Research ArticleInter-Organ Communication in Homeostasis and Disease

Selenoprotein P deficiency protects against immobilization-induced muscle atrophy by suppressing atrophy-related E3 ubiquitin ligases

Published Online:https://doi.org/10.1152/ajpendo.00270.2022

Abstract

The quality of skeletal muscle is maintained by a balance between protein biosynthesis and degradation. Disruption in this balance results in sarcopenia. However, its underlying mechanisms remain underinvestigated. Selenoprotein P (SeP; encoded by Selenop in mice) is a hepatokine that is upregulated in type 2 diabetes and aging and causes signal resistances via reductive stress. We created immobilized muscle atrophy model in Selenop knockout (KO) mice. Immobilization (IMM) significantly reduced cross-sectional areas and the size of skeletal muscle fibers, which were ameliorated in KO mice. IMM upregulated the genes encoding E3 ubiquitin ligases and their upstream FoxO1, FoxO3, and KLF15 transcription factors in the skeletal muscle, which were suppressed in KO mice. These findings suggest a possible involvement of SeP-mediated reductive stress in physical inactivity-mediated sarcopenia, which may be a therapeutic target against sarcopenia.

NEW & NOTEWORTHY Selenoprotein P (SeP) is a hepatokine that is upregulated in type 2 diabetes and aging and causes signal resistances via reductive stress. Immobilization (IMM) significantly reduced skeletal muscle mass in mice, which was prevented in SeP knockout (KO) mice. IMM-induced Foxos/KLF15-atrogene upregulation was suppressed in the skeletal muscle of KO mice. These findings suggest that SeP-mediated reductive stress is involved in and may be a therapeutic target for physical inactivity-mediated muscle atrophy.

INTRODUCTION

The skeletal muscle is one of the primary organs necessary to maintain energy homeostasis. The mass and functional capacity of the skeletal muscle are tightly regulated by physical activity, nutrient supply, and hormonal influence (13). Aging and physical inactivity reduce skeletal muscle mass and function, a condition termed sarcopenia. Sarcopenia is associated with physical disability, declining quality of life, and death (4, 5). Specifically, diabetes is a cause and consequence of sarcopenia (6); insulinopenia and muscle insulin resistance in diabetes accelerate sarcopenia, whereas reduced glucose transport and energy expenditure perturb glucose homeostasis. Albeit, how diabetes accelerates sarcopenia remains underinvestigated.

Muscle mass maintenance is balanced by protein biosynthesis and protein degradation (79). Physical activity is necessary for maintaining muscle mass in adults, and increased work can cause fiber hypertrophy (10). Conversely, disuse or denervation causes rapid atrophy (1113). These systemic catabolic states accompany transcriptional changes in genes, termed atrophy-related genes, which are involved in protein degradation (14), leading to the activation of the ubiquitin-proteasome system that catalyzes the bulk of muscle proteins, especially the myofibrillar components (1518).

Selenoprotein P (SeP, encoded by Selenop in mice) is a hepatokine that is upregulated in diabetes (19) and causes the pathology of diabetes and its complications, such as insulin resistance (20), insulin secretory failure (21), impaired vasculogenesis (22), and cardiac ischemia-reperfusion injury (23). In addition to diabetes, SeP is elevated in starvation (24), aging (25), and pulmonary arterial hypertension (26). In the skeletal muscle, SeP causes exercise and insulin resistance by inactivating the exercise-mediated AMP-activated protein kinase (AMPK; 27). Insulin resistance causes muscle protein degradation via suppressing PI3K/Akt signaling leading to activation of the ubiquitin-proteasome proteolytic pathway, and the lack of insulin receptors affects muscle growth and metabolism (2830). However, hitherto, the relationship between SeP and skeletal muscle mass has not been established; hence, this is the focus of this study. We investigated the significance of SeP in the mice immobilization (IMM)-mediated muscle atrophy model.

MATERIALS AND METHODS

Animal Care

The mouse studies were performed according to the Guidelines on the Care and Use of Laboratory Animals issued by Kanazawa University. The ethical committee approved the protocol of Kanazawa University (Approval No. AP183978). Selenop-KO mice were produced by homologous recombination using genomic DNA cloned from an Sv-129 P1 library, as previously described (31). All mice used here had a C57BL/6J genetic background; they were male and aged 10 wk. Mice were housed in a 12-h light/dark cycle and allowed free access to food and water.

Immobilization Procedure

The immobilization was conducted as previously reported (13). First, mice were anesthetized, and then both hindlimbs were immobilized with a 1.5-mL test tube and fixed with an adhesive bandage. The animals were free to move and ate and drank ad libitum. The immobilization procedure prevented the movement of the immobilized legs alone. After 3 and 14 days, the animals were euthanized, and the tibialis anterior (TA) and plantaris muscles were removed from both hindlimbs. A nonimmobilized group was used as a control.

RNA Isolation, cDNA Synthesis, and Real-Time PCR Analysis

Total RNA was extracted from the plantaris and TA muscles using RNeasy Fibrous Tissue Mini RNA Isolation Kit (QIAGEN, Cat. No. 74704) according to the manufacturer’s protocol. The reverse transcription of RNA was performed using a High-Capacity cDNA Reverse Transcription Kit (Thermo Fisher Scientific). Quantitative RT-PCR was performed using TaqMan probes (Actb, 4352340E; Gapdh, 4351309; Myh1, Mm01332489_m1; Myh2, Mm01332564_m1; Myh4, Mm01332518_m1; Myh7, Mm01319006_g1; Atrogin-1, Mm00499523_m1; Murf1, Mm01185221_m1; Foxo1, Mm00490671_m1; Foxo3, Mm01185722_m1) and StepOnePlus Real-Time PCR System (Life Technologies). The PCR conditions were set for one cycle at 50°C for 2 min and 95°C for 10 min, followed by 40 cycles at 95°C for 15 s and 60°C for 1 min.

