Skeletal muscle development 978-1-4939-7283-8, 1493972839, 978-1-4939-7282-1

Table of contents :
Front Matter ....Pages i-xii
Front Matter ....Pages 1-1
Bisulfite Sequencing for DNA Methylation Analysis of Primary Muscle Stem Cells (Kohei Miyata, Masashi Naito, Tomoko Miyata, Sho Mokuda, Hiroshi Asahara)....Pages 3-13
Whole Genome Chromatin IP-Sequencing (ChIP-Seq) in Skeletal Muscle Cells (Karl Kamhei So, Xianlu Laura Peng, Hao Sun, Huating Wang)....Pages 15-25
Analysis of RNA Expression in Adult Zebrafish Skeletal Muscle (Tamar E. Sztal, Peter D. Currie, Robert J. Bryson-Richardson)....Pages 27-35
Front Matter ....Pages 37-37
Targeted Lipidomic Analysis of Myoblasts by GC-MS and LC-MS/MS (Jordan Blondelle, Jean-Paul Pais de Barros, Fanny Pilot-Storck, Laurent Tiret)....Pages 39-60
Measuring Mitochondrial Substrate Utilization in Skeletal Muscle Stem Cells (C. Hai Ly, James G. Ryall)....Pages 61-73
Microcontact-Printed Hydrogel Microwell Arrays for Clonal Muscle Stem Cell Cultures (Victor M. Aguilar, Benjamin D. Cosgrove)....Pages 75-92
Isolation, Culture, and Differentiation of Fibro/Adipogenic Progenitors (FAPs) from Skeletal Muscle (Robert N. Judson, Marcela Low, Christine Eisner, Fabio M. Rossi)....Pages 93-103
Human Satellite Cell Isolation and Xenotransplantation (Steven M. Garcia, Stanley Tamaki, Xiaoti Xu, Jason H. Pomerantz)....Pages 105-123
Front Matter ....Pages 125-125
Application of Split-GFP Reassembly Assay to Study Myogenesis and Myofusion In Vitro (Manami Kodaka, Xiaoyin Xu, Zeyu Yang, Junichi Maruyama, Yutaka Hata)....Pages 127-134
Myogenic Maturation by Optical-Training in Cultured Skeletal Muscle Cells (Toshifumi Asano, Toru Ishizuka, Hiromu Yawo)....Pages 135-145
Fabrication of Micromolded Gelatin Hydrogels for Long-Term Culture of Aligned Skeletal Myotubes (Gio C. Suh, Archana Bettadapur, Jeffrey W. Santoso, Megan L. McCain)....Pages 147-163
Front Matter ....Pages 165-165
Quantification of Embryonic Myofiber Development by Immunofluorescence (Harika Nagandla, M. David Stewart)....Pages 167-176
How to Wire the Diaphragm: Wholemount Staining Methods to Analyze Mammalian Respiratory Innervation (Maximilian Michael Saller, Paolo Alberton, Andrea B. Huber, Rosa-Eva Huettl)....Pages 177-192
Front Matter ....Pages 193-193
Membrane Repair Assay for Human Skeletal Muscle Cells (Romain Carmeille, Coralie Croissant, Flora Bouvet, Anthony Bouter)....Pages 195-207
Cryoinjury Model for Tissue Injury and Repair in Bioengineered Human Striated Muscle (Richard J. Mills, Holly K. Voges, Enzo R. Porrello, James E. Hudson)....Pages 209-224
Back Matter ....Pages 225-226

Citation preview

Methods in Molecular Biology 1668

James G. Ryall Editor

Skeletal Muscle Development

METHODS

IN

MOLECULAR BIOLOGY

Series Editor John M. Walker School of Life and Medical Sciences University of Hertfordshire Hatfield, Hertfordshire, AL10 9AB, UK

For further volumes: http://www.springer.com/series/7651

Skeletal Muscle Development Edited by

James G. Ryall Department of Physiology, The University of Melbourne, Parkville, VIC, Australia

Editor James G. Ryall Department of Physiology The University of Melbourne Parkville, VIC, Australia

