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ABSTRACT Skeletal muscle mitochondria readily accumulate Ca2+ in response to SR store-releasing stimuli thanks to the activity of the mitochondrial calcium uniporter (MCU), the highly

selective channel responsible for mitochondrial Ca2+ uptake. MCU positively regulates myofiber size in physiological conditions and counteracts pathological loss of muscle mass. Here we show

that skeletal muscle-specific MCU deletion inhibits myofiber mitochondrial Ca2+ uptake, impairs muscle force and exercise performance, and determines a slow to fast switch in MHC

expression. Mitochondrial Ca2+ uptake is required for effective glucose oxidation, as demonstrated by the fact that in muscle-specific MCU−/− myofibers oxidative metabolism is impaired and

glycolysis rate is increased. Although defective, mitochondrial activity is partially sustained by increased fatty acid (FA) oxidation. In MCU−/− myofibers, PDP2 overexpression drastically

reduces FA dependency, demonstrating that decreased PDH activity is the main trigger of the metabolic rewiring of MCU−/− muscles. Accordingly, PDK4 overexpression in MCUfl/fl myofibers is

sufficient to increase FA-dependent respiration. Finally, as a result of the muscle-specific MCU deletion, a systemic catabolic response impinging on both liver and adipose tissue metabolism

occurs. You have full access to this article via your institution. Download PDF SIMILAR CONTENT BEING VIEWED BY OTHERS THE MITOCHONDRIAL ATP-DEPENDENT POTASSIUM CHANNEL (MITOKATP) CONTROLS

SKELETAL MUSCLE STRUCTURE AND FUNCTION Article Open access 17 January 2024 MCU-INDEPENDENT CA2+ UPTAKE MEDIATES MITOCHONDRIAL CA2+ OVERLOAD AND NECROTIC CELL DEATH IN A MOUSE MODEL OF

DUCHENNE MUSCULAR DYSTROPHY Article Open access 21 March 2024 THE MITOCHONDRIAL MRNA-STABILIZING PROTEIN SLIRP REGULATES SKELETAL MUSCLE MITOCHONDRIAL STRUCTURE AND RESPIRATION BY

EXERCISE-RECOVERABLE MECHANISMS Article Open access 13 November 2024 INTRODUCTION Mitochondrial Ca2+ uptake has pleiotropic roles. By buffering [Ca2+] increases, it contributes to the

regulation of Ca2+-dependent functions that take place in the cytosol. In addition, within the organelle, Ca2+ positively regulates key dehydrogenases of the TCA cycle thus promoting ATP

synthesis. On the other hand, excessive mitochondrial [Ca2+] has a pro-apoptotic function [1]. Mitochondria are placed in close contact with the endoplasmic reticulum/sarcoplasmic reticulum

(ER/SR) Ca2+ stores, thus sensing microdomains of high [Ca2+] that enable prompt Ca2+ uptake [2]. Studies on this mechanism have been greatly facilitated by the identification of the long

sought mitochondrial calcium uniporter (MCU), the highly selective channel responsible for Ca2+ accumulation by the organelle [3, 4]. MCU is composed of pore-forming subunits (MCU, MCUb)

associated with essential MCU regulator, and of regulatory proteins (MICU1–3) that, together, contribute to the fine-tuned regulation of the channel activity [5,6,7,8]. We previously

highlighted the positive role of mitochondrial Ca2+ uptake in controlling muscle trophism. Overexpression of MCU in the murine hindlimb triggers muscle hypertrophy and exerts a protective

effect against denervation-induced atrophy, while MCU silencing causes muscle atrophy [9]. Importantly, in trained elderly subjects the improvement of muscle function is associated with

increased MCU expression [10]. Moreover, the MCU complex has a peculiar composition in the skeletal muscle. Indeed, a muscle-specific MICU1 splicing variant (i.e., MICU1.1), by binding Ca2+

with higher affinity, confers myofiber mitochondria sustained Ca2+ uptake and ATP production required for contraction [11]. Finally, skeletal muscle force is decreased in total MCU−/− mice,

underlying the crucial role of mitochondrial Ca2+ uptake for energy production required for muscle work [12]. To further clarify the role of skeletal muscle mitochondrial Ca2+ uptake in

organ physiology and metabolism and to unveil potential systemic effects of skeletal muscle MCU deletion, we have generated both a constitutive and an inducible skeletal muscle-specific MCU

knockout mouse model. Locally, loss of MCU in skeletal muscle triggered decreased muscle performance, a fiber-type switch toward fast myosin heavy chains (MHCs), and a metabolic rewiring

toward preferential fatty acid (FA) oxidation. Systemically, a catabolic response impinging on liver and adipose tissue metabolism occurred. MATERIALS AND METHODS DNA PLASMIDS The following

plasmids have been previously described: mtGCaMP6f [13] and R-GECO1 [14]. Flag-PDK4 and Flag-PDP2 were amplified by PCR from mouse liver cDNA with the following primers and cloned in

pcDNA3.1: Flag-PDK4 primers: Fw: 5’-TAATGGATCCGCCACCATGAAGGCAGCCCGCTTCGTGATG-3’; Rev: 5’-TAATGAATTCTCACTTATCGTCGTCATCCTTGTAATCCACTGCCAGCTTCTCCTTCGC-3’; Flag-PDP2 primers: Fw:

5’-TAATGGATCCGCCACCATGTCAAGTACTGTGTCCTACTGGATCTTC-3’; and Rev: 5’-TAATGAATTCTCACTTATCGTCGTCATCCTTGTAATCACCCTCTTTACAGTAGGTATCAATTGA-3’. ANIMALS All animal experiments were approved and

performed in accordance with the Italian law D. L.vo n_26/2014. C57BL/6N-Mcutm1a mice were either produced by blastocyst microinjection of mutant embryonic cell lines or purchased from

International Mouse Phenotyping Consortium (Mcu HEPD0762_7_B01). The former were derived by microinjection into C57BL/6NCrl blastocysts of mutagenized embryonic stem (ES) cell line from the

EUCOMM consortium (C57BL/6N-Mcutm1a(EUCOMM)Hmgu) in our laboratory, the second was produced by the consortium using similar ES cells, but different clones, obtained in the same recombination

experiment. The mutant allele presents a cassette composed of a FRT site followed by lacZ sequence and a loxP site. This first loxP site is followed by neomycin under the control of the

human beta-actin promoter, SV40 polyA, followed by a FRT site and a second loxP site. The third loxP site is located downstream exon 5 of MCU gene (Ccdc109a). To generate the

C57BL/6N-Mcutm1c line (here referred to as MCUfl/fl line), C57BL/6N-Mcutm1a mice were crossed with an FLP-expressing line. FLP-recombinase excited both the LacZ and neomycin-resistance

cassettes restoring MCU expression. Homozygous MCUfl/fl mice were crossed with transgenic mice expressing the Cre recombinase under the control of the Mlc1f promoter (Mlc1f-Cre) [15] or the

Cre recombinase fused to the mutated ligand-binding domain of the human estrogen receptor under the HSA promoter (HSA-Cre-ERT2) [16]. MCUfl/fl mice from both procedures displayed similar

phenotypes after Cre-recombination. For the present work, MCUfl/fl mice derived from purchased C57BL/6N-Mcutm1a mice were used. In all, 1.5 mg/30 g body weight (BW) of tamoxifen

(Sigma-Aldrich) dissolved in 10% EtOH and 90% sunflower oil (Sigma-Aldrich) was intraperitoneally (i.p.) injected every 3 days for 6 weeks. In addition, the same animals were fed with a

tamoxifen-containing diet (Envigo TAM400/CreER) that allowed administration of tamoxifen of 40 mg/kg BW/die. In some key experiments, the Cre control groups were used (mlc1f-Cre and

HSA-Cre-ERT2 mice). All the mice were fed ad libitum and always sacrificed before 9:30 a.m. IN VIVO DNA TRANSFECTION OF MOUSE SKELETAL MUSCLE Adult male mice were used in all experiments.

