
==== Front
Mol Metab
Mol Metab
Molecular Metabolism
2212-8778
Elsevier

S2212-8778(24)00146-7
10.1016/j.molmet.2024.102015
102015
Original Article
Loss of mitochondria long-chain fatty acid oxidation impairs skeletal muscle contractility by disrupting myofibril structure and calcium homeostasis
Pereyra Andrea S. apereyr2@jh.edu
⁎⁎
Fernandez Regina F.
Amorese Adam
Castro Jasmine N.
Lin Chien-Te
Spangenburg Espen E.
Ellis Jessica M. ellisje18@ecu.edu
⁎
Brody School of Medicine at East Carolina University, Department of Physiology and East Carolina Diabetes and Obesity Institute, Greenville, NC, 27834, USA
⁎ Corresponding author. ellisje18@ecu.edu
⁎⁎ Corresponding author apereyr2@jh.edu
28 8 2024
11 2024
28 8 2024
89 1020156 6 2024
5 8 2024
19 8 2024
© 2024 The Author(s)
2024
https://creativecommons.org/licenses/by-nc-nd/4.0/ This is an open access article under the CC BY-NC-ND license (http://creativecommons.org/licenses/by-nc-nd/4.0/).
Objective

Abnormal lipid metabolism in mammalian tissues can be highly deleterious, leading to organ failure. Carnitine Palmitoyltransferase 2 (CPT2) deficiency is an inherited metabolic disorder affecting the liver, heart, and skeletal muscle due to impaired mitochondrial oxidation of long-chain fatty acids (mLCFAO) for energy production.

Methods

However, the basis of tissue damage in mLCFAO disorders is not fully understood. Mice lacking CPT2 in skeletal muscle (Cpt2Sk−/−) were generated to investigate the nexus between mFAO deficiency and myopathy.

Results

Compared to controls, ex-vivo contractile force was reduced by 70% in Cpt2Sk−/− oxidative soleus muscle despite the preserved capacity to couple ATP synthesis to mitochondrial respiration on alternative substrates to long-chain fatty acids. Increased mitochondrial biogenesis, lipid accumulation, and the downregulation of 80% of dystrophin-related and contraction-related proteins severely compromised the structure and function of Cpt2Sk−/− soleus. CPT2 deficiency affected oxidative muscles more than glycolytic ones. Exposing isolated sarcoplasmic reticulum to long-chain acylcarnitines (LCACs) inhibited calcium uptake. In agreement, Cpt2Sk−/− soleus had decreased calcium uptake and significant accumulation of palmitoyl-carnitine, suggesting that LCACs and calcium dyshomeostasis are linked in skeletal muscle.

Conclusions

Our data demonstrate that loss of CPT2 and mLCFAO compromise muscle structure and function due to excessive mitochondrial biogenesis, downregulation of the contractile proteome, and disruption of calcium homeostasis.

Highlights

• Long-chain acylcarnitines impede SR-mediated calcium dynamics in skeletal muscle.

• Mitochondrial overload in CPT2 deficiency disrupts muscle contractile machinery.

• Metabolic myopathies impact muscle structural integrity.

• CPT2 loss disrupts SR structure and intracellular calcium homeostasis.

Keywords

Fatty acid oxidation
CPT2
Muscle contraction
Palmitoyl-carnitine
Calcium
==== Body
pmc1 Introduction

Skeletal muscle is an energetically demanding tissue due to ATP-dependent muscle contraction. In high-intensity or prolonged physical activity, fatty acids are an essential substrate for mitochondria-mediated energy production. Thus, acquired and inherited defects in mitochondrial nutrient metabolism are expected to impact muscle contractility and force production. Disorders of mitochondrial fatty acid oxidation (FAODs) are a common cause of metabolic myopathy characterized by muscle weakness, rhabdomyolysis, and exercise intolerance [1]. Because the inability to oxidize fatty acids directly compromises the supply of reduction equivalents to the mitochondrial electron transport chain (ETC) and, ultimately, the production of ATP, FAODs are classified as energy deficiency disorders [2]. However, the mechanisms linking mitochondrial dysfunction and tissue pathology in FAODs are poorly understood.

We recently reported that in a model of FAOD that muscle bioenergetics was seemingly preserved through metabolic adaptations [3]. However, despite mitochondrial respiration being conserved or even increased for alternative substrates, exercise performance was sub-normal. Here, the impact of impaired mitochondrial long-chain FAO (mLCFAO) in skeletal muscle structure and function in the FAOD model of muscle Carnitine Palmitoyltransferase 2 (CPT2) deficiency, Cpt2Sk−/− mice, was investigated [3,4]. CPT2 is a critical, autonomous enzyme required for mLCFAO. It converts CPT1-generated acyl-carnitines into acyl-Coenzyme A esters in the mitochondrial matrix which are later utilized to fuel β-oxidation. The loss of CPT2 obliterates mitochondrial long-chain fatty acid oxidation in the muscle, with a significant accumulation of long-chain acylcarnitines.

In this study, ex vivo muscle force assessment revealed extreme weakness only in oxidative Soleus muscle, while contractility was nearly preserved in the highly glycolytic Extensor Digitorum Longus (EDL) muscle. Energetically, mitochondrial membrane potential and substrate-supported ATP production were either preserved or increased in Cpt2Sk−/− oxidative Soleus, indicating energetic sufficiency. We demonstrate a myriad of mechanisms linking FAOD to muscle dysfunction, including alterations in myofibril tethering, reduction in sarcomeric proteins, mitochondrial content overload, and calcium dyshomeostasis that is further exacerbated by the impact of long-chain acylcarnitines on calcium balance. These findings question the long-standing assumption that disorders of intermediate metabolism directly cause myopathy strictly due to energetic defects.

2 Results

2.1 Impaired long-chain fatty acid oxidation compromises contraction in a muscle-type-dependent manner

Reduced tolerance to physical activity is a common feature of fatty acid oxidation disorders, including CPT2 deficiency. As such, the skeletal muscle-specific loss of CPT2 in mice (Cpt2Sk−/−) results in reduced running distance at high [3] and low speed (Figure 1A). Specifically, control mice completed a 60-minute run at low speed with no signs of fatigue, covering 660 m (Figure 1A), whereas Cpt2Sk−/− mice could not complete the running challenge reaching exhaustion after ∼32 min at ∼300 m (Figure 1A). Because running performance is highly dependent on systemic aerobic capacity, we wanted to assess how performance was impaired in a muscle-specific manner rather than confounded by cardiovascular and respiratory system stress. Thus, we used the inverted screen test to evaluate grip strength against body weight. While body weight was not different between controls and Cpt2Sk−/− mice, Cpt2Sk−/− exhibited a significantly reduced holding impulse on the inverted screen by 45% (Figure 1B) suggesting intrinsic muscle weakness.Figure 1 Impaired long-chain fatty acid oxidation compromises contraction in a muscle-type-dependent manner. (A) Treadmill running performance in Cpt2Skf/f and Cpt2Sk−/− mice. (B) Inverted screen test performance. (C) Muscle wet mass in grams (g) for EDL and Soleus from Cpt2Skf/f and Cpt2Sk−/− mice. (D) Ex-vivo Extensor Digitorum Longus (EDL) muscle contraction. (E) Ex-vivo EDL muscle contraction normalized to muscle size. (F) Percentage change of ex-vivo contraction in Cpt2Sk−/− EDL muscle compared to control. (G) Ex-vivo soleus (SOL) muscle contraction. (H) Ex-vivo Soleus contraction normalized to muscle size. (I) Percentage change of ex-vivo contraction in Cpt2Sk−/− Soleus muscle compared to control. All data were generated in adult male mice and presented as mean ± SEM. N = 4–6. ∗P ≤ 0.05 by T-Test.

Figure 1

To isolate muscle-specific effects even further, ex-vivo analysis of contractile function was assessed in muscles containing either high percentages of glycolytic (EDL) or oxidative fibers (Soleus). Tissue wet weight was not different between control and CPT2-deficient muscles (Figure 1C). While the production of absolute force in response to electrical stimulation was not different between control and Cpt2Sk−/− EDL muscle, force normalized to muscle mass (i.e. specific force) was reduced by 10–15% in the Cpt2Sk−/− EDL at higher frequencies (Figure 1D–F). The oxidative soleus muscle of Cpt2Sk−/− mice demonstrated severe impairments in muscle force production across all stimulating frequencies. CPT2-deficient soleus was largely unresponsive across all stimulating frequencies, with absolute and specific force reduced by 60–70% compared to control muscle (Figure 1G–I). Together, these data suggest that loss of CPT2 and mitochondrial fatty acid oxidation in skeletal muscle reduces the capacity for physical activity and causes muscle weakness, particularly in oxidative muscles. Combined, these data demonstrate that although CPT2-deficient mice present overall muscle weakness and exercise intolerance, contractile capacity is impaired in a muscle-type-specific manner, predominately in oxidative muscles.

