Lipid accumulation and mitochondrial abnormalities are associated with fiber atrophy in the skeletal muscle of rats with collagen-induced arthritis

Journal Pre-proof Lipid accumulation and mitochondrial abnormalities are associated with fiber atrophy in skeletal muscle of rats with collagen-induced arthritis

Gaelle Vial, Cécile Coudy-Gandelon, Alexandre Pinel, Fabien Wauquier, C. Chevenet, Daniel Bechet, Yohan Wittrant, Veronique Coxam, Martin Soubrier, Anne Tournadre, Frederic Capel PII:

S1388-1981(19)30225-2

DOI:

https://doi.org/10.1016/j.bbalip.2019.158574

Reference:

BBAMCB 158574

To appear in:

BBA - Molecular and Cell Biology of Lipids

Received date:

22 March 2019

Revised date:

29 October 2019

Accepted date:

31 October 2019

Please cite this article as: G. Vial, C. Coudy-Gandelon, A. Pinel, et al., Lipid accumulation and mitochondrial abnormalities are associated with fiber atrophy in skeletal muscle of rats with collagen-induced arthritis, BBA - Molecular and Cell Biology of Lipids(2019), https://doi.org/10.1016/j.bbalip.2019.158574

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© 2019 Published by Elsevier.

Muscle abnormalities and arthritis

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Lipid accumulation and mitochondrial abnormalities are associated with fiber atrophy in skeletal muscle of rats with collagen-induced arthritis Gaelle Vial1, Cécile Coudy-Gandelon2, Alexandre Pinel2, Fabien Wauquier2, C Chevenet3, Daniel Bechet2, Yohan Wittrant2, Veronique Coxam2, Martin Soubrier1,2,

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Anne Tournadre1,2, Frederic Capel2

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1 : Service de rhumatologie, CHU de Clermont-Ferrand, 63003 Clermont-Ferrand 2 : Unité de Nutrition Humaine (UNH), INRA/Université Clermont Auvergne, 63009

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Clermont-Ferrand, France

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3 : Laboratoire d'Anatomie et de Cytologie Pathologique, CHU de Clermont-Ferrand,

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Hôpital Estaing, 63003 Clermont-Ferrand, France

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*Contact information for corresponding author : Frederic Capel, 28 place Henri Dunant - BP 38, UFR DE MEDECINE, UMR1019, Equipe ASMS ; 63001 Clermont-

Highlights:

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Ferrand Cedex 1, France. Tel.: +33 4 73 17 82 67; E-mail : [email protected]

Arthritis induces skeletal fiber atrophy Triglycerides accumulate in skeletal muscle from arthritic rats Alteration in skeletal muscle lipid metabolism are related to mitochondrial defects

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Abstract Rheumatoid arthritis (RA) has a negative impact on muscle mass, and reduces patient’s mobility and autonomy. Furthermore, RA is associated with metabolic comorbidities, notably in lipid homeostasis by unknown mechanisms. To understand the links between the loss in muscle mass and the metabolic abnormalities, arthritis was induced in male Sprague Dawley rats (n=11) using the collagen-induced arthritis

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model. Rats immunized with bovine type II collagen were compared to a control group of animals (n=11) injected with acetic acid and complete Freund’s adjuvant.

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The clinical severity of the ensuing arthritis was evaluated weekly by a semi-

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quantitative score. Skeletal muscles from the hind limb were used for the histological

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analysis and exploration of mitochondrial activity, lipid accumulation, metabolism and

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regenerative capacities A significant atrophy in tibialis anterior muscle fibers was observed in rats with arthritis despite a non-significant decrease in the weight of the

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muscles. Despite moderate inflammation, accumulation of triglycerides (P<0.05), reduced mitochondrial DNA copy number (P<0.05) and non-significant dysfunction in

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mitochondrial cytochrome c oxidase activity were found in the gastrocnemius muscle. Concomitantly, our results suggested an activation of the muscle specific E3 ubiquitin ligases MuRF-1 and MAFbx. Finally, the adipose tissue from the arthritic rats exhibited decreased PPARγ mRNA suggesting reduced adipogenic capacities. In conclusion, the reduced adipose tissue adipogenic capacity and skeletal muscle mitochondrial capacity are probably involved in the activation of protein catabolism, inhibition of myogenesis, accumulation of lipids and fiber atrophy in skeletal muscle during RA. Keywords Muscle, Lipids, Mitochondria, Sarcopenia, Arthritis 2

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Abbreviations 2M, alpha2-macroglobulin; cDNA, complementary DNA; CIA, collagen induced arthritis; COX, cytochrome c oxidase; CS, citrate synthase; CSA, cross section area; DAGs, diacylglycerols; FA, fatty acid; HAD, 3-hydroxyacyl-CoA dehydrogenase; mRNA, messenger RNA; mtDNA, mitochondrial DNA, nDNA, nuclear DNA; PCR,