ISSN 1064-3745 ISSN 1940-6029 (electronic) Methods in Molecular Biology ISBN 978-1-4939-7282-1 ISBN 978-1-4939-7283-8 (eBook) DOI 10.1007/978-1-4939-7283-8 Library of Congress Control Number: 2017950877 © Springer Science+Business Media LLC 2017 This work is subject to copyright. All rights are reserved by the Publisher, whether the whole or part of the material is concerned, specifically the rights of translation, reprinting, reuse of illustrations, recitation, broadcasting, reproduction on microfilms or in any other physical way, and transmission or information storage and retrieval, electronic adaptation, computer software, or by similar or dissimilar methodology now known or hereafter developed. The use of general descriptive names, registered names, trademarks, service marks, etc. in this publication does not imply, even in the absence of a specific statement, that such names are exempt from the relevant protective laws and regulations and therefore free for general use. The publisher, the authors and the editors are safe to assume that the advice and information in this book are believed to be true and accurate at the date of publication. Neither the publisher nor the authors or the editors give a warranty, express or implied, with respect to the material contained herein or for any errors or omissions that may have been made. The publisher remains neutral with regard to jurisdictional claims in published maps and institutional affiliations. Printed on acid-free paper This Humana Press imprint is published by Springer Nature The registered company is Springer Science+Business Media LLC The registered company address is: 233 Spring Street, New York, NY 10013, U.S.A.

Preface Skeletal muscle development (myogenesis) is a highly complex, yet carefully regulated process. The important role played by the myogenic regulatory factor (MRF) family of transcription factors (Myf5, MyoD, MyoG, and MRF4) and the paired homeobox proteins (Pax3/7) in regulating the processes of muscle cell progenitor proliferation and differentiation into mature muscle has been studied extensively. The Skeletal Muscle Development volume in the Methods in Molecular Biology series aims to present a wide range of techniques that go beyond the standard assays typically used to assess myogenesis, including assays to analyze skeletal muscle gene expression, proliferating muscle cells, the process of differentiation, muscle development (in vivo), and muscle repair. Parkville, VIC, Australia

James G. Ryall

v

Contents Preface . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Contributors. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

PART I

SKELETAL MUSCLE GENE EXPRESSION

1 Bisulfite Sequencing for DNA Methylation Analysis of Primary Muscle Stem Cells . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Kohei Miyata, Masashi Naito, Tomoko Miyata, Sho Mokuda, and Hiroshi Asahara 2 Whole Genome Chromatin IP-Sequencing (ChIP-Seq) in Skeletal Muscle Cells . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Karl Kamhei So, Xianlu Laura Peng, Hao Sun, and Huating Wang 3 Analysis of RNA Expression in Adult Zebrafish Skeletal Muscle. . . . . . . . . . . . . . . Tamar E. Sztal, Peter D. Currie, and Robert J. Bryson-Richardson

PART II

v ix

3

15

27

ASSAYS FOR PROLIFERATING SKELETAL MUSCLE CELLS

4 Targeted Lipidomic Analysis of Myoblasts by GC-MS and LC-MS/MS . . . . . . . 39 Jordan Blondelle, Jean-Paul Pais de Barros, Fanny Pilot-Storck, and Laurent Tiret 5 Measuring Mitochondrial Substrate Utilization in Skeletal Muscle Stem Cells . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 61 C. Hai Ly and James G. Ryall 6 Microcontact-Printed Hydrogel Microwell Arrays for Clonal Muscle Stem Cell Cultures . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 75 Victor M. Aguilar and Benjamin D. Cosgrove 7 Isolation, Culture, and Differentiation of Fibro/Adipogenic Progenitors (FAPs) from Skeletal Muscle . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 93 Robert N. Judson, Marcela Low, Christine Eisner, and Fabio M. Rossi 8 Human Satellite Cell Isolation and Xenotransplantation . . . . . . . . . . . . . . . . . . . . . 105 Steven M. Garcia, Stanley Tamaki, Xiaoti Xu, and Jason H. Pomerantz

PART III

ASSESSMENT OF MYOGENIC DIFFERENTIATION

9 Application of Split-GFP Reassembly Assay to Study Myogenesis and Myofusion In Vitro. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 127 Manami Kodaka, Xiaoyin Xu, Zeyu Yang, Junichi Maruyama, and Yutaka Hata 10 Myogenic Maturation by Optical-Training in Cultured Skeletal Muscle Cells . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 135 Toshifumi Asano, Toru Ishizuka, and Hiromu Yawo

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Contents

Fabrication of Micromolded Gelatin Hydrogels for Long-Term Culture of Aligned Skeletal Myotubes . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 147 Gio C. Suh, Archana Bettadapur, Jeffrey W. Santoso, and Megan L. McCain