Western Blotting

Muscles were collected and weighed. After that, they were immediately frozen with liquid nitrogen and stored at −80°C. Muscles were homogenized by tissue rupture in 1 mL of radioimmunoprecipitation assay (RIPA) buffer that contains protease and phosphatase inhibitors. Samples were rotated at 4°C for 2 h and centrifuged at 15,000 rpm for 30 min at 4°C. Supernatants were collected and stored at −80°C. Protein samples were subjected to SDS-PAGE and transferred to PVDF membranes using the iBlot Gel Transfer System (Life Technologies) or Immobilon-P transfer membrane. The membranes were blocked in PVDF blocking reagent for Can Get Signal (Toyobo Co. Ltd., Osaka, Japan) for 1 h at room temperature. Thereafter, the membranes were incubated with specific primary antibodies, washed, and then incubated with the secondary antibodies. Bands were visualized with the ECL Prime Western Blotting Detection System (GE Healthcare UK Ltd., Amersham BioSciences Place, Little Chalfont, UK) and LAS-3000 (Fujifilm, Tokyo, Japan), and ChemiDoc Touch Imaging System (Bio-Rad). The antibodies against phospho-p70S6K (T389; Cat. No. 9234, used at 1:1,000), p70S6K (Cat. No. 9202, used at 1:1,000), phospho-Akt (Cat. No. 9271, used at 1:1,000), Akt (Cat. No. 9272, used at 1:1,000), and HRP-linked rabbit (Cat. No. 7074, used at 1:1,000) were purchased from Cell Signaling Technology (Beverly, MA). The antibody against GAPDH (Cat. No. ab181602, 1:10,000) was purchased from Abcam (Cambridge, MA). The antibody against ubiquitinated protein (Cat. No.10201-2-AP, used at 1:1,000) was purchased from Proteintech.

Muscle Histology

Muscles were embedded in an optimum cutting temperature (OCT) solution (Tissue Tek), frozen in isopentane, cooled with liquid nitrogen, and stored at −80°C until analysis. Muscle cryosections (10 μm) were collected and stained with hematoxylin/eosin (H/E). Digital images were acquired using the OLYMPUS inverted research microscope. Fiber-type distribution and cross-sectional area were quantified using Image J software.

ELISA

After blood samples were taken, they were centrifuged at 8,000 g for 10 min at 4°C and serum samples were collected. Serum levels of SeP were measured as previously described (32). Serum levels of insulin were measured using Morinaga Ultra Sensitive Mouse/Rat Insulin ELISA Kit (Morinaga, Japan) according to the manufacturer’s protocol.

Statistical Analyses

All data were analyzed using the GraphPad Prism 8.2.0 version. Data involving more than two groups were assessed by analysis of variance.

RESULTS

General Characteristics of Selenop KO and WT Mice with and without IMM

To investigate whether Selenop deficiency affects muscle mass, we induced muscle atrophy by immobilizing the hindlimbs of mice. We distributed the mice into four groups: wildtype (WT) non-IMM, WT IMM, Selenop-knockout (KO) non-IMM, and Selenop-KO IMM. IMM for 3 days significantly reduced blood glucose levels and body weight of both WT and KO mice, with no significant difference between WT and KO mice (Fig. 1, A and B). IMM did not alter insulin levels in WT mice, whereas it reduced insulin levels in KO mice (Fig. 1C). Similarly, IMM did not alter SeP levels, which were reduced in KO mice (Fig. 1D). IMM reduced the weight of quadriceps, gastrocnemius, tibialis anterior (TA), and plantaris muscles in WT mice but not in KO mice (Fig. 1, EH). IMM did not affect extensor digitorum longus, and there was no significant difference between genotypes in soleus muscle (Fig. 1, I and J).

Figure 1.

Figure 1.General characteristics of Selenop-knockout (KO) and wild-type (WT) mice with and without IMM. The 10-wk-old male mice were immobilized for 3 days. Changes in casual blood glucose levels (A), bodyweight (B), serum levels of insulin (C), and serum levels of selenoprotein P (SeP; D). Quadriceps (E), gastrocnemius (F), tibialis anterior (G), plantaris (H), extensor digitorum longus (EDL; I), and soleus (J) weight after 3 days of IMM. n = 6 for each group. Data were analyzed by one-way ANOVA with post hoc Tukey’s test. *P < 0.05; **P < 0.01; ***P < 0.001; ****P < 0.0001. EDL, extensor digitorum longus; IMM, immobilization.


Muscle Fiber Size of Selenop-KO and WT Mice with and without IMM

Subsequently, we analyzed the muscle tissue in detail and found that IMM significantly reduced the TA cross-sectional area in WT and KO mice (Fig. 2A). IMM-induced reduction in the mean cross-sectional area was significantly preserved in IMM KO mice compared with that in WT mice (Fig. 2B). As shown in Fig. 2C, myofiber size distribution analyses confirmed that IMM shifted myofiber size to smaller, which was protected in KO mice. These results suggest that SeP accelerates IMM-induced sarcopenia.

Figure 2.

Figure 2.Muscle fiber size in the skeletal muscle of WT and Selenop-KO mice after IMM. Hematoxylin and eosin staining of the tibialis anterior muscle from 14-day IMM mice (A) for the calculation of mean cross-sectional area (B) and fiber distribution (C). Approximately 200 sections were measured for each sample. n = 3 for each group. The scale bar represents 200 µm. Data represent means ± SE (error bars). Data were analyzed by one-way ANOVA with post hoc Tukey’s test. *P < 0.05; **P < 0.01; ***P < 0.001. CSA, cross-sectional area; IMM, immobilization; KO, knockout; WT, wild-type.


Myosin Heavy Chain Expression during IMM in WT and Selenop-KO Mice

We extracted mRNA from the plantaris and TA muscles of 3-day IMM mice and examined the expression of the genes encoding myosin heavy chains. Essentially, there were no significant differences in the expression levels of myosin heavy chain genes between the genotypes (Fig. 3, AF), suggesting that Selenop deficiency does not affect muscle fiber switch.