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polymerase chain reaction; RA, rheumatoid arthritis; TAGs, triacylglycerols

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1 Introduction

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Rheumatoid arthritis (RA) induces alterations in body composition characterized by a

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loss in both muscle mass and strength, increased fat mass and intramuscular lipid

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infiltration, contributing to disability and decreased quality of life [1, 2]. Furthermore, RA also predisposes patients to metabolic syndrome, insulin resistance and

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cardiovascular disease [3]. The excess of adipose tissue and the related defects in its functions could mediate skeletal muscle impairments and the development of

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metabolic syndrome during RA. The infiltration of ectopic fat in skeletal muscle was found to be associated with metabolic abnormalities, reduced strength and lower scores in performance tests, increasing the incidence of mobility disability in the elderly [4]. Ectopic fat decreases muscle anabolic response to nutrients and insulin leading to a chronic imbalance in the rates of protein synthesis and breakdown which promotes skeletal muscle loss [5]. On the contrary, such a link was not observed when fatty acid (FA) oxidation capacity was stimulated by physical training [6]. Metabolic abnormalities are also closely linked to mitochondrial dysfunctions leading to impaired FA oxidation and energy production, inducing alterations in metabolic homeostasis [7] and protein anabolism [8]. A limitation in adipose tissue expansion 3

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capacity could also promote fat redistribution in other organs [9]. A chronic inflammatory state could stimulate muscle wasting because of the activation of proteolytic processes. It was found that two muscle-specific E3 ubiquitin ligases, muscle RING finger-1 (MuRF-1) and muscle atrophy F-Box (MAFbx ; also called atrogin-1) which play a major role in skeletal muscle atrophy, were increased with arthritis in a rodent model [10, 11]. In other catabolic conditions (cancer, inactivity, unloading) or when muscle cells are exposed to tumor necrosis factor alpha (TNF-α),

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activation of p38 mitogen-activated protein kinases (p38 MAPK) and eukaryotic

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translation initiation factor 2 alpha (eIF2α) proteins was found to control the activation

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of MuRF1 and MAFbx E3 ligases [12-14]. It remains unknown if these proteins are

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involved in muscle abnormalities during RA. The contribution of alterations in regenerating processes to muscle wasting in RA patients is also undetermined. Any

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defect in the activation, proliferation and differentiation of satellite cells to replace

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damaged fibers could affect muscle mass and function. The regeneration of skeletal muscle is controlled by myogenic differentiation factor D (MyoD) which is active in the

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early phase of this process and myogenin which is one of the master controllers of differentiation. MyoD and myogenin were found to be affected in catabolic states [15, 16] but other regulators, such as myogenic factor 5 (Myf5) and myogenic regulatory factor 4 (Mrf4) were also found to be involved [17]. To the best of our knowledge, no data on the role of these proteins are available in the context of RA. Consequently, the aim of this study was to evaluate alterations in skeletal muscle mass, mitochondrial function, ectopic fat accumulation and muscle metabolism in a rodent model of arthritis to better understand how targeted pharmaceutical or nutritional strategies could be proposed to RA patients. 2 Materials and methods 4

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2.1 Animals and experimental groups Male Sprague-Dawley rats of 12 weeks of age were randomly assigned to a control and a collagen-induced arthritis (CIA) group (n=11 in each group). Animals were kept at 20 °C, with a 12h/12h light–dark cycle and free access to food and water. All experiments were performed according to the guidelines for animal care and with the

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authorization from the local ethical committee (ref 6522 – 2016081016428147).

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Arthritis was induced with bovine type II collagen (CII; Chondrex Inc., Redmond, WA,

(CFA;

Sigma-Aldrich Saint-Quentin

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USA) dissolved in 0.1M acetic acid and emulsified with complete Freund’s adjuvant Fallavier,

France)

containing

inactivated

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Mycobacterium tuberculosis (1mg/mL). Equal volumes of CII and CFA were mixed to

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form the emulsion. For primary immunization, 400 μg CII was injected intradermally on both sides of the tail. Seven days after the first injection, a booster injection was

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performed with 200 μg CII emulsified in an equal volume of CFA. Control rats

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received injections of acetic acid + CFA. 2.2 Clinical evaluation of CIA

The clinical severity of arthritis was evaluated weekly according to the presence of oedema and/or erythema in the paws. The phenotypic changes in each paw were estimated according to a scale of 0 to 4 per paw. CIA was graded according to modifications in swelling and deformation: 0. no erythema or swelling; 1. erythema and mild swelling confined to one site; 2. two or more sites with erythema and mild swelling; 3. erythema and severe swelling of the whole paw; 4. severe swelling with ankylosis, deformation, and functional disturbance [18]. 2.3 Sacrifice and tissue sampling 5

