Targeting cPLA2 derived lipid hydroperoxides as a potential intervention for sarcopenia

Pharaoh, Gavin; Brown, Jacob L.; Sataranatarajan, Kavithalakshmi; Kneis, Parker; Bian, Jan; Ranjit, Rojina; Hadad, Niran; Georgescu, Constantin; Rabinovitch, Peter; Ran, Qitao; Wren, Jonathan D.; Freeman, Willard; Kinter, Michael; Richardson, Arlan; Van Remmen, Holly

doi:10.1038/s41598-020-70792-7

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Published: 18 August 2020

Targeting cPLA₂ derived lipid hydroperoxides as a potential intervention for sarcopenia

Gavin Pharaoh^1,2,
Jacob L. Brown²,
Kavithalakshmi Sataranatarajan²,
Parker Kneis²,
Jan Bian²,
Rojina Ranjit²,
Niran Hadad^3,4,
Constantin Georgescu⁵,
Peter Rabinovitch⁶,
Qitao Ran^7,8,
Jonathan D. Wren⁵,
Willard Freeman^1,3,4,5,
Michael Kinter²,
Arlan Richardson³ &
…
Holly Van Remmen^1,2,4,9

Scientific Reports volume 10, Article number: 13968 (2020) Cite this article

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Abstract

Defects in neuromuscular innervation contribute significantly to the age-related decline in muscle mass and function (sarcopenia). Our previous studies demonstrated that denervation induces muscle mitochondrial hydroperoxide production (H₂O₂ and lipid hydroperoxides (LOOHs)). Here we define the relative contribution of mitochondrial electron transport chain (ETC) derived H₂O₂ versus cytosolic phospholipase A₂ (cPLA₂) derived LOOHs in neurogenic muscle atrophy. We show that denervation increases muscle cPLA₂ protein content, activity, and metabolites downstream of cPLA₂ including LOOHs. Increased scavenging of mitochondrial H₂O₂ does not protect against denervation atrophy, suggesting ETC generated H₂O₂ is not a critical player. In contrast, inhibition of cPLA₂ in vivo mitigates LOOH production and muscle atrophy and maintains individual muscle fiber size while decreasing oxidative damage. Overall, we show that loss of innervation in several muscle atrophy models including aging induces generation of LOOHs produced by arachidonic acid metabolism in the cPLA₂ pathway contributing to loss of muscle mass.

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Introduction

The pathological age-related loss of skeletal muscle mass and function (sarcopenia) contributes to decreased quality of life and independence and increases the risk of injury and chronic disease^1,2. Preventing or reducing the effects of sarcopenia could increase quality of life, reduce risk of comorbidity development, and save billions of dollars in healthcare costs annually. Designing therapeutic interventions will remain difficult until the complex pathways and processes underlying sarcopenia are identified³.

Studies supporting a loss of innervation with age and a link between denervation and decreased muscle fiber size suggest that sarcopenia is a form of neurogenic atrophy^4,5,6. In addition to the loss of muscle mass, contractile force also declines with age⁷. Loss of muscle strength occurs more rapidly than can be explained by the loss of muscle mass alone, and evidence suggests that a reduction in muscle quality and neuromuscular junction signaling also play a role in the functional decline of aging muscle⁸. Our previous studies using mice lacking the cytosolic superoxide (O₂⁻) scavenger CuZn superoxide dismutase (CuZn SOD, Sod1^−/−) demonstrate that elevated oxidative stress leads to an accelerated sarcopenia in the Sod1^−/− mice that is characterized by denervation, muscle hydroperoxide production, and muscle atrophy^9,10,11. Returning expression of CuZnSOD specifically to motor neurons of the Sod1^−/− mice prevented denervation, muscle hydroperoxide production, and atrophy, supporting a link between loss of innervation, hydroperoxides, and muscle atrophy¹². We have also shown that loss of innervation to skeletal muscle directly induces basal hydroperoxide production from isolated mitochondria including both hydrogen peroxide (H₂O₂) and lipid hydroperoxides (LOOHs)¹¹. The magnitude of this hydroperoxide increase is correlated with the extent of muscle atrophy in several neurogenic atrophy conditions including aging¹¹. We identified generation of arachidonic acid (AA) by cytosolic phospholipase A₂ (cPLA₂) as a major source of LOOHs in denervation; however, whether increased hydroperoxide production contributes to neurogenic atrophy has not been determined¹³.

The goal of the current study is to define the role of hydroperoxides in neurogenic atrophy by (1) measuring the identity and source of released hydroperoxides (H₂O₂ vs LOOHs) and (2) testing whether inhibiting specific hydroperoxide generation in vivo can modulate downstream atrophy. We hypothesize that the increase in muscle hydroperoxides released following denervation is a causal component of neurogenic muscle atrophy and sarcopenia. To test this, we investigated the identity and source of hydroperoxide production from muscle fibers in several neurogenic atrophy conditions including aging, a mouse model of oxidative stress-induced atrophy (Sod1^−/−), surgical denervation (sciatic nerve transection), and a mouse model (SOD1^G93A) of Amyotrophic Lateral Sclerosis (ALS) using a combination of scavengers and small-molecule inhibitors. We also tested whether H₂O₂ or LOOHs are causal to neurogenic atrophy using genetic approaches to increase H₂O₂ scavenging and pharmacological interventions to inhibit cPLA₂ as a potential source of LOOHs in the sciatic nerve transection model.

We report that neurogenic atrophy primarily induces muscle LOOHs through cPLA₂ metabolism of AA and not mitochondrial H₂O₂. In vivo cPLA₂ inhibition mitigates denervation atrophy, while H₂O₂ scavenging does not. We identify the cPLA₂ pathway as a negative regulator of muscle mass in neurogenic atrophy and a potential target for therapeutic intervention in sarcopenia and other diseases of muscle wasting.

Results

Neurogenic atrophy induces muscle hydroperoxide production and atrophy

Aging in mice and humans is associated with an age-related loss of motor neurons and sarcopenia^4,5. Our previous studies show that loss of muscle mass in response to denervation is accompanied by increased mitochondrial generation of hydroperoxides^11,13. To determine the effect of hydroperoxide production on the loss of muscle mass during aging, we compared gastrocnemius muscle mass and basal hydroperoxide production in permeabilized gastrocnemius muscle fibers harvested from young, middle aged, and old C57BL/6 J mice. Figure 1a shows a loss of gastrocnemius muscle mass evident first at 26 months of age that continues into advanced age (32 months), while Fig. 1b shows a clear association between basal hydroperoxide production rates and loss of gastrocnemius mass (Fig. 1a,b). The increase in basal hydroperoxide production rate correlates with the amount of muscle atrophy in aging mice and in several other advanced atrophy models, including mice lacking the Nrf2 antioxidant response transcription factor (Nrf2^−/−), mice with accelerated neurogenic sarcopenia (Sod1^−/−), and mice with motor neuron disease (SOD1^G93A) (Fig. 1c). These data support a direct relationship between basal hydroperoxide production rate and neurogenic muscle atrophy.

