Loss of muscle mass and function with age is detrimental to health and quality of life (Evans, 2010; Larsson et al., 2019). Sarcopenia, muscle atrophy and weakness with aging, is due to a combination of inactivity, injury, surgery, and biological consequences of aging (Bonaldo and Sandri, 2013; Dolbow and Gorgey, 2016). A pharmacological therapy for muscle loss does not exist, and current diet or exercise therapeutic approaches are often ineffective or unfeasible. Oxidative stress has been implicated in muscle atrophy by accelerating proteolysis (Powers et al., 2011; Scicchitano et al., 2018), but the exact mechanism by which reactive oxygen species (ROS) contributes to the decrease in muscle mass and strength is not well understood.

Lipid hydroperoxide (LOOH) is a class of ROS molecules that has been implicated in cell damage, particularly as a trigger to induce ferroptosis, a non-apoptotic form of regulated cell death (Wiernicki et al., 2020; Yang et al., 2014). Lipid peroxidation is initiated by prooxidants such as hydroxyl radicals attacking the carbon-carbon double bond in fatty acids, particularly the polyunsaturated fatty acids (PUFAs) containing phospholipids (Bochkov et al., 2010). Lipid radicals (L•) created by this reaction rapidly react with oxygen to form a lipid peroxy-radical which subsequently reacts with another lipid to produce L• and LOOH, the former propagating lipid peroxidation. LOOH is the primary product of lipid peroxidation that forms secondary reactive lipid aldehydes such as 4-hydroxynonenal (4-HNE) and malondialdehyde (MDA), inducing carbonyl stress with high reactivity against biological molecules to promote cellular toxicity. The intracellular level of LOOH is endogenously suppressed by glutathione peroxidase 4 (GPx4) that catalyzes the reaction by which LOOH is reduced to its nonreactive hydroxyl metabolite (Anderson et al., 2018).

Despite the evidence for the role of LOOH-mediated cell damage and cell death, the biological consequence of LOOH accumulation in skeletal muscle is not well understood (Bhattacharya et al., 2009; Pharaoh et al., 2020). Below, we provide evidence that LOOH mediates loss of muscle mass and function associated with sarcopenia. An increase in muscle LOOH was a common feature with aging and disuse, and accumulation of LOOH in vitro and in vivo augmented muscle atrophy. We further show that genetic or pharmacological suppression of LOOH and their reactive lipid aldehydes is sufficient to prevent disuse-induced muscle atrophy, and to a greater extent in muscle contractile function, in young and old mice.

We first evaluated the changes in skeletal muscle LOOH with aging. In humans and in mice, aging promoted a reduction in the expression of Gpx4 in skeletal muscle (Figure 1A, B). To examine the changes in skeletal muscle LOOH landscape with age, we performed a comprehensive oxidolipidomic analysis in gastrocnemius muscle samples from young (4 months) and old (20 months) mice (Figure 1C, D). We detected over 300 species of oxidized lipids with an effect distribution that was highly class-dependent. Among these, age had the most robust effect on oxidized phosphatidylethanolamine (Figure 1C, red), a class of lipids that have been implicated as a potential lipid signal to induce ferroptosis (Kagan et al., 2017). Among the top ten oxidized lipid species whose abundance had the most robust fold-increase with age, six of them were oxidized phosphatidylethanolamine species (Figure 1D). These oxidized phosphatidylethanolamine species were also substantially more highly abundant compared to other oxidized lipids. LOOH can be indirectly assessed by quantifying lipid aldehyde adducts such as 4-HNE and MDA. We confirmed increased muscle 4-HNE and MDA with age (Figure 1E–G).

Figure 1 with 1 supplement see all

Download asset Open asset

Figure 1—source data 1

Original western blots.

https://cdn.elifesciences.org/articles/85289/elife-85289-fig1-data1-v2.zip

Download elife-85289-fig1-data1-v2.zip

Disuse promoted by inactivity, injury, or surgery is a major contributor to age-associated decline in muscle mass and function. Disuse also promotes skeletal muscle atrophy that is likely contributed by ROS (Powers et al., 2011). To model disuse atrophy, mice underwent a hindlimb unloading (HU) procedure as previously described (Eshima et al., 2020; Heden et al., 2019; Figure 1—figure supplement 1A–J). As expected, HU-induced muscle atrophy and weakness (Figure 1—figure supplement 1D, E) concomitant to reduction in body and lean mass (Figure 1—figure supplement 1B, C). Disuse robustly elevated muscle LOOH levels (Figure 1—figure supplement 1F, G) without significant changes in mitochondrial ROS production (Figure 1—figure supplement 1H-J). An increase in muscle LOOH preceded atrophy (Figure 1—figure supplement 1D, F), consistent with the notion that LOOH may trigger mechanisms to promote loss of muscle mass.

Next, we tested our hypothesis that LOOH contributes to muscle atrophy using C2C12 myotubes (Figure 2A). Lentivirus-mediated knockdown (KD) of GPx4 increased LOOH and markers of ferroptosis concomitant with a decrease in myotube diameter (Figure 2B–I). We also recapitulated these findings with erastin (a system X_c^- inhibitor that suppresses glutathione synthesis) (Figure 2B, C and Figure 2—figure supplement 1A–E) and RSL3 (GPx4 inhibitor) (Figure 2B, C and Figure 2—figure supplement 1F–I), commonly used acute pharmacological interventions to elevate intracellular LOOH. These data support the idea that LOOH reduces myotube size in a cell-autonomous manner.

Figure 2 with 1 supplement see all

Download asset Open asset

Figure 2—source data 1

Original western blots.

https://cdn.elifesciences.org/articles/85289/elife-85289-fig2-data1-v2.zip

Download elife-85289-fig2-data1-v2.zip

We then translated these findings in vivo with global heterozygous GPx4 knockout mice (Gpx4^+/-). Germline deletion of GPx4 is embryonically lethal (Yant et al., 2003), but Gpx4^+/- mice appear normal and do not have an observable muscle phenotype at baseline (Anderson et al., 2018; Katunga et al., 2015). We studied 4 months (young) and 20 months (old) Gpx4^+/- and wildtype littermates with or without HU (Figure 3—figure supplement 1A–D). In young mice, GPx4 haploinsufficiency augmented the loss in soleus mass induced by HU (Figure 3A and Figure 3—figure supplement 1E). However, muscle masses between old Gpx4^+/- and wildtype mice were not different. We interpret these findings to mean that disuse in old mice promotes an increase in LOOH that has already reached a maximally effective threshold with age such that GPx4 deletion had no further effect. In support of this, we saw no differences in 4-HNE or MDA levels between old Gpx4^+/- and wildtype mice (Figure 3B and Figure 3—figure supplement 1F, G). GPx4 haploinsufficiency did not alter force-generating capacity (Figure 3C and Figure 3—figure supplement 1H–J).