2.2 Loss of CPT2 does not impair the bioenergetic capacity of skeletal muscle mitochondria

CPT2 deficiency is an inborn error of metabolism traditionally classified as an energy disorder since it obliterates the mitochondria's capability to utilize long-chain fatty acids to support ATP synthesis. We have previously reported metabolic adaptations in soleus muscle, but not EDL muscle, that improve capacity for oxidation of non-fatty acid substrates as metabolic compensation [3]. Here, to reiterate any contribution of bioenergetic limitations to the reduced exercise capacity, respiration of mitochondria isolated from the red gastrocnemius (RGa) were determined when energized with pyruvate/malate, octanoylcarnitine/malate, palmitoylcarnitine, or succinate/rotenone across a range of free energy demands. Evaluation of oxygen consumption over time (JO2) across a range of energy demands and fueled by different substrates show that only respiration on palmitoyl-carnitine/malate is limited in RGa muscle mitochondria while oxidation of non-fatty acid substrates was preserved (Figure 2A–I). These data suggest relatively preserved mitochondrial oxidative capacity due to the loss of CPT2. An increased reliance on glucose of Cpt2Sk−/− compared to controls is demonstrated by modest hypoglycemia after a bout of exercise to exhaustion (Figure 2J); however, the relatively preserved glycemia values after high intensity exercise suggests a lack of overall energetic insufficiency.Figure 2 Bioenergetic capacity of CPT2 deficient skeletal muscle mitochondria. (A–D) Representative traces of oxygen consumption rates (JO2) in isolated mitochondria from Cpt2Skf/f and Cpt2Sk−/− red gastrocnemius muscle (RGa) energized with pyruvate (Pyr), octanoylcarnitine (OctC), palmitoylcarnitine (PC), or succinate (S) with rotenone (Rot) and malate (M) across energy demands control by phosphocreatine (PCr) and uncoupling induced by carbonyl cyanide p-trifluoromethoxyphenylhydrazone (FCCP). Substrate-supported oxygen consumption (JO2) under low (E) or high (F) energy demands and the fold-change from low to high energy (G) in isolated mitochondria from RGa. Comparison of mitochondrial oxygen consumption rates at low (H) or high (I) energy demands between genotypes for RGa muscle presented as Cpt2Sk−/− percentage change from control. (J) Blood glucose immediately after a high-intensity treadmill challenge. (K) Rate of ATP production (JATP) and (L) ATP production per oxygen molecule (P/O ratio) on different energetic substrates. Data were generated in adult male mice and presented as mean ± SEM. N = 5–6. ∗P ≤ 0.05 by T-Test.

Figure 2

To further explore bioenergetics, mitochondrial membrane potential (MP) was determined via TMRM fluorophore in control and Cpt2Sk−/− muscle as an indicator of how energy supply and Electron Transport Chain activity compare between genotypes. Mitochondria derived from white quadriceps and red gastrocnemius muscles of CPT2-deficient mice were able to generate membrane potential in a similar fashion to controls when energized with pyruvate/malate, octanoylcarnitine/malate, and succinate/rotenone and across a range of free energy demands (Figs. S1A and B). Unlike white Quadriceps and red gastrocnemius, mitochondria isolated from oxidative Soleus muscle displayed genotype effects, with Cpt2Sk−/− muscle having significant, or trending, higher TMRM intensity as percentage of alamethicin during pyruvate, octanoyl-carnitine, and succinate oxidation compared to control (Fig. S1C). These data agree with our previous report of preserved mitochondrial bioenergetic capacity in Cpt2Sk−/− mixed muscles, and compensatory increased respiration in the highly oxidative Cpt2Sk−/− soleus mitochondria with non-fatty acid substrates [3], suggesting substrate-supported oxygen consumption is sufficiently feeding the mitochondrial electron transport chain.

To directly assess energy production in skeletal muscle, the coupling of ATP phosphorylation to oxygen consumption during pyruvate, octanoyl-carnitine, and succinate oxidation (P/O ratio) under high energy demand conditions were determined. Here, mitochondrial ATP production (JATP) (Figure 2K) and ATP produced per oxygen molecule consumed (P/O ratio) (Figure 2L) were similar between Cpt2Sk−/− and control in both glycolytic white Quadriceps, mixed red gastrocnemius, and highly oxidative Soleus muscles except for a minor reduction in P/O ratio in Cpt2Sk−/− Soleus and red gastrocnemius stimulated with pyruvate. The discrepancy between higher membrane potential but lower P/O in soleus for pyruvate may reflect TMRM readout interference due to changes in calcium homeostasis, ion balance, acylcarnitine accumulation, or mitochondrial structure. Ultimately, these data suggests that the capacity to synthesize ATP in CPT2-deficient muscles is overall preserved. In agreement, blood glucose levels are not limiting immediately after a high-intensity treadmill challenge in Cpt2Sk−/− mice (Figure 2J). Although CPT2-deficient mice had lower post-exercise glycemia than control mice by 23%, the average absolute value remained well above 100 mg/dl, thus ruling out hypoglycemia as the cause for impaired exercise tolerance. These findings demonstrate that Cpt2Sk−/− oxidative muscles undergo metabolic adaptations that preserve mitochondrial bioenergetics, suggesting that energy insufficiency is not the sole cause of contractile impairment.

2.3 Loss of CPT2 deregulates the myofibril-sarcolemma-ECM anchorage system in Soleus muscle

For muscle contraction to be effective, the mechanical force generated by the myofibers must be laterally transmitted to the surrounding connective tissue and longitudinally to the myotendinous junction [5]. Hence, the myofibrils are properly anchored to the extracellular matrix (ECM) via cytosolic and transmembrane proteins [6] (Figure 3A). Proteome analysis was used to determine if loss of CPT2 affects proteins associated with the myofibril-sarcolemma-ECM anchorage system. Compared to control, Cpt2Sk−/− oxidative Soleus muscle had significant reductions in dystrophin and dystrophin-linked proteins which connect the myofibrils to the sarcolemma (Figures 3B,S1D). Proteins involved in cell-ECM interaction like laminin and biglycan were reduced in the Cpt2Sk−/−, as were ECM proteins such as collagens, prolargin, and asporin (Figures 3B,S1D). In agreement with the proteomics data, immunofluorescent histological detection of dystrophin protein in muscle sections revealed a non-uniform expression with faint or incomplete delineation of myofibers in Cpt2Sk−/− soleus muscle compared to control (Fig. S1E). This data demonstrates that loss of CPT2 in oxidative muscles negatively affects the structural integrity of myofibers by downregulating the myofibril's anchorage system, which, in turn, is expected to impair effective muscle contraction.Figure 3 In oxidative muscles, loss of CPT2 deregulates the myofibril-sarcolemma-ECM anchorage system and disrupts the contractile machinery. (A–C) Expression abundance heatmaps of proteins involved in the ECM-sarcolemma-myofibril anchorage system in Cpt2Skf/f and Cpt2Sk−/− soleus muscle; asterisk denotes significant differences (q < 0.1) between genotypes. (D) TEM images of Cpt2Skf/f and Cpt2Sk−/− Soleus muscle. Data was generated in adult male mice and presented as Log2 abundance. ∗adjusted p-value (q < 0.1) calculated by Benjamini Hochberg FDR correction. N = 6 for A-C. For (C) magnification 20× and scale bar = 200 μm.

Figure 3

2.4 Loss of CPT2 disrupts the contractile machinery in Soleus muscle

While several studies have focused on the metabolic consequences of inherited structural myopathies like Duchene's and Becker's Muscular Dystrophies [[7], [8], [9]], little is known about if and how metabolic myopathies can directly affect contractile proteins. Control and Cpt2Sk−/− muscle proteome analysis revealed that sarcomere proteins were significantly reduced in CPT2-deficient soleus (Figure 3A–C). Specifically, expression of scarcoglycans, laminins, troponins, and myosins, were significantly lower in Cpt2Sk−/− soleus compared to control (Figure 3A–C). Thin filaments are tethered together at the sarcomere's Z-disc by α-actinin. Compared to the control, slow isoform α-actinin 2 and Z-disc accessory and scaffold proteins were significantly downregulated in Cpt2Sk−/− soleus muscle (Figure 3A–C). In agreement, classical myopathic features confirmed the presence of ragged red fibers in Cpt2Sk−/− Soleus muscles compared to control (Fig. S2A) [3]. Staining revealed reduced dystrophin intensity, in agreement with proteomic data, in Cpt2Sk−/− Soleus muscle (Fig. S2B). Cpt2Sk−/− Soleus muscle showed a 10-fold increase in central nuclei, 1.44%, compared to 0.12% in control muscle (Fig. S2C). This modest increase in central nuclei of CP2Sk−/− mice was not considered severe muscle remodeling because levels of Creatine Kinase in the serum of CP2Sk−/− mice were not different from control (Fig. S2D). These data suggest that reduced expression of contractile machinery contributes to muscle weakness.

To understand the ultrastructural bases of CPT2-derived metabolic myopathy, TEM was performed with longitudinal sections revealing that compared to control, sarcomeres in Cpt2Sk−/− Soleus muscle were misaligned, lacking continuity of the Z-disk across myofibrils (Figure 3D, red arrowheads). Additionally, transverse sections demonstrated loss of regular spacing and organization of myofibrils in CPT2-deficient soleus compared to control (Figure 3D). These data suggest that impairing mLCFAO via loss of CPT2 compromises the muscle's elemental contractile structure -the sarcomere- via severe downregulation of critical proteins and structural disorganization.

TEM images also revealed high mitochondrial content in the Cpt2Sk−/− soleus muscle, for both subsarcolemmal and intermyofibrillar populations (Figure 3D), in agreement with our previous findings of increased mitochondrial protein and DNA [3]. Soleus muscle of control mice have intermyofibrillar mitochondria adjacent to the sarcomere I band forming a single row (Figure 3D, blue arrowheads), whereas Cpt2Sk−/− muscles display large clusters of mitochondria 3 to 5 rows thick that cover the sarcomere's total length (Figure 3D, red asterisks). The subsarcolemmal mitochondria in Cpt2Sk−/− Soleus muscle occupied over 10 rows adjacent to the plasma membrane compared to 4–5 rows in the control muscle (Figure 3D, yellow asterisks). Importantly, Cpt2Sk−/− mitochondria displayed preserved structure, well-defined, thick cristae (Figure 3D, white arrowheads), and a high abundance of native matrix granules (Figure 3D, black arrows), suggesting high abundance of healthy mitochondria. Coinciding with increased mitochondria, we observed mitophagy-related vacuoles at different maturity states in Cpt2Sk−/− Soleus muscle (Figure 3D, green arrows). Together, this data suggests that in oxidative muscles, loss of CPT2 coincides with downregulation of proteins involved in sarcomere anchorage, structure, and contraction, which is further compounded by excessive mitochondrial biogenesis that seemingly disrupts organization and reduces levels of the muscle's myofibril.