First, the animal was anesthetized. Hyaluronidase solution (2 mg/ml) (Sigma-Aldrich) was injected under the hindlimb footpad. After 30 min, 20 μg of plasmid DNA in 20 µl of physiological

solution was injected with the same procedure of the hyaluronidase. Then one gold-plated acupuncture needle was placed under the skin at heel, and a second one at the base of the toes,

oriented parallel to each other and perpendicular to the longitudinal axis of the foot and connected to the BTX porator (Harvard apparatus). The muscles were electroporated by applying 20

pulses, 20 ms each, 1 s of interval to yield an electric field of 100 V. Single fibers cultures were carried out 7–10 days later. MOUSE EXERCISE STUDIES For acute concentric exercise

studies, 3- or 6-month-old mice were acclimated to and trained on a 10° uphill LE8700 treadmill (Harvard apparatus) for 2 days. On day 1, mice ran for 5 min at 8 m/min and on day 2 mice ran

for 5 min at 8 m/min followed by another 5 min at 10 m/min. On day 3, mice were subjected to a single bout of running starting at the speed of 10 m/min. Forty minutes later, the treadmill

speed was increased at a rate of 1 m/min every 10 min for a total of 30 min and then increased at the rate of 1 m/min every 5 min until mice were exhausted. Exhaustion was defined as the

point at which mice spent >5 s on the electric shocker without attempting to resume running. Total running time and total running distance were recorded for each mouse. For eccentric

exercise training, 3-month-old mice were acclimated to and trained on a 10° downhill for 2 days. On day 1, mice ran for 5 min at 8 m/min and on day 2 mice ran for 5 min at 8 m/min followed

by another 5 min at 10 m/min. On days 3–5, mice were subjected to a single bout of running starting at the speed of 10 m/min. Forty minutes later, the treadmill speed was increased at a rate

of 1 m/min every 10 min for a total of 30 min and then increased at the rate of 1 m/min every 5 min until mice were exhausted. Exhaustion was defined as above. Total running time and total

running distance were recorded for each mouse. MUSCLE FORCE AND FATIGUE MEASUREMENTS To measure muscle force in living animals, the contractile performance of gastrocnemius muscle in vivo

was measured as described previously [17]. Briefly, anesthetized mice were placed on a thermostatically controlled table, keeping the knee stationary, and the foot firmly fixed to a

footplate, which was connected to the shaft of the motor of a muscle-lever system (305B, Aurora Scientific). Contraction was elicited by electrical stimulation of sciatic nerve.

Teflon-coated seven-stranded steel wires (AS 632, Cooner Sales) were implanted with sutures on either side of the sciatic nerve proximal to the knee before its branching. At the distal ends

of the two wires, the insulation was removed, and the proximal ends were connected to a stimulator (S88, Grass). To avoid recruitment of the dorsal flexor muscles, the common peroneal nerve

was cut. The fatigue protocol consists of a tetanic stimulation (frequency 100 Hz, 4–6 V) lasting 300 ms each second for a total time of 120 s. HISTOLOGY AND FLUORESCENCE MICROSCOPY For

fiber size measurements, 20 μm-thick cryosections were fixed in 4% formaldehyde for 20 min, quenched with 50 mM NH4Cl in phosphate-buffered saline (PBS) and blocked in PBS containing 0.5%

bovine serum albumin (BSA) for 20 min. Sections were then incubated with primary antibody anti-Laminin (Sigma-Aldrich) to label the sarcolemma for 1 h at 37 °C and washed 3 times in PBS.

Alexa Fluor 488-conjugated secondary antibody (Thermo Fisher Scientific) was used. Fiber size measurements were performed with the Fiji distribution of ImageJ [18]. For fiber-type

measurements, 20 μm-thick cryosections were blocked in M.O.M. working solution (Vector Laboratories). Sections were then incubated with primary antibody sc-71 (Hybridoma bank) 1:100 in 0.5%

BSA in PBS to label the type IIA myosins for 1 h at 37 °C and washed 3 times in PBS. Alexa Fluor 488-conjugated secondary antibody (Life Technologies) was used. Then sections were incubated

with primary antibody Bf-f3 (Hybridoma bank) 1:100 in 0.5% BSA in PBS to label the type IIB myosins for 1 h at 37 °C and washed 3 times in PBS. Alexa Fluor 555-conjugated secondary antibody

(Life Technologies) was used. Finally, sections were then incubated with primary antibody Bad5 (Hybridoma bank) 1:100 in 0.5% BSA in PBS to label the type I myosins for 1 h at 37 °C and

washed 3 times in PBS. Alexa Fluor 647-conjugated secondary antibody (Thermo Fisher Scientific) was used. Fiber-type analyses were performed with the Fiji distribution of ImageJ [18]. For

hematoxylin and eosin (H&E) staining, 20 μm-thick cryosections were stained using Rapid Frozen Sections H&E staining Kit (Bio-Optica) according to manufacturer’s instructions.

Briefly, cryosections were incubated in hematoxylin solution for 60 s. Then, after 3 washes, they were incubated in eosin solution for 30 s. Finally, cryosections were dehydrated. RNA

EXTRACTION, REVERSE TRANSCRIPTION, AND QUANTITATIVE REAL-TIME PCR Total RNA was extracted from tibialis anterior (TA) muscles, livers, and visceral adipose tissue (VAT) using the SV Total

RNA Isolation Kit (Promega) following the manufacturer’s instructions. The RNA was quantified with Nanodrop (Thermo Fisher Scientific). Complementary DNA was generated from 500 nmol of total

RNA with a cDNA Synthesis Kit SuperScript II (Thermo Fisher Scientific). Oligo(dT)12–18 primers (Thermo Fisher Scientific) were used as primer for first-strand cDNA synthesis with reverse

transcriptase. The obtained cDNA was analyzed by real-time PCR using the IQ5 thermocycler and the SYBR green chemistry (Bio-Rad). The primers were designed and analyzed with Primer3 [19].

The housekeeping gene Gapdh was used as an internal control for cDNA normalization. For quantification, expression levels were calculated by the ΔCt method. Real-time PCR primer sequences

were as follows: Mcu: Fw 5’-AAAGGAGCCAAAAAGTCACG-3’; Rv 5’-AACGGCGTGAGTTACAAACA-3’. Gapdh: Fw 5’-CACCATCTTCCAGGAGCGAG-3’; Rv 5’-CCTTCTCCATGGTGGTGAAGAC-3’. Pgc1α-tot: Fw

5’-CGCTGCTCTTGAGAATGGAT-3’; Rv 5’-CGCAAGCTTCTCTGAGCTTC-3’. Pgc1α-1: Fw 5’-GGACATGTGCAGCCAAGACTCT-3’; Rv 5’-CACTTCAATCCACCCAGAAAGCT-3’. Pgc1α-4: Fw 5’-TCACACCAAACCCACAGAAA-3’; Rv

5’-CTGGAAGATATGGCACAT-3’. G6pase: Fw 5’-ACGCCCGTATTGGTGGGTCCT-3’; Rv 5’-GCCAGAGGGACTTCCTGGTCCG-3’. Pck1: Fw 5’-CTGCATAACGGTCTGGACTTC-3’; Rv 5’-CAGCAACTGCCCGTACTCC-3’. Pnpla2: Fw:

5’-CTGAGAATCACCATTCCCACATC-3’; Rv: 5’-CACAGCATGTAAGGGGGAGA-3’. Lipe: Fw: 5’-AGGATCGAAGAACCGCAGTC-3’; Rv: 5’-GTCTTCTGCGAGTGTCACCA-3’. Abhd5: Fw: 5’-TGGTGTCCCACATCTACATCA-3’; Rv:

5’-CAGCGTCCATATTCTGTTTCCA-3’. Mgll: Fw: 5’-CACGTGGACACCATCCAGAA-3’; Rv: 5’-GCCACTAGGATGGAGATGGC-3’. Hmgcs2: Fw: 5’-CCGGTGTCCCGTCTAATGG-3’; Rv: 5’-GCAGATGCTGTTTGGGTAGC-3’. Bdh1: Fw:

5’-ATAGGGCCTGAGAGGGAAGG-3’; Rv: 5’-GCAGTACAAATGCATCCCGC-3’. IL-6: Fw: 5’-TAGTCCTTCCTACCCCAATTTCC-3’; Rv: 5’-TTGGTCCTTAGCCACTCCTTC-3’. WESTERN BLOTTING AND ANTIBODIES To monitor protein

levels, frozen muscles and frozen livers were pulverized by means of Qiagen Tissue Lyser and protein extracts were prepared in an appropriate buffer containing: muscle lysis buffer (50 mM

Tris pH 7.5, 150 mM NaCl, 5 mM MgCl2, 1 mM dithiothreitol, 10% glycerol, 2% sodium dodecyl sulfate (SDS), 1% Triton X-100, Complete EDTA-free protease inhibitor mixture (Roche), 1 mM PMSF, 1

 mM NaVO3, 5 mM NaF, and 3 mM β-glycerophosphate) or RIPA buffer for liver extracts (125 mM NaCl, 25 mM Tris-Cl pH7.4, 1 mM EGTA-Tris pH 7.4, 1% Triton-X100, 0.5% sodium deoxycholate, 0.1%

SDS, and Complete EDTA-free protease inhibitor mixture (Roche)). In all, 40 μg of total proteins were loaded, according to BCA quantification. Proteins were separated by SDS-polyacrylamide

gel electrophoresis, in commercial 4–12% acrylamide gels (Thermo Fisher Scientific) and transferred onto nitrocellulose membranes (Thermo Fisher Scientific) by wet electrophoretic transfer.

Blots were blocked 1 h at room temperature (RT) with 5% non-fat dry milk (Bio-Rad) in TBS-tween (0.5 M Tris, 1.5 M NaCl, 0.01% Tween) solution and incubated at 4 °C with primary antibodies.

Secondary antibodies were incubated 1 h at RT. The following antibodies were used: anti-phosphoAKT (1:1000, Cell Signaling), anti-AKT (1:1000, Cell Signaling) anti-ACTIN (1:20,000, Santa

Cruz), anti-FLAG (1:1000, Cell Signaling) anti-phosphoGSK3α/β (1:1000, Cell Signaling), anti-GSK3α/β (1:1000, Cell Signaling), anti-MCU (1:1000, Sigma-Aldrich), anti-phosphoPDH (1:5000,

Abcam), anti-PDH (1:1000, Cell Signaling) anti-GRP75 (1:5000, Santa Cruz), and anti-TOM20 (1:20,000, Santa Cruz). Secondary horseradish peroxidase-conjugated antibodies were purchased from

Bio-Rad and used at 1:5000 dilution. REAL-TIME IMAGING OF MITOCHONDRIAL AND CYTOSOLIC CA2+ IN FLEXOR DIGITORUM BREVIS (FDB) FIBERS REAL-TIME IMAGING FDB fibers were isolated 7–10 days after

in vivo transfection. Muscles were digested in collagenase A (4 mg/ml) (Roche) dissolved in Tyrode’s salt solution (pH 7.4) (Sigma-Aldrich) containing 10% fetal bovine serum (Thermo Fisher

Scientific). Single fibers were isolated, plated on laminin-coated glass coverslips, and cultured in Dulbecco’s modified Eagle’s medium (DMEM) with HEPES (42430 Thermo Fisher Scientific),

supplemented with 10% fetal bovine serum, containing penicillin (100 U/ml), streptomycin (100 μg/ml). Fibres were maintained in culture at 37 °C with 5% CO2. MITOCHONDRIAL AND CYTOSOLIC CA2+

MEASUREMENTS FDB muscles were electroporated with a plasmid encoding 4mtGCaMP6f together with R-GECO1. After single fiber isolation, real-time imaging was performed. During the experiments,

myofibers were maintained in Krebs-Ringer modified buffer (135 mM NaCl, 5 mM KCl, 1 mM MgCl2, 20 mM HEPES, 1 mM MgSO4, 0.4 mM KH2PO4, 1 mM CaCl2, 5.5 mM glucose, pH 7.4) at RT, in the

presence of 75 μM _N_-benzyl-P-toluenesulfonamide (BTS, Sigma-Aldrich) to avoid the fiber contraction. In all, 20 mM caffeine (Sigma-Aldrich) was added when indicated to elicit Ca2+ release

from intracellular stores. Experiments were performed on a Zeiss Axiovert 200 microscope equipped with a ×40/1.3 N.A. PlanFluor objective. Excitation was performed with a DeltaRAM V

high-speed monochromator (Photon Technology International) equipped with a 75 W xenon arc lamp. Images were captured with a high-sensitivity Evolve 512 Delta EMCCD (Photometrics). The system

is controlled by MetaMorph 7.5 (Molecular Devices) and was assembled by Crisel Instruments. 4mtGCaMP6f and R-GECO1 were alternatively excited every second at 490 and 560 nm, respectively,

and images were acquired through a dual band emission filter (520/40 and 630/60) (Chroma). Exposure time was set to 50 ms (4mtGCaMP6f) and 150 ms (R-GECO1). Acquisition was performed at

binning 1 with 200 of EM gain. Image analysis was performed with Fiji distribution of the ImageJ software [18]. Images were background subtracted and linear unmixing was performed to get rid

of bleed through of the two fluorochromes. Data are expressed as F/F0 where F0 is the mean intensity at the beginning of the experiment. CYTOSOLIC CA2+ MEASUREMENTS Fibers were dissected

and loaded with 2 μM fura-2/AM (Thermo Fisher Scientific) diluted in Krebs-Ringer modified buffer (described above) containing 0.02% pluronic acid for 20 min at 37 °C and then washed with

Krebs-Ringer modified buffer in presence of 75 μM BTS (Sigma-Aldrich) to avoid the fiber contraction. 20 mM caffeine (Sigma-Aldrich) was added when indicated to elicit Ca2+ release from

intracellular stores. Experiments were performed on a Zeiss Axiovert 200 microscope equipped with a ×40/1.3 N.A. PlanFluor objective. Excitation was performed with a DeltaRAM V high-speed

monochromator (Photon Technology International) equipped with a 75 W xenon arc lamp. Images were captured with a high-sensitivity Evolve 512 Delta EMCCD (Photometrics). The system is

controlled by MetaMorph 7.5 (Molecular Devices) and was assembled by Crisel Instruments. Images were collected by alternatively exciting the fluorophore at 340 and 380 nm and fluorescence

emission recorded through a 515/30 nm band-pass filter (Semrock). Exposure time was set to 50 ms. Acquisition was performed at binning 1 with 200 of electron-multiplying gain. Image analysis

was performed with Fiji distribution of the ImageJ software. Images were background subtracted. Changes in fluorescence (340/380 nm ratio) was expressed as _R_/R0, where _R_ is the ratio at

time _t_ and R0 is the ratio at the beginning of the experiment. OCR (OXYGEN CONSUMPTION RATE) MEASUREMENTS For OCR experiments, FDB fibers were isolated. Muscles were digested in

collagenase A (4 mg/ml) (Roche) dissolved in Tyrode’s salt solution (pH 7.4) (Sigma-Aldrich) containing 10% fetal bovine serum (Thermo Fisher Scientific). Single fibers were isolated, plated

on laminin-coated XF24 microplate wells and cultured in DMEM (D5030 Sigma-Aldrich), supplemented with 1 mM NaPyr, 5 mM glucose, 33 mM NaCl, 15 mg phenol red, 25 mM HEPES, and 1 mM of L-Glu.