Figure 3 with 2 supplements see all

Download asset Open asset

Figure 3—source data 1

Original western blots.

https://cdn.elifesciences.org/articles/85289/elife-85289-fig3-data1-v2.zip

Download elife-85289-fig3-data1-v2.zip

Because GPx4 is expressed globally, we also studied mice with skeletal muscle-specific tamoxifen-inducible GPx4 knockout (GPx4-MKO) (Figure 3D and Figure 3—figure supplement 2A; Yoo et al., 2012). Consistent with Gpx4^+/- mice, soleus muscles from GPx4-MKO mice were also more prone to developing disuse-induced atrophy (Figure 3E and F and Figure 3—figure supplement 2B–F) concomitant to elevated LOOH (Figure 3G and Figure 3—figure supplement 2G, H), suggesting that loss of GPx4 in muscle augments atrophy in a cell-autonomous manner. Histological analyses revealed that reduced muscle mass was consistent with reduced cross-sectional area (CSA) of myofibers regardless of fiber-type compositions (Figure 3H&I and Figure 3—figure supplement 2I, J). These data implicate that LOOH directly reduces muscle cell size in vivo.

GPx4 primarily neutralizes LOOH but it also exhibits some activity toward other peroxides (Imai and Nakagawa, 2003). To confirm that the effects of GPx4 deletion to promote atrophy is specific to LOOH, we diminished the ability of cells to incorporate PUFAs into phospholipids by deleting lysophosphatidylcholine acyltransferase 3 (LPCAT3) (Lee et al., 2021; Ferrara et al., 2021a; Ferrara et al., 2021b). LPCAT3 is an enzyme of Lands cycle that preferentially acylates lysophospholipids with PUFAs, and thus an essential component of ferroptosis (Kagan et al., 2017). Indeed, LPCAT3 KD rescued the increase in 4-HNE induced by GPx4 KD (Figure 4A–C and Figure 4—figure supplement 1A). Remarkably, deletion of LPCAT3 KD completely restored the reduction in myotube diameter induced by GPx4 KD (Figure 4D&E). Similarly, LPCAT3 deletion also prevented LOOH and cell death induced by erastin (Figure 4F–H and Figure 4—figure supplement 1B, C). These findings indicate that muscle atrophy induced by loss of GPx4 or erastin treatment is due to the accumulation of LOOH and not other peroxides.

Figure 4 with 1 supplement see all

Download asset Open asset

Figure 4—source data 1

Original western blots.

https://cdn.elifesciences.org/articles/85289/elife-85289-fig4-data1-v2.zip

Download elife-85289-fig4-data1-v2.zip

What is the mechanism by which LOOH promotes muscle atrophy? C2C12 myotubes were pretreated with bafilomycin A1 (BafA1) or MG132 prior to erastin incubation to determine whether LOOH increases protein degradation in a lysosomal- or proteasomal-dependent manner, respectively. Erastin-induced reduction in myotube diameter was suppressed with BafA1, but not with MG132 (Figure 5A, B and Figure 5—figure supplement 1A), suggesting that the lysosome mediates protein degradation by LOOH (Gao et al., 2018). We also reproduced these findings with RSL3 treatment (Figure 5—figure supplement 1B, C). How does LOOH, a lipid molecule, promote lysosomal degradation? Upstream of the lysosome, autophagosome formation is mediated by a lipidation of LC3 by ATG3 (Ichimura et al., 2000). Thus, we hypothesized that LOOH may affect the lipidation of LC3. Indeed, GPx4 KD drastically reduced the protein content of p62, LC3-I, and LC3-II (Figure 5C and Figure 5—figure supplement 1D–H), potentially suggesting that LOOH either accelerates lysosomal degradation or reduces synthesis of these proteins. To test this possibility, we performed a targeted deletion of ATG3 in vitro. Indeed, ATG3 KD completely rescued the reduction in myotube diameter induced by GPx4 KD (Figure 5D, E and Figure 5—figure supplement 1I–L).

Figure 5 with 1 supplement see all

Download asset Open asset

Figure 5—source data 1

Original western blots.

https://cdn.elifesciences.org/articles/85289/elife-85289-fig5-data1-v2.zip

Download elife-85289-fig5-data1-v2.zip

Leveraging these findings, we generated mice with skeletal muscle-specific tamoxifen-inducible ATG3 knockout (ATG3-MKO) (Figure 6A, B and Figure 6—figure supplement 1A) and studied them with or without HU (Figure 6—figure supplement 1B–E). Loss of ATG3 was protective from disuse-induced atrophy (Figure 6C and Figure 6—figure supplement 1F) in soleus and weakness (Figure 6D and Figure 6—figure supplement 1G) in soleus and EDL. The protective effect of muscle mass in soleus was likely explained by greater myofiber CSA (Figure 6E, F and Figure 6—figure supplement 1H). Thus, suppression of autophagy is sufficient to attenuate disuse-induced muscle atrophy and weakness.

Figure 6 with 1 supplement see all

Download asset Open asset

Figure 6—source data 1

Original western blots.

https://cdn.elifesciences.org/articles/85289/elife-85289-fig6-data1-v2.zip

Download elife-85289-fig6-data1-v2.zip

We initially hypothesized that lysosomal degradation mediates LOOH-induced protein degradation to contribute to muscle atrophy. However, further assessment of muscle LOOH illuminated a more complex interaction between the lysosome and LOOH (Chen et al., 2021; Gao et al., 2016). Unexpectedly, quantification of 4-HNE revealed that inhibition of the autophagy-lysosome axis by ATG3 deletion or BafA1 was sufficient to inhibit LOOH induced by GPx4 deletion, erastin, or RSL3 (Figure 7A–D and Figure 7—figure supplement 1A–D). These findings suggest that the autophagy-lysosome axis is essential for LOOH amplification, in addition to its potential role in mediating protein degradation downstream. Indeed, immunofluorescence experiments revealed that 4-HNE is highly co-localized to LAMP2 (Figure 7E and Figure 7—figure supplement 1E), consistent with the notion that the lysosome is necessary for LOOH propagation. To support this idea, an increase in LOOH by hydrogen peroxide or carmustine (agents that increase pan oxidative stress without acting on GPx4 directly) was completely inhibited by lysosomal inhibition (Figure 7F–H and Figure 7—figure supplement 1F, G). Together, these observations suggest that the propagation of LOOH may be mediated by the lysosome (Figure 7—figure supplement 1H).