Figure 3.

Figure 3.Expression of myosin heavy chains in the skeletal muscle of WT mice and Selenop-KO mice after IMM. Expression of genes encoding myosin heavy chains, Myh1, Myh4, and Myh7, in the plantaris (AC) and tibialis anterior (DF) muscles. n = 6 for each group. Data were analyzed by one-way ANOVA with post hoc Tukey’s test. ****P < 0.0001. KO, knockout; IMM, immobilization; TA, tibialis anterior; WT, wild-type.


Atrogene Expression during IMM in WT and Selenop-KO Mice

To further investigate the molecular mechanism underlying how IMM-induced muscle fiber atrophy was ameliorated in KO mice, we investigated protein degradation and protein biosynthesis. There was no significant difference in p70S6K, which is located downstream of the mechanistic target of rapamycin complex (mTORC) in the protein biosynthetic pathway (Fig. 4, A and B). Immobilization significantly elevated Akt phosphorylation, upstream of p70S6K, in the TA skeletal muscles of WT mice, but not in those of KO mice (Fig. 4, A and C).

Figure 4.

Figure 4.Protein levels in the skeletal muscles of WT and Selenop-KO mice after IMM. Representative Western blot data of indicated protein of tibialis anterior (A) and quantification data (B and C). n = 6 for WT non-IMM, n = 6 for WT IMM, n = 5 for Selenop-KO, and n = 6 for Selenop-KO IMM. Data were analyzed by one-way ANOVA with post hoc Tukey’s test. ***P < 0.001. KO, knockout; IMM, immobilization; WT, wild-type.


IMM upregulated the E3 ubiquitin ligase atrogenes, Trim63 (Fig. 5, A and C) and Fbxo32 (Fig. 5, B and D), encoding MuRF-1 and atrogin-1, respectively, in the plantaris (Fig. 5, A and B) and TA (Fig. 5, C and D) muscles of WT mice, which were suppressed in KO mice (Fig. 5, AD). IMM elevated ubiquitinated protein in the TA muscle of WT mice but not in those of KO mice (Fig. 5, E and F).

Figure 5.

Figure 5.Expression of atrophy-related genes in the skeletal muscle of WT mice and Selenop-KO mice after IMM. Expression of the Trim63 (A and C) and Fbxo32 (B and D) in the plantaris (A and B) and tibialis anterior (C and D) muscles from 3-day IMM mice. n = 6 for each group. Representative Western blot data of indicated protein of tibialis anterior (E) and quantification data (F). n = 6 for WT non-IMM, n = 6 for WT IMM, n = 5 for Selenop-KO, and n = 6 for Selenop-KO IMM. Data were analyzed by one-way ANOVA with post hoc Tukey’s test. *P < 0.05; **P < 0.01; ***P < 0.001; ****P < 0.0001. KO, knockout; IMM, immobilization; TA, tibialis anterior; Ub, ubiquitinated protein; WT, wild-type.


The transcription factors, FoxOs, are the master regulators of the muscle atrophy program and govern the expression of the E3-ligase atrogenes. The transcription and subsequent activities of FoxOs are controlled at multiple levels (15, 17, 33). IMM upregulated Foxo1 and Foxo3 in the plantaris muscle (Fig. 6, A and B) and TA muscle (Fig. 6, C and D), which were prevented in KO mice.

Figure 6.

Figure 6.Expression of Foxos, Klf15, and Il6 in the skeletal muscle of WT mice and Selenop-KO mice. Expression of the Foxo1 (A and C) and Foxo3 (B and D) and Klf15 (E and G) and Il6 (F and H) in the plantaris (A, B, E, and F) and tibialis anterior muscles (C, D, G, and H) from 3-day IMM mice. n = 6 for each group. Data were analyzed by one-way ANOVA with post hoc Tukey’s test. *P < 0.05; **P < 0.01; ***P < 0.001; ****P < 0.0001. KO, knockout; IMM, immobilization; TA, tibialis anterior; WT, wild-type.


The previous study shows that IMM causes muscle atrophy by upregulating Klf15 and its downstream Il6 (32). Consistently, IMM also upregulated the expression of Klf15 and Il6 in the plantaris muscle (Fig. 6, E and F) and TA muscle (Fig. 6, G and H) of WT mice, which were prevented in KO mice.

DISCUSSION

This study demonstrates that deficiency of hepatokine SeP protects against IMM-induced skeletal muscle atrophy by downregulating the transcription factors, Foxo1 and Foxo3, and their downstream E3-ubiquitin ligase atrogenes. Remarkably, immobilization for 3 days significantly reduced the hindlimb muscle weights in WT mice, which was significantly ameliorated in KO mice (Fig. 1, EH). After IMM, the cross-sectional areas of the TA muscle were less profoundly reduced in KO mice than in WT mice (Fig. 2, AC).

Hornberger et al. reported that transgenic mice expressing a dominantly acting mutant form of selenocysteine transfer RNA that globally reduces selenoproteins are elevated in the plantaris muscle mass compared with WT mice after synergist ablation, a model of exercise overload. The authors speculate that elevated phosphorylation of Akt–S6K at baseline, but not after synergist ablation may account for the enhanced response to synergist ablation in mice with globally reduced selenoproteins compared with WT mice (34). However, which selenoprotein is associated with skeletal muscle hypertrophy has not been characterized. In addition, the involvement of the protein degradation pathways deserves to be examined in the sarcopenia model. This study indicates that, among the selenoproteins, deficiency of SeP is sufficient to ameliorate IMM-mediated sarcopenia by inhibiting the protein degradation pathways. Given that it is overproduced from the liver in diabetes, SeP may be a therapeutic target against diabetes-associated sarcopenia.