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Animals were sacrificed by exsanguination (blood collection from the abdominal aorta) and cervical dislocation under isoflurane anesthesia 4 weeks after the booster injection. Skeletal muscles and perigonadal adipose tissue were harvested and weighed before storage for further analysis. One gastrocnemius muscle was frozen in liquid nitrogen before storage at -80 °C for lipid/protein/gene expression analyses and enzymatic measurements. One tibialis anterior muscle was fixed in Formalin (Sigma-Aldrich) and embedded in paraffin for histological examination. Tibialis

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anterior muscle from the second paw was immediately embedded in Optimal Cutting

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Temperature compound (OCT, CellPath, UK) and frozen in isopentane (Sigma-

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Aldrich) cooled with liquid nitrogen. Blood was collected in EDTA-coated tubes and

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2.4 Plasmatic parameter assays

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centrifuged at 1,000 g for plasma isolation before storage at -80 °C.

Plasma glucose, total cholesterol, triacylglycerol (kits from Thermo-Fisher); HDL

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cholesterol, non-esterified FAs (kits from Diasys, Grabels, France) and C-Reactive

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Protein (kit from ALPCO Diagnostics, Salem, NH, USA) were quantified using a Konelab system (Thermo-Fisher, Asnières sur Seine, France). Alpha-2-macroglobulin (α2M) was quantified using an enzyme-linked immunosorbent assay (ELISA) according to the manufacturer’s instructions. The α2M plasma level was determined based on the calibration curve obtained with the dilution of a standard solution provided in the kit; all measurements were performed in triplicate at a fixed absorbance (450 nm) using a microplate-reader (Epoch, BioTek, Colmar, France. 2.5 Histology

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Serial cross-sections (10 µm thick) of tibialis anterior muscle embedded in OCT were performed using a cryostat (Microm, Francheville, France) at −25 °C. Dried sections were incubated for 15 minutes in 10% Goat Serum (Sigma-Aldrich), washed in 1X PBS and then incubated for 1 hour at 37 °C in a humid chamber with anti-laminin antibody (Sigma-Aldrich) diluted in 10% goat serum (1:200) to delimit fibers. Slides were washed with 1X PBS and incubated with Alexa fluor 546 Goat anti Rabbit

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antibody (Invitrogen) diluted in 10% goat serum (1:400). Slides were washed with 1X

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PBS, then mounted in an aqueous medium, before image acquisition on a BX51 fluorescence microscope (Olympus) coupled to a DP72 camera. Approximately 800

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to 900 fibers were analysed from 6 to 7 views (n= 9 per group). Image analysis was

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performed using the Visilog 6.9 software (Noesis, France) for histomorphometric fiber

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characterization including cross sectional area (CSA), fiber diameter and variance coefficient of the fiber size. The variance coefficient of the muscle fiber size is defined

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as the standard deviation of the muscle fiber diameter/mean muscle fiber diameter x

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1000 [19]. A shape factor was calculated (perimeter2/4π CSA) to quantify fiber circularity, a value of 1.0 indicating a circle, and >1.0, an increasingly elongated ellipse [20]. Observations were also performed using the conventional hematoxylin and eosin staining.

2.6 Skeletal muscle lipid content Total lipid content was estimated in skeletal muscle after extraction using the Folch method [21]. The final lipid fraction was used for the quantification of neutral lipids, triacylglycerols (TAGs) and diacylglycerols (DAGs). The TAGs were measured using a colorimetric method (Diasys). The DAGs were quantified by Gas Chromatography 7

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coupled to a flame ionization detector (Thermo Electron Corporation; Waltham, MA) after the removal of phospholipids from the final lipid extract using CHROMABOND® SiOH columns (MACHEREY-NAGEL, 67722 Hoerdt, France). Neutral lipid extracts were suspended in ethyl acetate before injection. Analyses were performed with a DB-5MS capillary column (5 m/0.25 mm internal diameter/0.25 µm film thickness, Agilent Technologies), using hydrogen as a carrier gas (3 mL/min). DAGs were

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quantified using a C14 DAG as the internal standard.

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2.7 Mitochondrial enzymatic activities

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Gastrocnemius muscle (50 mg) was homogenized in Tris buffer (Tris-Base 10mM, mannitol 225mM, sucrose 75mM, EDTA 0.5M, pH 7.2) on ice. Citrate synthase (CS)

following

the

conversion

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5,5-dithiobis(2-nitrobenzoate)

(DTNB)

into

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by

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activity was determined spectrophotometrically according to the method of Srere [22]

trinitrobenzene (TNB) at 412 nm using a spectrophotometer. All measurements were

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performed in duplicate, using the same settings at 20 to 22 °C. The solubilized protein extracts of the homogenates were quantified in duplicate by using bicinchoninic acid reagents (Pierce, Rockford, IL) and bovine serum albumin (BSA) as the standard.