Figure 7 with 1 supplement see all

Download asset Open asset

Figure 7—source data 1

Original western blots.

https://cdn.elifesciences.org/articles/85289/elife-85289-fig7-data1-v2.zip

Download elife-85289-fig7-data1-v2.zip

Inhibition of autophagy suppressed lysosomal degradation and LOOH to attenuate muscle atrophy. We next tested whether suppression of LOOH would be sufficient to ameliorate skeletal muscle atrophy. We studied young (4 months) and old (20 months) global GPx4-overexpressing (GPx4Tg) mice (Ran et al., 2004) with or without HU (Figure 8—figure supplement 1A–E). Strikingly, both young and old GPx4Tg mice were resistant to disuse-induced muscle atrophy (Figure 8A and Figure 8—figure supplement 1F, G, greater effect in soleus than EDL). Perhaps even more impactful was the effect of GPx4 overexpression on skeletal muscle force-generating capacity such that, in both young and old, GPx4 overexpression robustly protected mice from muscle weakness induced by HU (Figure 8B and Figure 8—figure supplement 1H–J, soleus and EDL). These findings are in contrast to our experiments in Gpx4^+/- mice where muscle mass phenotype was only present in the young mice (Figure 3A) and no phenotype on muscle strength (Figure 3C). Consistent with the notion that GPx4 overexpression acts on LOOH, HU-induced increase in 4-HNE was completely suppressed in GPx4Tg mice (Figure 8C and Figure 8—figure supplement 1K). We also found that the protection from muscle atrophy was explained by greater myofiber CSA regardless of fiber type (Figure 8D&E and Figure 8—figure supplement 1L, M).

Figure 8 with 3 supplements see all

Download asset Open asset

Figure 8—source data 1

Original western blots.

https://cdn.elifesciences.org/articles/85289/elife-85289-fig8-data1-v2.zip

Download elife-85289-fig8-data1-v2.zip

Next, we explored opportunities to pharmacologically suppress LOOH to prevent muscle atrophy. Ferrostatin-1 inhibits the propagation of lipid peroxidation and is widely used to study LOOH (Dixon et al., 2012; Codenotti et al., 2018). Indeed, incubation of cells with ferrostatin-1 was sufficient to suppress LOOH induced by GPx4 KD (Figure 8F and Figure 8—figure supplement 2A) concomitant with protection from myotube atrophy (Figure 8G&H and Figure 8—figure supplement 2B, C). Nevertheless, ferrostatin-1 is currently not an FDA-approved drug with uncertainty surrounding safety. Thus, we tested L-carnosine, a dipeptide composed of beta-alanine and L-histidine that has the ability to scavenge reactive lipid aldehydes formed from LOOH (Cripps et al., 2017; Everaert et al., 2013). Rather than acting to suppress the lipid peroxidation process, L-carnosine binds to reactive lipid aldehydes to neutralize carbonyl stress. Similar to ferrostatin-1, L-carnosine was sufficient to suppress 4-HNE and rescue cell death induced by GPx4 KD (Figure 8—figure supplement 2D–F) or erastin (Figure 8—figure supplement 2G–I). Leveraging these data, we performed a preclinical trial for L-carnosine provided in drinking water ad lib (80 mM) in young wildtype C57BL6/J mice. L-carnosine treatment did not alter body mass, body composition, food intake, and water intake (Figure 8—figure supplement 3A–D), and successfully suppressed 4-HNE induced by HU (Figure 8—figure supplement 3E, F). Remarkably, mice provided with L-carnosine were partly protected from disuse-induced atrophy in soleus muscles (Figure 8—figure supplement 3G).

In humans, L-carnosine is rapidly degraded by a circulating carnosinase (Boldyrev et al., 2013) that may render oral carnosine treatment ineffective. In contrast, N-acetylcarnosine has a longer half-life and may be a more effective reagent in humans thus improving its translational potential. Similar to ferrostatin-1 and L-carnosine, N-acetylcarnosine also prevented 4-HNE and cell death induced by GPx4 KD (Figure 9—figure supplement 1A–C) or erastin (Figure 9—figure supplement 1D–F). Thus, we proceeded with a preclinical trial for N-acetylcarnosine in drinking water (80 mM, Figure 9A) in young (4 months, C57BL6/J; Jax colony) and old (20 months, C57BL/6; NIA rodent colony) wildtype mice. Similar to L-carnosine treatment, N-acetylcarnosine did not alter body mass, body composition, food intake, or water intake (Figure 9—figure supplement 2A–E), and successfully suppressed muscle 4-HNE (Figure 9B and Figure 9—figure supplement 2F, G). Strikingly, similar to our findings in GPx4Tg mice, N-acetylcarnosine ameliorated atrophy in soleus (Figure 9C and Figure 9—figure supplement 2H) and weakness in both soleus and EDL (Figure 9D and Figure 9—figure supplement 2I–K) in both young and old mice. Protection from muscle atrophy was similarly explained by greater myofiber CSA regardless of fiber type (Figure 9E, F and Figure 9—figure supplement 2L, M).

Figure 9 with 2 supplements see all

Download asset Open asset

Figure 9—source data 1

Original western blots.

https://cdn.elifesciences.org/articles/85289/elife-85289-fig9-data1-v2.zip

Download elife-85289-fig9-data1-v2.zip

The current findings demonstrate a novel mechanism that indicate LOOH as a key downstream molecule by which oxidative stress promotes muscle atrophy and weakness. Skeletal muscle LOOH was robustly upregulated with aging and disuse, and genetic or pharmacological neutralization of LOOH and their secondary reactive lipid aldehydes was sufficient to rescue muscle atrophy and weakness. In particular, N-acetylcarnosine treatment shows a potent effect in preserving muscle mass and strength with disuse in both young and old mice, informing the potential trial to utilize this compound to ameliorate loss of muscle function in humans.

During the preparation of this manuscript, Van Remmen and colleagues published a complementary study demonstrating that liproxstatin-1 can suppress denervation-induced skeletal muscle atrophy (Brown et al., 2022). Denervation and HU elicits different but overlapping response in myofibers, and our studies demonstrate that their effects to drive skeletal muscle atrophy might converge on lipid peroxidation. Like ferrostatin-1, liproxstatin-1 acts to suppress the propagation of lipid peroxidation rather than acting directly on LOOH. Nevertheless, in vivo liproxstatin-1 treatment was highly effective in suppressing denervation-induced LOOH as well as reactive lipid aldehydes 4-HNE, suggesting that targeting lipid peroxidation is likely an equally effective strategy to suppress LOOH production in skeletal muscle. Conversely, our data with GPx4 overexpression and N-acetylcarnosine treatment indicate that the effect of lipid peroxidation to promote muscle atrophy is mediated by LOOH and their lipid reactive aldehydes. Neither liproxstatin-1 nor ferrostatin-1 are currently FDA-approved, but it is worthwhile to consider these drugs along with N-acetylcarnosine as potential therapeutics to treat disuse-induced muscle atrophy.