We examined the molecular mechanisms underlying SeP-mediated skeletal muscle atrophy. Previously, we reported that the deficiency of SeP enhances exercise response in mice through the upregulation in the levels of reactive oxygen species (ROS) and the activities of AMPK in the skeletal muscles. The skeletal muscle shifts to the type 1 slow-twitch fiber in Selenop KO mice after chronic exercise (27). In this study, the fiber composition was not altered between the genotypes after IMM (Fig. 3), which is consistent with our previous findings indicating no fiber switch before exercise in Selenop KO mice (27).

The Akt/mTORC pathway is one of the crucial pathways for protein biosynthesis in the skeletal muscles (35, 36). Thus, activating the Akt/mTORC pathway prevents skeletal muscle atrophy in the rat’s tail suspension model (37). SeP impairs the insulin-induced Akt phosphorylation in the skeletal muscle (20). It is also shown that hindlimb casting-induced reduction in muscle mass is partially dependent on proteolysis but independent of protein biosynthesis (12). Consistently, our results showed that IMM did not alter the phosphorylation status of p70S6K, which is downstream of mTORC1 in WT mice. In addition, the phosphorylation status of p70S6K was not different between WT and KO after IMM (Fig. 4, A and B). Furthermore, the deficiency of SeP enhances exercise response in mice through the upregulation of ROS levels and AMPK activity in the skeletal muscles (27). AMPK negatively regulates mTORC1 and suppresses protein biosynthesis in the skeletal muscles (38). Therefore, activated AMPK-mediated mTORC1 inactivation in KO mice may neutralize the Akt-mediated activation of mTORC1-p70S6K axis. These results suggest that the protein biosynthetic pathway is not dominantly involved in the IMM-induced SeP-mediated skeletal muscle atrophy.

Next, we examined the involvement of the protein degradation pathways. MuRF1 and atrogin-1 have been well recognized as two muscle-specific E3 ubiquitin ligases that are transcriptionally upregulated in the skeletal muscles under atrophy-inducing conditions (39). In this study, IMM upregulated Trim63 and Fbxo32 encoding MuRF-1 and atrogin-1, respectively, in the skeletal muscle, which were rescued in KO mice (Fig. 5, AD). These atrophy-related E3 ligases are governed by the transcription factors FoxOs (15) along with the downstream KLF15 transcription factor (40). Specifically, activated FoxOs lead to the activation of ubiquitin ligases (28), whereas FoxOs expression is upregulated in muscle atrophy conditions (41). IMM induces skeletal muscle atrophy through the KLF15-IL-6 pathway (42). In this study, in concert with alterations in the atrogenes, Foxo1, Foxo3, Klf15, and Il6 were upregulated in the skeletal muscle after IMM in WT mice; however, these were suppressed in KO mice (Fig. 6). These results suggest that deficiency of SeP protects against the IMM-induced sarcopenia, at least partly, via the Foxos/Klf15-atrogene pathway.

Although the involvement of oxidative stress in the disuse muscle atrophy is still under debate, some studies have shown that oxidative stress occurs due to disuse (43). Importantly, disuse induces oxidative stress and upregulates glutathione peroxidase 1 (4446), whereas antioxidants protect mice against IMM-induced muscle mass reduction (4749). Excess ROS damages the skeletal muscles; ROS reduces muscle mass by stimulating the ubiquitin-conjugating activity and upregulating atrogene expression in C2C12 myotubes (44, 50). Currently, it remains unclear how IMM upregulates the Foxos/Klf15-atrogene pathway. In addition, ROS induces FoxO1 nuclear translocation (51) and also causes insulin resistance by impairing the insulin/Akt signaling pathway in the skeletal muscles (19, 5254), which further activates FoxOs. SeP functions as an antioxidant protein by its intrinsic thioredoxin domain and by distributing selenium to intracellular glutathione peroxidases (55, 56), which may suppress ROS-induced FoxO activation. In addition, SeP inactivates the exercise-mediated activation of AMPK, which activates histone deacetylase SIRT1; SIRT1 deacetylates and activates FoxO (57). Therefore, theoretically, it is possible that the deficiency of SeP further elevates the IMM-induced ROS, activates FoxOs, and worsens muscle atrophy. However, in this study, the Selenop-KO mice were protected against IMM-induced muscle atrophy. These findings suggest that reductive stress is involved in IMM-induced muscle atrophy. The unknown reductive targets of SeP that upregulate the atrophy-related genes should be identified in the future. Alternatively, Selenop mRNA may modify the target protein functions and upregulate the atrophy-related genes as we showed that Selenop mRNA negatively regulates retinoic acid-inducible gene I (RIG-I) functions, leading to impaired innate immunity (58).

In summary, deficiency of SeP protects against reduced skeletal muscle weights and fiber size induced by IMM through the suppression of atrophy-related genes and their key transcription factors, Foxo1, Foxo3, and Klf15. The current study suggests a possible involvement of reductive stress in the physical inactivity-mediated sarcopenia. Therefore, SeP may be a therapeutic target against sarcopenia in patients with type 2 diabetes.

DATA AVAILABILITY

Data will be made available upon reasonable request.

GRANTS

This work was supported, in part, by Grants-in-Aid from the Ministry of Education, Culture, Sports, Science and Technology (17H04199 and 18K19560) and Takeda Science Foundation (to T.T.).

DISCLOSURES

No conflicts of interest, financial or otherwise, are declared by the authors.

AUTHOR CONTRIBUTIONS

H.A., K.K., and T.T. conceived and designed research; H.A., K.K., K.T., T.A., L.Q., H.T., and K.H. performed experiments; H.A., K.K., K.T., K.H., and T.T. analyzed data; H.A., K.K., K-A.I., Y.I., H.G., Y.N., Y.T., H.T., and T.T. interpreted results of experiments; H.A., K.K., and T.T prepared figures; H.A., K.K., K-A.I., and T.T. drafted manuscript; K.K., H.T., and T.T. edited and revised manuscript; H.A., K.K., K-A.I., K.T., T.A., L.Q., Y.I., H.G., Y.N., Y.T., H.T., K.H., and T.T. approved final version of manuscript.