PART IV ASSESSING SKELETAL MUSCLE DEVELOPMENT IN VIVO 12

13

Quantification of Embryonic Myofiber Development by Immunofluorescence . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 167 Harika Nagandla and M. David Stewart How to Wire the Diaphragm: Wholemount Staining Methods to Analyze Mammalian Respiratory Innervation . . . . . . . . . . . . . . . . . . . . . . . . . . . . 177 Maximilian Michael Saller, Paolo Alberton, Andrea B. Huber, and Rosa-Eva Huettl

PART V 14

15

MUSCLE REPAIR ASSAYS

Membrane Repair Assay for Human Skeletal Muscle Cells . . . . . . . . . . . . . . . . . . . 195 Romain Carmeille, Coralie Croissant, Flora Bouvet, and Anthony Bouter Cryoinjury Model for Tissue Injury and Repair in Bioengineered Human Striated Muscle . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 209 Richard J. Mills, Holly K. Voges, Enzo R. Porrello, and James E. Hudson

Index . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

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Contributors VICTOR M. AGUILAR  Meinig School of Biomedical Engineering, Cornell University, Ithaca, NY, USA PAOLO ALBERTON  Experimental Surgery and Regenerative Medicine (ExperiMed), Department of General, Trauma and Reconstructive Surgery, Ludwig-MaximiliansUniversity (LMU), Munich, Germany HIROSHI ASAHARA  Department of Molecular and Experimental Medicine, The Scripps Research Institute, La Jolla, CA, USA; Department of Systems Bio Medicine, Tokyo Medical and Dental University, Tokyo, Japan TOSHIFUMI ASANO  Department of Cell Biology, Graduate School of Medical and Dental Sciences, Tokyo Medical and Dental University (TMDU), Tokyo, Japan JEAN-PAUL PAIS DE BARROS  Plateforme de Lipidomique-uBourgogne, INSERM UMR1231/LabEx LipSTIC, UFR des Sciences de Sante´ - Baˆtiment B3, Dijon, France ARCHANA BETTADAPUR  Laboratory for Living Systems Engineering, Department of Biomedical Engineering, USC Viterbi School of Engineering, University of Southern California, Los Angeles, CA, USA JORDAN BLONDELLE  Institut Mondor de Recherche Biome´dicale (IMRB), U955-E10 Biologie du Syste`me Neuromusculaire, Universite´ Paris-Est, Ecole Nationale Ve´te´rinaire d’Alfort (EnvA), Maisons-Alfort, France; Department of Cardiology, University of California, San Diego, La Jolla, CA, USA ANTHONY BOUTER  Institute of Chemistry and Biology of Membranes and Nano-objects, UMR 5248, CNRS, University of Bordeaux, Pessac, France FLORA BOUVET  Institute of Chemistry and Biology of Membranes and Nano-objects, UMR 5248, CNRS, University of Bordeaux, Pessac, France ROBERT J. BRYSON-RICHARDSON  School of Biological Sciences, Monash University, Melbourne, VIC, Australia ROMAIN CARMEILLE  Institute of Chemistry and Biology of Membranes and Nano-objects, UMR 5248, CNRS, University of Bordeaux, Pessac, France BENJAMIN D. COSGROVE  Meinig School of Biomedical Engineering, Cornell University, Ithaca, NY, USA CORALIE CROISSANT  Institute of Chemistry and Biology of Membranes and Nano-objects, UMR 5248, CNRS, University of Bordeaux, Pessac, France PETER D. CURRIE  Australian Regenerative Medicine Institute, Monash University, Melbourne, VIC, Australia CHRISTINE EISNER  Biomedical Research Centre, University of British Columbia, Vancouver, BC, Canada STEVEN M. GARCIA  Program in Craniofacial Biology, Division of Plastic and Reconstructive Surgery, Department of Surgery, Eli and Edythe Broad Center of Regeneration Medicine, University of California, San Francisco, San Francisco, CA, USA; Program in Craniofacial Biology, Division of Plastic and Reconstructive Surgery, Department of Orofacial Sciences, Eli and Edythe Broad Center of Regeneration Medicine, University of California, San Francisco, San Francisco, CA, USA