2.5 CPT2 loss in muscle increases phospholipid content and alters acyl-chain composition

Loss of CPT2 in muscle's mLCFAO pathway results in the accumulation of long-chain acylcarnitines, which may or may not flux into complex lipids such as phospholipids and triacylglycerol [10]. In the Soleus muscle, histological sections stained for complex lipids with Oil-Red-O (ORO), Sudan Black, and Nile Red showed increased dye intensity in all myofibers of Cpt2Sk−/− mice compared to control, suggesting higher lipid content across all fibers within the muscle (Figure 4A). The Cpt2Sk−/− EDL muscle showed increased lipophilic dye intensity compared to the control but only in a subset of myofibers of smaller diameter indicative of more oxidative fibers (Figure 4A). TAG quantification showed no change between Cpt2Sk−/− and control soleus (Figure 4B). Because these lipophilic stains are not exclusive to neutral lipids, such as TAGs, but also react to phospholipids (PLs) [11,12], direct injection mass spectrometry confirmed that Cpt2Sk−/− soleus muscle had increased phospholipid content for phosphatidylcholine (PC), phosphatidylethanolamine (PE), sphingomyelin (SM), and phosphatidylglycerol (PG) by ∼2 to 3-fold compared to controls (Figure 4C–F). However, phosphatidylinositol (PI), phosphatidylserine (PS), and ceramides were not different between genotypes (Figure 4G–I). Analysis of individual lipid species revealed that PLs containing 1–4 unsaturation were higher in Cpt2Sk−/− soleus muscle and PLs containing 5 or more unsaturations were lower in CPT2-deficient muscle compared to control (Fig. S3). Lipidomic analysis of isolated mitochondria showed that the most abundant cardiolipin species, CL(72:8) containing four linoleic-acid residues (18:2), was significantly elevated by 2.7-fold in Cpt2Sk−/− soleus muscle compared to control (Fig. S3). This large shift in CL(72:8) content, going from 20% to 60% of total CLs in Cpt2Sk−/− soleus muscle, resulted in subsequent relative decrease in many of the less abundant CL species enriched in polyunsaturated fatty acids, such as CL(76:12), CL(76:11), CL(72:6), CL(74:9) and CL(76:10) (Fig. S3). Together, these shifts in acyl content suggest increased incorporation of modestly saturated fatty acids into complex lipids, perhaps due to reduced flux of such lipids through fatty acid oxidation. Additionally, this data demonstrates that the lipophilic aggregates observed in muscle upon loss of CPT2 are not neutral lipids but phospholipids and cardiolipin reflecting increased membrane content due to mitochondrial biogenesis.Figure 4 Upon loss of CPT2, oxidative myofibers accumulate polar lipids. (A) Lipid visualization by Oil-Red-O (ORO) (upper 4 images), Sudan Black (middle 4 images), and Nile Red (lower 4 images) in soleus and EDL muscle. (B) Relative abundance of triacylglycerides (TAGs) in Soleus. (C–I) Relative abundance of phosphatidylcholine (PC), sphingomyelin (SM), phosphatidylethanolamine (PE), phosphatidylglycerol (PG), phosphatidylinositol (PI), phosphatidylserine (PS), and ceramides in Soleus muscles. Data was generated in adult male mice and presented as mean ± SEM. N = 3–4. ∗P ≤ 0.05 by T-Test. For (A) magnification 20× and scale bar = 200 μm.

Figure 4

Lipid accumulation is known to trigger deleterious immune or oxidative stress responses mostly classically associated with accumulation of TAGs and ceramides [13]. To determine if the accumulation of long-chain acylcarnitines and polar lipids was associated with stress responses, immunohistochemistry against the macrophage surface marker CD68 in muscle cross-sections was performed. There was no difference in macrophage infiltration between Cpt2Sk−/− and the control Soleus muscle (Fig. S4A, white arrowheads), suggesting no active inflammatory response. Additionally, several antioxidant and stress response proteins were upregulated in Cpt2Sk−/− Soleus, suggesting a protective adaptation (Fig. S4B). Specifically, members of the cytosolic and mitochondrial antioxidant systems and proteins that metabolize hydrogen peroxide and lipid hydroperoxides were up by ∼2-fold in Cpt2Sk−/− Soleus muscle compared to control (Fig. S4B). In summary, this data suggests that upon loss of CPT2, Soleus oxidative muscle uses its antioxidant systems to maintain redox homeostasis.

2.6 The loss of CPT2 impacts intracellular calcium homeostasis

Myofibril contractile synchronism is facilitated by a series of proteins that tether the SR to the membrane's T-tubules, facilitating the coupling of membrane depolarization to SR calcium release [14]. Disruptions in this highly organized structure impair muscle contraction [15,16]. In Cpt2Sk−/− Soleus muscle, numerous SR tethering proteins were significantly less abundant than in control (Figure 5A). Visualization of muscle ultrastructure by TEM demonstrated that myofibril bundles in Cpt2Sk−/− Soleus were surrounded by abnormal yet organized vesicle-like structures, not observed in control muscle and indicative of distended SR (Figure 5B, red arrows). This data suggests that loss of CPT2 in oxidative muscles leads to SR's structural abnormalities, likely due to a reduced tethering system.Figure 5 The loss of CPT2 disrupts SR structure and intracellular calcium homeostasis. (A) Expression abundance heatmaps of SR tethering proteins in Cpt2Skf/f and Cpt2Sk−/− soleus muscle; asterisk denotes significant differences (q < 0.1) between genotypes. (B) Transverse TEM images of Cpt2Skf/f and Cpt2Sk−/− Soleus, red arrows indicate dilated SR. (C) Expression abundance heatmaps of calcium handling proteins in Soleus muscle, asterisk denotes significant differences (q < 0.1) between genotypes. (D) Representative traces of muscle force production over time and at peak rate of relaxation in control and Cpt2Sk−/− Soleus muscle stimulated with 100 Hz. (E, F, G) Relative calcium uptake over time, area under the curve (AUC), and time to half-life (1/2 Life) in homogenate from Cpt2Skf/f and Cpt2Sk−/− Soleus muscle. N = 5–6. For (A) data are presented as Log2 abundance. ∗adjusted p-value (q < 0.1) calculated by Benjamini Hochberg FDR correction. For (D–G), data are presented as Mean ± SEM and ∗P ≤ 0.05 by T-Test.

Figure 5

Skeletal muscle contraction and relaxation relies on cytosolic calcium cycling by the concerted opposite actions of proteins mediating calcium release and uptake. Compared to the control, calcium release mediating proteins and cytosolic calcium-binding proteins were significantly reduced by ∼20% in Cpt2Sk−/− Soleus muscle (Figure 5C). On the contrary, the SR-resident calcium retentive protein Calsequestrin was upregulated in CPT2-deficient Soleus compared to the control, but sarcoendoplasmic reticulum Ca2+-ATPase, slow isoform (SERCA2) was not different between genotypes (Figure 5C). Several mitochondrial calcium homeostasis regulating proteins were upregulated in Cpt2Sk−/− Soleus (Figure 5C). Peak rate of relaxation [17] trended slower in Cpt2Sk−/− soleus muscle (Figure 5D), suggesting altered calcium homeostasis in CPT2-deficient Soleus. To determine if the alterations in the SR-bound calcium-handling machinery of Cpt2Sk−/− mice impacted calcium homeostasis, ATP-stimulated free calcium uptake in muscle homogenates of control and Cpt2Sk−/− mice, which is an indicator of SERCA activity [18], revealed a rightward shift in Cpt2Sk−/− Soleus muscle compared to control (Figure 5E). Area under the curve (AUC) and time to half-life (½ Life) of calcium uptake were significantly increased by 22% and 66%, respectively, in Cpt2Sk−/− Soleus (Figure 5F–G), suggesting a slower calcium clearance from the cytosol into the SR. This perturbance in calcium uptake aligns with the muscle force decline observed in CPT2-deficient Soleus muscle (Figure 1). On the contrary, the rate of calcium uptake was modestly reduced in CPT2-deficient EDL compared to control (Figs. S5A–C). The minimally disrupted calcium handling in Cpt2Sk−/− EDL agrees with the minimally affected force (Figure 1). In mitochondria, calcium retention capacity from Cpt2Sk−/− Soleus muscle was lower than control (Fig. S5D), an effect which was reversed by the addition of the calcineurin inhibitor, cyclosporin A (Fig. S5E). Sodium-stimulated calcium efflux via the Na/Ca exchanger revealed larger free calcium mitochondrial load in Cpt2Sk−/− Soleus muscle (Fig. S5F). Efflux was quickly reversed by addition of free calcium to the media demonstrating mitochondrial viability. Additionally, calcium uptake assays conducted in permeabilized muscle fiber bundles from Cpt2Sk−/− Soleus muscle revealed a similar decreased retention capacity when the SERCA inhibitor thapsigargin was present (Fig. S5H), suggesting impaired mitochondrial retention capacity. Taken together, these results suggest that increased expression of calcium transporters in mitochondria from Cpt2Sk−/− Soleus muscle (Figure 5C) favors enhanced calcium influx which is turn triggers the mPTP. Furthermore, loss of CPT2 in the Soleus muscle impairs calcium uptake by the SR without changes in SERCA2 levels, suggesting an activity-based inhibitory effect on SERCA-mediated calcium pumping.