Fibres were maintained for 2 h in culture at 37 °C in 5% CO2. To measure exogenous FA utilization, the fibers were cultured in DMEM (D5030 Sigma-Aldrich), supplemented with 0.1 mM NaPyr, 5 

mM glucose, 33 mM NaCl, 15 mg phenol red, 25 mM HEPES, 1 mM of L-Glu, 0.5 mM carnitine, and 100 μM palmitate:BSA. Fibres were maintained for 2 h in culture at 37 °C in 5% CO2. The rate of

oxygen consumption was assessed in real-time with the XF24 Extracellular Flux Analyzer (Agilent), which allows to measure OCR changes after up to four sequential additions of compounds.

Fibers were plated as reported above. A titration with the uncoupler FCCP was performed in order to utilize the FCCP concentration (0.6 μM) that maximally increases OCR. The results were

normalized for the fluorescence of Calcein (Sigma-Aldrich). Fibers were loaded with 2 μM Calcein for 30 min. Fluorescence was measured using a Perkin Elmer EnVision plate reader in well scan

mode using 480/20 nm filter for excitation and 535/20 nm filter for emission. GLYCOLYSIS RATE MEASUREMENTS Glycolysis was measured by monitoring the conversion of 5-3H-glucose to 3H2O, as

described previously [20]. Briefly, TA muscle fibers were incubated in a medium that contained [5-3H]-glucose (PerkinElmer) at a specific activity of 10 μCi. Labeled glucose metabolized by

glycolysis produces 3H2O that is released into the media. Following incubation for 3 h at 37 °C, media was transferred to uncapped tubes containing 0.5 ml of 0.2 M HCl. The tube was

transferred to a scintillation vial containing 1 ml of H2O such that the water in the vial and the contents of the tube were not allowed to mix. The vials were sealed, and diffusion was

allowed for 24 h. The amount of diffused 3H2O was determined by scintillation counting. LACTATE MEASUREMENTS Freshly isolated TA muscle fibers were cultured in DMEM (D5030 Sigma-Aldrich),

supplemented with 1 mM NaPyr, 10 mM glucose, 33 mM NaCl, 15 mg phenol red, 5 mM HEPES, and 1 mM of L-Glu. Once an hour, an aliquot of medium was collected and lactate concentration was

measured by means of the colorimetric L-Lactate Assay Kit (Abcam) according to the manufacturer’s instructions. MUSCLE GLYCOGEN AND FA CONTENT Glycogen amount was measured by means of the

colorimetric Glycogen Assay Kit II (Abcam) according to the manufacturer’s instructions. FA amount was measured by means of the Free FA Quantification Kit (Abcam) according to the

manufacturer’s instructions. BLOOD METABOLITE QUANTIFICATION Blood was collected from the orbital sinus in heparin-coated Pasteur pipettes and centrifuged immediately after collection. FAs

and β-hydroxy-butyrate were dosed using an automated spectrophotometer Cobas Fara II (Roche) according to the manufacturer’s instruction. Blood glucose levels were measured with an YSI 2300

STAT PlusTM glucose and lactate analyzer (YSI Life Sciences, Yellow Springs, OH) according to the manufacturer’s instruction. Lactate levels were measured using Stat Strip Xpress measuring

system (Nova Biomedical). GLUCOSE TOLERANCE TEST For the glucose tolerance test, mice were fasted for 5 h. Glucose (2 mg/g BW) was administered i.p. and the blood glucose levels were

followed during 2:30 h using ContourXT (Bayer). GLUCOSE UPTAKE MEASUREMENTS Glucose uptake in tissues was measured using a non-radioactive colorimetric method [21] by means of the

2-Deoxy-d-Glucose Uptake Measurement Kit (Cosmo Bio Co., LTD, Tokyo, Japan). Mice were fasted 5 h before the beginning of the experiment. Mice were then injected i.p. with a solution

containing d-glucose (1 g/kg) and 2-deoxy-d-glucose (0.027 g/kg). One hour after injection, mice were killed by cervical dislocation, and tissues (soleus, TA, and liver) were rapidly removed

and frozen in liquid nitrogen. Soleus and TA muscles and 150 mg of liver were used for the assay. Tissues were subsequently weighed and homogenized in 1 ml of Tris-HCl (10 mM, pH 8), with

TissueLyser II (Qiagen), by shaking at maximum speed for 3 min, and then heated at 95 °C for 15 min. Samples were then centrifuged at 16,000 × _g_ for 15 min at 4 °C to remove tissue debris.

Supernatants were finally collected, and a portion was diluted in 20 µl (final volume; 4 µl for soleus, 5 µl for TA, and 2 µl for liver) with the sample dilution buffer provided with the

kit. The assay was then performed following the manufacturer’s instructions. METABOLOMICS ANALYSIS Gastrocnemius muscles of adult mice were harvested and underwent untargeted metabolomics

analysis performed by Metabolon, Inc. Briefly, samples were prepared using the automated MicroLab STAR® system from Hamilton Company. The resulting extract was divided into five fractions:

two for analysis by two separate reverse phase (RP)/ultra-performance liquid chromatography tandem mass spectrometry (UPLC-MS/MS) methods with positive ion mode electrospray ionization

(ESI), one for analysis by RP/UPLC-MS/MS with negative ion mode ESI, one for analysis by hydrophiliic interaction liquid chromatography/UPLC-MS/MS with negative ion mode ESI, and one sample

was reserved for backup. INTERLEUKIN (IL)-6 ENZYME-LINKED IMMUNOSORBENT ASSAY IL-6 serum levels were measured by means of the mouse IL-6 Kit (Abcam) according to the manufacturer’s

instructions. STATISTICAL ANALYSIS OF THE DATA Statistical data are presented as mean ± SD, unless otherwise specified. Significance was calculated by Student’s _t_ test or Mann–Whitney

rank-sum test. RESULTS CONSTITUTIVE MUSCLE-SPECIFIC MCU DELETION CAUSES DECREASED MUSCLE PERFORMANCE AND FIBER-TYPE SWITCHING We have previously demonstrated that MCU silencing in skeletal

muscle causes decreased myofiber size and inhibition of trophic signaling routes [9]. To discern the contribution of skeletal muscle mitochondrial Ca2+ uptake to muscle physiology and

whole-body metabolism, we generated a skeletal muscle-specific MCU−/− mouse model. We crossed mice in which the exon 5 of the MCU gene is flanked by loxP sequences with mice expressing the

Cre recombinase under the control of the myosin-light chain 1f promoter (mlc1f-Cre) [15], which is expressed since embryogenesis (Fig. 1a). Expression of MCU in hindlimb muscles of adult

mlc1f-Cre::_MCU__fl/fl_ (skMCU−/−) mice was essentially absent indicating that efficient recombination occurred (Fig. 1b and S1A-B). MCU deletion is expected to blunt mitochondrial Ca2+

transients triggered by Ca2+ release from SR stores. To confirm this notion, FDB muscles were simultaneously transfected in vivo with 4mtGCaMP6f, a Ca2+-sensitive probe targeted to the

mitochondria matrix [13], together with R-GECO, a cytosolic Ca2+-probe [14]. Upon caffeine-induced Ca2+ release from the SR, MCUfl/fl myofibers showed robust cytosolic Ca2+ transients

followed by effective mitochondrial Ca2+ uptake (Fig. 1c). In contrast, skMCU−/− mitochondria were unable to take up Ca2+ despite normal cytosolic Ca2+ increases (Fig. 1c). Quantitative

analysis of cytosolic [Ca2+] highlighted a slight reduction of Ca2+ transients in skMCU−/− myofibers, suggesting a negative effect of chronic MCU deletion on global Ca2+ homeostasis (Fig. 