Cytochrome c oxidase (COX) activity was determined spectrophotometrically at 37 °C by monitoring the oxidation of cytochrome c (Sigma-Aldrich) at 550 nm. 3hydroxyacyl-CoA dehydrogenase (HAD) activity was determined by monitoring the oxidation rate of NADH into NAD+ in the presence of acetoacetyl-CoA at 340 nm using a spectrophotometer. All measurements were performed in duplicate. The solubilized protein extracts of the homogenates were quantified in duplicate by using

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bicinchoninic acid reagents (Pierce) and BSA as the standard. All activities were normalized to the total protein content. 2.8 mRNA level quantification Total RNA was extracted from 80-100 mg of gastrocnemius muscle and 250-300 mg of adipose tissue using TRIzol® reagent (Thermo Scientific) according to the manufacturer’s instructions. RNA quantification and integrity were verified by

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measuring the ratio of optical density at 260 nm and 280 nm and by agarose gel,

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respectively. cDNAs were synthesized from 2 µg of total RNA using a High Capacity

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cDNA Reverse Transcription Kit from Applied Biosystem (Thermo Scientific). The products of reverse transcription were used for Quantitative Real Time Polymerase

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Chain Reaction (qRT-PCR) using specific primers and Rotor-Gene SYBR Green

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PCR master mix on a Rotor-Gene Q system (Qiagen, Courtaboeuf, France). Messenger RNA (mRNA) quantification was assayed using the standard curve of

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native cDNA and serial dilutions. Primer sequences and PCR conditions are available

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upon request ([email protected]). Beta actin (b-actin) and 18S rRNA genes were used as housekeeping gene in skeletal muscle and adipose tissue, respectively. 2.9 Mitochondrial DNA quantification DNA was isolated from gastrocnemius muscle during the RNA extraction procedure by recovering the phenolic fraction which was mixed with back extraction buffer (guanidine thiocyanate 4M, sodium citrate 50mM, Tris base 1M). After spinning, DNA was precipitated using isopropanol and ethanol as described in the TRIzol® reagent datasheet. Using the calculation method described by Rooney et al. [23], the quantification of NADH dehydrogenase, subunit 1 (ND1) and β-actin, was used as the marker of mitochondrial DNA (mtDNA) and the nuclear DNA (nDNA) amount,

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respectively. Experiments were performed from 4 ng of total DNA using the RotorGene SYBR Green PCR master mix on a Rotor-Gene Q system. 2.10 Western blotting Frozen tissues were homogenized in lysis buffer (50 mM HEPES, 150 mM sodium chloride, 10 mM EDTA, 10 mM sodium pyrophosphate tetrabasic anhydrous, 25 mM β-glycerophosphate,

100

mM

sodium

fluoride,

10%

glycerol

anhydrous)

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supplemented with a cocktail of phosphatase inhibitors (Sigma Aldrich). Protein

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quantification was performed using a BCA protein assay kit (Pierce, Thermo

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Scientific) and a BSA calibration curve according to the manufacturer’s instructions. For immunoblotting, 20 to 30 micrograms of proteins were loaded for separation by

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SDS-PAGE electrophoresis and transfer onto PVDF membranes. The membranes

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were immunoblotted with the appropriate antibody to detect glyceraldehyde 3phosphate dehydrogenase (GAPDH) (Sigma), serine 473 phosphorylated protein

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kinase B (AKT) & total AKT (Cell signaling, Ozyme, Saint-Cyr-L’École, France),

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phosphorylated p38 MAPK & total p38 MAPK (Sigma), MAFbx and MuRF1 (ECM bioscience, Euromedex, Souffelweyersheim, France). Antibody binding was detected using HRP-conjugated secondary antibodies and ECL western blotting substrate (Thermo Scientific). Immunoblots were visualized using a chemiluminescence imaging system (MF ChemiBIS 2020, DNR bio imaging systems, Jerusalem, Israel) and quantified using Multi Gauge V3.2 software. 2.11 Statistics Results are expressed as mean values with standard error (SE). Data were compared using a student t test. P values < 0.05 were considered significant. All

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statistical analyses were performed using R software (R Foundation for Statistical Computing, Vienna, Austria). 3 Results

3.1 Animal and muscle weight Food consumption and weight variation were similar between the two groups during

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the 4-week experiment. Final body weight was similar between the groups (data not shown). Joint swelling was only observed in the hind limbs of the animals after the