ix

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Contributors

YUTAKA HATA  Department of Medical Biochemistry, Graduate School of Medical and Dental Sciences, Tokyo Medical and Dental University, Tokyo, Japan; Center for Brain Integration Research, Tokyo Medical and Dental University, Tokyo, Japan ANDREA B. HUBER  Department of Biosystems Science and Engineering (D-BSSE), Swiss Federal Institute of Technology, Zurich (ETHZ), Basel, Switzerland JAMES E. HUDSON  Cardiac Regeneration Laboratory, School of Biomedical Sciences, The University of Queensland, St Lucia, QLD, Australia ROSA-EVA HUETTL  Department of Stress Neurobiology and Neurogenetics, Max-Planck-Institute of Psychiatry, Munich, Germany TORU ISHIZUKA  Department of Developmental Biology and Neuroscience, Tohoku University Graduate School of Life Sciences, Sendai, Japan ROBERT N. JUDSON  Biomedical Research Centre, University of British Columbia, Vancouver, BC, Canada; STEMCELL Technologies Inc., Vancouver, BC, Canada MANAMI KODAKA  Department of Medical Biochemistry, Graduate School of Medical and Dental Sciences, Tokyo Medical and Dental University, Tokyo, Japan MARCELA LOW  Biomedical Research Centre, University of British Columbia, Vancouver, BC, Canada C. HAI LY  Stem Cell Metabolism and Regenerative Medicine Group, Basic and Clinical Myology Laboratory, The University of Melbourne, Parkville, VIC, Australia JUNICHI MARUYAMA  Department of Medical Biochemistry, Graduate School of Medical and Dental Sciences, Tokyo Medical and Dental University, Tokyo, Japan MEGAN L. MCCAIN  Laboratory for Living Systems Engineering, Department of Biomedical Engineering, USC Viterbi School of Engineering, University of Southern California, Los Angeles, CA, USA; Department of Stem Cell Biology and Regenerative Medicine, Keck School of Medicine of USC, University of Southern California, Los Angeles, CA, USA RICHARD J. MILLS  Cardiac Regeneration Laboratory, School of Biomedical Sciences, The University of Queensland, St Lucia, QLD, Australia KOHEI MIYATA  Department of Molecular and Experimental Medicine, The Scripps Research Institute, La Jolla, CA, USA TOMOKO MIYATA  Department of Molecular and Experimental Medicine, The Scripps Research Institute, La Jolla, CA, USA SHO MOKUDA  Department of Molecular and Experimental Medicine, The Scripps Research Institute, La Jolla, CA, USA HARIKA NAGANDLA  Department of Molecular and Cellular Biology, Baylor College of Medicine, Houston, TX, USA MASASHI NAITO  Department of Systems Bio Medicine, Tokyo Medical and Dental University, Tokyo, Japan XIANLU LAURA PENG  Department of Chemical Pathology, The Chinese University of Hong Kong, Hong Kong, China; Li Ka Shing Institute of Health Sciences, The Chinese University of Hong Kong, Hong Kong, China FANNY PILOT-STORCK  Institut Mondor de Recherche Biome´dicale (IMRB), U955-E10 Biologie du Syste`me Neuromusculaire, Universite´ Paris-Est, Ecole Nationale Ve´te´rinaire d’Alfort (EnvA), Maisons-Alfort, France JASON H. POMERANTZ  Program in Craniofacial Biology, Division of Plastic and Reconstructive Surgery, Department of Surgery, Eli and Edythe Broad Center of Regeneration Medicine, University of California, San Francisco, San Francisco, CA, USA; Program in Craniofacial Biology, Division of Plastic and Reconstructive Surgery,