To more directly interrogate the relationship between loss of innervation, muscle atrophy and hydroperoxide generation, we measured changes in hydroperoxide production rates and muscle mass in a model of sciatic nerve transection. This model is relevant to aging because the proportion of denervated fibers increases with age⁶. Denervation by surgical transection of the sciatic nerve was performed in one hindlimb (denervation) and compared to the effect of sham surgery in the contralateral hindlimb of adult, male C57BL/6 J mice. The sciatic nerve transection model allows us to measure time-dependent changes after loss of innervation in samples composed entirely of denervated fibers. We measured the muscle response to nerve transection at 1, 2, 4, and 7 days following surgery. Muscle mass declines significantly between 4 and 7 days after loss of innervation, while hydroperoxide production is increased between 2 and 4 days (Fig. 1d–f). By 7 days post denervation, muscle mass is decreased by 20% and hydroperoxide production rates are increased compared to sham in both permeabilized muscle fibers (1,468%) and isolated mitochondria (987%). These results are supported by a previous finding that basal hydroperoxide production is increased in denervated tibialis anterior (TA) muscle prior to atrophy¹⁴.

We also compared sex-specific effects of denervation on gastrocnemius muscle from female and male mice seven days after surgery. Gastrocnemius muscle from both sexes shows a significant decline in mass at 7 days post-denervation (Supplemental Fig. 1a). Male mice have a higher initial gastrocnemius mass and a greater loss of total muscle mass; however, the relative loss of mass is equivalent in both sexes (Supplemental Fig. 1b-c). The greater loss in muscle mass in male mice corresponds to higher hydroperoxide production rates in isolated mitochondria and permeabilized fibers from denervated muscles in male mice compared to female mice (Supplemental Fig. 1d-e).

Loss of innervation induces transcriptional changes

To understand the molecular events underlying the effects of denervation, we measured denervation induced gene expression changes in muscle using RNAseq analysis of denervated muscle samples across a time course following sciatic nerve transection. We found that an acute gene expression response to denervation begins within 12–24 h followed by a chronic response beginning between 2 and 4 days, which coincides with the observed increase in LOOH production by the cPLA₂ pathway (Fig. 2a). Further changes in gene expression increase up to 14 days (Fig. 2a,b). A similar number of genes are upregulated compared to the number that are downregulated at each time point during the chronic response from days 2 through 14 (Fig. 2a). Four genes are significantly upregulated throughout the entire time course (Arpp21, Gadd45a, Gdf5, and Myog), and 68 of the genes that are differentially regulated beginning at 2–4 days persist throughout the measured chronic phase (Fig. 2b). By day 14, 35 biological processes (including processes related to muscle cell function, metabolism, intracellular signaling, and ion transport) are significantly affected through overrepresented differentially expressed genes (Supplemental Files). 88 canonical pathways show significant change in activity including activation of Production of Nitric Oxide and Reactive Oxygen Species in Macrophages and ERK/MAPK Signaling and significantly different expression of Calcium, AMPK, NRF2 Antioxidant Transcription Factor, p53, and GADD45A signaling pathways (Supplemental Files). Signaling from ATF4 as an upstream regulator of gene expression changes was elevated within 2–4 days (Fig. 2c). ATF4 was previously associated with age-related muscle atrophy but our findings now suggest that loss of innervation to muscle fibers plays a key role in inducing this pathway¹⁵. Of the differentially regulated genes, we observed a significant enrichment of genes containing transcription factor binding sites in their promoter regions for MEF2 beginning at 2–4 days and SIX2 at 7 days (Fig. 2d). In addition to upregulation of LOOH production, loss of innervation is associated with sweeping chronic changes in gene expression.

Loss of innervation induces LOOH production through activation of the cytosolic phospholipase A₂ (cPLA₂) pathway

We previously reported that in addition to detecting hydrogen peroxide (H₂O₂), the Amplex Red probe interacts with lipid hydroperoxides (LOOHs)¹³. Invitrogen has since released an updated probe (Amplex UltraRed, Invitrogen A36006) with improved sensitivity and stability. We report that H₂O₂ and the LOOHs 13(S)-HpODE and 15(S)-HpETE, two metabolites enzymatically produced in vivo by 12/15-lipoxygenase, react with Amplex UltraRed to produce a concentration-dependent linear increase in fluorescent resorufin¹⁶. Notably, H₂O₂ produces a stronger increase in fluorescence at the same concentrations (Fig. 3a).

Our previous study using isolated mitochondria revealed that inhibition of cytosolic phospholipase A₂ (cPLA₂) with arachidonyl trifluoromethyl ketone (AACOCF₃), an inhibitor that binds to the cPLA₂ active site to block its activity, prevented the increase in Amplex UltraRed signal in response to denervation^13,17. However, the study by Bhattacharya et al. did not define the total hydroperoxide production in muscle fibers, whether LOOHs were being produced upstream or downstream of cPLA₂, or whether cPLA₂ produces LOOHs in other neurogenic atrophy models including aging. cPLA₂ is a membrane associated enzyme that cleaves AA from membrane phospholipids, thereby increasing AA levels that are in turn converted by enzymatic pathways to eicosanoids, lipid mediators of cell signaling¹⁸. LOOHs can be released from membranes by cytosolic phospholipase A₂ (cPLA₂) or formed downstream of cPLA₂ by metabolism of AA into LOOH intermediates^18,19,20. To define the molecular identity of hydroperoxides produced by denervation using permeabilized muscle fibers, we used Amplex UltraRed and ex vivo treatment with a combination of antioxidants (catalase, glutathione peroxidase, and superoxide dismutase) and the cPLA₂ inhibitor AACOCF₃. All conditions contained 2.5 U/ml of exogenous CuZn superoxide dismutase (Sod1) to convert any superoxide (O₂⁻) produced by the mitochondria to H₂O₂. First, to specifically scavenge H₂O₂ and not LOOHs produced downstream of cPLA₂, we treated denervated muscle fibers with supraphysiological concentrations of exogenous catalase. Treatment of denervated fibers with catalase decreased the hydroperoxide production rate but only up to ~ 30%, even at supraphysiological concentrations of catalase (Fig. 3b). We next treated denervated fibers with exogenous glutathione peroxidase 1 (GPX1), which scavenges both H₂O₂ and LOOHs²¹. Combined treatment of denervated fibers with GPX1 and catalase significantly decreased the hydroperoxide production rate by approximately 50% (Fig. 3c). In contrast, the CPLA₂ inhibitor AACOCF₃ decreased the hydroperoxide signal to a much greater extent than catalase or GPX (approximately 90%) (Fig. 3d) suggesting the greater proportion of the Amplex UltraRed signal is generated by LOOH formed downstream of cPLA₂ as opposed to H₂O₂.