ACKNOWLEDGMENTS

We thank Dr. Yu Hirata of Kobe University for technical advice on immobilization. We thank Maki Kawamura of Kanazawa University for technical support.

REFERENCES

  • 1. Sishi B, Loos B, Ellis B, Smith W, Du Toit EF, Engelbrecht AM. Diet-induced obesity alters signalling pathways and induces atrophy and apoptosis in skeletal muscle in a prediabetic rat model. Exp Physiol 96: 179–193, 2011. doi:10.1113/expphysiol.2010.054189.
    Crossref | PubMed | Web of Science | Google Scholar
  • 2. Braun TP, Marks DL. The regulation of muscle mass by endogenous glucocorticoids. Front Physiol 6: 1–12, 2015. doi:10.3389/fphys.2015.00012.
    Crossref | PubMed | Web of Science | Google Scholar
  • 3. Theilen NT, Jeremic N, Weber GJ, Tyagi SC. Exercise preconditioning diminishes skeletal muscle atrophy after hindlimb suspension in mice. J Appl Physiol (1985) 125: 999–1010, 2018. doi:10.1152/japplphysiol.00137.2018.
    Link | Web of Science | Google Scholar
  • 4. Cruz-Jentoft AJ, Sayer AA. Sarcopenia. Lancet 393: 2636–2646, 2019 [Erratum in Lancet 393: 2590, 2019]. doi:10.1016/S0140-6736(19)31138-9.
    Crossref | PubMed | Web of Science | Google Scholar
  • 5. Goodpaster BH, Park SW, Harris TB, Kritchevsky SB, Nevitt M, Schwartz AV, Simonsick EM, Tylavsky FA, Visser M, Newman AB. The loss of skeletal muscle strength, mass, and quality in older adults: the health, aging and body composition study. J Gerontol A Biol Sci Med Sci 61: 1059–1064, 2006. doi:10.1093/GERONA/61.10.1059.
    Crossref | PubMed | Web of Science | Google Scholar
  • 6. Mesinovic J, Zengin A, De Courten B, Ebeling PR, Scott D. Sarcopenia and type 2 diabetes mellitus: a bidirectional relationship. Diabetes Metab Syndr Obes 12: 1057–1072, 2019. doi:10.2147/DMSO.S186600.
    Crossref | PubMed | Web of Science | Google Scholar
  • 7. Piccirillo R, Demontis F, Perrimon N, Goldberg AL. Mechanisms of muscle growth and atrophy in mammals and Drosophila. Dev Dyn 243: 201–215, 2014. doi:10.1002/dvdy.24036.
    Crossref | PubMed | Web of Science | Google Scholar
  • 8. Egerman MA, Glass DJ. Signaling pathways controlling skeletal muscle mass. Crit Rev Biochem Mol Biol 49: 59–68, 2014. doi:10.3109/10409238.2013.857291.
    Crossref | PubMed | Web of Science | Google Scholar
  • 9. Schiaffino S, Dyar KA, Ciciliot S, Blaauw B, Sandri M. Mechanisms regulating skeletal muscle growth and atrophy. FEBS J 280: 4294–4314, 2013. doi:10.1111/febs.12253.
    Crossref | PubMed | Web of Science | Google Scholar
  • 10. Thomson DM. The role of AMPK in the regulation of skeletal muscle size, hypertrophy, and regeneration. Int J Mol Sci 19: 3125, 2018. doi:10.3390/ijms19103125.
    Crossref | PubMed | Web of Science | Google Scholar
  • 11. Jackman RW, Kandarian SC. The molecular basis of skeletal muscle atrophy. Am J Physiol Cell Physiol 287: C834–C843, 2004. doi:10.1152/ajpcell.00579.2003.
    Link | Web of Science | Google Scholar
  • 12. Krawiec BJ, Frost RA, Vary TC, Jefferson LS, Lang CH. Hindlimb casting decreases muscle mass in part by proteasome-dependent proteolysis but independent of protein synthesis. Am J Physiol Endocrinol Metab 289: E969–E980, 2005. doi:10.1152/ajpendo.00126.2005.
    Link | Web of Science | Google Scholar
  • 13. You JS, Anderson GB, Dooley MS, Hornberger TA. The role of mTOR signaling in the regulation of protein synthesis and muscle mass during immobilization in mice. Dis Model Mech 8: 1059–1069, 2015. doi:10.1242/dmm.019414.
    Crossref | PubMed | Web of Science | Google Scholar
  • 14. Gomes MD, Lecker SH, Jagoe RT, Navon A, Goldberg AL. Atrogin-1, a muscle-specific F-box protein highly expressed during muscle atrophy. Proc Natl Acad Sci USA 98: 14440–14445, 2001. doi:10.1073/pnas.251541198.
    Crossref | PubMed | Web of Science | Google Scholar
  • 15. Sandri M, Sandri C, Gilbert A, Skurk C, Calabria E, Picard A, Walsh K, Schiaffino S, Lecker SH, Goldberg AL. Foxo transcription factors induce the atrophy-related ubiquitin ligase atrogin-1 and cause skeletal muscle atrophy. Cell 117: 399–412, 2004. doi:10.1016/S0092-8674(04)00400-3.
    Crossref | PubMed | Web of Science | Google Scholar
  • 16. Bonaldo P, Sandri M. Cellular and molecular mechanisms of muscle atrophy. Dis Model Mech 6: 25–39, 2013. doi:10.1242/dmm.010389.
    Crossref | PubMed | Web of Science | Google Scholar
  • 17. Brocca L, Toniolo L, Reggiani C, Bottinelli R, Sandri M, Pellegrino MA. FoxO-dependent atrogenes vary among catabolic conditions and play a key role in muscle atrophy induced by hindlimb suspension. J Physiol 595: 1143–1158, 2017. doi:10.1113/JP273097.
