Skeletal Muscle Regeneration and Oxidative Stress Are Altered ... - Plos

RESEARCH ARTICLE

Skeletal Muscle Regeneration and Oxidative Stress Are Altered in Chronic Kidney Disease Keith G. Avin1,2*, Neal X. Chen2, Jason M. Organ3, Chad Zarse2, Kalisha O’Neill2, Richard G. Conway4, Robert J. Konrad4, Robert L. Bacallao2, Matthew R. Allen3, Sharon M. Moe2 1 Department of Physical Therapy, Indiana University School of Health and Rehabilitation Sciences, Indianapolis, IN, United States of America, 2 Division of Nephrology, Indiana University School of Medicine, Indianapolis, IN, United States of America, 3 Department of Anatomy & Cell Biology, Indiana University School of Medicine, Indianapolis, IN, United States of America, 4 Lilly Research laboratories, Eli Lilly and Company, Indianapolis, IN, United States of America

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* [email protected]

Abstract

OPEN ACCESS Citation: Avin KG, Chen NX, Organ JM, Zarse C, O’Neill K, Conway RG, et al. (2016) Skeletal Muscle Regeneration and Oxidative Stress Are Altered in Chronic Kidney Disease. PLoS ONE 11(8): e0159411. doi:10.1371/journal.pone.0159411 Editor: Niels Olsen Saraiva Câmara, Universidade de Sao Paulo, BRAZIL Received: November 23, 2015 Accepted: July 2, 2016 Published: August 3, 2016 Copyright: © 2016 Avin et al. This is an open access article distributed under the terms of the Creative Commons Attribution License, which permits unrestricted use, distribution, and reproduction in any medium, provided the original author and source are credited.

Skeletal muscle atrophy and impaired muscle function are associated with lower healthrelated quality of life, and greater disability and mortality risk in those with chronic kidney disease (CKD). However, the pathogenesis of skeletal dysfunction in CKD is unknown. We used a slow progressing, naturally occurring, CKD rat model (Cy/+ rat) with hormonal abnormalities consistent with clinical presentations of CKD to study skeletal muscle signaling. The CKD rats demonstrated augmented skeletal muscle regeneration with higher activation and differentiation signals in muscle cells (i.e. lower Pax-7; higher MyoD and myogenin RNA expression). However, there was also higher expression of proteolytic markers (Atrogin-1 and MuRF-1) in CKD muscle relative to normal. CKD animals had higher indices of oxidative stress compared to normal, evident by elevated plasma levels of an oxidative stress marker, 8-hydroxy-2' -deoxyguanosine (8-OHdG), increased muscle expression of succinate dehydrogenase (SDH) and Nox4 and altered mitochondria morphology. Furthermore, we show significantly higher serum levels of myostatin and expression of myostatin in skeletal muscle of CKD animals compared to normal. Taken together, these data show aberrant regeneration and proteolytic signaling that is associated with oxidative stress and high levels of myostatin in the setting of CKD. These changes likely play a role in the compromised skeletal muscle function that exists in CKD.

Data Availability Statement: All relevant data are provided within the paper. Funding: This work was supported by the National Institutes of Health (R01AR058005), http://www.nih. gov/.

Introduction

Competing Interests: The authors have read the journal's policy and have the following competing interests: Authors RJK and RGC are employed by and own stock in Eli Lilly. SMM and MRA have received consulting fees from Eli Lilly for unrelated projects. Commercial affiliations do not alter the

Chronic kidney disease (CKD) is a progressive disease that leads to increased inflammation, increased concentrations of detrimental uremic toxins, augmented hormonal status and an impaired musculoskeletal system [1]. These musculoskeletal deficits contribute to a lower health-related quality of life, greater disability, and reduced physical activity associated with increased risk of mortality [2–4]. Physical deficits associated with CKD are due in part to both

PLOS ONE | DOI:10.1371/journal.pone.0159411 August 3, 2016

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Skeletal Muscle Dysfunction in CKD

authors’ adherence to PLOS ONE policies on sharing data and materials.