To further define the source of the Amplex UltraRed signal in permeabilized fibers, we measured the effect of a series of titrations of mitochondrial electron transport chain (ETC) substrates and inhibitors on hydroperoxide production described in the methods. Adding the mitochondrial ETC inhibitors rotenone and antimycin A greatly increases hydroperoxide production (presumably as a result of O₂⁻ and H₂O₂ from ETC complexes) in permeabilized muscle fibers^22,23. We show here that both ETC inhibitors induced hydroperoxide production in both untreated sham and denervated fibers (Fig. 3e). Exogenous catalase and CuZnSOD completely scavenged the source of this signal while AACOCF₃ had no effect (Fig. 3e). These results demonstrate that the CuZnSOD and catalase concentrations efficiently scavenge mitochondrial generation of O₂⁻ and H₂O₂, and that AACOCF₃ does not inhibit mitochondrial generation of O₂⁻ and H₂O₂ or act as a direct antioxidant scavenger. Together, these data suggest that LOOHs are the primary component of the increased hydroperoxides observed after loss of innervation, their production or release requires cPLA₂ activity, and they are not produced from electron transport chain reactive oxygen species (ROS) generation.

To determine the direct scavenging potential in vitro for a number of oxidant scavengers and AACOCF₃, we generated a dose–response curve for H₂O₂ or the 12/15-lipoxygenase generated LOOH 15(S)-HpETE in the presence of several oxidative scavengers and AACOCF₃ using the Amplex UltraRed signal as a readout¹⁶. Glutathione (GSH) is used to regenerate the antioxidant capacity of GPX1, although it can also serve as a direct antioxidant for free radicals, reactive oxygen species including H₂O₂, and LOOHs²⁴. SS-31 is a mitochondrial targeted peptide that was originally designed as a mitochondrially-targeted antioxidant that directly scavenges via its tyrosine amino acid residue²⁵. SS-31 was previously reported to completely inhibit H₂O₂ chemiluminescence at 100 µM and decrease lipid peroxidation by ~ 60% at 1 µM in vitro, although these experiments were carried out with isolated mitochondria or over an incubation period in cell-free conditions^25,26. Liproxstatin-1 is a recently reported LOOH scavenger and functions as a positive control²⁷. At the concentration we used in our permeabilized fiber assays, catalase and GPX1 with or without reduced glutathione (GSH) directly scavenge H₂O₂. In contrast, we observe no direct scavenging of LOOHs by AACOCF₃ or SS-31 (Fig. 3f). We also report that Liproxstatin-1 functions as a scavenger of H₂O₂ in addition to LOOHs (Fig. 3f). For LOOHs, Liproxstatin-1, GPX1 with and without GSH, and GSH work as direct scavengers while SS-31 and AACOCF₃ do not (Fig. 3g). AACOCF₃ decreases LOOH production by inhibiting cPLA₂ without direct scavenging of LOOHs.

The LOOHs produced in skeletal muscle during loss of innervation could be produced by two processes that are upstream or downstream of cPLA₂: i.e., oxidative insult to membrane phospholipids creating LOOHs that are released by cPLA₂, or metabolism of cleaved AA into eicosanoids that can produce LOOHs as a byproduct through the lipoxygenase and cyclooxygenase pathways^19,20. We hypothesized that if denervated fibers exposed to cPLA₂ inhibition by AACOCF₃ treatment are able to produce hydroperoxides after addition of exogenous AA, this would be evidence that LOOH production is primarily occurring by metabolism of AA into eicosanoids. Exogenous addition of AA to sham and denervated muscle fibers treated with AACOCF₃ increased hydroperoxide production rates to levels comparable to untreated denervation fibers (Fig. 3h). This suggests the cPLA₂ pathway primarily produces LOOHs in denervated muscle downstream of cPLA₂ release of AA and provides evidence that denervation-induced hydroperoxides are produced enzymatically during eicosanoid metabolism of AA and not by the canonical release of LOOHs from membranes formed by oxidative insult. The increase in LOOH production in both sham and denervated muscle treated with AACOCF₃ after addition of AA suggests that AA release is the main regulatory step for LOOH production during eicosanoid metabolism.

Next, we used the permeabilized fiber protocol and treatment with either catalase or AACOCF₃ to identify and compare the relative composition of hydroperoxide production rates in several models of neurogenic atrophy including an aging time-course, accelerated sarcopenia with neuromuscular degeneration (Sod1^−/− mice), and end-stage ALS motor neuron disease (SOD1^G93A mice). Catalase or AACOCF₃ inhibited ~ 50–66% of hydroperoxides in sarcopenic aged mice (Fig. 3i). Age-related hydroperoxides are a mixture of LOOHs produced in the cPLA₂ pathway and O₂⁻/H₂O₂ likely produced by the ETC (Supplemental Fig. 2 Inset). In fibers from Sod1^−/−, SOD1^G93A, and denervated mice, the increased basal hydroperoxides are almost completely inhibited by the cPLA₂ inhibitor AACOCF₃ (Supplemental Fig. 2). AACOCF₃ inhibits significantly more of the basal hydroperoxides in these models than catalase and in total inhibits ~ 80–90% of hydroperoxides (Fig. 3i). LOOHs produced by the cPLA₂ pathway are induced in all tested models of neurogenic atrophy and are the primary component of basal hydroperoxides.

H₂O₂ scavengers do not protect against denervation-induced muscle hydroperoxide production or atrophy

To more specifically test the contribution of increased H₂O₂ to neurogenic atrophy pathogenesis, we used two transgenic mouse models (skmMCAT and skmPRDX3) that overexpress mitochondrial H₂O₂ scavengers in skeletal muscle. The skmMCAT model expresses mitochondrially-targeted human catalase (mCAT) in muscle²⁸. Catalase is an H₂O₂ scavenger normally localized to the peroxisome. The skmMCAT model uses a flox-stopped version of the construct to express catalase specifically in muscle²⁹. The skmPRDX3 model over expresses peroxiredoxin 3 (PRDX3) in muscle also using a flox-stopped transgene. Peroxiredoxin 3 is an endogenous mitochondrial H₂O₂ scavenger³⁰. Analysis of skmMCAT expression using Western blot, targeted mass-spectrometry, and qRT-PCR revealed a similar level of mCAT gene expression compared to mouse catalase and approximately a three–ninefold increase in total catalase content in the skeletal muscle of skmMCAT mice compared to controls (Supplemental Fig. 3a-d). mCAT expression did not change protein content of other antioxidant or metabolic enzymes measured by mass spectrometry (Supplemental Table 1). PRDX3 was overexpressed approximately two orders of magnitude in gastrocnemius muscles of the skmPRDX3 model (Supplemental Fig. 3e-f).