    Crossref | PubMed | Web of Science | Google Scholar
  • 18. Ji LL, Yeo D. Cellular mechanism of immobilization-induced muscle atrophy: a mini review. Sports Med Health Sci 1: 19–23, 2019. doi:10.1016/j.smhs.2019.08.004.
    Crossref | PubMed | Google Scholar
  • 19. Takamura T. Hepatokine selenoprotein P-mediated reductive stress causes resistance to intracellular signal transduction. Antioxid Redox Signal 33: 517–524, 2020. doi:10.1089/ars.2020.8087.
    Crossref | PubMed | Web of Science | Google Scholar
  • 20. Misu H, Takamura T, Takayama H, Hayashi H, Matsuzawa-Nagata N, Kurita S, Ishikura K, Ando H, Takeshita Y, Ota T, Sakurai M, Yamashita T, Mizukoshi E, Yamashita T, Honda M, Miyamoto K, Kubota T, Kubota N, Kadowaki T, Kim HJ, Lee IK, Minokoshi Y, Saito Y, Takahashi K, Yamada Y, Takakura N, Kaneko S. A liver-derived secretory protein, selenoprotein P, causes insulin resistance. Cell Metab 12: 483–495, 2010. doi:10.1016/j.cmet.2010.09.015.
    Crossref | PubMed | Web of Science | Google Scholar
  • 21. Mita Y, Nakayama K, Inari S, Nishito Y, Yoshioka Y, Sakai N, Sotani K, Nagamura T, Kuzuhara Y, Inagaki K, Iwasaki M, Misu H, Ikegawa M, Takamura T, Noguchi N, Saito Y. Selenoprotein P-neutralizing antibodies improve insulin secretion and glucose sensitivity in type 2 diabetes mouse models. Nat Commun 8: 1658, 2017. doi:10.1038/s41467-017-01863-z.
    Crossref | PubMed | Web of Science | Google Scholar
  • 22. Ishikura K, Misu H, Kumazaki M, Takayama H, Matsuzawa-Nagata N, Tajima N, Chikamoto K, Lan F, Ando H, Ota T, Sakurai M, Takeshita Y, Kato K, Fujimura A, Miyamoto K, Saito Y, Kameo S, Okamoto Y, Takuwa Y, Takahashi K, Kidoya H, Takakura N, Kaneko S, Takamura T. Selenoprotein P as a diabetes-associated hepatokine that impairs angiogenesis by inducing VEGF resistance in vascular endothelial cells. Diabetologia 57: 1968–1976, 2014. doi:10.1007/s00125-014-3306-9.
    Crossref | PubMed | Web of Science | Google Scholar
  • 23. Chadani H, Usui S, Inoue O, Kusayama T, Takashima SI, Kato T, Murai H, Furusho H, Nomura A, Misu H, Takamura T, Kaneko S, Takamura M. Endogenous selenoprotein P, a liver-derived secretory protein, mediates myocardial ischemia/reperfusion injury in mice. Int J Mol Sci 19: 878, 2018. doi:10.3390/ijms19030878.
    Crossref | PubMed | Web of Science | Google Scholar
  • 24. Takayama H, Misu H, Iwama H, Chikamoto K, Saito Y, Murao K, Teraguchi A, Lan F, Kikuchi A, Saito R, Tajima N, Shirasaki T, Matsugo S, Miyamoto K, Kaneko S, Takamura T. Metformin suppresses expression of the selenoprotein P gene via an AMP-activated kinase (AMPK)/FoxO3a pathway in H4IIEC3 hepatocytes. J Biol Chem 289: 335–345, 2014. doi:10.1074/jbc.M113.479386.
    Crossref | PubMed | Web of Science | Google Scholar
  • 25. Oo SM, Misu H, Saito Y, Tanaka M, Kato S, Kita Y, Takayama H, Takeshita Y, Kanamori T, Nagano T, Nakagen M, Urabe T, Matsuyama N, Kaneko S, Takamura T. Serum selenoprotein P, but not selenium, predicts future hyperglycemia in a general Japanese population. Sci Rep 8: 16727, 2018. doi:10.1038/s41598-018-35067-2.
    Crossref | PubMed | Web of Science | Google Scholar
  • 26. Kikuchi N, Satoh K, Kurosawa R, Yaoita N, Elias-Al-Mamun M, Siddique MAH, Omura J, Satoh T, Nogi M, Sunamura S, Miyata S, Saito Y, Hoshikawa Y, Okada Y, Shimokawa H. Selenoprotein P promotes the development of pulmonary arterial hypertension: Possible novel therapeutic target. Circulation 138: 600–623, 2018. doi:10.1161/CIRCULATIONAHA.117.033113.
    Crossref | PubMed | Web of Science | Google Scholar
  • 27. Misu H, Takayama H, Saito Y, Mita Y, Kikuchi A, Ishii KA, Chikamoto K, Kanamori T, Tajima N, Lan F, Takeshita Y, Honda M, Tanaka M, Kato S, Matsuyama N, Yoshioka Y, Iwayama K, Tokuyama K, Akazawa N, Maeda S, Takekoshi K, Matsugo S, Noguchi N, Kaneko S, Takamura T. Deficiency of the hepatokine selenoprotein P increases responsiveness to exercise in mice through upregulation of reactive oxygen species and AMP-activated protein kinase in muscle. Nat Med 23: 508–516, 2017. doi:10.1038/nm.4295.
    Crossref | PubMed | Web of Science | Google Scholar
  • 28. Wang X, Hu Z, Hu J, Du J, Mitch WE. Insulin resistance accelerates muscle protein degradation: activation of the ubiquitin-proteasome pathway by defects in muscle cell signaling. Endocrinology 147: 4160–4168, 2006. doi:10.1210/EN.2006-0251.
    Crossref | PubMed | Web of Science | Google Scholar
  • 29. O'Neill ED, Wilding JP, Kahn CR, Van Remmen H, McArdle A, Jackson MJ, Close GL. Absence of insulin signalling in skeletal muscle is associated with reduced muscle mass and function: evidence for decreased protein synthesis and not increased degradation. Age (Dordr) 32: 209–222, 2010. doi:10.1007/S11357-009-9125-0.