2.7 Long-chain acylcarnitines interfere with SR-mediated calcium dynamics in skeletal muscle

NanoDESI-based lipid imaging analyzed in alignment with myosin-based fiber identification in TA muscle demonstrated the localization of LCACs in more oxidative fiber types (Figure 6A). Palmitoyl-carnitine (C16:0) accumulated predominantly in type 2A while oleoyl-carnitine (C18:1) was abundantly present in oxidative type 2A and glycolytic type 2B and 2X fibers (Figure 6A). The Soleus muscle is predominantly composed with ∼60% type 2A fibers and ∼40% of the most oxidative type 1 fiber [3]. Whole tissue lipidomics confirmed that the oxidative Cpt2Sk−/− soleus muscle had higher accumulation of the saturated LCACs, stearoyl-carnitine (C18:0) and palmitoyl-carnitine (C16:0), compared to the TA (Figure 6B), suggesting oxidative muscles might be particularly prone to palmitoyl-carnitine accumulation. Indeed, comparison of LCAC levels between soleus and white quadriceps within the Cpt2Sk−/− mice, revealed that the Cpt2Sk−/− mice accumulate 23-fold higher C16:0-carnitine in oxidative muscle compared to glycolytic muscle (Figure 6C). These data suggest that although LCACs are elevated across CPT2-deficient muscles, palmitoyl-carnitine accumulation is greater in oxidative muscle.Figure 6 Long-chain acylcarnitines interfere with SR-mediated calcium dynamics in skeletal muscle. (A) Immunohistochemistry of myosin heavy chain type 2A (green), type 2B (red), type 2X (black), and dystrophin (yellow) in TA muscle of Cpt2Sk−/− mice. Palmitoyl-carnitine (C16:0) and oleoyl-carnitine (C18:1) detection by mass-spectroscopy-based lipid scanning in TA muscle of Cpt2Sk−/− mice. (B) Relative abundance of LCACs in TA and soleus (SOL) muscle of Cpt2Skf/f and Cpt2Sk−/− mice. (C) Abundance of palmitoyl-carnitine (C16:0) and oleoyl-carnitine (C18:1) relative to internal standards (IS) in Cpt2Sk−/− white quadriceps (WQuad) and soleus. (D) Representative traces of calcium uptake and release in isolated sarcoplasmic reticulum (SR) from control muscle in the absence of fatty acids, and in the presence of palmitoyl-carnitine (PC) or octanoyl-carnitine (OctC). (E) Quantitation of the calcium uptake and release slope over time relative to protein concentration. Data was generated in adult male mice. Data are presented as mean ± SEM and ∗P ≤ 0.05 by T-Test or 1-way ANOVA. N = 3–6.

Figure 6

Palmitoyl-carnitine interferes with calcium homeostasis in cardiac muscle by stimulating calcium release from the myocardial sarcoplasmic reticulum [19,20]. To test if excessive accumulation of palmitoyl-carnitine could impact calcium homeostasis in muscle, the impact of palmitoyl-carnitine on calcium uptake and release in isolated SR was determined. Pre-incubation of SR with 10 μM palmitoyl-carnitine did not impact calcium release, but reduced calcium uptake by ∼70% (Figure 6D,E). The selective SERCA inhibitor cyclopiazonic acid (CPA) inhibited calcium uptake similarly to palmitoyl-carnitine (Fig. S6A). Palmitoyl-CoA similarly inhibited SR calcium uptake, whereas free palmitate did not (Figs. S6B and C). Octanoyl-carnitine (C8-carn) (Figure 6D), a medium-chain acylcarnitine did not affect calcium uptake or release by the SR (Figure 6E). These data suggest calcium dynamics are regulated by long-chain fatty acids as carnitine and CoAs esters. Calcium uptake and release in SR isolated from Cpt2Sk−/− muscles was not different from controls (Figs. S6D and E), suggesting that the SR-bound machinery required for cytosolic calcium clearance is structurally preserved and functionally viable in CPT2-deficient muscles. These results suggest that disrupted calcium homeostasis in CPT2-deficient skeletal muscle, is mediated in part by the accumulation of long-chain palmitoyl-carnitine interference on SR calcium uptake.

3 Discussion

The loss of CPT2 and mLCFAO in soleus muscles causes a metabolic shift towards a glycolytic phenotype demonstrated by increased mitochondrial biogenesis and pyruvate and succinate oxidative capasity [3]. Here, we demonstrate respiration of red gastrocnemius Cpt2Sk−/− mitochondria is preserved across a range of energy demands and not different from control and that oxidative phosphorylation-coupled ATP production is preserved in Cpt2Sk−/− muscles. However, Cpt2Sk−/− soleus tethering system connecting myofibrils to the sarcolemma and the extracellular matrix was severely decreased. Reduced anchoring proteins included components of the dystrophin-node [6], transmembrane glycoproteins, laminin, and extracellular collagen I, VI, and XII. Mutations in these anchoring proteins cause congenital muscle myopathies with muscle weakness [[21], [22], [23]]. Specifically, mutations in dystrophin, utrophin, sarcoglycans, and laminin are the hallmark of inherited muscle disorders such as Duchenne's, Becker's, and Limb-Girdle muscular dystrophies and Sarcoglycanopathies [24] presenting with progressive muscle weakness, impaired exercise tolerance, and reduced mobility. Interestingly, several recent reports have demonstrated that the loss of structural membrane proteins disrupts cellular metabolism. For instance, loss of dystrophin in the mdx mouse is associated with reduced reliance on lipid substrates for energy production and increased oxidation of carbohydrates [[7], [8], [9]]. Similarly, in human patients affected by Duchene's and Becker's Muscular Dystrophy, muscles show differential expression of mitochondrial metabolic enzymes and respiratory complexes [25]. Thus, while evidence of how losing muscle structural proteins compromises metabolism continues to grow, data presented herein suggest that the reverse can also occur, wherein metabolic myopathies impact muscle structural integrity.

In addition to a reduced myofiber anchoring system, the deletion of CPT2 significantly decreased sarcomere-related proteins. Reduced contractile proteins, such as myosin, actin, troponin, and tropomyosin, are likely contributors to muscle dysfunction in Cpt2Sk−/− oxidative muscles. Similarly, loss of sarcomere accessory proteins can contribute to altered contractility. Precisely, the sarcomere misalignment observed in CPT2-deficient soleus muscle reflects the reduced levels of the Z-disc accessory proteins nebulin, titin, PDZ-LIM, and CapZ, and is a known structural link to muscle weakness. For example, mutations in titin cause an array of skeletal and cardiac myopathies characterized by loss of sarcomere scaffolding [26,27]. Structural abnormalities in sarcomeres and ineffective cross-bridging between filaments were also reported upon the downregulation of nebulin, CapZ, and PDZ-LIM proteins [[28], [29], [30]].

We have previously reported that CPT2-deficient muscles undergo significant mitochondrial biogenesis, visible as ragged-red fibers [3]. The excessive mitochondrial accumulation in Cpt2Sk−/− muscles likely contributes to enlarged myofibers, predominately in type IIa fibers [3]. Additionally, mitochondrial overload also likely mechanically displaces and compresses myofibrils in response to CPT2 deficiency, leading to excessive branching and further impairment of muscle contraction. Here, we further characterized myopathic features in CPT2 deficiency by evaluating central nuclei as an indicator of ongoing cycles of muscle damage and regeneration. Although 10-fold higher and statistically significant, central nuclei prevalence in mLC-FAO deficient muscles is minor compared to muscle dystrophies, such as DMD, which fluctuates between 60% and 100% of myofibers [[31], [32], [33]]. In alignment with the low prevalence of central nuclei as an indicator of scarce muscle damage–regeneration cycles, levels of Creatine Kinase in the serum of CP2Sk−/− mice were not elevated.

In skeletal muscle, the sarcoplasmic reticulum (SR) forms an interconnected network throughout the muscle cell. Junctophilins (JPHs) are structural proteins that stabilize functionally relevant contact sites between the SR and the sarcolemma where the voltage-gated calcium channel Cav1.1 interacts with the ryanodine receptor (RyR) [16]. Here, we demonstrate that junctophilins and other tether-associated proteins were downregulated in CPT2-deficient soleus muscle, which resulted in areas of dilated SR. Our findings agree with reports showing that junctophilin 1 knockout mice present with deformed triads, vacuolated SR regions, and muscle weakness [15], while decreased junctophilin 1 expression in vitro reduces calcium influx [15]. The effects of CPT2 deficiency on SR biology is not limited to disrupted anchoring to the sarcolemma and extends to SR calcium handling as demonstrated by the significant downregulation of proteins involved in calcium release such as RyR1 and the upregulation of proteins involved in calcium sequestration inside the SR such as Calsequestrin 2. Contrarily, but in agreement with increased mitochondrial content, proteins involved in mitochondrial calcium uptake like MCU, MCUR1, and VDAC are upregulated in CPT2-deficient Soleus muscle while cytosolic buffering of free calcium by binding proteins such as TNNC1, CAMK isoforms, and CALM2 seems to be compromised by downregulation in Cpt2Sk−/− Soleus compared to control. Further investigation is required to determine if changes in cytosolic and mitochondrial calcium levels can modulate rates of mitochondrial respiration in metabolic myopathies [34,35]. Several mitochondrial bioenergetic enzymes are regulated by calcium [36,37] and ETC complexes increase in the presence of calcium [38], an effect that may contribute to increased expression of glycolysis enzymes, TCA cycle enzymes, ETC complexes, and matrix dehydrogenases expression in CPT2 deficiency. While mitochondria's calcium affinity is low, its large abundance can play a significant role in calcium homeostasis in skeletal muscle. Mitochondria isolated from CPT2-deficient Soleus muscle have higher basal calcium load, impairing calcium retention during an experimental loading challenge and prematurely triggering the opening of the mPTP. Further studies to determine the exact calcium loading capacity in mitochondria derived from inherited FAODs are needed. Our findings suggest that impaired force production observed in Cpt2Sk−/− oxidative muscles is due, in part, to abnormal calcium handling during the contraction cycle. SERCA activity is fundamental in removing calcium from the cytosol to promote muscle relaxation. ATP-stimulated SR calcium uptake was slower in Cpt2Sk−/− oxidative Soleus muscle despite normal SERCA levels. This suggests that an allosteric modulation of the SERCA pump might contribute to delayed calcium clearance in CPT2-deficiency. Importantly, although ATP-synthetic capacity was preserved in Cpt2Sk−/− muscle mitochondria, it remains possible that ATP levels in CPT2-deficient Soleus muscle might be insufficient to support SERCA activity [39].