To measure the impact of H₂O₂ generation on muscle atrophy induced by denervation, we performed sciatic nerve transection on control, skmMCAT, and skmPRDX3 mice and analyzed muscle mass and hydroperoxide production rates in isolated mitochondria and permeabilized fibers from gastrocnemius muscles after seven days (Fig. 4a). We find that sham muscle fibers and isolated mitochondria from skmMCAT but not skmPRDX3 mice have decreased hydroperoxide production rates after addition of ETC inhibitors compared to wildtype mice as expected for the increased H₂O₂ scavenging potential in muscle from these mice (Fig. 4c,e). However, neither skmMCAT or skmPRDX3 mice exhibit decreased peroxide production rates in denervated muscle fibers or isolated mitochondria compared to wildtype mice (Fig. 4b,d). Importantly, increased scavenging of H₂O₂ in muscle of skmMCAT and skmPRDX3 mice does not protect the mice from denervation induced loss of muscle mass (Fig. 4f). skmPRDX3 mice do tend to show a lower percent of atrophy in the denervated muscle compared to sham control, however this was not statistically significant. Interestingly, the skmPRDX3 mice have significantly smaller sham muscles, and a linear regression analysis shows that the size of the sham muscle partially explains variations in percent atrophy of the denervated gastrocnemius muscle compared to sham (Fig. 4g,h). Muscle fibers from skmMCAT mice and isolated muscle mitochondria efficiently scavenged H₂O₂ under control conditions, but increased H₂O₂ scavenging was not sufficient to protect against denervation-induced muscle hydroperoxide production or atrophy. Together our evidence suggests that increased muscle mitochondrial H₂O₂ production is not causal for muscle atrophy in response to denervation.

The tetrapeptide SS-31 was initially reported to have direct H₂O₂ antioxidant scavenging and to a lesser extent LOOH scavenging²⁵. More recently, it has been suggested to exert its protective effects, including those in several age-related conditions, by virtue of its targeting to cardiolipin, with enhancement of ETC function, reduced ROS, and improved ATP production³¹. We treated wildtype mice for 10 days with daily injections of saline or SS-31 (3 mg/kg body weight). After 3 days of treatment, we performed sciatic nerve transection and analyzed muscle mass and hydroperoxide production 7 days later (Fig. 4i). SS-31 does not protect against muscle atrophy or increased basal hydroperoxide production in denervated muscle (Fig. 4j,k). In summary, the results of these three interventions suggest that H₂O₂ signaling is not required to induce muscle wasting, but instead denervation is primarily associated with increased LOOHs.

Denervation induces cPLA₂ activity and downstream eicosanoids in skeletal muscle resulting in LOOH production

We previously reported an increase in cPLA₂ protein content in denervated muscle compared to sham muscle¹³. In the current study, we measured cPLA₂ activity in gastrocnemius muscles from sham and denervated mice seven days after sciatic nerve transection. cPLA₂ activity is significantly elevated in denervated muscle fibers (Fig. 5a). We also measured cPLA₂ activity in gastrocnemius muscles from young 6–9 month and old 28 month wild-type, onset Sod1^−/−, and end-stage SOD1^G93A mice. cPLA₂ activity was significantly elevated in wasting muscle from SOD1^G93A mice but not sarcopenic or Sod1^−/− mice (Fig. 5b).

To better understand the nature of the LOOH response following loss of innervation, we used a targeted lipidomics approach to quantify the content of eicosanoids in gastrocnemius muscle from young control and seven-day post-denervation mice. We find that multiple eicosanoids downstream of cPLA₂ are increased in muscle from 7-day denervated mice (Fig. 5c). In the lipoxygenase pathway, phospholipid hydroperoxide glutathione peroxidase (GPX4) converts LOOH intermediates to hydroxylipids^18,20. Of the increased eicosanoids in denervation, 9 out of 16 are hydroxy lipids found downstream of LOOH intermediates in the lipoxygenase pathway.

We chose a targeted panel of eight eicosanoids based on our results in denervated muscle and measured their content in muscle from aged and SOD1^G93A mice. Seven out of the eight measured eicosanoids were upregulated in aged and SOD1^G93A gastrocnemius compared to control (Fig. 5d,e). We found significant upregulation of eicosanoids in all neurogenic atrophy models compared to control.

In vivo cPLA2₂ inhibition protects against denervation-induced muscle hydroperoxide production and atrophy

To determine in vivo if increased production of LOOHs generated by cPLA₂ plays a causal role in denervation atrophy, we performed sciatic nerve transection surgeries and treated mice with a daily ip injection of 9.5 mg/kg body weight AACOCF₃ or corn oil vehicle control for 7 days then measured the effect of cPLA₂ inhibition on muscle hydroperoxide production, atrophy, and individual fiber size (Fig. 6a). AACOCF₃ is a potent cell-permeable inhibitor of cPLA₂ and previous publications demonstrated this dose inhibited cPLA₂ in vivo in spinal cords³². Vehicle injected mice show a similar amount of atrophy compared to untreated controls (~ 20% atrophy in denervated muscle compared to sham at 7 days) (Fig. 6b). AACOCF₃-injected mice undergo significantly less atrophy (~ 17%) at 7 days, resulting in a rescue of 16% of atrophy (Fig. 6b). We have repeated this experiment in several cohorts of mice, and each cohort saw similar protection from atrophy (data not shown).

We measured basal hydroperoxide production in permeabilized fibers from sham and denervated gastrocnemius muscle with and without AACOCF₃ treatment. The standard protocol for fiber preparation found no difference in basal hydroperoxide production with AACOCF₃ treatment. However, the protection in atrophy in the treatment group suggested some drug effect. Because AACOCF₃ is a reversible inhibitor, we hypothesized it may be washed out during the dilution steps in the permeabilization protocol. We performed mechanical separation of fibers from vehicle and treated mice and immediately measured hydroperoxide production rates omitting the permeabilization and washing steps. No difference was found in fibers from vehicle-injected mice with or without permeabilization, therefore this protocol is suitable for measuring basal hydroperoxide production (Fig. 6c). Using this fiber preparation protocol, we observe a significant reduction in basal hydroperoxides (~ 34%) in denervated fibers from AACOCF₃-injected mice compared to control (Fig. 6c).

