micro-balance and digested at 120 °C overnight in HNO₃ (14 M). Cooled digests were dissolved in H₂O₂ (5%, final), evaporated to dryness at 120 °C and further dissolved in HNO₃ (0.5 M). Minor elements (Fe, Cu, Mn, Zn) were measured using inductively coupled plasma mass spectrometry (ICP-MS) (iCAP-Q, Thermo Scientific, Bremen, Germany) with In at 2 µg.L⁻¹ as internal standard for correction of the instrumental drift. The major element (P) was measured using inductively coupled plasma atomic emission spectroscopy (ICP-AES) (iCAP-PRO, Thermo Scientific, Bremen, Germany) with Sc at 1 mg.L⁻¹ as internal standard for correction of the instrumental drift. SPC33 MS multi-element and single-element standard solutions (SCP science, Courtaboeuf, France) were used for the quantification of minor and major elements respectively.

## Statistical analysis

Data were analyzed with GraphPad Prism version 8.4.3 for windows (GraphPad Software, San Diego, California USA). Independent samples t tests were used for comparisons between two groups. For comparisons of three groups, a one-way analysis of variance (ANOVA) was used with Tukey's post hoc multiple comparison test. For comparisons of four groups with two factors (genotype and treatment or genotype and gender), a two-way analysis of variance (ANOVA) was used with Bonferroni's post hoc multiple comparison test. The threshold for significance was established at P<0.05 for all comparisons.

## Results

### Loss of muscle mTOR alters iron levels in slow-oxidative muscle

For this study, mTORmKO mice were bred on a congenic C57BL/6J background. These mice exhibit a slower
progression of myopathy compared to the previously described F5; C57BL/6J background [27], with first mortality occurring at 34 weeks. Increased robustness of myopathic mice on a congenic C57BL/6J background has been previously documented [33–35].

To evaluate the impact of mTOR loss on muscle iron homeostasis, we measured iron content in the *soleus* (enriched in slow-oxidative fibers) and *plantaris* (PLA; enriched in fast-glycolytic fibers) muscles of 7- and 25-week-old (wk) mice. In 7-wk-old male mTORmKO mice, representing an early stage of myopathy, iron content in the *soleus* muscle was significantly reduced by

<table><caption>**Table 1** Metal imbalances in skeletal muscle of 7-wk-old mTORmKO male mice. Quantification of metals in *soleus* and *plantaris* muscles from 7-wk-old mTORmKO and control male mice by ICP-MS and ICP-AES. Metals were normalized to total dry weight of tissue, and to phosphorus (P). control *soleus*: N = 12, mTORmKO *soleus*: N = 8, control *plantaris*: N = 12, mTORmKO *plantaris*: N = 8. N represents the number of individual mice used in each group. Values represent the mean ± SEM. \*P ≤ 0.05; \*\*P ≤ 0.01; \*\*\*P ≤ 0.001; ns, not significant</caption><thead><tr><th>MALES</th><th>Metal</th><th>Control</th><th>mTORmKO</th><th>% of Control</th><th>p-Value</th></tr></thead><tbody><tr><td rowspan="10">SOLEUS</td><td>Fe<br>(µg/g)</td><td>121.80±3.7</td><td>87.0±3.5</td><td>72</td><td>###</td></tr><tr><td>Fe/P<br>(*10<sup>3</sup>)</td><td>15.0±0.5</td><td>11.7±0.4</td><td>78</td><td>###</td></tr><tr><td>Cu<br>(µg/g)</td><td>8.6±0.1</td><td>6.0±0.3</td><td>70</td><td>###</td></tr><tr><td>Cu/P<br>(*10<sup>4</sup>)</td><td>10.5±0.1</td><td>8.1±0.2</td><td>77</td><td>###</td></tr><tr><td>Mn<br>(µg/g)</td><td>1.0±0.1</td><td>0.7±0.0</td><td>64</td><td>###</td></tr><tr><td>Mn/P<br>(*10<sup>5</sup>)</td><td>12.8±0.8</td><td>9.1±0.4</td><td>71</td><td>##</td></tr><tr><td>Zn<br>(µg/g)</td><td>133.4±2.7</td><td>151.0±5.3</td><td>113</td><td>##</td></tr><tr><td>Zn/P<br>(*10<sup>3</sup>)</td><td>16.4±0.3</td><td>20.4±0.7</td><td>125</td><td>###</td></tr><tr><td>P<br>(µg/g)</td><td>8138.8±90.8</td><td>7413.0±221.5</td><td>91</td><td>##</td></tr><tr><td rowspan="10">PLAN-TARIS</td><td>Fe<br>(µg/g)</td><td>62.8±3.5</td><td>56.6±3.5</td><td>90</td><td>ns</td></tr><tr><td>Fe/P<br>(*10<sup>3</sup>)</td><td>7.2±0.4</td><td>6.1±0.3</td><td>85</td><td>ns</td></tr><tr><td>Cu<br>(µg/g)</td><td>5.1±0.1</td><td>4.5±0.1</td><td>89</td><td>##</td></tr><tr><td>Cu/P<br>(*10<sup>4</sup>)</td><td>5.8±0.1</td><td>4.9±0.2</td><td>84</td><td>###</td></tr><tr><td>Mn<br>(µg/g)</td><td>0.6±0.0</td><td>0.6±0.0</td><td>93</td><td>ns</td></tr><tr><td>Mn/P<br>(*10<sup>5</sup>)</td><td>7.4±0.2</td><td>6.5±0.5</td><td>87</td><td>ns</td></tr><tr><td>Zn<br>(µg/g)</td><td>57.9±1.1</td><td>69.0±2.8</td><td>119</td><td>###</td></tr><tr><td>Zn/P<br>(*10<sup>3</sup>)</td><td>6.7±0.2</td><td>7.5±0.3</td><td>112</td><td>#</td></tr><tr><td>P<br>(µg/g)</td><td>8717.5±101.3</td><td>9275.9±167.8</td><td>106</td><td>##</td></tr></tbody></table>

22–28% compared to controls, while the PLA showed no significant difference from controls (10–15% lower, *p* > 0.05; Table 1). This early iron deficiency in the *soleus* coincided with a reduction in iron-containing myoglobin (Fig. 1A, B). By 25 weeks, *soleus* iron levels in male mTORmKO mice showed no significant difference from controls (10% lower, *p* > 0.05; Table 2), despite persistent myoglobin downregulation. Similarly, PLA iron content remained unchanged compared to controls (10% lower, *p* > 0.05; Table 2). Notably, fast-glycolytic muscles did not exhibit reduced myoglobin content, as demonstrated by the *tibialis anterior* (TA), a mixed muscle with a predominance of fast-glycolytic fibers (Fig. 1A, B). Similar to their male counterparts, 25-wk-old female mTORmKO mice displayed near-normal iron levels in both *soleus* and PLA muscles (Table S3), indicating that mTOR loss similarly affects muscle iron regulation in both male and female mice. Thus, mTOR depletion disrupts iron homeostasis in the *soleus* muscle, with an initial deficiency followed by normalization of iron levels as the myopathy progresses. Analysis of other physiologically important biomaterials revealed increased zinc (Zn) levels, a characteristic of skeletal muscle atrophy models [36], and reduced copper (Cu) and manganese (Mn) levels in both muscle types at both ages (Tables 1 and 2 and S3).

To further investigate iron homeostasis in mTORmKO *soleus* muscles from 25-wk-old mice, we examined the distribution of iron by quantifying ferrous Fe(II) and ferric Fe(III) iron levels. This muscle exhibited an increase in Fe(III) content with a concomitant decrease in Fe(II) compared to controls (Fig. 1C). However, despite the shift towards Fe(III) in mutant *soleus* muscle, Perls’ staining did not reveal localized accumulation of ferric iron, in contrast to the spleen used as a positive control, where Fe(III) deposits, originating from red pulp macrophages, were clearly observed (Fig. 1D). Similarly, PLA and *gastrocnemius* (GC) from 25-wk-old mTORmKO mice showed no sign of localized iron overload.