Inhibition of the autophagy-lysosomal axis prevented myotube atrophy induced by erastin or GPx4 deletion, and muscle atrophy and weakness induced by HU in vivo. These findings are in contrast with previous studies that have shown inhibition of autophagy accelerates muscle mass (Carnio et al., 2014; Castets et al., 2013; Masiero et al., 2009). We do not interpret these differences as a discrepancy. In our studies, both inhibition of autophagy-lysosomal axis and induction of LOOH (in vitro or HU) were performed short term. It is likely that ATG3-MKO mice will eventually begin to undergo muscle atrophy similar to the muscle-specific ATG7 knockout model (Masiero et al., 2009). Importantly, inhibition of the autophagy-lysosomal axis not only inhibited muscle atrophy but also the accumulation of LOOH. The exact mechanism of the role of the lysosome in LOOH propagation is unclear, but our preliminary evidence shows LOOH accumulation occurring near the lysosome. Further studies are needed to clarify the exact nature of this interaction.

Sarcopenia is an age-associated decline in muscle mass and strength, that occurs due to a combination of inactivity, injury, and/or surgery, in addition to the biological consequences of aging itself. In the current study, mice were studied at 4 or 20 months of age. While not statistically compared directly (these experiments were not performed side-by-side), skeletal muscle mass at 20 months of age was not significantly lower compared to those at 4 months of age. Thus, the current data is unclear whether genetic or pharmacological suppression of LOOH prevents the loss of muscle mass due to the biological effect of aging in the absence of HU. We chose to study mice with 20 months of age for two reasons. First, mice greater than 20 months of age do not tolerate the HU intervention well, often resulting in their inability to consume food or water. Because disuse is an integral component of human aging, we wanted to study how muscles from old mice respond to disuse. This therefore compromised our ability to study sarcopenia without disuse. Second, while muscle mass was not diminished at 20 months of age, skeletal muscle force-generating capacity was lower in 20-month-old mice compared to the 4-month-old mice, particularly in the extensor digitorum longus (EDL) muscles. In GPx4Tg mice, age-associated decrease in muscle strength (in non-HU mice) appeared to be rescued, while short-term treatment with N-acetylcarnosine had no effect. We are currently following up on these results with a long-term treatment of N-acetylcarnosine with age that display the loss of muscle mass independent of HU to see if such intervention might alleviate the loss of muscle mass and strength due to the effect of aging per se. Studies in mice greater than 20 months of age would be necessary to more definitively show the role of LOOH in sarcopenia.

We initially set out to investigate the role of LOOH in age- and disuse-induced skeletal muscle atrophy, while measuring force-generating capacity as a secondary outcome. However, in all experimental models in which accumulation of muscle LOOH was suppressed (young and old GPx4Tg mice, young and old mice with N-acetylcarnosine, and young ATG3-MKO mice), force-generating capacity (i.e., specific force normalized to CSA) was more robustly rescued compared to skeletal muscle mass. This suggests the role of LOOH to induce muscle weakness independent of muscle atrophy, and likely independent of muscle protein degradation. While out of the scope for the current study, it would be important to determine whether reactive lipid aldehydes induced by aging or disuse preferentially bind to enzymes of skeletal muscle contraction or excitation-contraction coupling to compromise their activities. As described in the previous paragraph, aging promoted muscle weakness prior to atrophy. Similarly, muscle atrophy induced by cancer cachexia is also preceded by muscle weakness (Delfinis et al., 2022). Thus, these observations highlight the need to better study the mechanisms that regulate force-generating capacity independent of muscle mass.

In conclusion, we provide evidence that LOOH contributes to the loss of muscle mass and strength associated with age and disuse. Neutralization of LOOH, particularly their reactive lipid aldehyde byproducts, attenuates muscle atrophy and weakness. The mechanisms by which LOOH contributes to these phenotypes are not entirely clear, but they include protein degradation mediated by the autophagy-lysosomal axis, as well as loss in the force-generating capacity that is likely mediated by carbonyl stress. Last, but not least, these promising observations inform a potential clinical trial to test the efficacy of N-acetylcarnosine treatment in ameliorating muscle atrophy in humans.

All data generated or analyzed during this study are included in the manuscript.

1. 1. Anderson EJ
   2. Vistoli G
   3. Katunga LA
   4. Funai K
   5. Regazzoni L
   6. Monroe TB
   7. Gilardoni E
   8. Cannizzaro L
   9. Colzani M
   10. De Maddis D
   11. Rossoni G
   12. Canevotti R
   13. Gagliardi S
   14. Carini M
   15. Aldini G

   (2018) A carnosine analog mitigates metabolic disorders of obesity by reducing carbonyl stress

   The Journal of Clinical Investigation 128:5280–5293.

   https://doi.org/10.1172/JCI94307

   • PubMed
   • Google Scholar
2. 1. Bhattacharya A
   2. Muller FL
   3. Liu Y
   4. Sabia M
   5. Liang H
   6. Song W
   7. Jang YC
   8. Ran Q
   9. Van Remmen H

   (2009) Denervation induces cytosolic phospholipase A2-mediated fatty acid hydroperoxide generation by muscle mitochondria

   The Journal of Biological Chemistry 284:46–55.

   https://doi.org/10.1074/jbc.M806311200

   • PubMed
   • Google Scholar
3. 1. Bochkov VN
   2. Oskolkova OV
   3. Birukov KG
   4. Levonen A-L
   5. Binder CJ
   6. Stöckl J

   (2010) Generation and biological activities of oxidized phospholipids

   Antioxidants & Redox Signaling 12:1009–1059.

   https://doi.org/10.1089/ars.2009.2597

   • PubMed
   • Google Scholar
4. 1. Boldyrev AA
   2. Aldini G
   3. Derave W

   (2013) Physiology and pathophysiology of carnosine

   Physiological Reviews 93:1803–1845.

   https://doi.org/10.1152/physrev.00039.2012

   • PubMed
   • Google Scholar
5. 1. Bonaldo P
   2. Sandri M

   (2013) Cellular and molecular mechanisms of muscle atrophy

   Disease Models & Mechanisms 6:25–39.

   https://doi.org/10.1242/dmm.010389

   • PubMed
   • Google Scholar
6. 1. Brown JL
   2. Peelor FF
   3. Georgescu C
   4. Wren JD
   5. Kinter M
   6. Tyrrell VJ
   7. O’Donnell VB
   8. Miller BF
   9. Van Remmen H

   (2022) Lipid hydroperoxides and oxylipins are mediators of denervation induced muscle atrophy

   Redox Biology 57:102518.

   https://doi.org/10.1016/j.redox.2022.102518

   • PubMed
   • Google Scholar
7. 1. Cai J
   2. Pires KM
   3. Ferhat M
   4. Chaurasia B
   5. Buffolo MA
   6. Smalling R
   7. Sargsyan A
   8. Atkinson DL
   9. Summers SA
   10. Graham TE
   11. Boudina S