1d). However, basal cytosolic [Ca2+] was unaffected by MCU deletion (Fig. 1e). skMCU−/− TA muscles did not show any sign of damage (Figure S1C) but fiber size was reduced (Fig. 1f), although

at a milder degree than AAV-shMCU-treated muscles [9]. Moreover, phosphorylation levels of Akt and mRNA expression levels of PGC1α4 were decreased (Figure S1D-E). Overall, these results

indicate that skMCU−/− mice mirror the effects of MCU silencing [9], thus representing an ideal model for the analysis of both local and systemic physiologic and metabolic outcomes of

skeletal muscle mitochondrial Ca2+ uptake inhibition. Next, we evaluated muscle performance. In vivo analysis showed a decline in tetanic force of skMCU−/− compared to MCUfl/fl mice (Figure 

S1F). We then measured the distance run on a treadmill during a single bout of uphill strenuous exercise. In line with the constitutive MCU−/− mice [12], muscle-specific MCU deletion caused

a deficit in running capacity, both at 3 and 6 months of age (Fig. 1g and S1G-H), thus demonstrating that impaired exercise performance is caused by an intrinsic skeletal muscle defect. In

addition, a significant fiber-type switch was apparent, as opposed to the constitutive MCU ablation [12]. Indeed, quantitative analysis demonstrated reduced expression of the slow-twitch

type 1 MHC and an increase of the fast-twitch type 2A MHC in the soleus muscle of skMCU−/− mice (Fig. 1h). To assess the response to repeated isometric tetanic contractions, we performed 120

isometric contractions of a duration of 300 ms once every second. However, the relative decrease in force generation was not different between MCUfl/fl and skMCU−/− animals (Fig. 1i),

indicating that the differences in exercise performance is not due to a different resistance to fatigue. Finally, repeated bouts of exhausting downhill running in 3 consecutive days revealed

a progressive decrease in performance that was similar between MCUfl/fl and skMCU−/− mice, indicating that lengthening contraction-induced muscle damage occurs independently of MCU (Figure 

S1I). MUSCLE-SPECIFIC MCU DELETION TRIGGERS A METABOLIC SHIFT TOWARD FA UTILIZATION Mitochondrial Ca2+ uptake is expected to control oxidative metabolism [1]. To test whether decreased

mitochondrial Ca2+ accumulation influences skeletal muscle respiration, we measured OCR in freshly isolated FDB muscle myofibers. Myofibers were treated consecutively with oligomycin, to

measure ATP-linked respiration; FCCP, to measure maximal oxygen consumption; and finally with rotenone and antimycin, to verify the extent of non-mitochondrial respiration. skMCU−/− fibers

showed a decrease in the basal, maximal, and ATP-linked respiration compared to MCU-expressing fibers (Fig. 2a and S2A), in line with the role of mitochondrial Ca2+ uptake in regulating

oxidative metabolism. Both MCUfl/fl and skMCU−/− myofibers showed little spare respiratory capacity (Fig. 2a). We hypothesized that this effect might be due to the use of freshly isolated

myofibers that are still actively contracting. To verify this hypothesis, we measured OCR in myofibers treated with BTS to avoid fiber contraction [9]. In these conditions, myofibers display

large spare respiratory capacity, as expected (Figure S2B). Next, in skMCU−/− mice blood lactate levels were increased (Fig. 2b and S2C). These data suggest that the absence of

mitochondrial Ca2+ uptake drastically reduces pyruvate utilization by mitochondria, which is rather converted into lactate. To directly determine the proportion of the glycolytic rate, we

incubated MCUfl/fl and skMCU−/− myofibers with 5-3H-glucose, and we measured the β-emission of 3H2O produced by glycolysis in the cell medium, as reported [20]. skMCU−/− samples released

higher [3H2O] compared to MCUfl/fl fibers, demonstrating increased glycolysis (Fig. 2c). Accordingly, skMCU−/− TA muscle myofibers showed increased lactate production compared to MCUfl/fl

over time (Figure S2D). Mitochondrial Ca2+ uptake positively regulates the activity of three critical mitochondrial dehydrogenases. Isocitrate dehydrogenase and α-ketoglutarate dehydrogenase

are directly modulated by mitochondrial Ca2+, instead pyruvate dehydrogenase (PDH) is indirectly activated by Ca2+, which stimulates the activity of PDH phosphatase isoform 1 (PDP1) [22].

In line with the total MCU−/− mouse muscles [12] and the AAV-shMCU-infected muscles [9], in skMCU−/− muscles phosphorylation levels of PDH were increased, suggesting the inhibition of its

enzymatic activity (Fig. 2d and S2E). If pyruvate utilization is impaired in skMCU−/− muscles, in MCUfl/fl mitochondria inhibition of pyruvate uptake should mimic the effects of MCU

deletion, while it should have little effect in skMCU−/− myofibers. To test this hypothesis, we performed OCR measurements in myofibers treated with UK5099, an inhibitor of the mitochondrial

pyruvate carrier (MPC) [23,24,25,26]. MPC inhibition had small effect on basal OCR of skMCU−/− myofibers, while it reduced O2 consumption of MCUfl/fl fibers to the levels of untreated

skMCU−/− myofibers (Fig. 2e). These data demonstrating the aberrant glucose oxidation in skMCU−/− muscles suggest that other energy sources may be utilized by MCU-depleted mitochondria. FA

metabolism could represent such alternative [27]. FA concentration was indeed elevated in skMCU−/− muscles compared to skMCUfl/fl controls (Fig. 2f). To determine the genuine contribution of

FA oxidation to mitochondrial respiration, we measured OCR upon treatment with etomoxir, an inhibitor of carnitine palmitoyltransferase-1 and thus of FA oxidation. While in MCUfl/fl

myofibers, FA accounted for 40% of the basal respiration, in skMCU−/− fibers 80% of basal OCR was dependent on FA (Fig. 2g). Treatment with UK5099 followed by etomoxir further confirmed the

differential substrates utilization (Fig. 2h). In addition, acetyl-CoA carboxylase (ACC) phosphorylation was increased in skMCU−/− muscles, suggesting decreased ACC activity and thus FA

synthesis (Fig. 2i). These data demonstrate that MCU deletion impairs glucose oxidation and triggers a metabolic rewiring that favors FA utilization. Nonetheless, OCR measurements in the

presence of exogenous palmitate demonstrated that the overall capacity to oxidize FA is actually reduced in skMCU−/− myofibers compared to controls, because of the Ca2+-dependent regulation

of the above-mentioned tricarboxylic acid (TCA) cycle enzymes (Figure S2F). Accordingly, untargeted metabolomics analysis of MCUfl/fl and skMCU−/− muscles revealed significant changes in

lipid-related metabolites, while the other metabolic pathways were mostly unaffected. In particular, we detected increased levels of diacylglycerols, suggestive of elevated triacylglycerol

lipolysis, and acylcarnitines (Fig. 2j and Table 1). Altogether these data demonstrate that, despite lower absolute OCR values, β-oxidation in MCU−/− muscles accounts for more than half of

total OCR, in contrast with MCUfl/fl muscles, which mainly rely on glucose oxidation. PDH INHIBITION UNDERLIES THE METABOLIC SWITCH OF SKMCU− /− MYOFIBERS We wished to know whether the

increased PDH phosphorylation (Fig. 2d) is sufficient to trigger the metabolic switch of skMCU−/− muscles. To this aim, we performed two complementary experiments. First, we overexpressed

PDH kinase 4 (PDK4) in MCUfl/fl FDB muscles to increase PDH phosphorylation (Fig. 3a) and thus inhibit PDH activity. Similarly to MCU deletion, PDK4 overexpression caused a drastic increase

in FA dependency, as demonstrated by the great reduction in OCR upon etomoxir treatment (Fig. 3b), that was barely observed in control-transfected muscles. This result indicates that, in the

absence of pyruvate decarboxylation, FA metabolism contributes to replenish the mitochondrial acetyl-CoA pool. As complementary approach, we transiently overexpressed the Ca2+-independent

PDH phosphatase (PDP2) [22] in skMCU−/− FDB muscles to rescue PDH activity. PDP2 overexpression triggered PDH dephosphorylation (Fig. 3c) in line with increased PDH activation. This was

sufficient to rescue pyruvate oxidation in skMCU−/− myofibers, as demonstrated by the barely detectable effect of etomoxir treatment on basal OCR in PDP2-overexpressing skMCU−/− myofiber

compared to control-transfected fibers (Fig. 3d). These results conclusively demonstrate that loss of MCU causes a metabolic switch toward increased FA oxidation by decreasing PDH activity.