Fe(II) is the metabolically active form of iron that can promote the production of reactive oxygen species (ROS) via Fenton chemistry and oxidative damage. Reflecting the decrease in Fe(II) levels, the amount of malondialdehyde (MDA) and protein carbonyls, markers of lipid peroxidation and protein oxidation, respectively, were unchanged and lower in mTORmKO *soleus* muscles from 25-wk-old mice compared to controls (Fig. 1E, F). In line with the observations in *soleus* muscles, PLA muscles showed no signs of oxidative damage in the absence of mTOR.

Collectively, these data reveal that mTOR is essential for preserving iron balance in slow-oxidative muscles, but its loss does not induce oxidative damage.
<figure><img src="image_4.png" /><figcaption>**Fig. 1** Impact of muscle mTOR deficiency on iron distribution and oxidative stress. (A) Western blot showing myoglobin (Mb) protein levels in *soleus* from 7- and 25-wk-old male mice and TA muscle from 25-wk-old male mice (N=4). (B) Quantification of the western blot shown in (A) normalized to GAPDH protein level. (C) Concentrations of Fe(II) and Fe(III) in *soleus* muscles from 25-wk-old control and mTORmKO male mice (N=6). (D) Left panels: Representative Perls' staining of *soleus* (SOL), *plantaris* (PLA), and *gastrocnemius* (GC) muscles, along with spleen sections from 25-wk-old control and mTORmKO mice. Bar, 400 µm. Right panels: Higher magnification</figcaption></figure>

view of a region in the left panel. Bar, 100 µm. The staining reveals an absence of detectable iron deposits in mutant muscles, contrasting with the iron-rich red pulp of the spleen (black arrows) (N=3). (E) Malondialdehyde (MDA) concentration in *soleus* and PLA muscles from 25-wk-old control and mTORmKO male mice (N=5). (F) Protein carbonyl concentration in *soleus* and PLA muscles from 25-wk-old control and mTORmKO male mice (N=6). N represents the number of individual mice used in each group. Values represent the mean±SEM.
\#P≤0.05; \*\*P≤0.01; \*\*\*P≤0.001; ns, not significant

mTORmKO mice do not develop ISCU-like myopathy

Reduced levels of Fe(II) in mTORmKO *soleus* muscles
could potentially limit iron availability for ISC (iron-
sulfur cluster) protein synthesis, which are crucial for
mitochondrial respiration [1]. In addition, ISC biogenesis
relies on a set of ISC assembly proteins, including the
scaffold protein ISCU that has been proposed to be sta-
bilized by mTOR [21] and frataxin (FXN). Importantly, a
partial reduction in ISCU levels in mice heterozygous for
ISCU does not impair ISC biogenesis or mitochondrial
function [37], whereas severe ISCU deficiency is required
to induce mitochondrial myopathy [22]. Likewise,

significant FXN downregulation is necessary to trigger
mitochondrial dysfunction and muscle abnormalities [38].
Therefore, we asked whether the mitochondrial dysfunc-
tion in mTORmKO *soleus* muscle [27, 39] could be related
to defects in ISC assembly proteins and ISC biosynthesis.
The *soleus* muscle from 20-23-wk-old mTORmKO male
mice showed a 1.8-fold decrease in ISCU protein expres-
sion (Fig. 2A, B upper panel), associated with a 2.9-fold
reduction in *Iscu* mRNA levels (Fig. 2C). Expression of
FXN was also reduced at both the mRNA and protein lev-
els, by 1.6- and 2.7-fold, respectively. Conversely, protein
level of ISCU and FXN was not significantly reduced in
*soleus* muscle from 7-wk-old mTOR mutant mice (Figure
<table><caption>**Table 2** Metal imbalances in skeletal muscle of 25-wk-old mTORmKO male mice. Quantification of metals in *soleus* and *plantaris* muscles from 25-wk-old mTORmKO and control male mice by ICP-MS and ICP-AES. Metals were normalized to total dry weight of tissue, and to phosphorus (P). control *soleus*: *N* = 8, mTORmKO *soleus*: *N* = 5, control *plantaris*: *N* = 8, mTORmKO *plantaris*: *N* = 5. N represents the number of individual mice used in each group. Values represent the mean ± SEM. \*P ≤ 0.05; \*\*P ≤ 0.01; \*\*\*P ≤ 0.001; ns, not significant</caption><thead><tr><th>MALES</th><th>Metal</th><th>Control</th><th>mTORmKO</th><th>% of Control</th><th>p-Value</th></tr></thead><tbody><tr><td rowspan="12">SOLEUS</td><td>Fe (µg/g)</td><td>195.0±11.1</td><td>169.1±6.4</td><td>87</td><td>ns</td></tr><tr><td>Fe/P (*10<sup>3</sup>)</td><td>25.6±1.0</td><td>23.5±1.3</td><td>92</td><td>ns</td></tr><tr><td>Cu (µg/g)</td><td>8.9±0.5</td><td>4.4±0.5</td><td>49</td><td>###</td></tr><tr><td>Cu/P (*10<sup>4</sup>)</td><td>11.6±0.4</td><td>6.0±0.7</td><td>52</td><td>###</td></tr><tr><td>Mn (µg/g)</td><td>1.1±0.1</td><td>0.6±0.0</td><td>56</td><td>##</td></tr><tr><td>Mn/P (*10<sup>5</sup>)</td><td>13.9±1.2</td><td>8.1±0.4</td><td>59</td><td>##</td></tr><tr><td>Zn (µg/g)</td><td>142.3±6.1</td><td>238.6±19.5</td><td>168</td><td>###</td></tr><tr><td>Zn/P (*10<sup>3</sup>)</td><td>18.7±0.5</td><td>32.9±1.9</td><td>176</td><td>###</td></tr><tr><td>P (µg/g)</td><td>7604.6±272.6</td><td>7241.9±360.6</td><td>95</td><td>ns</td></tr><tr><td rowspan="12">PLAN-TARIS</td><td>Fe (µg/g)</td><td>101.6±5.6</td><td>91.5±5.2</td><td>90</td><td>ns</td></tr><tr><td>Fe/P (*10<sup>3</sup>)</td><td>12.0±0.6</td><td>10.8±0.6</td><td>90</td><td>ns</td></tr><tr><td>Cu (µg/g)</td><td>5.8±0.1</td><td>4.3±0.2</td><td>74</td><td>###</td></tr><tr><td>Cu/P (*10<sup>4</sup>)</td><td>6.9±0.2</td><td>5.1±0.3</td><td>74</td><td>###</td></tr><tr><td>Mn (µg/g)</td><td>0.8±0.0</td><td>0.5±0.0</td><td>71</td><td>##</td></tr><tr><td>Mn/P (*10<sup>5</sup>)</td><td>9.1±0.6</td><td>6.4±0.3</td><td>71</td><td>##</td></tr><tr><td>Zn (µg/g)</td><td>72.7±1.7</td><td>130.7±5.2</td><td>180</td><td>###</td></tr><tr><td>Zn/P (*10<sup>3</sup>)</td><td>8.6±0.2</td><td>15.4±0.5</td><td>179</td><td>###</td></tr><tr><td>P (µg/g)</td><td>8465.4±52.8</td><td>8505.4±128.3</td><td>100</td><td>ns</td></tr></tbody></table>

S1A, B). Downregulation of ISCU and/or FXN in older mTORmKO *soleus* muscle could potentially impair ISC biogenesis. To test this, and on the basis that ISC deficiency generally leads to ISC-apoproteins instability [40], we examined the abundance of several ISC-containing mitochondrial proteins, including NDUFS3, SDHB, ferrochelatase, and ACO2. Except for ACO2, all these proteins were downregulated in mutant muscle (Fig. 2A, B upper panel). However, the corresponding mRNAs were also downregulated (Fig. 2C), precluding definitive conclusion regarding the effect of mTOR deficiency on the stability of these proteins.