   (2018) Autophagy ablation in adipocytes induces insulin resistance and reveals roles for lipid peroxide and Nrf2 signaling in adipose-liver crosstalk

   Cell Reports 25:1708–1717.

   https://doi.org/10.1016/j.celrep.2018.10.040

   • PubMed
   • Google Scholar
8. 1. Carnio S
   2. LoVerso F
   3. Baraibar MA
   4. Longa E
   5. Khan MM
   6. Maffei M
   7. Reischl M
   8. Canepari M
   9. Loefler S
   10. Kern H
   11. Blaauw B
   12. Friguet B
   13. Bottinelli R
   14. Rudolf R
   15. Sandri M

   (2014) Autophagy impairment in muscle induces neuromuscular junction degeneration and precocious aging

   Cell Reports 8:1509–1521.

   https://doi.org/10.1016/j.celrep.2014.07.061

   • PubMed
   • Google Scholar
9. 1. Castets P
   2. Lin S
   3. Rion N
   4. Di Fulvio S
   5. Romanino K
   6. Guridi M
   7. Frank S
   8. Tintignac LA
   9. Sinnreich M
   10. Rüegg MA

   (2013) Sustained activation of mTORC1 in skeletal muscle inhibits constitutive and starvation-induced autophagy and causes a severe, late-onset myopathy

   Cell Metabolism 17:731–744.

   https://doi.org/10.1016/j.cmet.2013.03.015

   • PubMed
   • Google Scholar
10. 1. Chen X
    2. Yu C
    3. Kang R
    4. Kroemer G
    5. Tang D

    (2021) Cellular degradation systems in ferroptosis

    Cell Death and Differentiation 28:1135–1148.

    https://doi.org/10.1038/s41418-020-00728-1

    • PubMed
    • Google Scholar
11. 1. Codenotti S
    2. Poli M
    3. Asperti M
    4. Zizioli D
    5. Marampon F
    6. Fanzani A

    (2018) Cell growth potential drives ferroptosis susceptibility in rhabdomyosarcoma and myoblast cell lines

    Journal of Cancer Research and Clinical Oncology 144:1717–1730.

    https://doi.org/10.1007/s00432-018-2699-0

    • PubMed
    • Google Scholar
12. 1. Cripps MJ
    2. Hanna K
    3. Lavilla C
    4. Sayers SR
    5. Caton PW
    6. Sims C
    7. De Girolamo L
    8. Sale C
    9. Turner MD

    (2017) Carnosine scavenging of glucolipotoxic free radicals enhances insulin secretion and glucose uptake

    Scientific Reports 7:13313.

    https://doi.org/10.1038/s41598-017-13649-w

    • PubMed
    • Google Scholar
13. 1. Delfinis LJ
    2. Bellissimo CA
    3. Gandhi S
    4. DiBenedetto SN
    5. Garibotti MC
    6. Thuhan AK
    7. Tsitkanou S
    8. Rosa-Caldwell ME
    9. Rahman FA
    10. Cheng AJ
    11. Wiggs MP
    12. Schlattner U
    13. Quadrilatero J
    14. Greene NP
    15. Perry CG

    (2022) Muscle weakness precedes atrophy during cancer cachexia and is linked to muscle-specific mitochondrial stress

    JCI Insight 7:e155147.

    https://doi.org/10.1172/jci.insight.155147

    • PubMed
    • Google Scholar
14. 1. Dixon SJ
    2. Lemberg KM
    3. Lamprecht MR
    4. Skouta R
    5. Zaitsev EM
    6. Gleason CE
    7. Patel DN
    8. Bauer AJ
    9. Cantley AM
    10. Yang WS
    11. Morrison B
    12. Stockwell BR

    (2012) Ferroptosis: an iron-dependent form of nonapoptotic cell death

    Cell 149:1060–1072.

    https://doi.org/10.1016/j.cell.2012.03.042

    • PubMed
    • Google Scholar
15. 1. Dolbow DR
    2. Gorgey AS

    (2016) Effects of use and disuse on non-paralyzed and paralyzed skeletal muscles

    Aging and Disease 7:68–80.

    https://doi.org/10.14336/AD.2015.0826

    • PubMed
    • Google Scholar
16. 1. Eshima H
    2. Siripoksup P
    3. Mahmassani ZS
    4. Johnson JM
    5. Ferrara PJ
    6. Verkerke ARP
    7. Salcedo A
    8. Drummond MJ
    9. Funai K

    (2020) Neutralizing mitochondrial ROS does not rescue muscle atrophy induced by hindlimb unloading in female mice

    Journal of Applied Physiology 129:124–132.

    https://doi.org/10.1152/japplphysiol.00456.2019

    • PubMed
    • Google Scholar
17. 1. Evans WJ

    (2010) Skeletal muscle loss: cachexia, sarcopenia, and inactivity

    The American Journal of Clinical Nutrition 91:1123S–1127S.

    https://doi.org/10.3945/ajcn.2010.28608A

    • PubMed
    • Google Scholar
18. 1. Everaert I
    2. De Naeyer H
    3. Taes Y
    4. Derave W

    (2013) Gene expression of carnosine-related enzymes and transporters in skeletal muscle

    European Journal of Applied Physiology 113:1169–1179.

    https://doi.org/10.1007/s00421-012-2540-4

    • PubMed
    • Google Scholar
19. 1. Ferrara PJ
    2. Verkerke ARP
    3. Brault JJ
    4. Funai K

    (2018) Hypothermia decreases O2 cost for ex vivo contraction in mouse skeletal muscle

    Medicine and Science in Sports and Exercise 50:2015–2023.

    https://doi.org/10.1249/MSS.0000000000001673

    • PubMed
    • Google Scholar
20. 1. Ferrara PJ
    2. Rong X
    3. Maschek JA
    4. Verkerke AR
    5. Siripoksup P
    6. Song H
    7. Green TD
    8. Krishnan KC
    9. Johnson JM
    10. Turk J
    11. Houmard JA
    12. Lusis AJ
    13. Drummond MJ
    14. McClung JM
    15. Cox JE
    16. Shaikh SR
    17. Tontonoz P
    18. Holland WL
    19. Funai K

    (2021a) Lysophospholipid acylation modulates plasma membrane lipid organization and insulin sensitivity in skeletal muscle

    The Journal of Clinical Investigation 131:e135963.

    https://doi.org/10.1172/JCI135963

    • PubMed
    • Google Scholar
21. 1. Ferrara PJ
    2. Verkerke ARP
    3. Maschek JA
    4. Shahtout JL
    5. Siripoksup P
    6. Eshima H
    7. Johnson JM
    8. Petrocelli JJ
    9. Mahmassani ZS
    10. Green TD
    11. McClung JM
    12. Cox JE
    13. Drummond MJ
    14. Funai K