Denervation results in a significant decrease in average muscle fiber cross-sectional area (CSA) in gastrocnemius muscles from both vehicle- (29%) and AACOCF₃-treated (20%) mice (Fig. 6d–f). However, we observe a significant treatment effect in response to AACOCF₃, which is associated with an increase in individual denervated fiber CSA (27%) compared to vehicle treated mice (Fig. 6d–f). Denervated muscle contains an increased proportion of small muscle fibers compared to sham, but we observe that AACOCF₃ treatment increases the proportion of large muscle fibers in both sham and denervated muscles and increases average CSA of denervated fibers compared to vehicle control (Fig. 6e–f). Average CSA area is linearly correlated with gastrocnemius mass, which suggests that denervation primarily induces muscle atrophy through individual fiber atrophy that is partially rescued with AACOCF₃ treatment (Fig. 6g). We previously reported that mice lacking 12/15-lipoxygenase (Alox15^−/−) were protected from denervation atrophy³³. Here we report that the basal hydroperoxide production rate in Alox15^−/− denervated fibers is significantly decreased (~ 64%) (Fig. 6h). These experiments provide evidence that LOOHs produced by the cPLA₂ pathway play a causal role in neurogenic muscle atrophy and are a viable target to protect against loss of muscle mass in neurogenic atrophy.

AACOCF₃ treatment does not decrease cPLA₂ pathway protein content

We next measured protein content of cPLA₂, calcium-independent mitochondrial phospholipase A₂ (iPLA₂), and 12- and 12/15-lipoxygenase following denervation and cPLA₂ inhibition in vivo. We measured protein content of key components of the cPLA₂ pathway in both muscle whole homogenate and isolated mitochondria. Our analysis reveals that loss of innervation increases protein content of cPLA₂, iPLA₂, and 12- and 12/15-lipoxygenase in whole homogenate of the gastrocnemius muscle (Fig. 7a,b). In addition, AACOCF₃ treatment significantly increases 12- and 12/15-lipoxygenase in denervated muscle compared to vehicle-treated denervated muscle (Fig. 7a,b). cPLA₂ can be activated by increased intracellular calcium or phosphorylation by MAP kinase³⁴. We observe no difference in phosphorylated cPLA₂ content in whole homogenate. We also observe no difference in isolated mitochondrial content of cPLA₂, phosphorylated cPLA₂, iPLA₂, or 12- and 12/15-lipoxygenase. Denervation does not increase activation of cPLA₂ by phosphorylation (data not shown). Furthermore, AACOCF₃ treatment inhibits cPLA₂ activity but does not decrease expression or content of PLA₂ pathway enzymes. These results suggest that denervation induces an increase in PLA₂ pathway enzyme protein content, which results in the increased cPLA₂ activity we observe in denervation.

cPLA₂ inhibition mitigates oxidative stress in denervation

We measured several parameters to determine if LOOH production in the cPLA₂ pathway contributes to atrophy primarily through oxidative damage or eicosanoid signaling. F₂-isoprostanes are a biomarker of oxidative stress formed by non-enzymatic reaction of AA with a free radical³⁵. F₂-isoprostane content is increased in denervated muscle, but treatment with AACOCF₃ mitigates this increase (Fig. 8a). The cPLA₂ knockout mouse were previously identified to have cardiac fiber hypertrophy via modulating IGF-1 pathway signaling³⁶. We measured the content and phosphorylation of several proteins in the IGF-1 signaling pathway. Loss of innervation increases AKT protein content and MAP kinase phosphorylation (Fig. 8b,c). However, treatment with AACOCF₃ did not affect these changes after loss of innervation.

Discussion

In this work, we report a novel effect of lipid hydroperoxide generation in the initiation of skeletal muscle atrophy following loss of innervation. This is a new mechanism for induction of muscle atrophy by lipid mediators. Our studies further provide evidence for potential intervention and modulation of muscle atrophy by cPLA₂ pathway inhibition that could be relevant for sarcopenia and other diseases of muscle wasting including ALS and nerve injury. cPLA₂-derived LOOHs hold clinical relevance, as vastus lateralis biopsies from octogenarians also show LOOH production inhibited by AACOCF₃³⁷. These findings represent a paradigm shift in the understanding of ROS in muscle: previously research focused intensely on mitochondrial ETC-derived O₂⁻, H₂O₂, and lipid peroxidation as causal factors in sarcopenia and not LOOHs produced in the cPLA₂ pathway. Increased scavenging of muscle mitochondrial H₂O₂ did not protect against denervated muscle hydroperoxide production or atrophy, although whole body mCAT expression and SS-31 injection were previously found to protect aged muscle against H₂O₂ production or oxidative stress^28,38,39. While we have focused on basal hydroperoxide production primarily composed of LOOHs, a recent report shows that activated mitochondria from aged skeletal muscle produce higher concentrations of superoxide (O₂⁻) and H₂O₂ at physiological ADP concentrations, which may explain the mechanism of MCAT and SS-31 protection⁴⁰. In the mitochondrial fraction, we observe cPLA₂, iPLA₂, and 12- and 12/15-lipoxygenase proteins, and elevated LOOH production in denervated muscles. We previously reported that both the cPLA₂ inhibitor AACOCF₃ and the glutathione peroxidase mimetic, ebselen, decreased basal hydroperoxide production in isolated mitochondria from denervated muscle to a greater extent than catalase¹³. Together, these results suggest some parts of the eicosanoid pathway are taking place in the mitochondrial fraction of denervated muscles¹³.

cPLA₂ has been previously reported to be a negative regulator of cardiac and skeletal muscle hypertrophy³⁶. Under physiological conditions, cPLA₂ negatively regulates individual muscle fiber size by inhibiting IGF-1 signaling. cPLA₂ releases AA from membranes, which is metabolized enzymatically in the cyclooxygenase, lipoxygenase, cytochrome p450, and anandamide pathways and non-enzymatically into isoprostanes and nitroalkenes⁴¹. The cyclooxygenase (COX) and lipoxygenase (LOX) pathways both produce LOOHs as intermediates⁴¹. cPLA₂ and cyclooxygenase inhibitors are an ongoing topic of study for ALS treatment, although previous publications focus primarily on neuronal expression and inhibition^42,43. The 12/15-Lipoxygenase is implicated in the pathology of a range of diseases¹⁶. We previously reported that Alox15^−/− mice were partially protected from muscle loss after sciatic nerve transection, and here we show they also have significantly decreased skeletal muscle LOOH production in response to denervation³³. However, removing 12/15-lipoxygenase is not sufficient to completely prevent loss of muscle mass or hydroperoxide production. This suggests that multiple eicosanoid related enzymes are responsible for LOOH production in denervation and that there are multiple targetable steps in the pathway for intervention. While AACOCF₃ is a more potent inhibitor of cPLA₂ than iPLA₂ or sPLA₂, it also inhibits cyclooxygenase activity⁴⁴. Production of LOOHs by cyclooxygenases could explain the total effectiveness of AACOCF₃ ex vivo (~ 90%) compared to the partial effectiveness of 12/15-lipoxygenase knockout at inhibiting LOOH production (~ 64%)⁴⁴. By identifying the other responsible enzymes and developing or testing inhibitors, we can more specifically inhibit LOOH production. LOOHs could cause atrophy through oxidative stress and signaling changes. Denervation increases F₂-isoprostane content that is blunted with cPLA₂ inhibition by AACOCF₃. We previously reported denervation induces proteasome-mediated protein degradation³³. However, deletion of 12/15-lipoxygenase protects against this induction in protein degradation, which suggests that cPLA₂ regulates muscle size partially through downstream metabolites affecting proteostasis. Overall, we show that the cPLA₂ pathway negatively regulates muscle mass. Enzymes in the cPLA₂ pathway could be targeted using small molecule inhibitors to treat several atrophy conditions including aging, nerve injury, and ALS.