To further assess the impact of mTOR deficiency on ISC metabolism, we analyzed the lipoylation status of the E2 subunit of pyruvate dehydrogenase (PDH), a critical ISC-dependent process known to be sensitive to ISC defects [40]. Lipoic acid (LA) biogenesis is catalyzed by LA synthase (LIAS), which requires an ISC for its activity. We observed a 1.4-fold decrease in lipoylated PDH(E2) levels in *soleus* muscle of mTORmKO mice compared with controls. However, this difference was no longer observed after normalization to the total amount of PDH(E2), whose expression was reduced to a similar extent (Fig. 2A, B, lower panel), likely due to transcriptional downregulation of *Pdh(e2)* (Fig. 2C). Supporting the idea that muscle mTOR is not required for ISC biogenesis or integrity, the TA muscle from 20-23-wk-old mTORmKO mice exhibited no downregulation of ISC assembly proteins (ISCU and FXN), ISC-containing mitochondrial proteins, or PDH lipoylation (Figure S2A, B).

Therefore, our data strongly suggest that muscle mTOR is not directly involved in ISC biogenesis or integrity. Instead, the observed mitochondrial changes at the mRNA and protein levels in mutant *soleus* muscle are likely a consequence of mitochondrial remodeling during myopathy progression.

## Loss of mTOR alters iron-related gene expression profile in skeletal muscle

To understand how mTOR deficiency impacts iron metabolism, we examined the expression of key iron-related biomarkers in the *soleus* and TA muscles from 20-23-wk-old male mice. In the mutant *soleus* muscle, we observed a marked 3.2-fold decrease in TFR1 protein expression, which was associated with a corresponding reduction in *Tfr1* mRNA levels (Fig. 3A-C). This downregulation of *Tfr1* likely involves a mechanism independent of TTP/ZFP36, as *Ttp/Zfp36* mRNA levels remained unchanged (Fig. 3C). Conversely, expression of FTL and FTH increased by 3.9- and 4.3-fold, respectively (Fig. 3A, B). FPN displayed a similar trend (3.1-fold), although with some inter-individual variability (Fig. 3A, B). The increase in FTL, FTH and FPN protein content was associated with a reduction in the corresponding mRNAs (Fig. 3C), reflecting a post-transcriptional regulatory mechanism.

To confirm that these changes were not merely a late-stage effect of the dystrophy, we examined younger mice. *Soleus* muscles from mTORmKO mice at 7 weeks of age already displayed reduced TFR1 protein expression (1.7-fold) and increased FTL and FPN levels (2.6- and 2.4-fold, respectively) (Figure S1A, B). Moreover, the mildly affected TA muscle from 20-23-wk-old mTORmKO mice showed similar impact on iron-related molecules, although *Ttp/Zfp36* mRNA levels were increased in this tissue (Figure S3A-C). TFR1 downregulation in mTORmKO muscle
<figure><img src="image_5.png" /></figure>

<figure><img src="image_6.png" /></figure>

<figure><img src="image_7.png" /><figcaption>**Fig. 2** Prolonged loss of mTOR in the *soleus* muscle does not com-
promise ISC biogenesis and integrity. (A) Immunoblots of *soleus*
muscles from mTORmKO and control mice are shown for ISCU,
FXN, NDUFS3, SDHB, FECH, ACO2, LA-PDH(E2), PDH(E2)
and GAPDH (N=4). (B) Upper and lower panels are densitometric
quantifications of the western blots shown in (A) for the ISC-con-</figcaption></figure>

aligns with previous findings demonstrating mTOR-medi-
ated TFR1 mRNA and protein expression regulation [19].
Intriguingly, the post-transcriptional upregulation of ferri-
tin and FPN is a signature typically induced by iron excess,
while the mRNA downregulation of *Ftl*, *Fth* and *Fpn* con-
tradicts this response.

taining mitochondrial proteins compared to GAPDH, and PDH(E2)
lipoylation, respectively. (C) Relative mRNA levels encoding the pro-
teins shown in (A) in *soleus* muscles from control and mTORmKO
mice (N=6). N represents the number of individual male mice, aged
20–23 weeks, used in each group. Values represent the mean±SEM.
\*P≤0.05; \*\*P≤0.01; \*\*\*P≤0.001; ns, not significant

Collectively, these data show that mTOR depletion
in skeletal muscle disrupts the expression profile of iron-
related genes, independently of overall iron content and
muscle pathology. This finding suggests a direct role for
mTOR in maintaining proper iron-related gene expression
in skeletal muscle.
<figure><img src="image_8.png" /><figcaption>**Fig. 3** Sustained mTOR depletion in *soleus* muscle alters expression of iron biomarkers. (A) Western blot showing protein levels of TFR1, FTL, FTH, FPN and GAPDH in *soleus* muscles from control and mTORmKO mice (N=4). (B) Densitometric measurements of TFR1, FTL, FTH, and FPN compared to GAPDH (in arbitrary units) in *soleus* muscles from both control and mTORmKO mice (N=4). (C) qPCR</figcaption></figure>

analysis revealed the relative mRNA levels of *Tfr1*, *Ttp/Zfp36*, *Ftl*, *Fth* and *Fpn* in *soleus* muscles from control and mTORmKO mice (N=6). N represents the number of individual male mice, aged 20–23 weeks, used in each group. Values represent the mean±SEM. \*P≤0.05; \*\*P≤0.01; \*\*\*P≤0.001; ns, not significant

## Absence of muscle mTOR impairs the HIFs and NRF2 iron-regulatory pathways

The mTOR pathway has been shown to enhance HIFs and NRF2 activities [23–25], two transcription factors that directly control the expression of iron metabolism genes [10, 11]. Specifically, HIFs enhance the transcription of *TFR1* and *FPN*, while NRF2 increases the transcription of *FTL*, *FTH*, and *FPN*. To elucidate the mechanisms underlying the downregulation of the *Tfr1*, *Ftl*, *Fth*, and *Fpn* mRNAs in mTORmKO mouse muscles, we investigated the impact of loss of mTOR on HIFs and NRF2 signaling. In *soleus* muscle from 20-23-wk-old male mice, *HIF2α* was markedly suppressed both at the protein (3.0-fold; Fig. 4A, B) and the mRNA (3.6-fold; Fig. 4C) levels. *Hif1α* mRNA expression

decreased (1.6-fold; Fig. 4C), although HIF1α protein was undetectable by western blot. Consistently, several HIF target genes were downregulated in mTORmKO *soleus* muscles. This includes, in addition to *Tfr1* (Fig. 3A-C), the iron genes *Ncoa4* and *Dmt1*, as well as the proangiogenic factors *Vegf-a* and *Vegf-b*, and superoxide dismutase 2 (*Sod2*) (Fig. 4C). In the mutant TA muscle, *Hif2α* mRNA and HIF target genes were also diminished (Figure S4A). Furthermore, mTOR deficiency in *soleus* muscle led to a marked reduction in NRF2 protein expression (5.1-fold; Fig. 4A, B) and, to a lower extent, in its mRNA (1.5-fold; Fig. 4D). The drop in NRF2 levels was expectedly associated with reduced mRNA and protein expression of NAD(P)H-quinone oxidoreductase-1 (NQO1) (Fig. 4A, B,D), a well-established indicator of NRF2 activity in mouse muscle [41]. Similarly,
A

Soleus

B

C

D

<caption>**Fig. 4** Long-term mTOR deficiency in *soleus* muscle alters HIFs and NRF2 signaling. (A) Immunoblotting showing protein levels of HIF2α, NRF2, NQO1 and GAPDH in *soleus* muscles from control and mTORmKO mice (N=4). (B) Densitometric quantifications of HIF2α, NRF2, NQO1 relative to GAPDH (arbitrary units) in *soleus* muscles from control and mTORmKO mice (N=4). (C) Relative mRNA levels of *Hif2α* and *Hif1α* and of several HIF-target genes</caption>

including *Vegf-a*, *Vegf-b*, *Sod2*, *Dmt1* and *Ncoa4* in *soleus* muscles from control and mTORmKO mice (N=6). (D) Relative mRNA levels of *Nrf2* and of several NRF2-target genes including *Nqo1*, *Sod1*, *Gpx4*, *catalase*, *Herc2*, *Hmox1* and *Gclc* in *soleus* muscles from control and mTORmKO mice (N=6). N represents the number of individual male mice, aged 20–23 weeks, used in each group. Values represent the mean±SEM. \*P≤0.05; \*\*P≤0.01; \*\*\*P≤0.001; ns, not significant