Preclinical modeling of FAODs by reducing FAO enzymes demonstrates muscle weakness, exercise intolerance, and elevated circulating LCACs as hallmarks of these metabolic myopathies. Importantly, different long-chain acylcarnitines accumulate in different FAODs, such as CPTII, CACT, MTP, and VLCAD deficiency [40] and plasma levels of these biomarkers have been reported to fluctuate in a conserved manner during metabolic perturbances such as fasting, exercise, and infection [41,42,[42], [42], [43], [44], [45]]. Here, the muscle-specific loss of CPT2 demonstrates tissue-autonomous muscle weakness, suggesting that muscle outcomes observed in human patients are a direct result of muscle FAO metabolic activity rather than systemic impacts on muscle physiology.

The accumulation of lipid species such as TAGs, DAGs, and ceramides results in muscle oxidative stress and dysfunction, as is speculated for CPT1b disorders [46,47]. However, Cpt2Sk−/− muscle does not accumulate toxic lipid species, but do accumulate acylcarnitines, the substrate of CPT2, as well as phospholipids and cardiolipin most likely due to increased mitochondrial content. Differences in enzymatic substrates between FAODs explain the stark contrast in lipid species accumulation. Specifically, the loss of CPT1 leads to the accumulation of its substrate, acyl-CoAs which are toxic and upon accumulation are metabolically cleared by incorporation into complex lipids, such as TAGs. However, the substrate for CPT2 is acylcarnitines, not acyl-CoAs, and the resulting toxic impact of acylcarnitine accumulation has remained unclear [[48], [49], [50], [51]]. We report acylcarnitine accumulation in muscle and plasma of Cpt2Sk−/− despite intact FAO in all non-muscle tissues, including the liver. These data suggest that the high levels of plasma acylcarnitines used as biomarkers to diagnose individuals with FAODs are likely significantly contributed from muscle metabolism. Regarding the potential toxicity of acylcarnitines, we found no indication of neutral lipid toxicity or immune cell infiltration in CPT2-deficient muscle. However, long-chain acylcarnitines have been shown to alter calcium dynamics in cardiac muscle under pathological conditions such as ischemia-reperfusion [18,19]. Similarly, induced pluripotent stem cell-derived cardiomyocytes from patients with VLCAD deficiency had abnormal action potentials and increased intracellular calcium levels [52]. Correlating lipid levels and contractile dysfunction in skeletal myopathies suggests an association between LCACs accumulation and calcium dyshomeostasis [19], but the molecular mechanism remains unclear. Here, we demonstrated that exposing isolated skeletal muscle SR fractions to palmitoyl-carnitine reduces SERCA-mediated calcium uptake by 70%, and in a similar fashion to the selective SERCA inhibitor. This finding suggests that the accumulation of palmitoyl-carnitine in CPT2-deficient Soleus muscle may limit free cytosolic calcium, thus compromising muscle force production and leading to overall weakness in the Cpt2Sk−/− mice. Furthermore, palmitoyl-carnitine has been shown to interact directly with the RyR, leading to a leaky SR [53]. Although we have not tested whether this mechanism is present in CPT2-deficient mice, a leaky RyR could potentiate the palmitoyl-carnitine-mediated SERCA impairment observed in Cpt2Sk−/− Soleus, resulting in further muscle contractile dysfunction. On the contrary, we ruled out medium-chain acylcarnitines as contributors to mitochondrial dysfunction and calcium dyshomeostasis in CPT2-deficient muscles by demonstrating that isolated SR exposed to high concentrations of the 8-carbon-long octanoyl-carnitine does not affect calcium uptake or release.

In summary, loss of CPT2 greatly compromises muscle force production in oxidative soleus muscles through several likely mechanisms including deregulated myofibril tethering, reduced sarcomeric protein content, excessive mitochondrial content, and calcium dyshomeostasis. Furthermore, this work shows that palmitoyl-carnitine interferes with calcium uptake by the SR protein SERCA. Whether adaptation to CPT2 deficiency in preclinical animal modeling is translatable to human patients remains to be investigated. It is important to highlight that the bioenergetic impact of impaired mLCFAO in muscle is likely extensive, yet several additional pathophysiological mechanisms of contractile dysfunction are demonstrated with potentially relevant therapeutic implications.

4 Methods

4.1 Animal model

Skeletal muscle-specific carnitine palmitoyltransferase 2 deficient mice (Cpt2Sk−/−) were generated as described [3,4,54]. The conditional knock out is under the control of the human alpha-skeletal actin promoter (The Jackson Laboratory; Stock No: 006149). Control mice were littermates lacking the Cre transgene. Mice had free access to water and standard chow (PicoLab 5053, Lab Diets) in a pathogen-free housing under 12-h light–dark cycles. All procedures were performed in adult male mice (3–8-month-old) and were approved by the Institutional Animal Care and Use Committee of East Carolina University (Assurance A3469-01).

4.2 Physical activity

Four-month-old male mice were acclimated to the treadmill for two days by alternating 5-minute intervals of no speed, walking speed (3 m/min), and low running speed (10 m/min). The low-intensity exercise protocol was initiated at a speed of 8 m/min and transitioned to a final speed of 12 m/min within the initial 5 min of the trial. Gentle running encouragement was used, and mice were run until exhaustion or for 60 min, whichever occurred first. All acclimation and running sessions were performed during the dark cycle. Blood glucose immediately post-exercise was determined using a glucometer (NovaMax, Billerica, MA). The inverted screen test was adapted from Deacon [55]. A screen of wire mesh of 1 mm diameter surrounded by wooden beading was suspended 14 inches above a surface conditioned with pads and soft bedding. A mouse was placed in the center of the screen and the screen was flipped. Time until fall was recorded. Mice were returned to home cages for at least 5 min before repeating the test. A minimum of 3 trials per mouse were obtained and averaged. Body weight was recorded immediately before the beginning of the assay. Holding impulse was calculated as holding time in seconds multiplied by body weight in grams.

4.3 Muscle contraction

Ex-vivo electrophysiological analysis of muscle contractility was performed utilizing a force-frequency protocol as previously described [56]. EDL and soleus muscle were dissected, freed of any remaining connective and fat tissue, and prepared for mounting by tying sutures to the proximal and distal tendons. Muscles were then immediately transferred to a chamber containing oxygenated (95% O2/5% CO2) Krebs Ringer Buffer (KRB—[mM] 115 NaCl, 2.5 KCl, 1.8 CaCl2, 2.2 Na2HPO4, 0.85 NaH2PO4) at room temperature and tied to the force transducer. Optimal resting tension (Lo) was investigated and set for each muscle. After a 10-minute equilibration, the force-frequency response was determined by stimulating the muscles 60 s apart at 10, 20, 40, 60, 80, 100, and 120 Hz. After 1 min of resting, fatigue resistance was assessed by stimulating the muscles at 30 Hz every 2 s for 600 s for 300 contractions. Once contraction protocols were finalized, the muscle L0 was determined using digital calipers, and its tendon-free, wet mass was recorded. Specific muscle force in Newtons per square centimeter (N/cm2) was calculated using predetermined equations [57].

4.4 Mitochondrial bioenergetics

Mitochondria were isolated from white quadriceps, red gastrocnemius, and soleus muscles by trypsin-mediated digestion of connective tissue and subsequent differential centrifugation in KMEM buffer (KCl 100 mM, MOPS 50 mM, EGTA 1 mM, MgSO4 5 mM, BSA 0.2% pH 7.1) and yield was determined using the BCA method as previously described [3]. Because the soleus muscle and the red gastrocnemius are relatively small compared to the other muscles (≈8 mg) tissue from 3 to 4 littermate mice with the same genotype were pooled to secure sufficient isolated mitochondria for all downstream applications and considered as one biological replicate. A total of 4–6 biological replicates were analyzed per muscle type and genotype.

High-resolution respirometry was performed on freshly isolated mitochondria using an Oroboros® device as reported previously [58]. Oxygen consumption (JO2) was measured in ATP containing buffer Z (K-MES 105 mM, KCl 30 mM, KH2PO4 10 mM, MgCl2 5 mM, EGTA 1 mM, BSA 2.5 g/L, 5 mM creatinine, 500 mM ATP, 2000u/mL CK, 750 mM Tris-PCr, pH 7.1) using 25–50 μg of isolated mitochondria in each chamber depending on muscle type and substrate. After baseline mitochondrial respiration was established, malate (2 mM, Sigma M1296) was added to maintain the TCA cycle flux. Substrates were added in the following concentrations: octanoylcarnitine 0.2 mM (Sigma 50892), palmitoylcarnitine 0.02 mM (Sigma P1645), pyruvate 5 mM (Combi-Blocks, QA1116), or succinate 10 mM (Fisher BP336) plus rotenone 0.005 mM (Sigma R8875). A modified version of the creatine kinase clamp technique was used to determine oxygen consumption in response to changes in ATP free energy. Here, the respiration assay buffer Z was supplemented with 20 U/mL of creatine kinase (CK), 5 mM ATP and 1 mM Tris-Phosphocreatine (PCr) to stimulate maximal demand for ATP synthesis. Sequential additions of PCr to 6, 9, 15, 21, 24 and 30 mM allowed for gradual reduction of the ATP demand state, as described before [59]. After minimal respiration was achieved, 3 mM FCCP (Sigma C2920) was added to the chamber to determined uncoupled respiration rates.