    Crossref | PubMed | Google Scholar
  • 30. Long YC, Cheng Z, Copps KD, White MF. Insulin receptor substrates Irs1 and Irs2 coordinate skeletal muscle growth and metabolism via the Akt and AMPK pathways. Mol Cell Biol 31: 430–441, 2011 [Erratum in Mol Cell Biol 37: e00232-17, 2017]. doi:10.1128/MCB.00983-10.
    Crossref | PubMed | Web of Science | Google Scholar
  • 31. Hill KE, Zhou J, McMahan WJ, Motley AK, Atkins JF, Gesteland RF, Burk RF. Deletion of selenoprotein P alters distribution of selenium in the mouse. J Biol Chem 278: 13640–13646, 2003. doi:10.1074/JBC.M300755200.
    Crossref | PubMed | Web of Science | Google Scholar
  • 32. Oo SM, Oo HK, Takayama H, Ishii KA, Takeshita Y, Goto H, Nakano Y, Kohno S, Takahashi C, Nakamura H, Saito Y, Matsushita M, Okamatsu-Ogura Y, Saito M, Takamura T. Selenoprotein P-mediated reductive stress impairs cold-induced thermogenesis in brown fat. Cell Rep 38: 110566, 2022. doi:10.1016/j.celrep.2022.110566.
    Crossref | PubMed | Web of Science | Google Scholar
  • 33. Bodine SC, Latres E, Baumhueter S, Lai VK, Nunez L, Clarke BA, Poueymirou WT, Panaro FJ, Na E, Dharmarajan K, Pan ZQ, Valenzuela DM, Dechiara TM, Stitt TN, Yancopoulos GD, Glass DJ. Identification of ubiquitin ligases required for skeletal muscle atrophy. Science 294: 1704–1708, 2001. doi:10.1126/science.1065874.
    Crossref | PubMed | Web of Science | Google Scholar
  • 34. Hornberger TA, McLoughlin TJ, Leszczynski JK, Armstrong DD, Jameson RR, Bowen PE, Hwang ES, Hou H, Moustafa ME, Carlson BA, Hatfield DL, Diamond AM, Esser KA. Selenoprotein-deficient transgenic mice exhibit enhanced exercise-induced muscle growth. J Nutr 133: 3091–3097, 2003. doi:10.1093/jn/133.10.3091.
    Crossref | PubMed | Web of Science | Google Scholar
  • 35. Mirzoev T, Tyganov S, Vilchinskaya N, Lomonosova Y, Shenkman B. Key markers of mTORC1-dependent and mTORC1-independent signaling pathways regulating protein synthesis in rat soleus muscle during early stages of hindlimb unloading. Cell Physiol Biochem 39: 1011–1020, 2016. doi:10.1159/000447808.
    Crossref | PubMed | Web of Science | Google Scholar
  • 36. Shimkus KL, Shirazi-Fard Y, Wiggs MP, Ullah ST, Pohlenz C, Gatlin DM III, Carroll CC, Hogan HA, Fluckey JD. Responses of skeletal muscle size and anabolism are reproducible with multiple periods of unloading/reloading. J Appl Physiol (1985) 125: 1456–1467, 2018. doi:10.1152/JAPPLPHYSIOL.00736.2017/ASSET/IMAGES/LARGE/ZDG0101827700009.JPEG.
    Link | Web of Science | Google Scholar
  • 37. Bodine SC, Stitt TN, Gonzalez M, Kline WO, Stover GL, Bauerlein R, Zlotchenko E, Scrimgeour A, Lawrence JC, Glass DJ, Yancopoulos GD. Akt/mTOR pathway is a crucial regulator of skeletal muscle hypertrophy and can prevent muscle atrophy in vivo. Nat Cell Biol 3: 1014–1019, 2001. doi:10.1038/ncb1101-1014.
    Crossref | PubMed | Web of Science | Google Scholar
  • 38. Bolster DR, Crozier SJ, Kimball SR, Jefferson LS. AMP-activated protein kinase suppresses protein synthesis in rat skeletal muscle through down-regulated mammalian target of rapamycin (mTOR) signaling. J Biol Chem 277: 23977–23980, 2002. doi:10.1074/jbc.C200171200.
    Crossref | PubMed | Web of Science | Google Scholar
  • 39. Komatsu R, Okazaki T, Ebihara S, Kobayashi M, Tsukita Y, Nihei M, Sugiura H, Niu K, Ebihara T, Ichinose M. Aspiration pneumonia induces muscle atrophy in the respiratory, skeletal, and swallowing systems. J Cachexia Sarcopenia Muscle 9: 643–653, 2018. doi:10.1002/JCSM.12297.
    Crossref | PubMed | Web of Science | Google Scholar
  • 40. Oyabu M, Takigawa K, Mizutani S, Hatazawa Y, Fujita M, Ohira Y, Sugimoto T, Suzuki O, Tsuchiya K, Suganami T, Ogawa Y, Ishihara K, Miura S, Kamei Y. FOXO1 cooperates with C/EBPδ and ATF4 to regulate skeletal muscle atrophy transcriptional program during fasting. FASEB J 36: e22152, 2022. doi:10.1096/fj.202101385RR.
    Crossref | PubMed | Web of Science | Google Scholar
  • 41. Kamei Y, Miura S, Suzuki M, Kai Y, Mizukami J, Taniguchi T, Mochida K, Hata T, Matsuda J, Aburatani H, Nishino I, Ezaki O. Skeletal muscle FOXO1 (FKHR) transgenic mice have less skeletal muscle mass, down-regulated type I (slow twitch/red muscle) fiber genes, and impaired glycemic control. J Biol Chem 279: 41114–41123, 2004. doi:10.1074/jbc.M400674200.
    Crossref | PubMed | Web of Science | Google Scholar
  • 42. Hirata Y, Nomura K, Kato D, Tachibana Y, Niikura T, Uchiyama K, Hosooka T, Fukui T, Oe K, Kuroda R, Hara Y, Adachi T, Shibasaki K, Wake H, Ogawa W. A Piezo1/KLF15/IL-6 axis mediates immobilization-induced muscle atrophy. J Clin Invest 132: 1–13, 2022. doi:10.1172/JCI154611.