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immunization with collagen. The mean arthritis score in CIA was 2.7/8. Absolute

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gastrocnemius and the tibialis anterior muscle weights tended to decrease in the CIA

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group compared to the control group (2.28 ± 0.49 g versus 2.58 ± 0.28 g, P=0.11 and

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0.71 ± 0.11 g versus 0.76 ±0.06, P=0.26 for gastrocnemius and tibialis anterior, respectively). The extensor digitorum longus muscle weight was significantly lower in

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the CIA group (0.18 ± 0.03 g) than in the control group (0.20 ± 0.02 g, P=0.05). CIA did not modify the plasma glucose levels, nor the lipid profile (data not shown). No

groups.

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differences were observed in the epididymal adipose tissue weight between the

3.2 Effect of CIA on plasma α2M α2M is a typical acute-phase protein which could be considered as an inflammatory marker linked to sarcopenia in rats [24]. We observed a non-significant increase (P=0.17) in the concentration of α2M in the plasma from the CIA animals (1.73 ± 0.12 mg/ml compared to the controls (1.51 ± 0.10 mg/ml). 3.3 Muscle histology CSA, perimeter, diameter and shape were assessed for 800 to 1,000 muscle fibers per tibialis anterior muscle. Muscle sections were reduced in the CIA group (Figure 11

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1A)

compared

to

the

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group.

Quantitative

analysis

showed

a

nonhomogeneous distribution of muscle fiber CSAs with a predominance of small fibers in the CIA group (Figure 1B). As shown in Figure 1C, mean diameter was also significantly decreased in the rats with CIA (195.1 ± 19.5 µm and 62.1 ± 6.2 µm) compared to the control group (224.8 ± 29.2 µm and 71.6 ± 9.3 µm, respectively P<0.05). Even if the fibers were smaller in the CIA animals, the variance coefficient of fiber size was not different between the groups.

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To quantify fiber shape, and therefore evaluate deformation, a shape factor was

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used. As shown in Figure 1D, no difference was observed between the mean shape

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factor in the control group compared to the CIA group (1.43 ± 0.09 vs 1.41 ± 0.05,

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respectively, P=0.58). The tibialis anterior muscle sections were also stained with hematoxylin and eosin to evaluate inflammatory infiltration, necrosis and regeneration

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but no differences were observed between the two groups.

3.4 Muscle molecular markers

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We looked into the potential molecular pathways involved in fiber atrophy in a muscle with very similar properties (more than 80% of IIx IIb fibers) and thus focused on the gastrocnemius muscle (Figure 2A). CIA induced a significant increase in the ubiquitin ligase MAFbx mRNA levels (+42% in the CIA group vs the control group, P<0.05), whereas no difference was observed in the MuRF1 mRNA levels. The mRNA levels of MyoD were reduced in the CIA group compared to the control group (-18%, P<0.05). The mRNA levels of myf5, myostatin and myogenin were not affected by CIA. The protein levels of muscle-specific E3 ubiquitin ligases were also evaluated, revealing an increase in MuRF1 protein although MAFbx was unaffected (Figure 2B, 2C). The control of muscle protein metabolism is dependent on the activation of the

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insulin dependent pathway and the phosphorylation of Akt proteins, but no difference was observed in the amount of phosphorylated Akt between the CIA and control rats. We next evaluated the activation of the key regulators of E3 ligases, but no change in the phosphorylation of p38 MAPK and eIF2a proteins was found (Figure 2D, 2E).

3.5 Muscle cytokine expression The mRNA levels of different cytokines were evaluated to assess the effects of CIA

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on the muscular inflammatory cytokine profile (Figure 3). The gene expression of

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TNFα and anti-inflammatory cytokines, such as interleukin 10 and 15 (IL-10, IL-15),

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were similar between the two groups. On the contrary, the IL-6 mRNA levels were

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reduced in the muscles from the animals with CIA compared to the control rats (-35%

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3.6 Mitochondrial function

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in the CIA group compared to the control group, P<0.05).

mtDNA copy number, COX and CS activities were evaluated in gastrocnemius

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muscle as an index of mitochondrial functionality and density (Figure 4). The mtDNA/nDNA ratio was reduced in the rats with CIA (- 27% in the CIA group compared to the control group, P<0.05, Figure 4A). Mitochondrial COX activity was also decreased by 20% in muscles from the CIA group compared to the control, but the variation did not reach statistical significance (P=0.18, Figure 4B). No difference in CS activity was observed (Figure 4C). In direct line with the reduced mtDNA amount, the ratio of COX/CS activities also tended to be lower with CIA (P<0.1 vs group control, Figure 4D). The mRNA levels of peroxisome proliferator-activated receptor gamma coactivator 1-alpha (PGC1α) and the mitochondrial transcription

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factor A (TFAM), involved in the regulation of muscle mitochondrial biogenesis, were not affected by CIA (data not shown).