The transmembrane electric potential was determined using a QuantaMaster Spectrofluorometer (Horiba Scientific) as described here [58]. Briefly, 20 μg of isolated mitochondria were suspended in 200 μL of buffer Z at 37 °C (K-MES 105 mM, KCl 30 mM, KH 2 PO 4 10 mM, MgCl 2 5 mM, EGTA 1 mM, BSA 2.5 g/L, pH 7.1) and supplemented with 0.2 μM tetramethylrhodamine methyl ester (TMRM), 5 mM creatine, 20 U/mL of creatine kinase (CK), 5 mM ATP, and 1 mM Tris-Phosphocreatine (PCr) to stimulate maximal demand for ATP synthesis. Membrane potential was supported by mitochondrial respiration on the following substrates: octanoylcarnitine 0.2 mM (Sigma 50892) plus malate (2 mM, Sigma M1296), pyruvate 5 mM (Combi-Blocks, QA1116) plus malate (2 mM, Sigma M1296), or succinate 10 mM (Fisher BP336) plus rotenone 0.005 mM (Sigma R8875). Sequential additions of PCr to 6, 9, 15, 21, 24, and 30 mM allowed a gradual reduction of the ATP demand state. The ΔGATP at each titration point was calculated using the online bioenergetics calculator tool described in this reference [60]. Next, 25 μM oligomycin (Tocris 4110), 10 mM potassium cyanide (Sigma 60178), and 0.03 mg/mL alamethicin (Sigma A4665) were sequentially added to simulate conditions of maximum, minimum, and abolished membrane potential. The fluorescence ratio of the following excitation/emission parameters [Ex/Em, (572/590)/(551/590)] was determined for each step of the assay and compared to the values recorded upon the addition of alamethicin (theoretically null membrane potential).

The rate of ATP synthesis (JATP) in relation to oxygen consumption (JO2) in isolated mitochondria was assessed via glucose-6-phosphate dehydrogenase (G6PDH)-mediated NADP + reduction to NADPH coupled to hexokinase (HK)-mediated ATP hydrolysis as described here [61]. Briefly, 7.5 μg mitochondria were resuspended in 1.5 mL buffer Z at 37 °C and supplemented with 0.1 mM P1,P5-di(adenosine-5′) pentaphosphat (AP5A, Sigma D4022), 1 U/mL HK (Sigma H4502), 2.5 U/mL G6PDH (Sigma G5885), 2.5 mM d-Glucose (Sigma G5767) and 2.5 mM NADP+ (Sigma 481971) and respiration was stimulated with either pyruvate/malate, octanoylcarnitine/malate or succinate/rotenone at the concentrations described above plus 200 μM ADP. NADPH auto-fluorescence and O2 concentration were measured continuously using a customized system that integrates monochromatic fluorescence (FluoroMax-3, Horiba Jobin Yvon, Edison, NJ) with high-resolution respirometry (Oroboros Oxygraph 2k, Innsbruck, Austria). The ATP synthesis as a function of oxygen consumption (P/O ratio) was calculated as JATP divided by JO2 for each substrate.

4.5 Proteomics

TMT quantitative proteomics was performed in soleus muscle from 5-month-old male control and Cpt2Sk−/− mice as previously described [3,58]. In brief, muscles were lysed in ice-cold 8 M Urea Lysis Buffer supplemented with a protease inhibitor, snap-froze at −80 °C and freeze-thaw 3 times before sonicating with a probe sonicator in three 5 s bursts set at an amplitude of 30 (Q Sonica, Newtown, CT). Samples were cleared by centrifugation at 10,000×g for 10 min at 4 °C. Samples were subsequently reduced and alkylated. Initial digestion was performed with Lys C (Thermo Fisher) 1:100 w/w; followed by overnight incubation with trypsin (Promega, Madison, WI) 50:1 w/w, protein:enzyme at 32 °C. Samples were acidified to 0.5% TFA and centrifuged at 10,000×g for 10 min at 4 °C to pellet insoluble material. The supernatant containing soluble peptides was desalted on a 50 mg tC18 SEP-PAK solid phase extraction column (Waters, Milford, MA) and eluted. The 1.5 mL eluate was frozen and dried down via SpeedVac.

TMT labeling was performed as described [3,62]. Briefly, dried-down peptides were re-suspended in triethylammonium bicarbonate (TEAB), mixed with a unique 10-plex Tandem Mass Tag (TMT) reagent (Thermo Fisher), and shaken for 4 h. Following quenching with 50% hydroxylamine, samples were frozen and placed in SpeedVac. Samples were re-suspended in 0.5% TFA and again subjected to solid phase extraction but with a 10 mg tC18 SEP-PAK SPE column (Waters). The multiplexed peptide sample was subjected to high pH reversed-phase fractionation (Thermo Fisher; Catalog #84868) into 8 fractions. Following elution, fractions were frozen and placed in SpeedVac.

nLC-MS/MS was performed as described [3,62]. In brief, peptide fractions were quantitated and suspended in 0.1% formic acid at a concentration of 0.25 μg/μL. NanoLC-MS/MS analysis was performed using an UltiMate 3000 RSLCnano system (Thermo Fisher) coupled to a Q Exactive PlusHybrid Quadrupole-Orbitrap mass spectrometer (Thermo Fisher) via nanoelectrospray ionization source. MS1 was done at 70,000 resolution, with an AGC target of 1 × 10E6 ions and a maximum IT of 60 ms. MS2 spectra were collected by data-dependent acquisition (DDA) of the top 20 most abundant precursor ions with a charge greater than 1 per MS1 scan, with dynamic exclusion enabled for 30 s. Precursor ions were filtered with a 1.0 m/z isolation window and fragmented with a normalized collision energy of 30. Similarly, MS2 scans were done at 17,500 resolution, AGC target of 1 × 10E5 ions, and a maximum IT of 60 ms. Raw data was analyzed with Proteome Discoverer 2.2 (PDv2.2) set at default parameters. Data were searched against both the full mouse proteome, as well as the Mito Carta 3.0 database. Peptide spectrum matches (PSMs) were grouped into unique peptides while maintaining a 1% FDR at the peptide level. Peptides were grouped into proteins using the rules of strict parsimony, and proteins were filtered to 1% FDR using the Protein FDR Validator node of PD2.2. Results were exported as tab delimited.txt. files and statistically analyzed based on a previously described workflow [62]. First, M2 reporter (TMT) intensities were added up together for each TMT channel, and each channel's sum was divided by the average of all channels' sums, resulting in channel-specific loading control normalization factors to correct for any deviation from equal protein input in the 10-plex experiments. Reporter intensities for proteins were divided by the loading control normalization factors for each respective TMT channel. All loading control-normalized reporter intensities were converted to log2 space, and the average value from the 10 samples per kit was subtracted from each sample-specific measurement to normalize the relative measurements to the mean of each kit. Data from each kit were then combined for statistical comparisons. For comparisons, condition average, standard deviation, p-value (p, two-tailed student's t-test, assuming equal variance), and adjusted p-value (Padjusted, Benjamini Hochberg FDR correction) were calculated. A total of 6 biological replicates were analyzed genotype. The entire proteome data set has been made available online using accession number ‘PXD030896’ for Proteome Xchange [63]and accession number ‘JPST001445’ for jPOST Repository [64].

4.6 Histology

Soleus, EDL, and TA muscles from 5-month-old male mice were collected immediately after euthanasia, covered with a thin layer of Optimal Cutting Temperature compound (O.C.T., Fisher Scientific), flash-frozen in liquid nitrogen-chilled isopentane, and stored at −80 °C until further processing. Serial cross-sections of 10 μm thickness were obtained in positively charged glass slides (Fisherbrand™ Superfrost™ Plus Microscope Slides).

General histological characteristics were visualized via Hematoxylin and Eosin staining. Briefly, sections were fixed in 4% PFA, immersed in Mayer's Hematoxylin (Sigma MHS16-500 ML) for 15 min, placed in circulating lukewarm tap water until blue colors appeared, and then immersed in Eosin Y (Sigma HT110132-1L) for 20 s. Sections were then dehydrated in increasing concentrations of ethanol and xylene and mounted with Cytoseal (Thermo Scientific 8310-16). Gomori Trichrome stain (Engel-Cunningham modification) was performed in serial cross-sections as previously described [3]. Neutral and polar lipid content in muscle was visualized by staining soleus and EDL sections with Oil Red O (Sigma O9755), Sudan Black B (Sigma 199664), and Nile Red (Sigma 72485). Sections were fixed in 4% PFA for 10 min and washed in distilled water for 5 min twice. Specific stains were conducted as follows: Oil Red O [65] and Sudan Black B [66] were prepared fresh by dissolving 0.7 g of powder stain into 100 mL of propylene glycol (Fisher Scientific P355-4) with constant stirring and heated to 100 °C for 5 min. The solutions were then filtered, cooled down, and filtered again before use. Sections were first placed in two changes of 100% propylene glycol for 5 min and then placed in either the Oil Red O solution for 30 min or the Sudan Black B solution for 7 min with agitation. Then, sections were rinsed in 85% propylene glycol and distilled water for 3 min each before mounting. The Nile Red stock solution (1 mg/mL) was prepared in acetone and then diluted 10 times in PBS to obtain a working solution [11]. Muscle sections were incubated for 15 min at room temperature in this solution and briefly washed in PBS before mounting. Sections stained with Oil Red O, Sudan Black, and Nile Red were mounted using a glycerol-based media (Sigma GG1) pre-heated at 40 °C before use. Images were acquired with a ZEISS Axio Observer microscope at 20× magnification except for immunofluorescence and Nile Red images that were captured using an EVOS FL auto microscope at 10× and 20× (Life Technologies, Inc.).