muscle loss (atrophy) and muscle weakness [5, 6]. Unfortunately, little is known about the cellular mechanisms underlying skeletal muscle changes in CKD. Muscle dysfunction in CKD may be accelerated by either increased catabolism, decreased protein synthesis or impaired regeneration. However, it is not clear which are the overriding factors that sway the balance between muscle production and loss in CKD. A number of studies and reviews postulate oxidative stress as a major contributor of muscle atrophy [7, 8]. Oxidative stress is the result of accumulated endogenous reactive oxygen species (ROS); ROS can amass from dysfunctional mitochondria and increased NADPH oxidases (NOX). Specifically, oxidative stress can lead to atrophy by activating autophagy pathways through forkhead transcription factor (FoxO) 3-mediated transcription factors, Atrogin-1 and muscle ring finger protein 1 (MuRF-1) [9, 10]. Oxidative stress may also affect skeletal muscle through the myostatin pathway [11]. Myostatin, (growth differentiation factor 8) regulates muscle atrophy via activation of proteolytic pathways and impaired muscle regeneration [12]. Muscle regeneration is an organized process that, in response to a harmful stimulus, activates quiescent muscle stem (satellite) cells to differentiate and form myotubes and subsequent myofibers [13]. Impaired regenerative processes and increased catabolism have been studied in mouse models of kidney injury [14]. However, it is not clear how these processes may be altered in a slow progressing, naturally occurring CKD model, which may better capture the progressive nature of human CKD. We have previously published data demonstrating that by 35 weeks of age, the Cy/+ (CKD) rat has developed progressive, significant azotemia, hyperphosphatemia, secondary hyperparathyroidism, and markedly elevated FGF23, which result in kidney function equal to approximately 15% of kidney function in the normal littermates (NLs) [15]. We recently published that CKD rats demonstrate significantly reduced muscle fiber cross sectional area indicative of atrophy and peak isometric torque during ankle dorsiflexion [16]. In the current study, we tested the hypothesis that in CKD there is increased oxidative stress and myostatin levels that together could explain altered skeletal muscle regeneration and catabolic signaling.

Methods Animal model and tissue harvest We used a naturally occurring rat model of Chronic Kidney Disease-Mineral Bone Disorder (CKD-MBD); the Cy/+ rat model (CKD rat) transmits cystic kidney disease as an autosomal dominant trait with slow progressing CKD due to a missense mutation in the gene Anks6 (samcystin) [17]. The CKD rat with slowly progressive azotemia results in terminal uremia by 40 weeks and development of all three manifestations of CKD-MBD (i.e. biochemical abnormalities, extraskeletal calcification, and abnormal bone) [15]. Weaned rats were housed in open top, shoebox cages, and had free access to tap water and standard chow until they were 24 weeks old when they were switched from a standard pellet rat chow to a diet of 18% caseinbased protein, 0.7% phosphorus, 0.7% calcium, 5% fat (Harlan Teklan TD.04539), until sacrifice, which leads to more reproducible phenotype [18]. Male Cy/+ rats (hereafter called CKD rat), and normal littermates (NL) rat; (n = 6–8 each group) were sacrificed at 35 weeks with pentobarbital (50 mg/kg intraperitoneal) and blood was collected for oxidative stress markers and myostatin assays. The extensor digitorum longus (EDL) was collected and stored at –80°C for RNA and protein isolation. To preserve the middle third of the left EDL for histology, the muscle was placed on a piece of corkboard, in optimal cutting temperature compound (OCT) and frozen in liquid nitrogen chilled 2-methylybutane for 45 seconds; then stored at -80°C until analysis. All procedures were reviewed and approved by the Indiana University School of Medicine Institutional Animal Care and Use Committee, which adheres to the Guide for the


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Ethical Treatment of Animals to minimize pain and suffering. (http://grants.nih.gov/grants/ olaw/Guide-for-the-Care-and-Use-of-Laboratory-Animals.pdf)