    (2021b) Low lysophosphatidylcholine induces skeletal muscle myopathy that is aggravated by high-fat diet feeding

    FASEB Journal 35:e21867.

    https://doi.org/10.1096/fj.202101104R

    • PubMed
    • Google Scholar
22. 1. Gao M
    2. Monian P
    3. Pan Q
    4. Zhang W
    5. Xiang J
    6. Jiang X

    (2016) Ferroptosis is an autophagic cell death process

    Cell Research 26:1021–1032.

    https://doi.org/10.1038/cr.2016.95

    • PubMed
    • Google Scholar
23. 1. Gao H
    2. Bai Y
    3. Jia Y
    4. Zhao Y
    5. Kang R
    6. Tang D
    7. Dai E

    (2018) Ferroptosis is a lysosomal cell death process

    Biochemical and Biophysical Research Communications 503:1550–1556.

    https://doi.org/10.1016/j.bbrc.2018.07.078

    • PubMed
    • Google Scholar
24. 1. Heden TD
    2. Johnson JM
    3. Ferrara PJ
    4. Eshima H
    5. Verkerke ARP
    6. Wentzler EJ
    7. Siripoksup P
    8. Narowski TM
    9. Coleman CB
    10. Lin C-T
    11. Ryan TE
    12. Reidy PT
    13. de Castro Brás LE
    14. Karner CM
    15. Burant CF
    16. Maschek JA
    17. Cox JE
    18. Mashek DG
    19. Kardon G
    20. Boudina S
    21. Zeczycki TN
    22. Rutter J
    23. Shaikh SR
    24. Vance JE
    25. Drummond MJ
    26. Neufer PD
    27. Funai K

    (2019) Mitochondrial PE potentiates respiratory enzymes to amplify skeletal muscle aerobic capacity

    Science Advances 5:eaax8352.

    https://doi.org/10.1126/sciadv.aax8352

    • PubMed
    • Google Scholar
25. 1. Ichimura Y
    2. Kirisako T
    3. Takao T
    4. Satomi Y
    5. Shimonishi Y
    6. Ishihara N
    7. Mizushima N
    8. Tanida I
    9. Kominami E
    10. Ohsumi M
    11. Noda T
    12. Ohsumi Y

    (2000) A ubiquitin-like system mediates protein lipidation

    Nature 408:488–492.

    https://doi.org/10.1038/35044114

    • PubMed
    • Google Scholar
26. 1. Imai H
    2. Nakagawa Y

    (2003) Biological significance of phospholipid hydroperoxide glutathione peroxidase (PHGPx, GPX4) in mammalian cells

    Free Radical Biology & Medicine 34:145–169.

    https://doi.org/10.1016/s0891-5849(02)01197-8

    • PubMed
    • Google Scholar
27. 1. Johnson JM
    2. Verkerke ARP
    3. Maschek JA
    4. Ferrara PJ
    5. Lin C-T
    6. Kew KA
    7. Neufer PD
    8. Lodhi IJ
    9. Cox JE
    10. Funai K

    (2020) Alternative splicing of UCP1 by non-cell-autonomous action of PEMT

    Molecular Metabolism 31:55–66.

    https://doi.org/10.1016/j.molmet.2019.10.007

    • PubMed
    • Google Scholar
28. 1. Kagan VE
    2. Mao G
    3. Qu F
    4. Angeli JPF
    5. Doll S
    6. Croix CS
    7. Dar HH
    8. Liu B
    9. Tyurin VA
    10. Ritov VB
    11. Kapralov AA
    12. Amoscato AA
    13. Jiang J
    14. Anthonymuthu T
    15. Mohammadyani D
    16. Yang Q
    17. Proneth B
    18. Klein-Seetharaman J
    19. Watkins S
    20. Bahar I
    21. Greenberger J
    22. Mallampalli RK
    23. Stockwell BR
    24. Tyurina YY
    25. Conrad M
    26. Bayır H

    (2017) Oxidized arachidonic and adrenic PEs navigate cells to ferroptosis

    Nature Chemical Biology 13:81–90.

    https://doi.org/10.1038/nchembio.2238

    • PubMed
    • Google Scholar
29. 1. Katunga LA
    2. Gudimella P
    3. Efird JT
    4. Abernathy S
    5. Mattox TA
    6. Beatty C
    7. Darden TM
    8. Thayne KA
    9. Alwair H
    10. Kypson AP
    11. Virag JA
    12. Anderson EJ

    (2015) Obesity in a model of GPX4 haploinsufficiency uncovers a causal role for lipid-derived aldehydes in human metabolic disease and cardiomyopathy

    Molecular Metabolism 4:493–506.

    https://doi.org/10.1016/j.molmet.2015.04.001

    • PubMed
    • Google Scholar
30. 1. Larsson L
    2. Degens H
    3. Li M
    4. Salviati L
    5. Lee YI
    6. Thompson W
    7. Kirkland JL
    8. Sandri M

    (2019) Sarcopenia: aging-related loss of muscle mass and function

    Physiological Reviews 99:427–511.

    https://doi.org/10.1152/physrev.00061.2017

    • PubMed
    • Google Scholar
31. 1. Lee J-Y
    2. Kim WK
    3. Bae K-H
    4. Lee SC
    5. Lee E-W

    (2021) Lipid metabolism and ferroptosis

    Biology 10:184.

    https://doi.org/10.3390/biology10030184

    • PubMed
    • Google Scholar
32. 1. Mahmassani ZS
    2. Reidy PT
    3. McKenzie AI
    4. Stubben C
    5. Howard MT
    6. Drummond MJ

    (2019) Age-Dependent skeletal muscle transcriptome response to bed rest-induced atrophy

    Journal of Applied Physiology 126:894–902.

    https://doi.org/10.1152/japplphysiol.00811.2018

    • PubMed
    • Google Scholar
33. 1. Masiero E
    2. Agatea L
    3. Mammucari C
    4. Blaauw B
    5. Loro E
    6. Komatsu M
    7. Metzger D
    8. Reggiani C
    9. Schiaffino S
    10. Sandri M

    (2009) Autophagy is required to maintain muscle mass

    Cell Metabolism 10:507–515.

    https://doi.org/10.1016/j.cmet.2009.10.008

    • PubMed
    • Google Scholar
34. 1. McCarthy JJ
    2. Srikuea R
    3. Kirby TJ
    4. Peterson CA
    5. Esser KA

    (2012) Inducible CRE transgenic mouse strain for skeletal muscle-specific gene targeting

    Skeletal Muscle 2:8.