Immunohistochemical detection of dystrophin (Thermo Fisher PA5-32388), smooth muscle actin (SMA, Dako 1A4), macrophage surface marker CD68 (Invitrogen 14-0681-82), MyHC-2A (type IIa fibers; DSHB SC-71) and MyHC-2B (type IIb fibers; DSHB BF–F3) was performed by fixating muscle sections with 4% PFA for 10 min, washing in PBS for 5 min twice, blocking with 5% goat serum (Abcam ab7481) for 45 min at room temperature and overnight incubation at 4 °C with primary antibodies. The following secondary antibodies were used to visualize the different targets upon a 60-minute incubation at room temperature: Alexa fluor 647 goat anti-mouse IgG1 (A-21236) for dystrophin, Alexa fluor 568 goat anti-mouse IgG2a (Invitrogen A-21134) for SMA, Alexa Fluor 488 goat anti-rat (Invitrogen A-11006) for CD68, Alexa Fluor 488 IgG1 (A-21121) for MyHC IIa and Alexa Fluor 546 IgM (A-21045) for MyHC IIb. Coverslip was mounted with FluoroQuest™ with DAPI for nuclei visualization (AAT Bioquest), and images were acquired using an EVOS FL auto microscope and software (Life Technologies, Inc.). A total of 3 biological replicates were analyzed per muscle type and genotype.

Transmission Electron Microscopy observations were performed on 4-month-old male control and CPT2-deficient soleus muscles in both sagittal and longitudinal orientation. Samples were prepared as follows: muscles were quickly excised from the mouse under anesthesia, transferred to glass vials with 10 mL of fixative containing 2.5% glutaraldehyde (EMS), 4% formaldehyde (EMS), 0.1 M sodium cacodylate, 2 mM CaCl2) for 2 h with very gentle rocking at room temperature. Then, muscles were rinsed 3× in the same buffer for 15 min each, cut into 3 pieces of approximately 2 mm3 each (distal, medial, proximal), post-stained with reduced 1% osmium tetroxide (EMS) in 1.2% potassium ferrocyanide (Sigma) for 1 h, washed in buffer (3 × 15 min), stained en block with aqueous 1% uranyl acetate (EMS) for 40 min, washed with milli-Q water (3 × 15 min) dehydrated in graded acetone series until absolute, infiltrated in Epon 812 resin (EMS), and polymerized at 60C for 2 days. Ultrathin sections of 60 nm were obtained using a diamond knife (Diatom) on a ultramicrotome (Leica UC7), placed on 300 mesh copper grids and finally stained with uranylLess (EMS) for 5 min. Sections were observed on a Zeiss Libra 120 operating at 80 kV and photographed with a Gatan UltraScan 4000 CCD camera in full resolution, using the energy filter slit of 20 nm.

4.7 Lipid imaging

Mass spectrometry-based lipid imaging was performed as described [3]. Briefly, 10-μm-thick TA muscle sections from 4-month-old male mice were mounted onto a glass slide and subjected to nanospray desorption electrospray ionization (nano-DESI) as described [67]. Nano-DESI MSI experiments were performed on a Q Exactive HF Orbitrap mass spectrometer (Thermo Electron, Bremen, Germany) equipped with a custom-designed nano-DESI source. Mass spectra were acquired in positive mode in the range of m/z 150–1000 with a mass resolution of m/Δm = 60,000 at 412 m/z. Imaging data were acquired at a scan rate of 40 μm/s. Data processing was performed using Peak-by-Peak software (Spectroswiss, Lausanne, Switzerland) for two biological replicates per genotype.

4.8 Lipidomics

High-throughput lipid profiling was performed as previously described [68]. Briefly, direct injection mass spectrometry (DIMS) was used to identify phospholipids, ceramides, and acylcarnitines in skeletal muscles from control and Cpt2Sk−/− mice. Bligh and Dyer Method [69] was used for tissue lipid extraction and both, the lipid and the polar phases were dried separately, resuspended in ACN:MeOH:NH4Ac and directly injected via a micro-autosampler (G1377A) into a triple quadrupole mass spectrometer (QQQ6410 from Agilent Technologies, San Jose, CA). The equipment was operated in the positive ion mode and equipped with an ESI ion source. Raw data was collected in Multiple Reaction Monitoring mode and ion intensities were further normalized to milligrams of tissue or to total ion count.

Total triacylglyceride (TAG) levels in skeletal muscle tissue were determined using a colorimetric assay that detects free glycerol upon TAG enzymatic hydrolysis (Sigma TR0100). In brief, soleus muscle was homogenized in molecular biology grade water and then a mixture of chloroform:methanol (2:1) was added to the homogenate for lipid extraction. Samples were spun down at 1700 rpm at 4 °C for 5 min and the lower phase was recovered and dried down. Lipid extraction was resuspended in a mixture of tert-butanol: methanol:triton X-100 (3:1:1) and allowed to incubate for 1 h at room temperature. TAG levels were quantitated at 540 nm following the manufacturer's instructions.

4.9 Calcium assays

Calcium uptake by the sarcoplasmic reticulum in muscle homogenates was determined as described here [18]. Briefly, soleus and EDL muscles from 6-month-old male control and Cpt2Sk−/− mice were collected and homogenized in sample buffer (250 mM sucrose, 5 mM HEPES, 0.2 mM phenylmethylsulfonyl fluoride and 0.2% [w/v] NaN3, pH 7.5) using glass conical homogenizers to a 1:10 [w/v] ratio. Then, 20 μL of muscle homogenate was mixed with 200 μL of calcium uptake buffer (200 mM KCl, 20 mM HEPES, 10 mM NaN3, 5 μM TPEN, 15 mM MgCl2, pH 7), and 4 μL of the cell-impermeant calcium fluorophore Fluo-4 (Thermo F14200), vortexed and transferred to an all-black 96-well plate in duplicate. Plate reader (Molecular Devices) settings were as follows: Ex/Em 490/535, 37 °C, shaking, and data collection every 30 s. The baseline was registered for 10 min, followed by adding 4 μL of 250 mM ATP to trigger uptake. Upon signal plateau, 10 μL of 50 mM EGTA was added to determine the minimum signal value, followed by 40 μL of 100 mM CaCl2 to establish the maximum signal value. The area under the curve and the time to half-life were calculated.

Calcium uptake in isolated sarcoplasmic reticulum from 6-month-old male control and Cpt2Sk−/− mice was measured as described here [70,71]with some modifications. Briefly, sarcoplasmic reticulum fractions were isolated from a mixture of Tibialis Anterior (TA), white quadriceps (WQuad), and Gastrocnemius (GA) muscles homogenized in 10 mL of ice-cold isolation buffer (20 mM HEPES, 0.2% NaN3, 0.2 mM PMSF, pH 6.8) using a Tissue Tearor™ at medium setting. Homogenates were then centrifuged at 2,600 g for 7 min at 4 °C to remove debris. The supernatant was collected and filtered through double gauze prior to centrifuge at 10,500 g for 7 min. The resulting pellet was stored at −80 °C in suspension buffer (20 mM HEPES, 0.2% NaN3, 0.2 mM PMSF, 300 mM Sucrose, 140 mM KCl, pH 6.8). The supernatant was transferred to a new tube, and 600 KCl was added and allowed to rest on ice for 30 min with the occasional gentle mix. Following, the supernatant was ultracentrifuged at 50,000 g for 60 min in a fixed-angle rotor. The recovered pellet was resuspended in suspension buffer and stored at −80 °C until calcium uptake and release assays were performed. Protein was quantitated using Pierce™ BCA assay (Thermo Scientific A55864).

SR-mediated calcium uptake and release was determined using a QuantaMaster Spectrofluorometer (Horiba Scientific) set at 37 °C. Calcium sensor Indo-1, pentapotassium salt (Sigma I1202) was used and samples were excited at 350 μm. Changes in the 468:402 emission ratio were collected every second. Briefly, 10ug of SR protein was resuspended in 200uL of assay buffer (1 mM KCl, 20 mM HEPES, 1 mM MgCl2, 7.5 mM Na4P2O7 · 10H2O, 1 mM ATP, 1.5 μM Indo-1 pentapotassium salt, pH 7) with or without fatty acids as follows: 10 μM palmitic acid (Sigma P0500), 10 μM palmitoyl-carnitine (Sigma P1645), 5 μM palmitoyl-CoA (Sigma P9716), 20 μM octanoyl-carnitine (Sigma 50892). The baseline signal was recorded for about 5 min or until equilibrium was achieved, and then, 40 μM CaCl2 was added to initiate calcium uptake. Upon emission ratio plateaus, 141 μM AgNO3 was added to stimulate calcium release. The purity of isolated SR was assessed by calcium uptake and release assay in the presence of 50uM of the SERCA inhibitor, cyclopiazonic acid (CPA, Sigma C1530). At least 3 biological samples were tested per condition. Changes in intensity signal over time (slope) were calculated and normalized to protein concentration.