RNA isolation and real time PCR Total RNA from normal and CKD EDL was isolated using miRNeasy Mini Kit (Qiagen) according to the manufacturer’s instruction. Total RNA was eluted from the column in RNasefree water and stored at –80°C. The gene and miRNA expression was determined by real time PCR using TaqMan miRNA assays (Applied Biosystems, Foster City, CA). Target-specific PCR primers (Pax-7, MyoD, Myostatin, Myogenin, Atrogin 1, MuRF-1, miR-29b, Activin 2b, SOD1, and SOD-2) were obtained from Applied Biosystems. Real-time PCR amplifications were performed using TaqMan miRNA Assays (TaqMan MGP probes, FAM dye-labeled) using Applied Biosystems ViiA 7 Real-Time PCR systems (Applied Biosystems). The cycle number at which the amplification plot crosses the threshold was calculated (CT), and the ΔΔCT method was used to analyze the relative changes in gene expression and normalized by β-actin or U6 (RNA and miRNA, respectively).

Western blot Western blotting was performed as previously described [19]. In brief, the EDL from NL and CKD were homogenized and the total tissue protein lysates were stored at -20C. The expression of Nox4 was measured using antibody against Nox4 (1:300 dilution; Novus Biologicals, Littleton, CO). Nuclear and cytosolic protein was isolated using Cayman’s Nuclear Extraction kit (Cayman Chemical Company, Ann Arbor, MI) according to the manufacturer’s instructions. Nrf2 was measured in the nuclear fraction, and the major regulator of Nrf2, Keap1 (Kelch-like ECH-associated protein I) was measured in the cytosolic fraction with antibody against Nrf2 or Keap 1(1:1000, Santa Cruz Biotechnology, Santa Cruz, CA) overnight at 4°C followed by incubating with peroxidase conjugated secondary antibody (1:5000 dilution), and immunodetection with the Enhanced Chemiluminescence Prime Western Blot Detection Reagent (Amersham, Piscataway, NJ). The band intensity was analyzed by ChemiDoc MP Imaging System (Imaging Lab 4.0, Bio-Rad, Richmond, CA) and normalized to total protein expression using Ponceau S (Santa Cruz Biotechnology, Santa Cruz, CA).

Plasma oxidative stress assay and myostatin assay Plasma levels of an oxidative stress marker, 8-hydroxy-2' -deoxyguanosine (8-OHdG), were measured using a DNA damage ELISA kit (Enzo Life Sciences, Farmingdale, NY). Plasma myostatin levels were measured using a dual-monoclonal sandwich immunoassay developed by Eli Lilly and Company (Indianapolis, IN). Briefly, a myostatin ELISA was performed using Mesoscale Discovery (MSD) plates with streptavidin-coated and pre-blocked wells that were incubated for 1-hour with biotinylated anti-myostatin capture antibody. Afterward, wells were aspirated and washed three times with TBST (Tris buffered saline containing 50 mmol/L Tris pH 7.40, 150 mmol/L NaCl, with 0.5 mL Tween 20/L). Next, 100 μL of recombinant myostatin standards (varying concentrations of myostatin protein in assay buffer consisting of 50 mmol/ L HEPES, pH 7.40, 150 mmol/L NaCl, 10 mL/L Triton X-100, 5 mmol/L EDTA,5 mmol/L EGTA, and 0.1 mg/ml Heterophilic Blocking Reagent I (Scantibodies Laboratory Inc, Santee, CA)) were added to the wells to generate a standard calibration curve. Plasma samples were diluted in assay buffer and added to their respective wells, and plates were incubated for 1 hour at room temperature. Following aspiration, wells were washed 3 times with TBST, and 100 μL of conjugate antibody (ruthenium-labeled anti-myostatin detection antibody) were added to the wells for a 1-hour incubation at room temperature. Following aspiration, wells were washed


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3 times with TBST, and the plate was developed using a MSD reader, which recorded ruthenium electrochemiluminescence. MesoScale Discovery (MSD) software and SigmaPlot version 8.0 were used for fitting the ELISA calibration curves as well as for final determination of serum levels.

Transmission electron microscopy (TEM) A small section (