Materials and methods

Animals and animal models

All animal experiments were conducted in accordance with the guidelines for the care and use of laboratory animals of the University of Oklahoma Health Sciences Center (OUHSC) and Oklahoma Medical Research Foundation (OMRF). The study was approved by the Institutional Animal Care and Use Committees at the OUHSC and the OMRF. All experiments used mice from strains on a C57BL/6 J genetic background currently housed in the mouse colony of Dr. Van Remmen at the OMRF or Veterans Affairs (VA) Medical Center. Male 4 month old wild-type and 24 month old wild-type and Nrf2^−/− (B6.129X1-Nfe2l2^tm1Ywk/J) gastrocnemius mass and basal hydroperoxide data was previously reported²². End-stage (135–152 days) SOD1^G93A mouse [B6-Tg(SOD1-G93A)1Gur/J(G93AGur1)] gastrocnemius mass and basal hydroperoxide data for vehicle and catalase or AACOCF₃ treatment was previously reported²³. Sod1^−/− mice were previously described⁴⁵. mCAT-flox mice were previously described²⁹. PRDX3tg-flox mice have a transgene that consists of CAG promoter, an LSL-STOP cassette flanked by loxP sites, and a human PRDX3 cDNA. In those mice, activation of Cre removes the LSL-STOP cassette, leading to expression of PRDX3. mCAT-flox or PRDX3tg-flox specifically target expression to muscle using a Cre-Lox system driven by human α-skeletal actin (HSA) directed Cre expression to make skmMCAT and skmPRDX3 lines respectively⁴⁶. The transgenes contain a stop sequence at the start of the coding sequence flanked by LoxP sites. When the Cre is expressed in skeletal muscle fibers, the stop sequence is removed and the transgene is translated⁴⁷. Alox15^−/− mice were acquired from The Jackson Laboratory (B6.129S2-Alox15^tm1Fun/J) and were previously described³³. Mice were caged in a pathogen free environment with free access to standard chow and water and maintained on a 12 h light/dark cycle. Mice were euthanized using a CO₂ chamber and cervical dislocation. Hind limb muscles were dissected, weighed, and flash frozen in liquid nitrogen for biochemical analysis or used fresh for isolated mitochondria and permeabilized muscle fiber experiments. All experiments were carried out in the gastrocnemius muscle unless specified otherwise.

Sciatic nerve transection surgeries and drug treatment regimens

Sciatic nerve transection and sham surgeries were performed as previously described on 6–8 month old C57BL/6 J, skmMCAT, skmPRDX3, and Alox15^−/− mice^11,13,33. The mice were anesthetized in the induction chamber with isoflurane (2–3 units) then maintained under anesthesia using a mouse nose cone. The lateral thigh and buttock from the sciatic notch to the knee were shaved, and 100% ethanol and 2% chlorhexidine was used to clean the skin surface. A small incision was made from the sciatic notch to the knee (approximately 1 cm), and the sciatic nerve was exposed. The nerve was cut and a 5 mm piece removed. The ends of the nerve were folded back and sutured to prevent nerve regrowth. On the other leg, the nerve was exposed but not cut and removed. This limb served as a sham-operated control. The incision on each leg was closed using the PDS suture 5–0 and skin glue, and the mouse was placed on a warm heating pad until recovered. The mice were treated with 9.5 mg/kg ketaprofen dissolved in saline for three days following surgery to minimize pain. The mice were euthanized and dissected at 0.5, 1, 2, 4, 7, or 14 days after the surgery.

A cohort of male control mice were treated with arachidonyl trifluoromethyl ketone (AACOCF₃; Enzo BML-ST335-0,050) or vehicle (corn oil) following sciatic nerve transection surgery. AACOCF₃ was dissolved in DMSO (25 mg/ml) and then diluted in corn oil to 4 mM. Mice were treated with intraperitoneal (ip) injections of vehicle control (corn oil) or 9.5 mg/kg AACOCF₃ once daily beginning with the first injection at the time of surgery and continuing until euthanasia seven days after surgery. Another cohort of mice was treated with SS-31 or vehicle control. Mice were treated daily with ip injections of 3 mg/kg SS-31 or vehicle (saline) control based on previous publications demonstrating efficacy of this dose in mouse skeletal muscle^38,48. Sciatic nerve transection and sham surgeries were performed on the fourth day of SS-31 or vehicle injection treatment and mice were euthanized seven days later.