3.7 Intramuscular lipid content and lipid metabolism The quantification of the neutral lipid species in gastrocnemius muscle revealed a significant 1.5 fold increase in TAGs in the muscles from the rats with CIA compared to the controls (P=0.05, Figure 5A), whereas the content of DAGs remained

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unchanged (Figure 5B). The accumulation of TAGs was associated with an increased

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expression of FA transporters in gastrocnemius muscle. Indeed, the mRNA levels of

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the membrane-associated fatty acid transport protein 1 (FATP1) were increased by

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35% in the CIA group compared to the control (P<0.05). The mRNA levels of CD36, a FA transporter involved in the uptake of FAs, tended to be higher in the CIA group P=0.25

vs

control).

The

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(+18%,

mitochondrial

transporter,

carnitine

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palmitoyltransferase 1B (CPT1b), also exhibited an increase in its mRNA in the CIA group (+27%, P<0.05). To further explore cellular FA flux, we evaluated the activity of

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mitochondrial HAD, a key enzyme of the β-oxidation cycle, as an index of the FA oxidative capacity in skeletal muscle. The activity of HAD tended to be lower in the CIA group by 24% compared to the control group (Figure 4E, P=0.15). As dysregulations in lipid metabolism and insulin resistance could be closely related, we quantified the mRNA of sterol regulatory element-binding transcription factor 1 (Srebf1), a crucial transcriptional regulator of lipid metabolism. As shown in Figure 5, Srebf1 mRNA levels were decreased in the muscle from the animals with CIA (-37% vs controls, P<0.05). As suggested by the decrease in the mRNA level of proliferator-activated receptor gamma PPARγ (-22%, P<0.05) and Srebf1 (-26%, P<0.1) in the CIA group compared

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to the control, the adipogenic capacity in adipose tissue was diminished by CIA (not shown).

4 Discussion RA is a chronic inflammatory disease of the musculoskeletal system which increases the risk of mortality by about 50%, mainly as a consequence of cardio-metabolic

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comorbidities [25]. Changes in body composition related to decreased mobility, reduced physical activity, inflammation and insulin resistance are observed in the

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early stages of the disease [26]. Skeletal muscle is a source of power and strength

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and thus a key determinant for locomotion. It is also a crucial organ involved in the

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control of energy metabolism. Consequently, skeletal muscle mass and function

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could be considered as a predictive index of the clinical outcome in chronic diseases. Even if still debated, ectopic intramuscular lipid accumulation might be causally linked

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to the development of insulin resistance and skeletal muscle atrophy [9]. A mechanistic hypothesis could involve an increased uptake and a reduced

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mitochondrial oxidation rate of FA. In aging and obesity, the ectopic deposition of lipids in skeletal muscle affects the metabolic regulatory pathways, inducing insulin and muscle anabolic resistance, which are the key mechanisms leading to sarcopenia [5]. To the best of our knowledge, this association between alterations in muscle mass or function, lipid metabolism and mitochondrial dysfunctions has never been studied in the context of RA. In the present study, rats with collagen-induced arthritis did not exhibit any loss in body weight or reduction in food intake. Despite a moderate severity of the disease, the skeletal muscles from the hind limb tended to be atrophied in the animals with CIA. This observation was associated with a reduction in fiber diameter and a higher 15

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proportion of small-sized fibers. However, no modification in fiber morphology was observed. Interestingly, two recent studies have put forward that muscle fibers were atrophied in humans with RA and osteoarthritis [27, 28]. However, no healthy, wellmatched subjects were included in these studies [27, 28]. Muscle injury induces a regenerative process through a well-controlled activation, differentiation and fusion of satellite cells to generate new myofibers [29]. Yet, we

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observed in muscle from arthritic rats, a decrease in the mRNA level of MyoD, a basic helix-loop-helix transcription factor, which is critical for myogenesis and

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differentiation. The mRNA levels of other molecular regulators of differentiation

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(Myogenin) or proliferation (Myf5) were not modified. The mRNA level of myostatin, a

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negative regulator of these processes, remained stable. The activation of protein

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catabolism in muscle from arthritic rats was suggested by the increase in MAFbx mRNA levels. MAFbx is a E3 ubiquitin ligase implicated in muscle atrophy during

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various catabolic states [11] and has been described as being up-regulated in the skeletal muscle from rats with a strong induction of CIA [10]. We also identified a