    https://doi.org/10.1186/2044-5040-2-8

    • PubMed
    • Google Scholar
35. 1. Pharaoh G
    2. Brown JL
    3. Sataranatarajan K
    4. Kneis P
    5. Bian J
    6. Ranjit R
    7. Hadad N
    8. Georgescu C
    9. Rabinovitch P
    10. Ran Q
    11. Wren JD
    12. Freeman W
    13. Kinter M
    14. Richardson A
    15. Van Remmen H

    (2020) Targeting cPLA2 derived lipid hydroperoxides as a potential intervention for sarcopenia

    Scientific Reports 10:13968.

    https://doi.org/10.1038/s41598-020-70792-7

    • PubMed
    • Google Scholar
36. 1. Powers SK
    2. Smuder AJ
    3. Criswell DS

    (2011) Mechanistic links between oxidative stress and disuse muscle atrophy

    Antioxidants & Redox Signaling 15:2519–2528.

    https://doi.org/10.1089/ars.2011.3973

    • PubMed
    • Google Scholar
37. 1. Ran Q
    2. Liang H
    3. Gu M
    4. Qi W
    5. Walter CA
    6. Roberts LJ
    7. Herman B
    8. Richardson A
    9. Van Remmen H

    (2004) Transgenic mice overexpressing glutathione peroxidase 4 are protected against oxidative stress-induced apoptosis

    The Journal of Biological Chemistry 279:55137–55146.

    https://doi.org/10.1074/jbc.M410387200

    • PubMed
    • Google Scholar
38. 1. Reidy PT
    2. McKenzie AI
    3. Brunker P
    4. Nelson DS
    5. Barrows KM
    6. Supiano M
    7. LaStayo PC
    8. Drummond MJ

    (2017) Neuromuscular electrical stimulation combined with protein ingestion preserves thigh muscle mass but not muscle function in healthy older adults during 5 days of bed rest

    Rejuvenation Research 20:449–461.

    https://doi.org/10.1089/rej.2017.1942

    • PubMed
    • Google Scholar
39. 1. Scicchitano BM
    2. Pelosi L
    3. Sica G
    4. Musarò A

    (2018) The physiopathologic role of oxidative stress in skeletal muscle

    Mechanisms of Ageing and Development 170:37–44.

    https://doi.org/10.1016/j.mad.2017.08.009

    • PubMed
    • Google Scholar
40. 1. Tanner RE
    2. Brunker LB
    3. Agergaard J
    4. Barrows KM
    5. Briggs RA
    6. Kwon OS
    7. Young LM
    8. Hopkins PN
    9. Volpi E
    10. Marcus RL
    11. LaStayo PC
    12. Drummond MJ

    (2015) Age-related differences in lean mass, protein synthesis and skeletal muscle markers of proteolysis after bed rest and exercise rehabilitation

    The Journal of Physiology 593:4259–4273.

    https://doi.org/10.1113/JP270699

    • PubMed
    • Google Scholar
41. 1. Verkerke ARP
    2. Ferrara PJ
    3. Lin C-T
    4. Johnson JM
    5. Ryan TE
    6. Maschek JA
    7. Eshima H
    8. Paran CW
    9. Laing BT
    10. Siripoksup P
    11. Tippetts TS
    12. Wentzler EJ
    13. Huang H
    14. Spangenburg EE
    15. Brault JJ
    16. Villanueva CJ
    17. Summers SA
    18. Holland WL
    19. Cox JE
    20. Vance DE
    21. Neufer PD
    22. Funai K

    (2019) Phospholipid methylation regulates muscle metabolic rate through Ca2+ transport efficiency

    Nature Metabolism 1:876–885.

    https://doi.org/10.1038/s42255-019-0111-2

    • PubMed
    • Google Scholar
42. 1. Wiernicki B
    2. Dubois H
    3. Tyurina YY
    4. Hassannia B
    5. Bayir H
    6. Kagan VE
    7. Vandenabeele P
    8. Wullaert A
    9. Vanden Berghe T

    (2020) Excessive phospholipid peroxidation distinguishes ferroptosis from other cell death modes including pyroptosis

    Cell Death & Disease 11:922.

    https://doi.org/10.1038/s41419-020-03118-0

    • PubMed
    • Google Scholar
43. 1. Yang WS
    2. SriRamaratnam R
    3. Welsch ME
    4. Shimada K
    5. Skouta R
    6. Viswanathan VS
    7. Cheah JH
    8. Clemons PA
    9. Shamji AF
    10. Clish CB
    11. Brown LM
    12. Girotti AW
    13. Cornish VW
    14. Schreiber SL
    15. Stockwell BR

    (2014) Regulation of ferroptotic cancer cell death by GPX4

    Cell 156:317–331.

    https://doi.org/10.1016/j.cell.2013.12.010

    • PubMed
    • Google Scholar
44. 1. Yant LJ
    2. Ran Q
    3. Rao L
    4. Van Remmen H
    5. Shibatani T
    6. Belter JG
    7. Motta L
    8. Richardson A
    9. Prolla TA

    (2003) The selenoprotein GPx4 is essential for mouse development and protects from radiation and oxidative damage insults

    Free Radical Biology & Medicine 34:496–502.

    https://doi.org/10.1016/s0891-5849(02)01360-6

    • PubMed
    • Google Scholar
45. 1. Yoo S-E
    2. Chen L
    3. Na R
    4. Liu Y
    5. Rios C
    6. Van Remmen H
    7. Richardson A
    8. Ran Q

    (2012) Gpx4 ablation in adult mice results in a lethal phenotype accompanied by neuronal loss in brain

    Free Radical Biology & Medicine 52:1820–1827.

    https://doi.org/10.1016/j.freeradbiomed.2012.02.043

    • PubMed
    • Google Scholar

Author details

1. Hiroaki Eshima

   1. Diabetes and Metabolism Research Center, University of Utah, Salt Lake City, United States
   2. Molecular Medicine Program, University of Utah, Salt Lake City, United States
   3. Department of International Tourism, Nagasaki International University, Nagasaki, Japan

   Contribution

   Conceptualization, Data curation, Formal analysis, Funding acquisition, Investigation, Visualization, Methodology, Writing – original draft, Project administration

   Contributed equally with

   Justin L Shahtout

   Competing interests

   No competing interests declared

   "This ORCID iD identifies the author of this article:" 0000-0003-4492-5892

2. Justin L Shahtout

   1. Diabetes and Metabolism Research Center, University of Utah, Salt Lake City, United States
   2. Department of Physical Therapy & Athletic Training, University of Utah, Salt Lake City, United States

   Contribution

   Conceptualization, Data curation, Formal analysis, Validation, Investigation, Visualization, Methodology, Writing – original draft, Project administration, Writing – review and editing