Contributors

xi

Department of Orofacial Sciences, Eli and Edythe Broad Center of Regeneration Medicine, University of California, San Francisco, San Francisco, CA, USA ENZO R. PORRELLO  Cardiac Regeneration Laboratory, School of Biomedical Sciences, The University of Queensland, St Lucia, QLD, Australia FABIO M. ROSSI  Biomedical Research Centre, University of British Columbia, Vancouver, BC, Canada JAMES G. RYALL  Stem Cell Metabolism and Regenerative Medicine Group, Basic and Clinical Myology Laboratory, Department of Physiology, School of Biomedical Sciences, The University of Melbourne, Parkville, VIC, Australia MAXIMILIAN MICHAEL SALLER  Experimental Surgery and Regenerative Medicine (ExperiMed), Department of General, Trauma and Reconstructive Surgery, Ludwig-Maximilians-University (LMU), Munich, Germany JEFFREY W. SANTOSO  Laboratory for Living Systems Engineering, Department of Biomedical Engineering, USC Viterbi School of Engineering, University of Southern California, Los Angeles, CA, USA KARL KAMHEI SO  Department of Chemical Pathology, The Chinese University of Hong Kong, Hong Kong, China; Li Ka Shing Institute of Health Sciences, The Chinese University of Hong Kong, Hong Kong, China M. DAVID STEWART  Department of Biology and Biochemistry, University of Houston, Houston, TX, USA; Department of Stem Cell Engineering, Texas Heart Institute at St. Luke’s Episcopal Hospital, Houston, TX, USA GIO C. SUH  Laboratory for Living Systems Engineering, Department of Biomedical Engineering, USC Viterbi School of Engineering, University of Southern California, Los Angeles, CA, USA; Department of Stem Cell Biology and Regenerative Medicine, Keck School of Medicine of USC, University of Southern California, Los Angeles, CA, USA HAO SUN  Department of Chemical Pathology, The Chinese University of Hong Kong, Hong Kong, China; Li Ka Shing Institute of Health Sciences, The Chinese University of Hong Kong, Hong Kong, China TAMAR E. SZTAL  School of Biological Sciences, Monash University, Melbourne, VIC, Australia STANLEY TAMAKI  Program in Craniofacial Biology, Division of Plastic and Reconstructive Surgery, Department of Surgery, Eli and Edythe Broad Center of Regeneration Medicine, University of California, San Francisco, San Francisco, CA, USA; Program in Craniofacial Biology, Division of Plastic and Reconstructive Surgery, Department of Orofacial Sciences, Eli and Edythe Broad Center of Regeneration Medicine, University of California, San Francisco, San Francisco, CA, USA LAURENT TIRET  Institut Mondor de Recherche Biome´dicale (IMRB), U955-E10 Biologie du Syste`me Neuromusculaire, Universite´ Paris-Est, Ecole Nationale Ve´te´rinaire d’Alfort (EnvA), Maisons-Alfort, France HOLLY K. VOGES  Cardiac Regeneration Laboratory, School of Biomedical Sciences, The University of Queensland, St Lucia, QLD, Australia HUATING WANG  Li Ka Shing Institute of Health Sciences, The Chinese University of Hong Kong, Hong Kong, China; Department of Orthopaedics and Traumatology, The Chinese University of Hong Kong, Hong Kong, China

xii

Contributors

XIAOTI XU  Program in Craniofacial Biology, Division of Plastic and Reconstructive Surgery, Department of Surgery, Eli and Edythe Broad Center of Regeneration Medicine, University of California, San Francisco, San Francisco, CA, USA; Program in Craniofacial Biology, Division of Plastic and Reconstructive Surgery, Department of Orofacial Sciences, Eli and Edythe Broad Center of Regeneration Medicine, University of California, San Francisco, San Francisco, CA, USA XIAOYIN XU  Department of Medical Biochemistry, Graduate School of Medical and Dental Sciences, Tokyo Medical and Dental University, Tokyo, Japan; Department of Breast Surgery, The Second Affiliated Hospital of Wenzhou Medical University, Wenzhou, China ZEYU YANG  Department of Ultrasound, Shengjing Hospital of China Medical University, Shenyang, China HIROMU YAWO  Department of Developmental Biology and Neuroscience, Tohoku University Graduate School of Life Sciences, Sendai, Japan

Part I Skeletal Muscle Gene Expression

Chapter 1 Bisulfite Sequencing for DNA Methylation Analysis of Primary Muscle Stem Cells Kohei Miyata, Masashi Naito, Tomoko Miyata, Sho Mokuda, and Hiroshi Asahara Abstract In skeletal muscle, DNA methylation contributes to the suppression of gene expression in several biological processes and diseases. A protocol for the detection of methylated cytosine was thus established based on methylation-sensitive enzymes, immunoprecipitation, and bisulfite conversion. DNA methylation analysis, with bisulfite conversion and sequencing, enables the quantification of methylation at each single base position. Here, we describe a basic method of bisulfite sequencing that can be used to analyze local DNA methylation status to confirm genome-wide DNA methylation analysis or correlation of gene expression regulatory mechanisms. Key words Epigenetics, DNA methylation, Methylcytosine, Bisulfite conversion, Bisulfite sequencing analysis