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higher MuRF1 protein level in the rats with CIA. Like MAFbx, MuRF1 is a E3 ubiquitin ligase, implicated in the degradation of structural proteins. MAFbx is involved in the degradation of regulatory proteins such as eukaryotic initiation factor 3 (eIF-3) or MyoD [30, 31]. Unfortunately, immunoblotting with commercial antibodies did not allow us to detect MyoD in the protein extracted from skeletal muscles. These findings suggest that a higher catabolism of both structural and regulatory proteins could be involved in fiber atrophy with CIA. The molecular regulations that we observed did not appear to be related to an overactivation of p38 MAPK, nor eIF2α. The decreases in muscle mass and fiber size in the CIA rats were not associated with local inflammation. Despite the swelling of the paws, no evident infiltration of 16

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inflammatory cells, necrotic or regenerative fibers was noted during histological observations. At the systemic level, we detected a non-significant increase in the level of circulating α2M. This protein is an acute phase protein and a typical marker of inflammation in rats. The slight increase in its plasma concentration suggested that the peak was reached earlier, probably after 48 to 72 h following injection [32]. It has been demonstrated that IL-6 is involved in the synthesis of acute phase proteins in rats [33]. This may corroborate our hypothesis as we did not observed any increase

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in the mRNA levels of pro-inflammatory markers, such as IL-6 and TNF-α in skeletal

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muscle. On the contrary, the IL-6 mRNA levels were lower in the skeletal muscle

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from rats with CIA. Considering the role of IL-6 in arthritis, the decrease in its gene

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expression could appear surprising. However, a differential regulation of IL-6 and TNF-α expression, a role of IL-6 in intracellular calcium flux [34] and energy sensing

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in skeletal muscle [35], have been previously demonstrated. Further work is

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necessary to decipher the role of IL-6 in inflammation and metabolism locally in skeletal muscle. It is well accepted that abnormalities in skeletal muscle mass and

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metabolism are related to insulin resistance. The activity of the insulin dependent cellular signaling in our model did not seem to be affected because Akt activation was similar between the CIA and control animals. However, no nutritional stimulus was induced before tissue sampling, limiting the exploration of insulin-dependent activation of Akt. In accordance with a plausible insulin resistance, we observed a decreased srebf1 mRNA level in the muscle and adipose tissue from rats with CIA. Such a decrease was also detected in these tissues in diabetic patients [36]. The variations in srebf1 expression suggested that alterations occurred at the level of lipid metabolism. Consistent with this, an accumulation of intramuscular triacylglycerol was observed in rats with CIA. This accumulation could be a consequence of an

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increased FA uptake involving FA transporters such as CD36 or FATP1 and/or a lower oxidation leading to their incorporation into TAG [37]. This uptake could be at least partially linked to a reduced adipogenic capacity in adipose tissue, as suggested by the decrease in PPARγ gene expression in the adipose tissue from arthritic rats. Despite this, no changes were detected in circulating lipids or glucose between the control and CIA animals. To further explain the accumulation of lipids in skeletal muscles, mitochondrial defects have been identified in association with TAG

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accumulation in muscles and insulin resistance in old subjects [38] or diabetic

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patients [7]. Hence, the activity of HAD, one of the main enzymes of the

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mitochondrial b-oxidation pathway, tended to be reduced in skeletal muscle with

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arthritis. On the contrary, the mRNA levels of Cpt-1b, an enzyme involved in FA translocation into mitochondria for their β-oxidation, were elevated in the CIA rats.

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This increase could be a consequence of a compensatory mechanism which uses

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intracellular FA, involving a higher transcriptional activity of PPARs. In line with this, we also found higher mRNA levels of CD36 and FATP1 which are also known to be

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regulated by the transcription factors of the PPAR family [39]. There was a strong tendency for mitochondrial COX to be reduced in skeletal muscle in the CIA rats. An unchanged CS activity indicated no difference in mitochondrial density but indicated a specific decline in COX activity per mitochondria in the CIA group. This decline could be directly linked to alterations at the mitochondrial genome level, as suggested by the lower levels of mtDNA in skeletal muscle in the CIA group. Mitochondrial genome encodes specific proteins of the respiratory chain complexes, including 3 subunits of COX. Alterations induced by CIA appeared to be restricted to the mtDNA copy number because the mRNA levels of TFAM and PGC1α were not different between the control and CIA groups. TFAM and PGC1α are two key

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regulators of the expression of mitochondrial proteins encoded by the mitochondrial and nuclear genomes, respectively. The limitation of the study was the moderate activity of the disease and the lack of assessment of animal locomotor activity during the protocol. But, it was unlikely that a decreased locomotor activity was involved in the observations because the activity of the disease was only moderate, suggesting a small impact on rat mobility. In our

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study, moderate inflammatory joint involvement was sufficient in itself to induce skeletal muscle abnormalities. Furthermore, the effect of disuse in experimental

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arthritis has previously been found not to be a major determinant of muscle atrophy

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[40]. Finally, we performed histologic and metabolic analyses on two different

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muscles. However, the tibialis anterior and the gastrocnemius muscle have a very

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similar fiber pattern with over 80% being composed of glycolytic fibers. Further work is required to explore the contribution of adipose tissue metabolism and adipokine

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production. Furthermore, a more specific analysis of tissular lipidome could help to

5 Conclusion

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identify which reactive lipid mediators are involved in the observed alterations.