<figure><img src="image_9.png" /></figure>

<figure><img src="image_10.png" /></figure>

<figure><img src="image_11.png" /></figure>

<figure><img src="image_12.png" /></figure>
mRNA levels of several NRF2 target genes were decreased in mutant *soleus* muscles, including the iron-regulating gene *HERC2*, along with superoxide dismutase 1 (*Sod1*), glutathione peroxidase 4 (*Gpx4*), and *catalase*, which are involved in oxidative stress responses (Fig. 4D). Notably, *Fech* and *Fxn*, which were found to be downregulated in the mutant muscle (Fig. 2C) are also direct transcriptional targets of NRF2 [42, 43]. Conversely, the NRF2 targets heme oxygenase 1 (*Hmox1*) and glutamate-cysteine ligase catalytic subunit (*Gclc*) remained unchanged (Fig. 4D), consistent with previous reports of minimal impact of NRF2 deficiency on these antioxidant genes in mouse skeletal muscle [44]. The mutant TA muscle showed similar but less pronounced effects on NRF2 signaling (Figure S4B-D). Importantly, *soleus* muscles from 7-wk-old mTORmKO mice exhibited strong downregulation of NRF2 and its target NQO1 (Figure S1A, B). This early-onset effect suggests that mTOR directly regulates NRF2 signaling in skeletal muscle.

These results indicate that loss of muscle mTOR dampens the activity of both HIFs and NRF2, which likely contribute to the observed decrease in mRNA levels of key iron-related genes.

## Muscle mTOR deficiency mitigates the inhibition of ferritin synthesis by the IRPs and promotes NCOA4-mediated ferritinophagy

We found that mTOR deficiency in skeletal muscle is associated with increased ferritin protein content and NRF2 downregulation. Interestingly, NRF2 controls both ferritin synthesis and degradation by positively regulating the transcription of the E3 ubiquitin ligase *HERC2* [18]. Specifically, HERC2 degrades FBXL5, which itself targets IRP2 and apo-IRP1 for degradation [15], thus leading to IRP-mediated ferritin translation repression. Additionally, HERC2 promotes the degradation of NCOA4 [18], the cargo receptor recruiting ferritin to autophagosomes for processing, thus preventing ferritinophagy. Given the downregulation of NRF2 (Fig. 4A, B, S1A, B and S4B, C) and the concurrent decrease in *Herc2* mRNA expression in mTORmKO muscle (Fig. 4D and S4D), we assessed the impact of mTOR deficiency on HERC2 downstream targets involved in both ferritin synthesis and processing.

We observed an approximately 2-fold increase in FBXL5 protein expression in mTORmKO *soleus* muscle from 20-23-wk-old male mice (Fig. 5A, B) despite a drop in *Fbxl5* mRNA levels (Fig. 5C). Consistent with the increase in FBXL5, we found a significant reduction in IRP1 and IRP2 protein levels by 2.0- and 3.2-fold, respectively (Fig. 5A, B). Similar results were obtained in TA muscle from age-matched mTORmKO mice (Figure S5A-C). These results suggest that IRP downregulation results from increased

expression of the FBXL5 ubiquitin ligase, with the partial suppression of *Aco1* and *Ireb2* mRNAs potentially contributing to this effect (Fig. 5C). To assess the functional consequences of altered IRP levels, we evaluated the IRP RNA-binding activity of mTORmKO muscles to the IRE site of ferritin using RNA electrophoretic mobility shift assay (REMSA). Notably, REMSA in control *soleus* muscle did not detect significant IRE-binding under native conditions, possibly due to the predominance of IRP1 in its ACO1 form in this iron-rich muscle and the limited sensitivity of the fluorescent probe method. Conversely, under native conditions, REMSA revealed significantly lower IRE-binding in mTORmKO TA and GC muscles compared to controls (Fig. 5D), indicating suppression of IRP function. Furthermore, under reducing conditions, which convert ACO1 to its IRE-binding form (IRP1), IRE-binding was markedly increased in mTORmKO TA and GC muscles. These results suggest that a substantial fraction of the remaining IRP1 in these mutant muscles functioned as cytosolic aconitase.

In addition to the changes in the FBXL5-IRP axis, mTOR deficiency in *soleus* muscle also raised NCOA4 protein levels (1.8-fold; Fig. 5A, B) which is predicted to promote ferritinophagy, despite a reduction in *Ncoa4* mRNA expression (Fig. 4C). In TA muscle of mTORmKO mice, NCOA4 protein expression was also increased (Figure S5A, B) with a concurrent reduction in its mRNA levels (Figure S4A).

Importantly, significantly increased NCOA4 protein levels (1.7-fold, *p*<0.05) and a non-significant trend toward elevated FBXL5 levels (1.5-fold, *p*=0.1), accompanied by decreased IRP2 protein expression, were observed in *soleus* muscles from 7-wk-old mTORmKO mice (Figure S1A, B). These early-onset changes are consistent with the observed NRF2 downregulation at this stage (Figure S1A, B).

Thus, mTOR deficiency and the ensuing decrease in NRF2 activity may affect ferritin content in skeletal muscle by two apparently antagonistic HERC2-dependent mechanisms: derepression of ferritin translation via the FBXL5-IRP axis and stimulation of ferritinophagy via NCOA4.

## Ferritinophagy in *soleus* muscle is hindered by sustained mTOR deficiency

Our findings reveal that mTOR deficiency in muscle upregulates NCOA4 protein, a key mediator of ferritinophagy (Figs. 5A, B, S1A, B and S5A, B). Importantly, while loss of muscle mTORC1 in mTORC1mKO mice leads to sustained induction of autophagy, the flux through the autophagy pathway declines with age due to alterations in the final stages of autophagic degradation linked to chronic AKT/PKB activation [28, 29]. Since autophagy inhibition has been shown to suppress ferritinophagy [45], this age-related decline in autophagy flux may limit the effectiveness of
<figure><img src="image_13.png" /></figure>

<figure><img src="image_14.png" /></figure>

**Fig. 5** Prolonged loss of muscle mTOR leads to reduced inhibitory IRP activity on ferritin synthesis and NCOA4 upregulation. (A) Immunoblotting analysis of *soleus* muscles from control and mTORmKO mice showing the expression levels of FBXL5, IRP1, IRP2, NCOA4 and GAPDH (N=3–4). (B) Densitometric quantifications of FBXL5, IRP1, IRP2 and NCOA4 relative to GAPDH (arbitrary units) in *soleus* muscles from control and mTORmKO mice (N=3–4). (C) Relative mRNA levels of *Fbxl5*, *Aco1* and *Ireb2* in *soleus* muscles from control and mTORmKO mice (N=6). (D) Upper panel: Analysis of IRE-binding activities to the IRE of ferritin-H in TA and *gastrocnemius* (GC)

NCOA4-mediated ferritin degradation in mTOR-deficient muscle.

To investigate the age-dependent changes in ferritinophagy, we examined the *soleus* muscle from 25-wk-old female mTORmKO mice. Similar to their male counterparts, these mice exhibited impaired NRF2 signaling, as evidenced by NQO1 downregulation, FBXL5 elevation associated with IRP2 suppression and subsequent FTL upregulation, as well as increased NCOA4 levels (Fig. 6A: lanes 1–3