1

Introduction To date, there are three established strategies to detect DNA methylation [1]. Methylation-sensitive enzymatic digestion was a simple way to define methylated DNA, but it is no longer widely used because of its sequence restriction and low working efficacy. Realtime PCR, following digestion with methylation-specific enzymes, enables the quantification of the methylation status of a few methylated-CpG regions; however, it cannot define the specific methylated CpG position. Several recent reports have combined real-time PCR with DNA microarray (or sequencing) technologies to perform genome-wide screening or profiling of DNA methylation; however, this enzymatic strategy is not compatible with highthroughput analysis [2]. On the other hand, combining highthroughput approaches with immunoprecipitation, which captures sheared DNA fragments around the methylated cytosine using antimethyl-CpG binding protein (MeCP2) or anti-5-methyl cytosine antibodies, could achieve the desired result. However, each of these

James G. Ryall (ed.), Skeletal Muscle Development, Methods in Molecular Biology, vol. 1668, DOI 10.1007/978-1-4939-7283-8_1, © Springer Science+Business Media LLC 2017

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Kohei Miyata et al.

CH3 5’ 3’

CG GC

CG GC

GC ACCT CGTGGA

3’ 5’

CH3 CH3 5’

CG

UG

G U AUU T

3’

3’

GC

GU

UGTGGA

5’

CH3 5’ 3’

CG GC

TG AC

G TAT T T C ATA A A

5’ 3’

CG GC

CA GT

3’ 5’ AC AC C T TGTGG A

3’ 5’

Cycle sequence NGS COBRA

Fig. 1 Methylation analysis based on Bisulfite conversion

approaches remains limited by an inability to identify the exact quantity or position of methylated cytosine. Bisulfite conversion is regarded as the gold standard in defining DNA methylation status. Treatment of DNA with sodium bisulfite converts unmethylated cytosine to uracil, which is subsequently converted to thymine during PCR amplification, while methylcytosine remains unchanged (Fig. 1) [3, 4]. In this regard, several commercial kits have proven useful and show high conversion rates. Leontiou et al. examined the performance of four such kits and accordingly ranked them by performance [5]. The MethylEdge Bisulfite Conversion System (Promega) emerged as the recommended kit to use due to the results obtained for bisulfite amplicon Sanger sequencing, but no significant difference in conversion rate was observed [5]. Coupling bisulfite conversion with DNA sequence analysis enables the qualitative and quantitative detection of methylcytosine at a single base-pair resolution. Following bisulfite conversion, DNA is analyzed with next-generation transcriptomics to validate the genome-wide DNA methylation profile, and the regional methylation status established by Sanger sequencing. DNA methylation status can also be analyzed by treating bisulfite-converted DNA

Bisulfite Sequencing

5

with restriction enzymes, more commonly referred to as Combined Bisulfite Restriction Analysis (COBRA). Digestion of PCRamplified DNA with BstUI (which digests the sequence “CGCG”), followed by an analysis of DNA fragmentation with electrophoresis, is used to evaluate the approximate rate of cytosine methylation [6]. Using primary muscle progenitor cells, referred to as satellite cells (SCs), we demonstrate an example of the bisulfite sequencing method based on Sanger sequencing. Bisulfite-converted SC DNA was PCR amplified and subcloned into a cloning vector. Sanger sequence analysis was subsequently used to show the DNA methylation status at each single base position. The procedure contains DNA preparation (Subheading 3.1), bisulfite conversion (Subheading 3.2), PCR and subcloning (Subheading 3.3), and colonydirect sequencing (Subheading 3.4). Each bisulfite sequencing step does not contain any unique technique; thus, numerous alternative methods or protocols can be substituted. The isolation and culture of muscle SCs is described in “Note 1.”

2 2.1

Materials DNA Preparation

1. AllPrep DNA/RNA Micro Kit (Qiagen). 2. AccuBlue High Sensitivity dsDNA Quantitation Kit (Biotium). 3. β-mercaptoethanol, ethanol, TE buffer. 4. Fluorometer.

2.2 Bisulfite Conversion

1. EpiTect Plus DNA Bisulfite Kit (Qiagen). 2. Thermal cycler. 3. Ethanol.

2.3 Primer Design and PCR

1. GO Taq master mix (Promega). 2. PCR primer. 3. Plasmid with analyzing sequence. 4. Agarose, TAE buffer. 5. Thermal cycler.

2.4 Subcloning and Sequencing

1. pGEM-T Easy Vector System (Promega). 2. LB agar plate, with Ampicillin. 3. Competent cells, SOC medium. 4. 2% X-gal, dilute with dimethylformamide (DMF). 5. GO Taq master mix (Promega). 6. M13 forward (
20) primer, M13 reverse primer.