The present study demonstrated, for the first time, an association between lipid accumulation, mitochondrial dysfunctions, protein catabolism and the atrophy of muscle fibers in a model of arthritis. It suggests that dedicated strategies should be employed to prevent skeletal muscle loss in patients with RA in order to reduce the impact of metabolic abnormalities involving alterations in lipid metabolism. Acknowledgements

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We are grateful to Jean Paul Rigaudière, Chrystèle Jouve and Patrice Lebecque for

their expert technical assistance to the study. We also thank the staff of the Installation Expérimentale de Nutrition for providing everyday care to the animals. Funding The animal study was supported by a public grant from FEDER- Région Auvergne attributed to the Unité de Nutrition Humaine. This research did not receive any

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specific grant from funding agencies in the commercial, or not-for-profit sectors.

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Financial Disclosures : The authors have no financial support or other financial

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Figure legends Figure 1: Effect of collagen-induced arthritis on skeletal muscle histology A: Representative images of the tibialis anterior section from control and CIA groups (amplification x 100) B: Fiber cross-sectional area distribution of tibialis anterior from control and collagen-

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induced arthritis (CIA) rats; n= 9 per group.

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C: Fiber diameter distribution of tibialis anterior from control and collagen-induced

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arthritis (CIA) rats; n= 9 per group.

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arthritis (CIA) rats; n= 9 per group.

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D: Fiber shape index distribution of tibialis anterior from control and collagen-induced

Figure 2: A: mRNA expression of MAFbx, Murf1, MyoD, myostatin and myogenin in

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gastrocnemius muscle. Expression quantification was performed in the control and

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the collagen-induced arthritis (CIA) groups using β-actin as the housekeeping gene. B: Expression of MuRF1 and MAFbx proteins in gastrocnemius muscle in CIA and control groups (Gapdh protein was used as loading control). C example of immunoblot analysis of MuRF1 and MAFbx protein in the CIA and control groups. D: Phosphorylation of Akt, p38 MAPK and eiF2a proteins in gastrocnemius muscle in CIA and control groups (total Akt, total p38 and Gapdh were used as loading control, respectively). E: example of immunoblot analysis of Akt, p38 MAPK and eiF2a proteins in the CIA and control groups. Data are shown as the mean ± SE of 9-11 animals per group. *P<0.05 compared with control rats, as assessed by t test.

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Figure 3: mRNA expression of cytokines in gastrocnemius muscle. Expression quantification was performed in the control and the collagen-induced arthritis (CIA) groups using β-actin as the housekeeping gene. Data are shown as the mean ± SE of 9-11 animals per group. *P<0.05 compared with the control rats, as assessed by t test.

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Figure 4: Exploration of mitochondrial density and enzymatic activities in

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gastrocnemius muscle. Measurements were performed in the control and the

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collagen-induced arthritis (CIA) groups. A mitochondrial (mtDNA) to total DNA ratio. B cytochrome c oxidase activity (COX). C citrate synthase activity (CS). D COX to CS

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ratio. E 3-hydroxyacyl-CoA dehydrogenase (HAD) activity. Data are shown as the

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assessed by t test.

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mean ± SE of 9-10 animals per group. *P<0.05 compared with the control rats, as

Figure 5: A Content of triacylglycerol (TAGs) in gastrocnemius muscle. B Content of

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diacylglycerol (DAGs) in gastrocnemius muscle. Measurements were performed in the control and the collagen-induced arthritis (CIA) groups. Data are shown as the mean ± SE of 9-10 animals per group. *P<0.05 compared with the control rats, as assessed by t test.

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Author’s contribution All authors were involved in drafting the article or revising it critically for important intellectual content. All authors approved the final version of the article to be published. Contribution to study conception and design. Vial, Wauquier, Wittrant, Soubrier,

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Coxam,Tournadre, Capel.

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Contribution to the acquisition of data. Vial, Pinel, Wauquier, Coudy-Gandelon,

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Chevenet, Capel.

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Contribution to the analysis and interpretation of data. Vial, Bechet, Tournadre,

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Capel.

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Figure 1

Figure 2

Figure 3

Figure 4

Figure 5