Muscle fiber permeabilization and hydroperoxide production rate measurement

Preparation of permeabilized muscle fiber bundles with saponin and measurement of hydroperoxide production with Amplex UltraRed using the Oroboros Oxygraph-2k (O2k, OROBOROS Instruments, Innsbruck, Austria) with fluorometer were performed as previously described^22,23. Amplex UltraRed measures hydroperoxide production (both hydrogen peroxide and lipid hydroperoxides). Horseradish peroxidase catalyzes a reaction between a hydroperoxide group and Amplex UltraRed to produce the fluorescent compound resorufin. The rate of resorufin production is detected as the change in fluorescence intensity. We performed a preliminary experiment adding increasing concentrations of GPX1 to a denervated permeabilized fiber and found 10 U/ml GPX1 more efficiently inhibited hydroperoxide production than lower or higher doses (data not shown). In Fig. 3c, the denervated gastrocnemius was removed from each animal then separated into four different permeabilized fiber bundles per animal. These bundles were untreated, treated with catalase, treated with GPX1, or treated with catalase and GPX1, respectively. Separate animals were used for the experiment in Fig. 3d. From the sham surgery leg, one control permeabilized gastrocnemius fiber bundle was prepared. From the denervated leg, three permeabilized gastrocnemius fiber bundles were prepared per animal. These were untreated (vehicle), treated with catalase, or treated with AACOCF_3, respectively. Permeabilized fibers were treated with 2000 U/ml catalase or 10 U/ml GPX1 during 30 min saponin permeabilization, 3 × 5 min wash steps, and in the O2k chamber during measurement. Permeabilized fibers were treated with 20 µM arachidonyl trifluoromethyl ketone (AACOCF₃; Cayman Chemicals, Ann Arbor, MI) during 30 min saponin permeabilization and 3 × 5 min wash steps, but not in the O2k chamber during measurement. The addition of substrates and inhibitors during measurements was performed as previously described^22,23. A dose response of Amplex UltraRed/resorufin fluorescence to H₂O₂ and lipid hydroperoxide (13(S)-HpODE and 15(S)-HpETE) concentrations was determined with and without the addition of substrates and inhibitors at the specified concentrations for H₂O₂ and 15(S)-HpETE (Fig. 3b,c). The supraphysiological catalase concentration increases background auto-oxidation of Amplex UltraRed, which increases background fluorescence rate. Auto-oxidation is accounted for in all samples by subtracting its rate for experimental measurements, however this induces increased variability in measurements with catalase. The actual inhibition of basal hydroperoxides by catalase in each model may be lower than that presented here.

Mitochondrial isolation and hydroperoxide production rate

Mitochondria were isolated using the Chappell–Perry method, and hydroperoxide production rate was measured using Amplex Red fluorescent rate measurements in a plate-based fluorometer as previously described in gastrocnemius muscles^11,13.

Cytosolic phospholipase A₂ (cPLA₂) activity assay

cPLA₂ activity was assessed using the Abcam Cytosolic Phospholipase A₂ Assay Kit (ab133090) according to the manufacturer's instructions. 50 mg of gastrocnemius muscle was homogenized on ice in a glass homogenizer (size 21) in 500 μl homogenization buffer (50 mM HEPES, 1 mM EDTA, pH 7.4). The homogenate was centrifuged at 10,000 × g for 15 min at 4 °C. The supernatant was removed and filtered using Amicon 30 kDA filter (UFC803024) at 4,000 × g for 20 min at 4 °C. Protein concentration was determined using the Bradford assay. Samples were incubated with bromoenol lactone for 15 min at RT to inhibit iPLA₂. 100 μg protein was loaded in triplicate for each sample. The assay was initiated with substrate solution and incubated for 60 min at RT. Enzyme catalysis with ended with DTNB/EGTA and absorbance was read at 414 nm and converted to enzyme activity using the manufacturer’s calculations.

Eicosanoid measurements by targeted lipidomics

Snap frozen gastrocnemius from 4 to 6 month old young control, 4–6 month old 7 day denervated, 28 month old aged, and end-stage SOD1^G93A male mice were sent to the LIPID MAPS Lipidomics Core, https://www.ucsd-lipidmaps.org. at the University of California, San Diego. Eicosanoids were analyzed by LC–MS as previously described⁴⁹. The data were analyzed using a Python 3 script. The script used the Bartlett variance test to analyze for unequal variance of each eicosanoid between control and experimental group. If variance was equal, each eicosanoid was compared with two-tailed student’s t-test. If variance was unequal, each eicosanoid was compared with Welch’s t-test. Benjamini–Hochberg FDR correction (q < 0.05) was used to determine statistical significance compared to control. Principal component analysis (PCA) plot were generated using ClustVis with default settings⁵⁰.

Individual muscle fiber cross sectional area

Fiber size was measured as previously described⁵¹. Gastrocnemius muscle was sectioned during sacrifice, mounted in optimum cutting temperature compound (OCT), and flash frozen in liquid nitrogen-cooled isopentane. Sections (10 μm) were mounted and stained with hematoxylin and then with Aeosin using a Leica ST5020 Multistainer Leica Microsystems, Wetzlar, Germany). Images were visualized and captured with Zeiss Axiovert 200 M microscope, Zeiss Axiocam MRC camera, and Zeiss AxioVision software V4.8.2.0 (Carl Zeiss AG, Oberkochen, Germany) at 10 × magnification. Cross-sectional area (CSA) was measured using ImageJ software for approximately 500 fibers per sample⁵².

Western blot analysis

Western blots were performed as previously described using standard techniques²³. Imaging was performed with G:BOX imaging system (Syngene) and quantified using GeneTools software (Syngene). Information on antibodies is available in Supplemental Table 2.

Quantitative real-time polymerase chain reaction (RT-PCR)

RT-PCR was performed as previously described⁵³. Total RNA was extracted from gastrocnemius using TRIzol reagent (Invitrogen, Carlsbad, CA, United States). Equal amounts of extracted RNA (1 μg) were converted to first strand cDNA using a cDNA synthesis kit (Bio-Rad, Herculus, CA, United States). Human catalase and mouse catalase mRNA levels were measured using Assay on Demand Hs00156308_m1 and Mm00437992_m1 (Applied Biosystems) as previously described²⁸. RT-PCR was performed in Quant Studio 6 (Applied Biosystems, Foster City, CA, United States). The ΔΔC_t method was used to calculate relative mRNA expression.

Lipid peroxidation by F₂-isoprostanes

Levels of F₂-isoprostanes in 100–150 mg of sham and denervated gastrocnemius muscles after seven days of denervation and treatment with vehicle or AACOCF₃ were determined by gas chromatography–mass spectrometry as previously described^22,54. The level of F₂-isoprostanes in muscle tissues was expressed as nanograms of 8-Iso-PGF_2α per gram of muscle mass.

Quantification of protein abundance using amino acid sequence

We used targeted quantitative mass spectrometry to measure protein abundance of metabolic and antioxidant proteins in control and skmMCAT gastrocnemius muscles as previously described^22,23,55. Skyline was used to monitor and process data from each peptide⁵⁶.

RNA extraction, library construction, and RNA sequencing (RNAseq)

Using TRIzol reagent (Invitrogen, CA, USA), total RNA was extracted from sham and denervated gastrocnemius that were collected at 0.5, 1, 2, 4, 7, and 14 days after sciatic nerve transection. The RNA integrity and concentration were first assessed using an Agilent Bioanalyzer (Agilent Technologies, Santa Clara, CA) to meet the experimental standard. The RNA samples were given to OMRF Clinical Genomics Center. Libraries were prepared using the TruSeq Stranded mRNA Library Kit (Illumina). The libraries were then sequenced on an Illumina NextSeq 500 to produce paired-end 75 base pair reads. The data collected were given to Discovery Bioinformatics Core at the Oklahoma Nathan Shock Center of Excellence in the Biology of Aging for data analysis.