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7. ExoSAP-IT For PCR Product Clean-Up (Affymetrix). 8. BigDye Terminator v3.1 Cycle sequencing Kit (Thermo Fisher Scientific). 9. T7 primer. 10. Water bath (42  C), Incubator (37  C). 11. ABI PRISM 3100 (Applied Biosystems).

3

Methods DNA Preparation

To validate the gene expression profile of the harvested sample, extracting genomic DNA and total RNA from the same sample is highly recommended. For satellite cell culture conditions and isolation, see Note 1. All the centrifuge steps in this section were performed at room temperature.

3.1.1 Cell Collection and Lysis

1. Add 10 μL of β-mercaptoethanol to 1 mL of Buffer RLT Plus.

3.1

2. Discard cell culture medium and add 350 μL of Buffer RLT Plus directly to cells on culture dish. 3. Scrape the cells with chip, and harvest the entire cell suspension into a 1.5 mL tube. 4. Pass the lysate a minimum of five times through a blunt 20gauge needle fitted to an RNase-free syringe. 5. Transfer the homogenized lysate to an AllPrep DNA spin column on a 2 mL collection tube and centrifuge for 30 s at 10,000 rpm ( 8,000  g). 6. Use the flow-through for RNA purification and store the column at 4  C on a new 2 mL collection tube for DNA purification later.

3.1.2 Extraction of DNA and RNA

1. Add 350 μL of 70% ethanol to the flow-through and mix well by pipetting. 2. Transfer the mixture into an RNeasy MinElute spin column. 3. Centrifuge for 15 s at 10,000 rpm ( 8,000  g). Discard the flow-through. 4. Add 700 μL of Buffer RW1 to the column. Centrifuge for 15 s at 10,000 rpm ( 8,000  g). Discard the flow-through. 5. Add 500 μL of Buffer RPE to the column. Centrifuge for 15 s at 10,000 rpm ( 8,000  g). Discard the flow-through. 6. Add 500 μL of 80% ethanol to the column. Centrifuge for 15 s at 10,000 rpm ( 8,000  g). Discard the flow-through. 7. Centrifuge for 5 min at 10,000 rpm ( 8,000  g) to completely remove excess buffer or ethanol.

Bisulfite Sequencing

7

8. Place the column in a new 1.5 mL collection tube and add 14 μL RNase-free water directly to the membrane. 9. Centrifuge for 1 min at 10,000 rpm ( 8,000  g) to elute the RNA. 10. Add 500 μL of Buffer AW1 to the AllPrep DNA spin column. Centrifuge for 15 s at 10,000 rpm ( 8,000  g). Discard the flow-through. 11. Add 500 μL of Buffer AW2 to the column. Centrifuge for 15 s at 14,000 rpm. Discard the flow-through. 12. Centrifuge for 2 min at 14,000 rpm (20,000  g) to remove buffer or ethanol completely. 13. Place the column in a new 1.5 mL collection tube and add 50 μL of EB buffer directly to the membrane. 14. Centrifuge for 1 min at 10,000 rpm ( 8,000  g) to elute the DNA. 15. Add the eluted DNA into the same column again. Centrifuge for 1 min at 10,000 rpm ( 8,000  g) to elute the DNA. 3.1.3 DNA Quantification

Measuring genomic DNA concentration was performed using a fluorescence-based method owing to its high sensitivity. In this section, we describe the method used for the AccuBlue High Sensitivity dsDNA Quantitation Kit (Biotium). 1. Warm the Quantification and 100 Enhancer solutions to room temperature and invert gently. 2. Dilute the sample 1:10 by adding 2 μL of sample to 18 μL of TE buffer. 3. Prepare the dsDNA concentration standards in TE buffer at 10, 8, 4, 2, 1, and 0.5 ng/μL. 4. To prepare the Working solution, add 2 μL of 100 Enhancer solution to 198 μL of Quantification solution in each well. 5. Add 10 μL of standards and samples into each well, and mix well by pipetting up and down. 6. Incubate at room temperature for 10 min in the dark. 7. Measure fluorescence using a microplate reader set to 485 nm excitation/530 nm emission maxima or other filter combination for detecting green fluorescence (e.g., FITC filter set).