MUSCLE-SPECIFIC MCU DELETION IMPINGES ON SYSTEMIC METABOLISM To verify whether altered energy substrate metabolism due to the loss of skeletal muscle MCU triggers systemic adaptations, we

measured blood concentration of key metabolites. Blood glucose levels were lower in skMCU−/− mice compared to MCUfl/fl (Fig. 4a and S3A). These data suggest that, despite defective glucose

oxidation, skMCU−/− muscles are perfectly capable to take up glucose. Accordingly, skMCU−/− TA and soleus muscles accumulated 2-deoxy-d-glucose even more efficiently than MCUfl/fl muscles

(Fig. 4b). In line with defective carbohydrates utilization, in skMCU−/− muscles glycogen levels were increased (Fig. 4c). In light of the role of skeletal muscle activity in controlling

systemic metabolism, we hypothesized that muscle-specific deletion of MCU would impinge on hepatic glucose metabolism. Indeed, the liver of skMCU−/− mice accumulated less glucose than

MCUfl/fl mice (Fig. 4d), pointing to a systemic metabolic adaptation in which the liver might contribute to euglycemia by counteracting the excessive muscle glucose uptake. The mRNA levels

of the gluconeogenesis enzymes glucose-6-phosphatase (G6pase) and phosphoenolpyruvate carboxykinase 1 (Pck1) were upregulated in skMCU−/− liver compared to MCUfl/fl controls (Fig. 4e). At

the same time, liver glycogen content was decreased (Fig. 4f). Glycogen synthase activity was presumably inhibited, in line with decreased phosphorylation, and thus increased activation of

glycogen synthase kinase 3β (GSK3β). GSK3 is a target of Akt that, coherently, was also dephosphorylated in skMCU−/− livers (Fig. 4g and S3B). To verify whether a general catabolic response

occurs, we analyzed liver lipases' expression. Hormone-sensitive lipase (Lipe) and monoglyceride lipase (Mgll) mRNA levels were increased in skMCU−/− liver compared to controls, and

1-acylglycerol-3-phosphate O-acyltransferase (ABHD5) and patatin-like phospholipase domain-containing 2 (Pnpla2) had an increasing tendency (Fig. 4h). Liver catabolism triggered by

muscle-restricted MCU deletion was finally assessed by increased ketogenesis, as demonstrated by increased expression of ketogenic enzymes, i.e., 3-hydroxybutyrate dehydrogenase 1 (Bdh1) and

hydroxymethylglutaryl-CoA synthase (Hmgcs2) (Fig. 4i), and of blood β-hydroxybutyric acid levels (Fig. 4j). Finally, blood FA levels were higher in skMCU−/− compared to MCUfl/fl samples

(Fig. 4k). The latter result may be suggestive not only of a hepatic response but could also implicate adipose tissue remodeling. While BW was unchanged (Figure S3C), VAT weight was

decreased in skMCU−/− relative to control mice (Fig. 4l) and the expression of Lipe and Pnpla2 lipases was increased (Fig. 4m), indicating increased adipose tissue catabolism. We wondered

whether these systemic responses are solely due to increased skeletal muscle glucose demand or whether muscle-specific cytokines (myokines) are secreted in response to MCU deficiency. In

particular, IL-6 has been shown to be produced by skeletal muscle during exercise and to enhance FA oxidation, insulin-stimulated glucose uptake, and to contribute to hepatic glucose

production during exercise [28]. IL-6 levels were increased in skMCU−/− serum and muscle samples compared to controls (Figure S3D-E). These data indicate that combined effects might be

responsible for the systemic phenotype of skMCU−/− mice. However, a comprehensive analysis of the myokinome will shed further light. Overall, these data reveal that the skeletal

muscle-restricted defect in mitochondrial metabolism is partially compensated through both (i) the exploitation of metabolic flexibility within the defective tissue (shift toward FA

oxidation) and (ii) the triggering of a whole-body rewiring of metabolic pathways. MCU DELETION IN ADULT SKELETAL MUSCLE CAUSES ATROPHY, IMPAIRS EXERCISE PERFORMANCE, AND TRIGGERS A

FIBER-TYPE SWITCH Constitutive gene deletion could trigger adaptive developmental responses that may hinder a peculiar phenotype. To overcome this issue, we developed an inducible

muscle-specific MCU knockout mouse (HSA-Cre-ERT2::_MCU_fl/fl), in which the MCUfl/fl mouse is mated with the HSA-Cre-ERT2 mouse, i.e., a mouse carrying the Cre recombinase fused to a mutated

ligand-binding domain of the human estrogen receptor α and controlled by the human skeletal actin promoter [16]. Upon tamoxifen treatment, the Cre enzyme translocates into the nucleus . The

HSA-Cre-ERT2::_MCU_fl/fl mouse, treated with tamoxifen during adulthood, was named iskMCU−/− (Fig. 5a). However, even long-term tamoxifen administration was only partially effective in

deleting MCU. While in some muscles MCU synthesis was inhibited, either totally or partially, in others (e.g., the diaphragm) it was unaffected (Fig. 5b and S4A). Coherently, fiber size

reduction was proportional to the degree of MCU downregulation (Fig. 5c). Nevertheless, performance of iskMCU−/− mice on an uphill exhaustion exercise protocol was significantly reduced

(Fig. 5d). Finally, in the soleus muscle, the expression of slow-twitch type 1 MHC was decreased, whereas that of type 2A MHC expression was increased (Fig. 5e). MCU DELETION IN ADULT

SKELETAL MUSCLE TRIGGERS LOCAL AND SYSTEMIC METABOLIC ADAPTATIONS Similarly to skMCU−/− mice, basal, maximal, and ATP-linked OCR of iskMCU−/− FDB myofibers were reduced compared to control

fibers (Fig. 6a and S5A). p-PDH in FDB muscle was increased (Fig. 6b). In agreement with impaired pyruvate utilization, the drastic reduction of basal OCR triggered by UK5099-dependent MPC

inhibition in MCUfl/fl myofibers (Figs 2e and 6c) was not observed in iskMCU−/− fibers (Fig. 6c). In addition, FA dependency of basal OCR was double in iskMCU−/− fibers compared to controls

(Fig. 6d). Although fasting glycemia was unaltered (Fig. 6e and S5B), blood lactate levels were increased in iskMCU−/− mice relative to controls (Fig. 6f and S5C), coherently with impaired

glucose oxidation. In addition, similarly to skMCU−/− mice, loss of MCU in adult skeletal muscle caused systemic metabolic effects, as demonstrated by the increased blood FA (Fig. 6g) and

ketone bodies (Fig. 6h). The incomplete MCU deletion prevented any further biochemical and physiological analysis, although the metabolic adaptations were anyway partially evident.

DISCUSSION The role of Ca2+ uptake by the skeletal muscle mitochondria was recently investigated [9, 10, 12], but outstanding questions still remain. First, the causes underlying the reduced

exercise performance consequent to MCU deletion are unclear. In our study, deletion of MCU specifically in the skeletal muscle impaired exercise performance, demonstrating that the

decreased running capacity is solely due to a skeletal muscle defect. In vivo measurements demonstrated reduced tetanic force of skMCU−/− muscles. However, the strength developed at

frequencies recruited during treadmill exercise is unaffected by loss of MCU. In addition, our results exclude the contribution of impaired fatigue resistance to the skMCU−/− phenotype.