RNA-seq data processing followed the guidelines and practices of the ENCODE and modENCODE consortia regarding proper experimental replication, sequencing depth, data and metadata reporting, and data quality assessment (https://www.encodeproject.org/documents/cede0cbe-d324-4ce7-ace4-f0c3eddf5972/) as previously described⁵⁷. Raw sequencing reads (in a FASTQ format) were trimmed of residual adaptor sequences using Scythe software. Low quality bases at the beginning or the end of sequencing reads were removed using sickle then the quality of remaining reads was confirmed with FastQC. Further processing of quality sequencing reads was performed with utilities provided by the Tuxedo Suite software. Reads were aligned to the Mus musculus genome reference (GRCm38/mm10) using the TopHat component, then cuffquant and cuffdiff were utilized for gene-level read counting and differentially expression analysis. A false discovery rate threshold of 0.05 was used as selection criteria for differentially expressed genes between pairs of time points. Functional analysis to find overrepresented functional sets (GO, KEGG pathways) was performed using specialized R Bioconductor packages. Ingenuity Pathway Analysis (IPA, QIAGEN, Redwood City CA, https://www.qiagenbioinformatics.com/products/ingenuitypathway-analysis) was used to explore significant gene networks and pathways interactively.

Promoter motif analysis in differentially expressed genes was performed using Hypergeometric Optimization of Motif EnRichment (HOMER) version 4.9.1⁵⁸. Motifs enrichment was conducted in gene’s proximal promoter (400 bp upstream and 100 upstream). Venn diagram was drawn in Microsoft Powerpoint. Venn diagram was drawn in Microsoft Powerpoint.

Statistical analyses

The results were analyzed using Microsoft Office Excel and GraphPad Prism 7.0b for Mac OS X (GraphPad Software, La Jolla, CA) unless otherwise specified. Statistical tests were determined according to experimental design as described in figure legends. Significantly different variation between groups was identified by Bartlett’s test. For pairwise comparisons, unpaired two-tailed t-test with p < 0.05 were used to determine significance for pairs with equal variance. For multiple groups, ordinary one-way ANOVA or ordinary two-way ANOVA and appropriate post hoc test with adjusted p < 0.05 determined significance for comparisons with equal variance. Statistical significance was determined by Welch’s ANOVA test with Dunnett’s T3 post hoc test for groups with unequal variance. Linear regression was used for correlation analysis.

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Acknowledgements

The authors would like to thank the Integrative Redox Biology Core, the Multiplexing Protein Quantification Core, and the Discovery Bioinformatics Core at the Oklahoma Nathan Shock Center of Excellence in the Biology of Aging, the Imaging Core Facility and the Clinical Genomics Center at the Oklahoma Medical Research Foundation, and the LIPID MAPS Lipidomics Core at the University of California, San Diego (UCSD) for providing core services. Thank you to Pavithra Premkumar, Brittany Forbes, Sara Jane Koehler, and Ashley Murphy for general lab assistance. SS-31 tetrapeptide was kindly provided by Zoltan Ungvari. Funding was provided by the National Institute on Aging (NIA) Grants (R01AG050676, P01AG051442, and T32AG052363), American Federation for Aging Research (AFAR) Scholarship for Research in the Biology of Aging, the John and Mildred Carson Oklahoma Medical Research Foundation (OMRF) Pre-doctoral Scholarship, and United States Department of Veterans Affairs (VA) Merit Award (I01BX002595) and VA Senior Research Career Scientist Award (12F-RCS-011).

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Physiology Department, University of Oklahoma Health Sciences Center, Oklahoma City, OK, USA
Gavin Pharaoh, Willard Freeman & Holly Van Remmen
Aging and Metabolism Research Program, Oklahoma Medical Research Foundation, Oklahoma City, OK, USA
Gavin Pharaoh, Jacob L. Brown, Kavithalakshmi Sataranatarajan, Parker Kneis, Jan Bian, Rojina Ranjit, Michael Kinter & Holly Van Remmen
Reynolds Oklahoma Center on Aging, University of Oklahoma Health Sciences Center, Oklahoma City, OK, USA
Niran Hadad, Willard Freeman & Arlan Richardson
Oklahoma Center for Neuroscience, University of Oklahoma Health Sciences Center, Oklahoma City, OK, USA
Niran Hadad, Willard Freeman & Holly Van Remmen
Genes and Human Disease Research Program, Oklahoma Medical Research Foundation, Oklahoma City, OK, USA
Constantin Georgescu, Jonathan D. Wren & Willard Freeman
Department of Pathology, University of Washington, Seattle, WA, USA
Peter Rabinovitch
Department of Cell Systems and Anatomy, UT Health San Antonio, San Antonio, TX, USA
Qitao Ran
South Texas Veterans Health Care System, San Antonio, TX, USA
Qitao Ran
Oklahoma City VA Medical Center, Oklahoma City, OK, USA
Holly Van Remmen

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Gavin Pharaoh
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Jan Bian
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Michael Kinter
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Contributions

G.P. and H.V.R. created the experimental design. G.P. performed experiments, statistical analysis, and data visualization in Figs. 1a–f, 2, 4, 5a,b, 6a–g, 8b,c, Supplemental Fig. 1, Supplemental Fig. 2, and Supplemental Fig. 3a-b. G.P. analyzed and plotted experiments in Figs. 3a–c, 5c–e, 6h, 7a,b, 8a, Supplemental Fig. 3e-f, and Supplemental Tables 1–2. J.L.B. performed the CSA experiment and measured hydroperoxide production in Alox15^−/− mice. K.S. performed a portion of the sciatic nerve transection surgeries in the time course and prepared samples for RNAseq and targeted lipidomics. P.K. performed F₂-isoprostane analysis. J.B. performed western blots in Fig. 7. R.R. performed isolated mitochondria experiments. N.H. and W.F. performed HOMER motif enrichment analysis. C.G. and J.D.W. analyzed the RNAseq. P.R. and Q.R. provided mCAT-flox and PRDX3-flox models as well as manuscript input. A.R. provided regular feedback on project direction and manuscript input. M.K. performed targeted protein mass-spectrometry experiments. G.P. and H.V.R. wrote the manuscript with editing by J.L.B.

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Correspondence to Holly Van Remmen.

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Pharaoh, G., Brown, J.L., Sataranatarajan, K. et al. Targeting cPLA₂ derived lipid hydroperoxides as a potential intervention for sarcopenia. Sci Rep 10, 13968 (2020). https://doi.org/10.1038/s41598-020-70792-7

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Received: 18 December 2019
Accepted: 23 July 2020
Published: 18 August 2020
DOI: https://doi.org/10.1038/s41598-020-70792-7

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