<figure><img src="image_15.png" /></figure>

<figure><img src="image_16.png" /></figure>

muscles from control and mTORmKO mice. The fluorescent probe (Alexa Fluor® 488) was incubated with muscle extracts in native or reducing conditions with 2-ME. Lower panel: quantification of IRP binding to the probe in mutant muscles compared to controls under native conditions. REMSA was performed three times for each condition (control and KO) using TA and GC muscles. In each experiment, two muscles (either TA or GC) from two different mice were used per condition (N=6). N represents the number of individual male mice, aged 20–23 weeks, used in each group. Values represent the mean±SEM. \*P≤0.05; \*\*P≤0.01; \*\*\*P≤0.001; ns, not significant

vs. 7–9, 6B). At this age, the mutant *soleus* muscles displayed molecular markers indicative of autophagy induction coupled with blockade, mirroring the phenotype observed in aged mTORC1mKO mice [28], and characterized by autophagosome accumulation. Specifically, we observed increased MAP1LC3B/LC3B lipidation, evidenced by elevated LC3B-II/-I and LC3B-II/GAPDH ratios (Fig. 6A: lanes 1–3 vs. 7–9, 6B), along with increased p62/SQSTM1 levels (Fig. 6A: lanes 1–3 vs. 7–9, 6B) and the formation
of p62 aggregates (Fig. 6C). Furthermore, immunofluores-
cence analysis revealed numerous FTL puncta colocalizing
with p62 within mutant muscle fibers (Fig. 6C), strongly
indicating active FTL recruitment and accumulation in
autophagosomes. Similarly, mTORmKO TA muscle dis-
played significant FTL and p62 colocalization (Figure S6A).
This contrasted with control soleus and TA muscle fibers,
which showed minimal FTL puncta and FTL/p62 associa-
tion (Fig. 6C, S6A). Interestingly, FTL showed little asso-
ciation with LAMP1-positive endolysosomal compartments
in mTORmKO soleus fibers (Fig. 6D), suggesting impaired
ferritin processing via ferritinophagy. In contrast to the find-
ings in mutant soleus muscle from 25-wk-old mice, FTL/
p62 and FTL/LAMP1 colocalization in soleus muscle from
7-wk-old mTORmKO mice was comparable to controls,
with low levels of colocalization, consistent with functional
autophagy at this younger age (Figure S6C, D).

Collectively, these observations suggest that elevated
muscle FTL in young mTORmKO mice primarily reflects
increased synthesis, whereas in older mice, defective fer-
ritinophagy further contributes to FTL accumulation.

We next tested whether spermidine, a known autophagy
inducer [46], could counteract the age-related accumulation
of autophagosomal FTL. We administered 3mM spermidine
to 22-wk-old female mice for 3 weeks. Notably, spermidine
supplementation has been shown to rescue autophagy in
dystrophic skeletal muscle of collagen VI-null (col6a1-/-) mice [30]. Given the pronounced dystrophic phenotype of
the soleus in more aged mTORmKO mice, we expected a
greater response to spermidine in this muscle. Supporting
this hypothesis, spermidine significantly reduced FTL levels
in the mTORmKO soleus muscle (2.1-fold), with no effect
on FTL levels in control mice (Fig. 6A, B). This reduction
in FTL was not attributable to changes in NRF2 signaling
impacting FTL synthesis or autophagosomal targeting, as
evidenced by comparable levels of NQO1, NCOA4, FBXL5,
and IRP2 in both treated and untreated mTORmKO mice
(Fig. 6A: lanes 7–9 vs. 10–12, 6B). In control mice, sper-
midine had no significant impact on autophagy markers in
control mice (Fig. 6A: lanes 1–3 vs. 4–6, 6B). However, in
mutant soleus, it led to partially normalized LC3B-II/-I and
LC3B-II/GAPDH ratios (Fig. 6A: lanes 1–3 vs. 7–12, 6B),
suggesting improved autophagy flux. Despite this improve-
ment, p62 levels remained elevated (Fig. 6A: lanes 7–9 vs.
10–12, 6B) with persistent aggregate formation (Fig. 6E).
This sustained p62 accumulation may be attributed to con-
tinued autophagy induction, incomplete autophagy recovery,
and slow turnover following aggregation [47]. Importantly,
FTL reduction in treated mTORmKO soleus muscle corre-
lated with decreased FTL/p62 colocalization (Fig. 6E) and
increased FTL association with LAMP1-positive endolyso-
somal compartments (Fig. 6F).

These findings strongly support the hypothesis that
impaired autophagy, linked to prolonged mTOR depletion
in skeletal muscle, contributes to ferritin accumulation.

Spermidine treatment is associated with normalized AKT/FOXO signaling but fails to induce autophagy-related genes in mTORmKO soleus muscle after 3 weeks

Spermidine induces autophagy through multiple mecha-
nisms, including modulation of AKT/PKB-FOXO signaling
[30, 46]. Therefore, we investigated the effects of spermi-
dine treatment on this signaling pathway in soleus muscle
from 25-wk-old female mTORmKO mice. Consistent with
our previous findings [27], the mutant soleus muscle dem-
onstrated feedback AKT/PKB activation, as evidenced by
an 8.6-fold increase in the pAKT<sup>S473</sup>/AKT ratio compared
to control muscle (Fig. 7A: lanes 1–3 vs. 7–9, 7B). This
was accompanied by increased phosphorylation of the
AKT/PKB substrate FOXO1 on Ser256, as well as elevated
FOXO1 total protein levels, resulting in a 1.4-fold increase
in the pFOXO1/FOXO1 ratio. Spermidine administration
to mTORmKO mice significantly downregulated AKT<sup>S473</sup>
phosphorylation, reducing the pAKT<sup>S473</sup>/AKT ratio by
5-fold (Fig. 7A: lanes 7–9 vs. 10–12, 7B). In line with
reduced AKT/PKB activity, the pFOXO1<sup>S256</sup>/FOXO1 ratio
was decreased by 1.5-fold, reaching control levels (Fig. 7A:
lanes 7–9 vs. 10–12, 7B). Interestingly, spermidine admin-
istration to control mice led to a mild increase in both
AKT<sup>S473</sup> phosphorylation and AKT protein levels, leaving
the pAKT<sup>S473</sup>/AKT ratio unchanged (Fig. 7A: lanes 1–3
vs. 4–6, 7B). While phosphorylation of FOXO1 on Ser256
remained unaltered, its protein levels increased, resulting in
a 1.7-fold reduction in the pFOXO1<sup>S256</sup>/FOXO1 ratio.

FOXO3, another substrate of AKT/PKB, is a crucial tran-
scriptional regulator of autophagy in skeletal muscle [48,
49]. To assess FOXO3 activity in mutant muscles, we exam-
ined its nuclear localization, which is indicative of its active
state. In soleus muscle fibers from 25-wk-old mTORmKO
mice, FOXO3 fluorescence intensity in myonuclei was
significantly reduced compared to controls (Fig. 7C), with
similar results observed in mTORmKO TA muscle fibers
(Figure S6B). In contrast, FOXO3 levels in soleus myo-
nuclei of 7-wk-old mTORmKO mice were comparable to
controls (Figure S6E), in line with a progressive downregu-
lation of FOXO3 activity resulting from chronic mTOR defi-
ciency. Spermidine treatment was associated with increased
nuclear FOXO3 levels in the soleus muscle from 25-wk-
old mTORmKO mice, although not reaching control lev-
els (Fig. 7D), consistent with its attenuation of AKT/PKB
signaling. To further investigate autophagy following sper-
midine treatment, we analyzed mRNA levels of autophagy
A **Soleus**

<figure><img src="image_17.png" /></figure>

<figure><img src="image_18.png" /></figure>

<figure><img src="image_19.png" /></figure>

<figure><img src="image_20.png" /></figure>

<figure><img src="image_21.png" /></figure>

<figure><img src="image_22.png" /></figure>

<figure><img src="image_23.png" /></figure>

<figure><img src="image_24.png" /></figure>

<figure><img src="image_25.png" /></figure>

<figure><img src="image_26.png" /></figure>
**Fig. 6** Impaired autophagy associated with chronic mTOR deficiency leads to autophagosomal FTL accumulation in *soleus* muscle. (A) Western blot analysis of NQO1, FBXL5, IRP2, FTL, NCOA4, LC3B lipidation, p62 and GAPDH in protein extracts from *soleus* muscles from control and mTORmKO female mice that were untreated or *per os* spermidine (Spd, 3 mM)-treated for 3 weeks (N=3). (B) Densitometric quantifications of NQO1, FBXL5, IRP2, FTL, NCOA4, LC3B-II vs. LC3B-I, and of LC3B-II and p62 vs. GAPDH (arbitrary units) of untreated (-) and *per os* spermidine (Spd)-treated (+) control and mTORmKO female mice (N=3). (C–F) Left panels: Isolated fibers of *soleus* muscles from control (CTL) and mTORmKO (mTOR-) female mice, untreated (C and D) or *per os* spermidine (Spd)-treated (E and F), were double-stained with FTL (in red) and either p62 (C and E; in green) or LAMP1 (D and F; in green) (N=3; x=19–47). Nuclei were labeled with DAPI (in blue in merged panels). White arrowheads points at colocalized puncta detected on the merged images as yellow dots. Dashed lines delimit muscle fibers. Scale bars: 25 µm; zoom sections (Inset 1–4 and 1’-4’). Right panels: colocalization quantification (in arbitrary units, n=5–10). N represents the number of individual 25-wk-old female mice used in each group; x is the total number of dissociated fibers counted; n is total number of confocal images counted. Data presented as mean±SEM. \*\#\#/\#\*P≤0.05; \*\*\#\#/\#\*\*P≤0.01; \*\*\#\#\#/\#\*\*\*P≤0.001; ns, not significant. \* is mTORmKO vs. treated mTORmKO, \# is control vs. mTORmKO, \#\* is control vs. treated control, \#\*\# is treated control vs. treated mTORmKO