Coherently with the decrease in exercise performance, a slow-to-fast MHC shift characterizes both skMCU−/− and iskMCU−/− soleus muscles, thus excluding a compensatory effect occurring during

development. Since muscle remodeling toward faster MHCs occurs in disuse conditions [29], one possible explanation is that the inefficient substrate oxidation of MCU−/− muscles impairs

muscle activity, which in turn determines MHC expression remodeling. The shift in MHC expression is coherent with a metabolic switch toward decreased oxidative metabolism (Fig. 7a). In line

with this, we measured a major OCR decrease in MCU-deleted fibers, indicating that MCU loss can limit the supply of reducing equivalents to the electron transport chain. This effect is

partially mitigated by upregulation in lipid catabolism. Similarly, inhibition of MPC has been shown to trigger a profound metabolic rewiring and to boost FA oxidation in muscle cells [27],

thus suggesting that the impairment in pyruvate catabolism is the main trigger of the ongoing metabolic rewiring. To confirm this notion, we here show that reduced PDH activity is _per se_

sufficient to shift substrate preference from carbohydrates toward lipids. Nonetheless, the reduced maximal OCR in PDK4-overexpressing fibers suggests that optimal glucose oxidation is

required to sustain high-energy demand, which is not otherwise met by the alternative FA metabolism. Accordingly, overexpression of the Ca2+-independent PDP2 isoform in skMCU−/− fibers was

sufficient to inhibit the preferred lipid oxidation. Altogether, these data indicate that MCU controls muscle substrate preference and warrants a high level of plasticity, by impinging on

the activity level of PDH. In the absence of MCU, pyruvate to acetyl-CoA conversion slows down, thus limiting the TCA supply, and rewiring of cell metabolism toward preferential lipid

catabolism partially compensates this defect. However, MCU loss causes an additional bioenergetics defect, because mitochondrial Ca2+ level is a positive allosteric regulator of two

additional TCA cycle enzymes (i.e., isocitrate and α-ketoglutarate dehydrogenases). OCR measurements in the presence of palmitate, directly demonstrated that, although MCU−/− muscles mostly

rely on β-oxidation to sustain respiration, their overall capacity to oxidize FA is actually reduced. Thus, in MCU−/− myofibers, the flux of FA-derived acetyl-CoA through the TCA cycle is

slowed down because of isocitrate and α-ketoglutarate dehydrogenase inhibition, decreasing the availability of reducing equivalents fuelling basal respiration. Overall, the consequences of

MCU deletion on mitochondria function in the skeletal muscle are apparent already in basal conditions, although metabolic rewiring mitigates potential more severe phenotype. In light of the

function of skeletal muscle activity on whole-body metabolism, the metabolic defect of MCU-deleted muscles translates into a systemic catabolic response (Fig. 7b). Whether this is a direct

effect of altered serum metabolites concentrations, or rather circulating factors (i.e., myokines) play a role, is not clear yet. By means of the iskMCU−/− mouse, we wished to uncover

potential phenotypic features otherwise lost by developmental adaptations. However, complete MCU deletion was achieved only in some muscles. Nonetheless, iskMCU−/− mouse phenotype was

similar to skMCU−/−, suggesting that either embryonic compensations are negligible or that complete MCU ablation in the adult would be required to uncover further defects. Finally, our

models are in accordance with observations made by cardiomyocyte-specific conditional MCU deletion [30, 31] in terms of lack of overt basal phenotype and absolute requirement of effective

mitochondrial Ca2+ uptake to match the metabolic output with increased energy demand. However, contrary to skMCU and iskMCU muscles, baseline p-PDH was unaltered in MCU_fl/fl-MCM_ hearts

[30, 31], maybe reflecting either the lower MCU currents or the high dependency on FA oxidation of cardiac mitochondria. Overall, our data indicate that a complex metabolic rewiring occurs

in skMCU−/− muscles. Mitochondrial Ca2+ uptake is required for efficient glucose oxidation and sustains rapid flux through the TCA cycle. The impairment in muscle performance of skMCU−/−

mice is mainly due to reduced glucose oxidation consequent to inhibition of PDH and thus TCA cycle activity, which is accompanied by fiber-type remodeling toward faster MHCs. Systemic

adaptations may also contribute to the decreased exercise capacity. Metabolic flexibility in terms of increased FA oxidation at least partially sustains muscle activity and compensates for

the impairment in carbohydrates' utilization. On the one hand, this metabolic rewiring contributes to explain the mild phenotype of MCU-deficient mouse models. On the other, despite

these profound metabolic adaptations, the selective loss of MCU in skeletal muscle still shows a negative impact on muscle strength and performance, thus further underlining the

physio-pathological relevance of mitochondrial Ca2+ homeostasis. REFERENCES * Rizzuto R, De Stefani D, Raffaello A, Mammucari C. Mitochondria as sensors and regulators of calcium signalling.

Nat Rev Mol Cell Biol. 2012;13:566–78. Article  CAS  Google Scholar  * Rizzuto R, Brini M, Murgia M, Pozzan T. Microdomains with high Ca2+close to IP3-sensitive channels that are sensed by

neighboring mitochondria. Science. 1993;262:744–7. Article  CAS  Google Scholar  * De Stefani D, Raffaello A, Teardo E, Szabò I, Rizzuto R. A forty-kilodalton protein of the inner membrane

is the mitochondrial calcium uniporter. Nature. 2011;476:336–40. Article  Google Scholar  * Baughman JM, Perocchi F, Girgis HS, Plovanich M, Belcher-Timme CA, Sancak Y, et al. Integrative

genomics identifies MCU as an essential component of the mitochondrial calcium uniporter. Nature. 2011;476:341–5. Article  CAS  Google Scholar  * Raffaello A, De Stefani D, Sabbadin D,

Teardo E, Merli G, Picard A, et al. The mitochondrial calcium uniporter is a multimer that can include a dominant-negative pore-forming subunit. EMBO J. 2013;32:2362–76. Article  CAS  Google

Scholar  * Sancak Y, Markhard ALL, Kitami T, Kovacs-Bogdan E, Kamer KJJ, Udeshi NDD, et al. EMRE is an essential component of the mitochondrial calcium uniporter complex. Science.

2013;342:1379–82. Article  CAS  Google Scholar  * Perocchi F, Gohil VM, Girgis HS, Bao XR, McCombs JE, Palmer AE, et al. MICU1 encodes a mitochondrial EF hand protein required for

Ca2+uptake. Nature. 2010;467:291–6. Article  CAS  Google Scholar  * Plovanich M, Bogorad RL, Sancak Y, Kamer KJ, Strittmatter L, Li AA, et al. MICU2, a paralog of MICU1, resides within the

mitochondrial uniporter complex to regulate calcium handling. PLoS ONE. 2013;8:e55785. Article  CAS  Google Scholar  * Mammucari C, Gherardi G, Zamparo I, Raffaello A, Boncompagni S,

Chemello F, et al. The mitochondrial calcium uniporter controls skeletal muscle trophism in vivo. Cell Rep. 2015;10:1269–79. Article  CAS  Google Scholar  * Zampieri S, Mammucari C,

Romanello V, Barberi L, Pietrangelo L, Fusella A, et al. Physical exercise in aging human skeletal muscle increases mitochondrial calcium uniporter expression levels and affects mitochondria

dynamics. Physiol Rep. 2016;4:e13005. Article  Google Scholar  * Vecellio Reane D, Vallese F, Checchetto V, Acquasaliente L, Butera G, De Filippis V, et al. A MICU1 splice variant confers

high sensitivity to the mitochondrial Ca2+ uptake machinery of skeletal muscle. Mol Cell. 2016;64:760–73. Article  CAS  Google Scholar  * Pan X, Liu J, Nguyen T, Liu C, Sun J, Teng Y, et al.