    Crossref | PubMed | Web of Science | Google Scholar
  • 43. Pellegrino MA, Desaphy JF, Brocca L, Pierno S, Camerino DC, Bottinelli R. Redox homeostasis, oxidative stress and disuse muscle atrophy. J Physiol 589: 2147–2160, 2011. doi:10.1113/jphysiol.2010.203232.
    Crossref | PubMed | Web of Science | Google Scholar
  • 44. Powers SK, Smuder AJ, Judge AR. Oxidative stress and disuse muscle atrophy: cause or consequence? Curr Opin Clin Nutr Metab Care 15: 240–245, 2012. doi:10.1097/mco.0b013e328352b4c2.
    Crossref | PubMed | Web of Science | Google Scholar
  • 45. Kondo H, Miura M, Itokawa Y. Oxidative stress in skeletal muscle atrophied by immobilization. Acta Physiol Scand 142: 527–528, 1991. doi:10.1111/j.1748-1716.1991.tb09191.x.
    Crossref | PubMed | Google Scholar
  • 46. Powers SK, Kavazis AN, DeRuisseau KC. Mechanisms of disuse muscle atrophy: role of oxidative stress. Am J Physiol Regul Integr Comp Physiol 288: R337–R344, 2005. doi:10.1152/ajpregu.00469.2004.
    Link | Web of Science | Google Scholar
  • 47. Min K, Smuder AJ, Kwon OS, Kavazis AN, Szeto HH, Powers SK. Mitochondrial-targeted antioxidants protect skeletal muscle against immobilization-induced muscle atrophy. J Appl Physiol (1985) 111: 1459–1466, 2011. doi:10.1152/japplphysiol.00591.2011.
    Link | Web of Science | Google Scholar
  • 48. Talbert EE, Smuder AJ, Min K, Kwon OS, Szeto HH, Powers SK. Immobilization-induced activation of key proteolytic systems in skeletal muscles is prevented by a mitochondria-targeted antioxidant. J Appl Physiol (1985) 115: 529–538, 2013. doi:10.1152/japplphysiol.00471.2013.
    Link | Web of Science | Google Scholar
  • 49. Powers SK. Can antioxidants protect against disuse muscle atrophy? Sports Med 44: 155–165, 2014. doi:10.1007/s40279-014-0255-x.
    Crossref | PubMed | Web of Science | Google Scholar
  • 50. Li YP, Chen Y, Li AS, Reid MB. Hydrogen peroxide stimulates ubiquitin-conjugating activity and expression of genes for specific E2 and E3 proteins in skeletal muscle myotubes. Am J Physiol Cell Physiol 285: C806–C812, 2003. doi:10.1152/ajpcell.00129.2003.
    Link | Web of Science | Google Scholar
  • 51. Frescas D, Valenti L, Accili D. Nuclear trapping of the forkhead transcription factor FoxO1 via Sirt-dependent deacetylation promotes expression of glucogenetic genes. J Biol Chem 280: 20589–20595, 2005. doi:10.1074/jbc.M412357200.
    Crossref | PubMed | Web of Science | Google Scholar
  • 52. Houstis N, Rosen ED, Lander ES. Reactive oxygen species have a causal role in multiple forms of insulin resistance. Nature 440: 944–948, 2006. doi:10.1038/nature04634.
    Crossref | PubMed | Web of Science | Google Scholar
  • 53. Yaribeygi H, Farrokhi FR, Butler AE, Sahebkar A. Insulin resistance: review of the underlying molecular mechanisms. J Cell Physiol 234: 8152–8161, 2019. doi:10.1002/JCP.27603.
    Crossref | PubMed | Web of Science | Google Scholar
  • 54. Powers SK, Schrager M. Redox signaling regulates skeletal muscle remodeling in response to exercise and prolonged inactivity. Redox Biol 54: 102374, 2022. doi:10.1016/J.REDOX.2022.102374.
    Crossref | PubMed | Web of Science | Google Scholar
  • 55. Burk RF, Hill KE, Motley AK. Selenoprotein metabolism and function: evidence for more than one function for selenoprotein P. J Nutr 133, Suppl 1: S1517–S1520, 2003. doi:10.1093/JN/133.5.1517S.
    Crossref | PubMed | Web of Science | Google Scholar
  • 56. Zhang Y, Roh YJ, Han SJ, Park I, Lee HM, Ok YS, Lee BC, Lee SR. Role of selenoproteins in redox regulation of signaling and the antioxidant system: a review. Antioxidants (Basel) 9: 383, 2020. doi:10.3390/antiox9050383.
    Crossref | PubMed | Web of Science | Google Scholar
  • 57. Cantó C, Gerhart-Hines Z, Feige JN, Lagouge M, Noriega L, Milne JC, Elliott PJ, Puigserver P, Auwerx J. AMPK regulates energy expenditure by modulating NAD+ metabolism and SIRT1 activity. Nature 458: 1056–1060, 2009. doi:10.1038/nature07813.
    Crossref | PubMed | Web of Science | Google Scholar
  • 58. Murai K, Honda M, Shirasaki T, Shimakami T, Omura H, Misu H, Kita Y, Takeshita Y, Ishii KA, Takamura T, Urabe T, Shimizu R, Okada H, Yamashita T, Sakai Y, Kaneko S. Induction of selenoprotein P mRNA during hepatitis C virus infection inhibits RIG-I-mediated antiviral immunity. Cell Host Microbe 25: 588–601.e7, 2019. doi:10.1016/j.chom.2019.02.015.
    Crossref | PubMed | Web of Science | Google Scholar

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