markers regulated by FOXO3, including *p62/SQSTM1*, *LC3B*, *Beclin 1*, *Bnip3* and *Cathepsin L* (Fig. 7E). In line with observations in mTORC1mKO muscle [28], transcript levels of these genes were markedly reduced in mTORmKO *soleus* muscles compared to controls. However, the expression levels of these genes remained low in this mutant muscle despite elevated FOXO3 nuclear localization following spermidine treatment. This observation aligns with previous work, which reported that spermidine normalized AKT-FOXO signaling and autophagy in col6a1⁻/⁻ skeletal muscle without upregulating FOXO target genes under fed conditions [30]. Thus, spermidine treatment was associated with normalized AKT-FOXO signaling, potentially creating a permissive environment for autophagy, as proposed by Chrisam et al. [30].

Following the observed increase in ferritinophagy after 3-week oral spermidine treatment, we examined its potential to ameliorate established myopathic features in mTORmKO mice. However, analysis of several parameters revealed a limited impact on the overall myopathic phenotype. The whole-body weight in 25-wk-old female mTORmKO mice showed no significant improvement (Figure S7A). Histological analysis of muscle cross-sections revealed no significant reduction in the percentage of fibers with centralized nuclei in both *soleus* and PLA muscles of spermidine-treated compared to non-treated mTORmKO mice (Figure S7B-E). In addition, mononuclear cells were still detectable in treated mTORmKO *soleus* muscle (Figure S7B). The treated mTORmKO muscle still displayed a prevalence of small fibers comparable to those in untreated mTORmKO muscle. Unexpectedly, the percentage of very small muscle

fibers (below 250 µm²) increased with treatment in both mutant *soleus* and PLA muscles (Figure S7F, G). Furthermore, glycogen content, which is markedly increased in the mutant PLA and GC muscles as shown by PAS staining, remained significantly elevated in treated mTORmKO mice (Figure S7H). This indicates that spermidine supplementation failed to normalize this metabolic alteration, which has been proposed to result from AKT/PKB activation [27, 39]. These results suggest that the 3-week spermidine treatment, while effective in restoring ferritinophagy, was insufficient to ameliorate the overall myopathic phenotype in mTORmKO mice.

## Discussion

Our study reveals a complex interplay between mTOR signaling and iron metabolism, demonstrating that constitutive mTOR deletion in mouse skeletal muscle results in distinct iron regulation patterns across different muscle types. In slow-twitch mutant *soleus* muscles, iron levels were initially reduced but increased as myopathy progressed, remaining within the control range. In contrast, fast-twitch mutant muscles, which showed less severe pathology, maintained near-normal levels. This pattern aligns with previous studies showing that iron accumulation occurs specifically in the severely affected muscles of dystrophic mouse models [6].

Regardless of the absolute iron content, mTORmKO muscles exhibit a paradoxical molecular signature of iron regulation (Fig. 8), reflecting features of both depletion (NCOA4-mediated ferritinophagy induction, *ferritin* and *Fpn* mRNA downregulation) and overload (enhanced FBXL5-mediated IRP inhibition, reduced TFR1 expression, increased ferritin and FPN protein synthesis, *Ncoa4* mRNA downregulation, and inhibited ferritin degradation at a more advanced age). Although our study uses whole-muscle homogenates, multiple lines of evidence suggest that the observed molecular signature is primarily intrinsic to the non-regenerating mTORmKO muscle fibers, rather than being confounded by infiltrating cells or the regenerative process. First, this signature is consistently detected in *soleus* muscles at both early and advanced stages of disease progression, as well as in the mildly affected TA muscle. Second, regenerating muscles display a distinct iron molecular signature compared to mTORmKO muscles. For example, acutely injured muscle fibers treated with cardiotoxin (CTX) typically show unchanged *Tfr1* mRNA levels but elevated TFR1 protein [50], while TFR1 protein remains unaffected in muscle from dystrophin-null mice (*mdx*), a mouse model for Duchenne Muscular Dystrophy [6]. Additionally, TFR1 is highly expressed in proliferative satellite cells [51]. Furthermore, CTX-injured
A Soleus

<figure><img src="image_27.png" /></figure>

B

Control
mTORmKO

<figure><img src="image_28.png" /></figure>

<figure><img src="image_29.png" /></figure>

<figure><img src="image_30.png" /></figure>

C DAPI FOXO3 merged

<figure><img src="image_31.png" /></figure>

<figure><img src="image_32.png" /></figure>

<figure><img src="image_33.png" /></figure>

D DAPI FOXO3 merged

<figure><img src="image_34.png" /></figure>

<figure><img src="image_35.png" /></figure>

<figure><img src="image_36.png" /></figure>

<figure><img src="image_37.png" /></figure>

▼ Control

▽ mTORmKO

☐ mTORmKO + Spd
**Fig. 7** 3-week spermidine treatment normalizes AKT/FOXO signaling without restoring autophagy-related gene expression in mTORmKO *soleus* muscle. (A) Western blot analysis of pAKT1<sup>S473</sup>, AKT, pFOXO1<sup>S256</sup>, FOXO1, FOXO3, and GAPDH in protein extracts from *soleus* muscles of 25-wk-old control and mTORmKO female mice that were untreated or *per os* spermidine (Spd, 3 mM)-treated for 3 weeks (N=3). (B) Densitometric quantifications of pAKT1<sup>S473</sup> vs. AKT, pFOXO1<sup>S256</sup> vs. FOXO1 and of FOXO1, FOXO3 vs. GAPDH (arbitrary units) of untreated (-) and *per os* spermidine (Spd)-treated (+) control and mTORmKO female mice (N=3). (C-D) Left panels: Isolated fibers of *soleus* muscles from control (CTL) and mTORmKO (mTOR-) female mice, untreated (C) or *per os* spermidine (Spd)-treated (D), were stained with FOXO3 (in green) (N=3; x=139–515). Nuclei were labeled with DAPI (in red). Thin dashed lines delimit muscle fibers, and thick dashed lines delimit nuclei. Scale bars: 25 µm; zoom sections (Inset 1–2 and 1’-2’). Right panels: quantification of nuclear FOXO3 (in arbitrary units, n=10–15). (E) qPCR analysis revealed the relative mRNA levels of *p62*, *LC3B*, *Beclin1*, *Bnip3* and *CtsL* in *soleus* muscles from control, untreated mTORmKO and Spd-treated mTORmKO mice (N=4–5). N represents the number of individual 25-wk-old female mice used in each group; x is the total number of nuclei counted; n is total number of confocal images counted. Data presented as mean±SEM. \*\#\Delta/\#\%P\le0.05; \*\*\#\Delta/\#\%\#P\le0.01; \*\*\#\#\Delta/\#\#\%\#P\le0.001; ns, not significant. \# is mTORmKO vs. treated mTORmKO, \#\# is control vs. mTORmKO, \#\#\# is control vs. treated control, \#\#\#\# is treated control vs. treated mTORmKO, \#\#\#\#\# is Control vs. treated mTORmKO

muscles show induced *ferritin* and *Fpn* mRNA expression originating largely from infiltrating immune cells [50], whereas the opposite is observed in mTORmKO muscles. Finally, our immunofluorescence staining confirms cell-autonomous accumulation of FTL protein within mutant muscle fibers. Although our evidence strongly supports an intrinsic molecular signature in mTORmKO muscle fibers, we cannot completely rule out any influence from infiltrated macrophages on the ferritin and FPN levels detected by western blot in aged mTORmKO soleus muscle [50]. However, the absence of Perls' staining-detectable ferric iron deposits in this muscle suggests minimal impact of macrophage-mediated iron handling, possibly due to the strong myoglobin downregulation in mTORmKO soleus (see below).