The physiological role of mitochondrial calcium revealed by mice lacking the mitochondrial calcium uniporter. Nat Cell Biol. 2013;15:1464–72. Article  CAS  Google Scholar  * Tosatto A,

Sommaggio R, Kummerow C, Bentham RB, Blacker TS, Berecz T, et al. The mitochondrial calcium uniporter regulates breast cancer progression via HIF-1α. EMBO Mol Med. 2016;8:569–85. Article 

CAS  Google Scholar  * Zhao Y, Araki S, Wu J, Teramoto T, Chang Y-F, Nakano M, et al. An expanded palette of genetically encoded Ca2+ indicators. Science (80-). 2011;333:1888–91. Article 

CAS  Google Scholar  * Bothe GW, Haspel JA, Smith CL, Wiener HH, Burden SJ. Selective expression of Cre recombinase in skeletal muscle fibers. Genesis. 2000;26:165–6. Article  CAS  Google

Scholar  * Schuler M, Ali F, Metzger E, Chambon P, Metzger D. Temporally controlled targeted somatic mutagenesis in skeletal muscles of the mouse. Genesis. 2005;41:165–70. Article  CAS 

Google Scholar  * Blaauw B, Canato M, Agatea L, Toniolo L, Mammucari C, Masiero E, et al. Inducible activation of Akt increases skeletal muscle mass and force without satellite cell

activation. FASEB J. 2009;23:3896–905. Article  CAS  Google Scholar  * Schindelin J, Arganda-Carreras I, Frise E, Kaynig V, Longair M, Pietzsch T, et al. Fiji: an open-source platform for

biological-image analysis. Nat Methods. 2012;9:676–82. Article  CAS  Google Scholar  * Rozen S, Skaletsky H. Primer3 on the WWW for general users and for biologist programmers. Methods Mol

Biol. 2000;132:365–86. CAS  PubMed  Google Scholar  * TeSlaa T, Teitell MA. Techniques to monitor glycolysis. Methods Enzymol. 2014;542:91–114. Article  CAS  Google Scholar  * Saito K, Lee

S, Shiuchi T, Toda C, Kamijo M, Inagaki-Ohara K, et al. An enzymatic photometric assay for 2-deoxyglucose uptake in insulin-responsive tissues and 3T3-L1 adipocytes. Anal Biochem.

2011;412:9–17. Article  CAS  Google Scholar  * Denton RM. Regulation of mitochondrial dehydrogenases by calcium ions. Biochim Biophys Acta Bioenerg. 2009;1787:1309–16. Article  CAS  Google

Scholar  * Gray LR, Sultana MR, Rauckhorst AJ, Oonthonpan L, Tompkins SC, Sharma A, et al. Hepatic mitochondrial pyruvate carrier 1 is required for efficient regulation of gluconeogenesis

and whole-body glucose homeostasis. Cell Metab. 2015;22:669–81. Article  CAS  Google Scholar  * McCommis KS, Chen Z, Fu X, McDonald WG, Colca JR, Kletzien RF, et al. Loss of mitochondrial

pyruvate carrier 2 in the liver leads to defects in gluconeogenesis and compensation via pyruvate-alanine cycling. Cell Metab. 2015;22:682–94. Article  CAS  Google Scholar  * Bricker DK,

Taylor EB, Schell JC, Orsak T, Boutron A, Chen Y-C, et al. A mitochondrial pyruvate carrier required for pyruvate uptake in yeast, drosophila, and humans. Science. 2012;337:96–100. Article 

CAS  Google Scholar  * Herzig S, Raemy E, Montessuit S, Veuthey J-L, Zamboni N, Westermann B, et al. Identification and functional expression of the mitochondrial pyruvate carrier. Science.

2012;337:93–96. Article  CAS  Google Scholar  * Vacanti NM, Divakaruni AS, Green CR, Parker SJ, Henry RR, Ciaraldi TP, et al. Regulation of substrate utilization by the mitochondrial

pyruvate carrier. Mol Cell. 2014;56:425–35. Article  CAS  Google Scholar  * Muñoz-Cánoves P, Scheele C, Pedersen BK, Serrano AL. Interleukin-6 myokine signaling in skeletal muscle: a

double-edged sword? FEBS J. 2013;280:4131–48. Article  Google Scholar  * Ciciliot S, Rossi AC, Dyar KA, Blaauw B, Schiaffino S. Muscle type and fiber type specificity in muscle wasting. Int

J Biochem Cell Biol. 2013;45:2191–9. Article  CAS  Google Scholar  * Kwong JQ, Lu X, Correll RN, Schwanekamp JA, Vagnozzi RJ, Sargent MA, et al. The mitochondrial calcium uniporter

selectively matches metabolic output to acute contractile stress in the heart. Cell Rep. 2015;12:15–22. Article  CAS  Google Scholar  * Luongo TS, Lambert JP, Yuan A, Zhang X, Gross P, Song

J, et al. The mitochondrial calcium uniporter matches energetic supply with cardiac workload during stress and modulates permeability transition. Cell Rep. 2015;12:23–34. Article  CAS 

Google Scholar  Download references ACKNOWLEDGEMENTS Research was supported by fundings from the European Union (ERC mitoCalcium, no. 294777 to RR), the Italian Ministry of Education,

University, and Research (PRIN 2015W2N883_003 to CM), the French Muscular Dystrophy Association AFM-Téléthon (18857 to CM), the Italian Telethon Foundation (GGP16029 to RR), and the Italian

Association for Cancer Research (IG 18633 to RR). We thank Margherita Franceschini for assistance with Ca2+ measurements, Elisabetta Iori for assistance in metabolites measurements, and the

laboratory of Jorge Ruas for sharing primer sequences. AUTHOR CONTRIBUTIONS GG performed most of the experiments, LN and BB performed force measurements, SC performed in vivo

2-deoxy-d-glucose uptake experiments, PBr and PBo performed microinjections for the production of the MCUfl/fl mouse, GPF supervised metabolites measurements, DDS supervised metabolomics

experiments, RR and CM conceived the research, and CM directed the research and wrote the paper. AUTHOR INFORMATION AUTHORS AND AFFILIATIONS * Department of Biomedical Sciences, University

of Padova, 35131, Padova, Italy Gaia Gherardi, Leonardo Nogara, Bert Blaauw, Diego De Stefani, Rosario Rizzuto & Cristina Mammucari * Department of Medicine, University of Padova, 35128,

Padova, Italy Stefano Ciciliot & Gian Paolo Fadini * Venetian Institute of Molecular Medicine, 35129, Padova, Italy Stefano Ciciliot, Gian Paolo Fadini & Bert Blaauw * Department of

Molecular Medicine, University of Padova, 35131, Padova, Italy Paola Braghetta & Paolo Bonaldo Authors * Gaia Gherardi View author publications You can also search for this author

inPubMed Google Scholar * Leonardo Nogara View author publications You can also search for this author inPubMed Google Scholar * Stefano Ciciliot View author publications You can also search

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can also search for this author inPubMed Google Scholar * Paola Braghetta View author publications You can also search for this author inPubMed Google Scholar * Paolo Bonaldo View author

publications You can also search for this author inPubMed Google Scholar * Diego De Stefani View author publications You can also search for this author inPubMed Google Scholar * Rosario

Rizzuto View author publications You can also search for this author inPubMed Google Scholar * Cristina Mammucari View author publications You can also search for this author inPubMed Google

Scholar CORRESPONDING AUTHORS Correspondence to Rosario Rizzuto or Cristina Mammucari. ETHICS DECLARATIONS CONFLICT OF INTEREST The authors declare that they have no conflict of interest.

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Gherardi, G., Nogara, L., Ciciliot, S. _et al._ Loss of mitochondrial calcium uniporter rewires skeletal muscle metabolism and substrate preference. _Cell Death Differ_ 26, 362–381 (2019).

https://doi.org/10.1038/s41418-018-0191-7 Download citation * Received: 15 February 2018 * Revised: 07 August 2018 * Accepted: 10 August 2018 * Published: 19 September 2018 * Issue Date:

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