   Contributed equally with

   Hiroaki Eshima

   Competing interests

   No competing interests declared

   "This ORCID iD identifies the author of this article:" 0000-0003-4111-4293

3. Piyarat Siripoksup

   1. Diabetes and Metabolism Research Center, University of Utah, Salt Lake City, United States
   2. Department of Physical Therapy & Athletic Training, University of Utah, Salt Lake City, United States

   Contribution

   Conceptualization, Data curation, Formal analysis, Funding acquisition, Validation, Investigation, Writing – review and editing

   Competing interests

   No competing interests declared

4. MacKenzie J Pearson

   Sciex, Framingham, United States

   Contribution

   Data curation, Formal analysis, Investigation, Methodology

   Competing interests

   is affiliated with Sciex. The author has no financial interests to declare

5. Ziad S Mahmassani

   1. Diabetes and Metabolism Research Center, University of Utah, Salt Lake City, United States
   2. Molecular Medicine Program, University of Utah, Salt Lake City, United States
   3. Department of Physical Therapy & Athletic Training, University of Utah, Salt Lake City, United States

   Contribution

   Data curation, Formal analysis, Validation, Investigation, Methodology

   Competing interests

   No competing interests declared

6. Patrick J Ferrara

   1. Diabetes and Metabolism Research Center, University of Utah, Salt Lake City, United States
   2. Molecular Medicine Program, University of Utah, Salt Lake City, United States
   3. Department of Nutrition & Integrative Physiology, University of Utah, Salt Lake City, United States

   Contribution

   Data curation, Formal analysis, Funding acquisition, Validation, Investigation, Methodology

   Competing interests

   No competing interests declared

7. Alexis W Lyons

   Diabetes and Metabolism Research Center, University of Utah, Salt Lake City, United States

   Contribution

   Data curation, Formal analysis, Validation, Investigation

   Competing interests

   No competing interests declared

8. John Alan Maschek

   1. Diabetes and Metabolism Research Center, University of Utah, Salt Lake City, United States
   2. Department of Nutrition & Integrative Physiology, University of Utah, Salt Lake City, United States
   3. Metabolomics Core Research Facility, University of Utah, Salt Lake City, United States

   Contribution

   Data curation, Formal analysis, Validation, Methodology

   Competing interests

   No competing interests declared

9. Alek D Peterlin

   1. Diabetes and Metabolism Research Center, University of Utah, Salt Lake City, United States
   2. Department of Nutrition & Integrative Physiology, University of Utah, Salt Lake City, United States

   Contribution

   Data curation, Formal analysis, Funding acquisition, Validation

   Competing interests

   No competing interests declared

   "This ORCID iD identifies the author of this article:" 0000-0002-2837-7446

10. Anthony RP Verkerke

    1. Diabetes and Metabolism Research Center, University of Utah, Salt Lake City, United States
    2. Department of Nutrition & Integrative Physiology, University of Utah, Salt Lake City, United States

    Contribution

    Data curation, Formal analysis, Funding acquisition, Validation, Investigation, Methodology

    Competing interests

    No competing interests declared

11. Jordan M Johnson

    1. Diabetes and Metabolism Research Center, University of Utah, Salt Lake City, United States
    2. Department of Nutrition & Integrative Physiology, University of Utah, Salt Lake City, United States

    Contribution

    Data curation, Formal analysis, Funding acquisition, Validation, Methodology

    Competing interests

    No competing interests declared

12. Anahy Salcedo

    Diabetes and Metabolism Research Center, University of Utah, Salt Lake City, United States

    Contribution

    Data curation, Formal analysis

    Competing interests

    No competing interests declared

13. Jonathan J Petrocelli

    1. Diabetes and Metabolism Research Center, University of Utah, Salt Lake City, United States
    2. Department of Physical Therapy & Athletic Training, University of Utah, Salt Lake City, United States

    Contribution

    Data curation, Formal analysis, Methodology

    Competing interests

    No competing interests declared

14. Edwin R Miranda

    1. Diabetes and Metabolism Research Center, University of Utah, Salt Lake City, United States
    2. Molecular Medicine Program, University of Utah, Salt Lake City, United States

    Contribution

    Formal analysis, Methodology, Writing – review and editing

    Competing interests

    No competing interests declared

15. Ethan J Anderson

    Fraternal Order of Eagles Diabetes Research Center, University of Iowa, Iowa City, United States

    Contribution

    Conceptualization, Resources, Funding acquisition, Methodology, Writing – review and editing

    Competing interests

    No competing interests declared

16. Sihem Boudina

    1. Diabetes and Metabolism Research Center, University of Utah, Salt Lake City, United States
    2. Molecular Medicine Program, University of Utah, Salt Lake City, United States
    3. Department of Nutrition & Integrative Physiology, University of Utah, Salt Lake City, United States

    Contribution

    Resources, Methodology

    Competing interests

    No competing interests declared

17. Qitao Ran

    Department of Cell Systems and Anatomy, The University of Texas Health Science Center at San Antonio, San Antonio, United States

    Contribution

    Resources, Investigation, Methodology, Writing – review and editing

    Competing interests

    No competing interests declared

18. James E Cox

    1. Diabetes and Metabolism Research Center, University of Utah, Salt Lake City, United States
    2. Metabolomics Core Research Facility, University of Utah, Salt Lake City, United States
    3. Department of Biochemistry, University of Utah, Salt Lake City, United States

    Contribution

    Funding acquisition, Investigation, Methodology, Writing – review and editing

    Competing interests

    No competing interests declared

19. Micah J Drummond

    1. Diabetes and Metabolism Research Center, University of Utah, Salt Lake City, United States
    2. Molecular Medicine Program, University of Utah, Salt Lake City, United States
    3. Department of Physical Therapy & Athletic Training, University of Utah, Salt Lake City, United States

    Contribution

    Conceptualization, Data curation, Formal analysis, Supervision, Funding acquisition, Investigation, Methodology, Writing – original draft, Project administration, Writing – review and editing

    Competing interests

    No competing interests declared

20. Katsuhiko Funai

    1. Diabetes and Metabolism Research Center, University of Utah, Salt Lake City, United States
    2. Molecular Medicine Program, University of Utah, Salt Lake City, United States
    3. Department of Physical Therapy & Athletic Training, University of Utah, Salt Lake City, United States
    4. Department of Nutrition & Integrative Physiology, University of Utah, Salt Lake City, United States

    Contribution

    Conceptualization, Resources, Data curation, Formal analysis, Supervision, Funding acquisition, Validation, Investigation, Visualization, Methodology, Writing – original draft, Project administration, Writing – review and editing

    For correspondence

    kfunai@utah.edu

    Competing interests

    No competing interests declared

    "This ORCID iD identifies the author of this article:" 0000-0003-3802-4756

• 1,860

  views

• 340

  downloads

• 9

  citations

Views, downloads and citations are aggregated across all versions of this paper published by eLife.