Instead, our findings indicate that mTOR depletion directly perturbs key intracellular iron regulatory mechanisms in mTORmKO muscles. This includes disruptions in the NRF2/NFE2L2 pathway, HIF axis, and AKT/PKB-mediated autophagy signaling (Fig. 8), with impaired NRF2 signaling playing a crucial role, likely due to loss of mTORC2 [24]. While iron homeostasis has not been extensively studied in muscle-specific NRF2 knockout models to date, our observations align with findings from NRF2-deficient ovarian cell lines (NRF2KO cells). Indeed, these NRF2KO cells exhibit reduced *ferritin* mRNA levels coupled with enhanced ferritin synthesis via FBXL5-mediated suppression of IRP activity, ferritinophagy induction via NCOA4 upregulation, and autophagosomal ferritin accumulation due to impaired autophagy flux [18]. Conversely, the

downregulation of *TFR1* mRNA and protein as well as *Fpn* mRNA appear to be specific to muscle mTOR depletion.

On one hand, NRF2KO cells exhibit only mild downregulation of *Tfr1* mRNA with unaffected protein levels. This observation is surprising, as IRP inhibition in these cells would typically be expected to downregulate TFR1 mRNA and protein levels. Our finding of significant TFR1 downregulation in mTOR-deficient muscle aligns with previous studies demonstrating TFR1 sensitivity to pharmacological mTOR inhibition [19]. Mechanistically, mTOR inhibition with rapamycin (a partial mTORC1 inhibitor) or Torin1 (inhibiting both mTORC1 and mTORC2) has been shown to induce *Tfr1* mRNA destabilization via TTP/ZFP36 induction in various cell types and tissues [19]. Nonetheless, another study demonstrated that Torin1 had no effect on *Ttp/Zfp36* mRNA induction in MEFs in the presence of serum, and even reduced protein expression [52].

Interestingly, we observed *Ttp/Zfp36* induction in mTOR-deficient TA muscle but not in the more dystrophic *soleus*, which is surprising given previous reports of increased TTP/ZFP36 expression in regenerating muscle [53, 54]. Importantly, TTP/ZFP36 regulation may be context-dependent. *TTP/ZFP36* transcription is influenced by multiple signaling pathways and transcription factors beyond mTOR, including AMPK, Smad3 (TGF-β signaling), p38 MAPK, HDAC, p53, and c-MYC [52, 55]. Differential activation or inhibition of these pathways in mTORmKO *soleus* versus TA muscle may contribute to the observed muscle-specific *Ttp/Zfp36* expression patterns. Alternatively, *Tfr1* mRNA downregulation in mTORmKO muscle may be attributed to altered expression or activity of mTOR-regulated transcription factors [56] that have been demonstrated to directly activate *Tfr1* expression. These include HIF [12] and PGC-1α [57], both of which are downregulated in mTOR mutant *soleus* muscle (this study and [27]), as well as YY1 [58] and c-MYC [59]. Additionally, reduced IRP activity due to impaired NRF2 signaling in mTOR-deficient muscle may contribute to decreased *Tfr1* mRNA stability [9], although this mechanism is seemingly not involved in NRF2KO cells [18].

On the other hand, contrary to the upregulation of *Fpn* mRNA observed in NRF2KO cells, we detected *Fpn* mRNA downregulation in mTORmKO muscle. Notably, *Fpn* mRNA regulation by NRF2 can vary depending on cell type and physiological conditions [10], and may rely on HIF signaling [14] which is impaired in mTORmKO muscle. Furthermore, our findings of FPN protein upregulation contrast with prior studies using rapamycin, which reported reductions in both FPN mRNA and protein levels in cells and mouse hearts [19]. *Fpn* mRNA downregulation under rapamycin conditions may result from the inhibition of the rapamycin-sensitive HIF signaling pathway. Conversely,
<figure><img src="image_38.png" /><figcaption>**Fig. 8** Schematic model: Impact of mTOR depletion on iron homeostasis signaling in skeletal muscle. - mTOR depletion disrupts NRF2 and HIF signaling, leading to the mRNA downregulation of key iron-related genes, including *Tfr1*, *Ftl*, *Fpn* and *Herc2*; HIF-mediated *Ncoa4* mRNA regulation is omitted for clarity. - *Tfr1* mRNA expression may be further suppressed through dual mechanisms: induction of TTP/ZFP36, an mRNA-destabilizing protein, and downregulation of the IRP system, which typically stabilizes *Tfr1* transcripts under low-iron conditions. - Reduced NRF2 activity lowers HERC2 levels, an E3 ubiquitin ligase for both FBXL5 (another E3 ubiquitin ligase) and NCOA4 (a cargo receptor for FTL). - Elevated FBXL5 suppresses</figcaption></figure>

FPN protein downregulation in this context is driven by the activation of the IRP system, which results from cellular iron deficiency caused by TFR1 downregulation. In contrast, our findings suggest that FPN protein upregulation in mTOR mutant muscle occurs via suppression of the IRP axis. This suppression is linked to NRF2, which is itself under the control of the rapamycin-resistant mTORC2 pathway [24]. Additionally, Bayeva et al. demonstrated that Torin1, a dual inhibitor of mTORC1 and mTORC2, reduced *Fpn* mRNA levels. However, the impact of this treatment on FPN protein levels was not assessed in their study. It is plausible that Torin1, by inhibiting mTORC2 and consequently suppressing both NRF2 and the IRP system, could potentially enhance FPN synthesis despite reduced mRNA levels.

The accumulation of autophagosomal ferritin in mTORmKO muscles, resulting from both enhanced feritinophagy and impaired autophagy flux, also parallels observations made in NRF2KO cells. In NRF2KO cells, strong NCOA4 induction promotes the autophagosomal

IRP expression and activity, enhancing FTL and FPN protein translation. - Increased NCOA4, combined with autophagy induction caused by mTOR depletion, promotes FTL recruitment to autophagosomes for ferritinophagy. - Chronic feedback AKT/PKB activation due to mTOR depletion triggers FOXO-mediated autophagy blockade associated with autophagosomal FTL accumulation. - Spermidine (Spd) treatment normalizes AKT-FOXO signaling, increases endolysosomal FTL and reduces total FTL levels. The molecular details underlying Spd's effect on endolysosomal FTL remains to be elucidated. TX, transcriptional regulation; PTR, post-transcriptional regulation; TL, translational regulation; PTM, post-translational modification

recruitment of apoferritin over holoferritin [18]. However, the nature of ferritin accumulating in autophagosomes in mTORmKO muscles remains unclear. The partial NRF2 downregulation, compared to its complete absence in NRF2KO cells, may differentially affect NCOA4 induction and ferritin recruitment. While the observed shift from Fe(II) to Fe(III) in mTORmKO *soleus* muscles suggests iron sequestration within ferritin, potentially as holoferritin, the precise localization, whether autophagosomal, cytosolic, or a combination of both, remains undefined. However, the absence of detectable ferric iron deposits following Perls' Prussian Blue staining argues against substantial accumulation of autophagosomal holoferritin. Further investigation employing highly sensitive methodologies is required to fully elucidate the nature and subcellular distribution of ferritin accumulation in mTORmKO *soleus* muscles.