Calcium uptake in isolated mitochondria was determined via Calcium Green-5N fluorophore protocol using a QuantaMaster Spectrofluorometer (Horiba Scientific) set at 37 °C, with Excitaton/Emission:506/532, 3 s signal integration [72]. Mitochondria from Soleus muscle were isolated as described for the respiration assays and resuspended in buffer containing 125 mM KCl, 5 mM HEPES, 2 mM KH2PO4, 1 mM MgCl2, pH 7.1 [73]. Then, calcium uptake was determined in 20 μg of freshly isolated mitochondria in 200 μL of calcium assay buffer (250 mM sucrose, 10 mM tris–HCl, 20 mM tris-base, 10 mM KH2PO4, 0.5 mg/mL BSA, 20 mM creatinine, 40 μM EGTA, 5 mM 2-DeOxyGlucose, 2 U/ml Hexokinase, 1 μM Ca5N) energized with 10 mM glutamate + 1 mM malate and in the presence of 0.05 mM ADP. 35 μM of CaCl2 was initially added and then subsequent 20 μM additions were used to load the mitochondria until mPTP opening was triggered. A final 400 μM addition was used to ensure uptake was indeed terminated. Additionally, and to test baseline calcium load in isolated mitochondria, subsequent additions of 10 mM NaCl were introduced to freshly isolated mitochondria in calcium assay buffer to induce calcium efflux via NCLX, as previously described [74].

Calcium uptake in muscle fiber bundles dissected from Soleus muscle was preformed as described above for isolated mitochondria with the following modifications: before calcium uptake protocol, bundles weighing ≈0.3 mg were permeabilized in the presence of saponin for 15–30 min on rocker at 4 °C (7.2 mM K2EGTA, 2.77 mM CaK2EGTA, 20 mM imidazole, 20 mM taurine, 5.7 mM ATP, 14.3 mM PCr, 6.56 mM MgCl2–6H2O, 50 mM K-MES, 30 μg/mL saponin, pH 7.1) and then washed for 10 min in buffer (250 mM sucrose, 10 mM tris–HCl, 20 mM tris-base, 10 mM KH2PO4, 0.5 mg/mL BSA) supplemented with 1 mM EGTA and then for another 10 min in buffer supplemented with 5 mM blebbistatin (Sigma B0560). Calcium uptake was assessed with 2 μM thapsigarin (Sigma T9033) to differentiate SR versus mitochondrial calcium uptake.

4.10 Creatine Kinase determination

Muscle-derived Creatine Kinase in serum was measured using a commercially available colorimetric assay (Stanbio 22-022-630) per the manufacturer's instructions.

4.11 Statistical analysis

Data are presented as mean ± SEM, unless otherwise specified. Statistical analysis and figures were generated using Excel or GraphPad Prism version 8.0.0 for Windows (GraphPad Software) except for proteomics data. Data were compared using unpaired Student's t -test and one-way or two-way ANOVA followed by multiple comparison analysis. The significance level was set at P < 0.05.

4.12 Illustrations

Illustrations were created using BioRender™ with permission to publish.

Funding

This work was supported by the 10.13039/100000002 National Institutes of Health Grant R01-DK125812 (to J.M.E.).

CRediT authorship contribution statement

Andrea S. Pereyra: Writing – review & editing, Writing – original draft, Visualization, Supervision, Methodology, Investigation, Formal analysis, Data curation, Conceptualization. Regina F. Fernandez: Investigation, Formal analysis. Adam Amorese: Visualization, Investigation. Jasmine N. Castro: Investigation. Chien-Te Lin: Investigation, Formal analysis. Espen E. Spangenburg: Resources, Methodology, Formal analysis. Jessica M. Ellis: Writing – review & editing, Writing – original draft, Visualization, Validation, Supervision, Resources, Project administration, Methodology, Funding acquisition, Conceptualization.

Declaration of competing interest

The authors declare that they have no known competing financial interests or personal relationships that could have appeared to influence the work reported in this paper.

Appendix A Supplementary data

The following is the Supplementary data to this article:Multimedia component 1

Supplemental Figure 1. Loss of CPT2 disrupts myofibril structure with overall preserved mitochondrial bioenergetics. (A-C) Membrane potential of isolated mitochondria presented as % of alamethicin (Ala) of (A) white quadriceps, (B) red gastrocnemius, and (C) soleus muscles of Cpt2Sk−/− and Cpt2Skf/f mice exposed to pyruvate + malate (PYR/M), octanoylcarnitine + malate (OCT/M), or succinate + Rotanone (S/ROT) across a range of free energy (ΔGATP) and followed by the addition of Complex V inhibitor oligomycin (Oligo), Complex IV inhibitor cyanide (CN) and pore-forming peptide alamethicin (Ala). (D) Illustration depicting the main components of the myofibril-sarcolemma-extra cellular matrix (ECM) anchorage system. The illustration was created with BioRender. (E) Immunofluorescent detection of dystrophin (yellow) and nuclei (blue) in Soleus. Magnification 20× and scale bar = 200 μm. N = 3–6. Data are presented as mean ± SEM and ∗P ≤ 0.05 by T-Test.

Supplemental Figure 2. CPT2-deficient muscles display myopathic features. (A) Soleus and EDL muscle cross-sections from Cpt2Skf/f and Cpt2Sk−/− mice stained with Hematoxylin & Eosin. (B) Top row: representative images of immunodetection of dystrophin (yellow), and nuclei (blue) in Soleus muscle. Red circles indicate myofibers with central nuclei. Bottom row: representative images of modified Gomori-stained Soleus muscle showing mitochondria (dark red) and myoplasma (light blue). Yellow circles indicate ragged myofibers. (C) Quantitation of myofibers with central nuclei in Soleus muscle. Between 800 and 900 myofibers per biological sample were screened for central nuclei (red circles in figure) and three biological replicates per genotype were analyzed in total. (D) Levels of Creatine Kinase in serum of control (Skf/f) and CPT2-deficient (Sk−/−) mice as measured by colorimetric assay. Data is presented as mean ± SEM and ∗P ≤ 0.05 by T-test. Scale bar = 100 μm. N = 3.

Supplemental Figure 3. Phospholipid acyl composition alterations in soleus muscle by loss of CPT2. Relative abundance of individual species of phosphatidylcholine (PC), sphingomyelin (SM) phosphatidylethanolamine (PE), phosphatidylserine (PS), phosphatidylinositol (PI) and cardiolipin (CL) as determined by DIMS in soleus muscle of 2-month-old, male, Cpt2Sk−/− and Cpt2Skf/f mice. Data are presented as mean ± SEM and ∗P ≤ 0.05 by T-Test. N = 4.

Supplemental Figure 4. CPT2 deficiency is not associated with oxidative damage in skeletal muscle. (A) Detection of resident macrophages (CD68, green), dystrophin (yellow), and nuclei (blue) in soleus muscle of Cpt2Skf/f and Cpt2Sk−/− mice by immunofluorescence. (B) Abundance of proteins involved in protection against oxidative damage in soleus. Data were generated in adult male mice and presented as Log2 abundance. ∗adjusted p-value (q < 0.1) calculated by Benjamini Hochberg FDR correction. N = 6 for (A), = 3 for (B) and (C).

Supplemental Figure 5. Loss of mLCFAO alters calcium homeostasis. (A) Relative calcium uptake over time in EDL muscle homogenate of Cpt2Skf/f and Cpt2Sk−/− mice. (B) Area under the curve (AUC) and (C) time to half-life of calcium uptake shown in (A). (D) Representative traces of calcium uptake in isolated mitochondria from Soleus muscle from Cpt2Skf/f and Cpt2Sk−/− mice energized with glutamate/malate. (E) Same as (D) but pre-incubated with the mPTP inhibitor cyclosporin A (CsA). (F) Representative traces of calcium efflux in isolated mitochondria from Soleus muscle from Cpt2Skf/f and Cpt2Sk−/− mice energized with glutamate/malate and stimulated with NaCl. (G) Same as (F) but followed by calcium uptake stimulated by addition of free Ca+2. (H) Representative traces of calcium uptake in permeabilized fiber bundles from control (Skf/f) and CPT2-deficient (Sk−/−) Soleus muscles in the presence of the SERCA inhibitor, thapsigargin. Data were generated in adult male mice. For (A to C) data and are presented as mean ± SEM and ∗P ≤ 0.05 by T-Test. n = 5–6 for A-C, n = 1 for D-I.

Supplemental Figure 6. Calcium uptake and release is not compromised in isolated SR of CPT2-deficient muscles. (A) Representative traces of calcium uptake and release in isolated SR from control muscle (Skf/f) in the absence of fatty acids and in the presence of palmitoyl-carnitine (PC) and the SERCA inhibitor cyclopiazonic acid (CPA). (B) Representative traces of calcium uptake and release in isolated sarcoplasmic reticulum (SR) from control muscle (Skf/f) in the absence of fatty acids and in the presence of palmitic acid (PA), and palmitoyl-CoA (PCoA). (C) Quantitation of the calcium uptake and release slope from (B) over time relative to protein concentration. (D) Representative traces of calcium uptake and release in isolated SR from control (Skf/f) and CPT2-deficient (Sk−/−) muscles in the absence of fatty acids. (E) Quantitation of the calcium uptake and release slope from (B) over time relative to protein concentration. Data was generated in adult male mice. Data are presented as mean ± SEM and ∗P ≤ 0.05 by 1-way ANOVA, n = 3–4.

Multimedia component 1

Data availability

Data will be made available on request.

Acknowledgments

The authors thank Dr. Julia Laskin of Purdue University for lipid imaging by Nano-DESI.

Appendix A Supplementary data to this article can be found online at https://doi.org/10.1016/j.molmet.2024.102015.
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