Noteworthy, the mechanisms underlying impaired autophagy differ between mTORmKO muscles and NRF2KO cells. In NRF2KO cells, impaired autophagy
is attributed to mTOR activation [18]. In contrast, in mTORmKO muscles, autophagy flux impairment is thought to result from chronic AKT/PKB activation [28, 29] caused by disruption of the well-established mTORC1/S6K negative feedback loop, along with the activity of an unidentified kinase independent of mTORC2 [27]. Sustained AKT/PKB signaling has been consistently associated with autophagy inhibition in various skeletal muscle models, including wild-type [48, 60], col6a1⁻/⁻ [30] and *mdx* mice, as well as individuals with Duchenne Muscular Dystrophy (DMD) [61]. AKT/PKB can inhibit autophagy in skeletal muscle through multiple downstream targets. Specifically, AKT/PKB-mediated phosphorylation and inhibition of the transcription factors FOXO suppresses the expression of key autophagy-related genes [30, 48], while AKT/PKB-mediated activation of mTORC1 blocks autophagy initiation [62]. Importantly, AKT/PKB has also been demonstrated to negatively regulate autophagy and lysosomal biogenesis in contexts of mTORC1 deficiency or inhibition [63–65].

Using spermidine treatment, a well-established autophagy inducer [46], we show that impaired autophagy contributes to ferritin accumulation in mTORmKO *soleus* muscle. Three-week spermidine administration was associated with increased ferritin processing through the endolysosomal pathway, concurrent with normalization of AKT/PKB-FOXO signaling. This observation aligns with prior findings demonstrating that spermidine rescues autophagy by modulating AKT/PKB hyperactivation in muscle from myopathic col6a1⁻/⁻ mice [30]. Interestingly, despite this effect, spermidine did not induce transcriptional activation of FOXO target genes in the mTORmKO *soleus* muscle. Similar findings were reported by Chrisam et al., where FOXO target gene activation under basal conditions was absent and only observed after 24 h of fasting. While spermidine has been suggested to create a permissive environment for autophagy activation by normalizing AKT-FOXO signaling [30], these results also point to the involvement of additional mechanisms. For instance, spermidine is known to inhibit EP300, a lysine acetyltransferase that acetylates and inactivates several autophagy-related proteins, including Beclin 1 and LC3, thereby repressing autophagy [46]. This suggests that EP300 inhibition or other post-translational modifications may play a more direct role in autophagy reactivation during spermidine treatment under basal conditions. Furthermore, future studies should investigate whether the observed increase in endolysosomal ferritin following spermidine treatment leads to its degradation or sorting for secretion. Notably, ferritin is known to be exported from cells via secretory pathways, such as the multivesicular body–exosome pathway, which plays a crucial role in maintaining tissue iron homeostasis [9].

Elevated ferritin levels have also been reported in skeletal muscles of aged mice and severely affected dystrophic muscles from *mdx* and dystrophin/utrophin-null mice [3, 6]. These models additionally exhibit significant iron overload and oxidative stress. Although iron content in mTORmKO *soleus* muscle normalizes as the myopathy progresses, it does not reach levels of iron overload compared to controls. This may be attributed to the lower expression of hem-containing myoglobin and TFR1, which collectively contribute to a reduced overall iron content in these muscles. In contrast, aged and *mdx* muscles maintain sustained TFR1 expression, potentially accounting for their excessive iron accumulation [3, 6]. These observations suggest that modulating mTOR activity to target TFR1 in aged or dystrophic models could potentially mitigate muscle iron overload.

Altered NRF2 signaling may also contribute to the dysregulation of ferritin in these models. While NRF2 signaling is not inherently impaired in *mdx* muscles, it may be insufficiently activated under stress conditions [66]. Aged muscles, however, exhibit a decline in NRF2 activity [67]. Ferritin accumulation and iron overload in aged and *mdx* muscles may also result from disrupted autophagy flux [61, 68], potentially leading to defective ferritinophagy [3, 6, 69]. In this sense, Alves et al. propose that interventions stimulating autophagy, such as rapamycin treatment and calorie restriction, could improve skeletal muscle iron homeostasis [3]. However, our study indicates that AKT/PKB feedback activation upon mTOR inhibition contributes to autophagy-ferritinophagy blockade. Instead, our findings suggest that spermidine supplementation could potentially restore ferritinophagy and alleviate impaired muscle iron handling in the context of autophagy blockade in dystrophic muscle. Nonetheless, a 3-week spermidine treatment was insufficient to significantly improve the overall myopathic phenotype in mTORmKO mice and even increased the percentage of small muscle fibers, aligning with Iwata et al.'s findings [70]. While spermidine may enhance satellite cell activation and early-stage myofiber formation [71], mTOR absence likely prevents these nascent fibers from maturing by impairing protein synthesis-dependent growth [72]. Additionally, iron recycling through macrophages, crucial for muscle regeneration and toxicity prevention [50], may be compromised in mutant *soleus* muscle due to myoglobin deficiency, a primary macrophage iron source during regeneration. Reduced TFR1 expression in regenerating fibers may further hinder iron uptake, essential for efficient muscle regeneration [50]. These mTOR-dependent defects in protein synthesis and iron homeostasis likely prevent significant improvement in *soleus* muscle, even if spermidine alleviates impaired ferritinophagy. Future studies should investigate whether earlier and longer spermidine treatment could mitigate myopathy progression in this model.
Finally, our study suggests that the mitochondrial respiratory defects in the *soleus* muscle of mTORmKO mice [27] are unlikely to be primarily driven by impairments in ISC biogenesis or integrity. While a minor role for mTOR in ISCU stability cannot be excluded [21], the lack of abnormal iron deposits within mTORmKO muscle fibers, a hallmark feature of ISCU myopathy [22] supports this conclusion. Instead, the downregulation of mitochondrial proteins, including ISC assembly proteins and ISC-containing proteins, in mutant *soleus* muscle is more likely linked to an overall reduction in mitochondrial content caused by decreased PGC-1α expression and mitochondrial remodeling associated with the myopathy [27, 39].

In conclusion, our study demonstrates that the loss of muscle mTOR disrupts essential intracellular mechanisms governing iron metabolism. Moreover, spermidine supplementation may represent a potential strategy to address impaired ferritinophagy in skeletal muscle.

**Supplementary Information** The online version contains supplementary material available at https://doi.org/10.1007/s00018-025-05695-9.

**Acknowledgements** We acknowledge the contribution of SFR Biosciences (Université Claude Bernard Lyon1, CNRS UAR3444, Inserm US8, ENS de Lyon), AniRA-PBES and AniRA-AGC. We thank the CIQLE microscopy facilities of Lyon for access to microscopes.

**Author contributions** Y-G.G. and A.C-D. conceived the project. A.C-D, A.O, H.P, P.L, B.G. and Y-G.G. designed the experiments, developed the methodology, analyzed and interpreted data. A.C-D, A.O, F.P, V.M. performed molecular biology procedures. S.M. performed histological procedures. C.A. and T.D-D. participated to the generation of mTORmKO mice, mouse breeding and animal welfare management. A.O. and P.L. performed immunofluorescence staining. A.L. and V.B. performed measurements of minor and major elements. L.S. supervised the work and secured funding. Y-G.G. wrote the paper with input from all authors.

**Funding** This work was supported by the Association Française contre les Myopathies (AFM) through MyoNeurALP strategic alliance, the INSERM, the CNRS, and the University of Lyon.

**Data availability** All analyzed data during this study are included in this published article and its supplementary information files.

## Declarations

**Ethics approval** Experimental protocols involving animals were performed in accordance with national and European legislation on animal experimentation and were approved by the local animal ethics committee (Apafis#18195; Ceccapp-ENS_2013_013; Ceccapp-ENS_2011_030).

**Consent to participate** Not applicable.

**Consent for publication** Not applicable.

**Conflict of interest** The authors declare no conflict of interest.

**Open Access** This article is licensed under a Creative Commons Attribution-NonCommercial-NoDerivatives 4.0 International License, which permits any non-commercial use, sharing, distribution and reproduction in any medium or format, as long as you give appropriate credit to the original author(s) and the source, provide a link to the Creative Commons licence, and indicate if you modified the licensed material. You do not have permission under this licence to share adapted material derived from this article or parts of it. The images or other third party material in this article are included in the article's Creative Commons licence, unless indicated otherwise in a credit line to the material. If material is not included in the article's Creative Commons licence and your intended use is not permitted by statutory regulation or exceeds the permitted use, you will need to obtain permission directly from the copyright holder. To view a copy of this licence, visit http://creativecommons.org/licenses/by-nc-nd/4.0/.

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