3.2 Bisulfite Conversion

For bisulfite conversion, an EpiTect Plus DNA Bisulfite Kit was used. Methylated mouse genomic DNA as a control for bisulfite conversion can be purchased from several sources; however, unmethylated mouse genomic DNA is not available commercially. In place of an unmethylated DNA control, the plasmid DNA used to subclone the target DNA sequence was used as a positive control for bisulfite conversion. In the preparation for Subheading 3.2.2,

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30 mL of 100% ethanol was added to Buffer BW and 27 mL of 100% ethanol was added to Buffer BD. Buffer BD and the MinElute DNA spin column are stored at 4  C. Based on the cell culture size, low genomic DNA yields were acquired from in vitro cultured SCs in the 8-well chamber slide. A single bisulfite reaction typically made use of 300 ng of genomic DNA. 3.2.1 Bisulfite Reaction

1. Dissolve the Bisulfite Mix in 800 μL of RNase-free water, vortex, and warm to 60  C until completely dissolved. 2. Prepare the bisulfite reactions in 200 μL PCR tubes as below: DNA sample (20 ng/μL)

15 μL

RNase free water

25 μL

Bisulfite mix

85 μL

DNA protect buffer

15 μL

Total volume

140 μL

3. Perform Bisulfite reaction by thermal cycler using the following protocol: Denature

95  C

5 min

Incubation

60  C

25 min

Denaturation

95  C

5 min

Incubation

3.2.2 Purify Single Strand DNA (ssDNA)



85 min



60 C

Denaturation

95 C

5 min

Incubation

60  C

175 min

Hold

4 C

1. Transfer the bisulfite reactions into 1.5 mL tubes. 2. Add 310 μL of Buffer BL to each sample. Vortex the solutions and spin down. 3. Add 250 μL of 100% ethanol to each sample. Vortex the solutions for 15 s and spin down. 4. Transfer the entire mixture into the MinElute DNA spin column. 5. Centrifuge the spin columns at 10,000 rpm ( 8,000  g) for 1 min and discard the flow-through. 6. Add 500 μL of Buffer BW to each column and centrifuge at 10,000 rpm ( 8,000  g) for 1 min. Discard the flow-through. 7. Add 500 μL of Buffer BD to each column. Close the lid completely and incubate for 15 min at room temperature. 8. Centrifuge at 10,000 rpm ( 8,000  g) for 1 min. Discard the flow-through.

Bisulfite Sequencing

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9. Add 500 μL of Buffer BW to each column and centrifuge at 10,000 rpm ( 8,000  g) for 1 min. Discard the flowthrough. 10. Repeat step 9. 11. Add 250 μL of 100% ethanol to each column and centrifuge at 14,000 rpm (20,000  g) for 1 min. Discard the flow-through. 12. Place the columns into new 2 mL collection tubes and centrifuge at 14,000 rpm (20,000  g) for 1 min to remove any residual liquid. 13. Place the spin columns with open lids into clean 1.5-mL microcentrifuge tubes and incubate at 60  C for 5 min in a heating block. 14. Place the spin columns into new 1.5 mL tubes. 15. Add 15 μL of Buffer EB directly onto the center of each spincolumn membrane. 16. Incubate the columns at room temperature for 1 min. 17. Centrifuge for 1 min at 10,000 rpm ( 8,000  g) to elute the ssDNA. 3.3 PCR and Subcloning

3.3.1 PCR

For the bisulfite sequence, primer design is most important to obtain reliable data. Since bisulfite-converted DNA was fragmented and the cytosine converted to thymine (except within CpG sites), the following tips are recommended: (1) amplicon should be short (around 100 bp); (2) CpG must be avoided; and (3) non-CpG cytosine should be contained. MethPrimer (http://www.urogene.org/ methprimer/) is widely known as a useful and convenient web tool to design primers for DNA methylation analysis (see Notes 2–4) [7]. High-fidelity enzymes are also appropriate to amplify for methylation analysis, as PCR errors can distort the results. However, high-fidelity enzymes based on Pfu polymerase failed to amplify the bisulfiteconverted DNA in many cases. To overcome this, the analyzed sequence was amplified by PCR and subcloned into a TA-vector (see Notes 5 and 6). 1. Prepare the PCR mixture (total volume 20 μL) as below: GO Taq master mix

10 μL

5 μM forward primer

1 μL

5 μM reverse primer

1 μL

Water

6 μL

Bisulfite-converted DNA

2 μL

2. Perform the PCR on a thermal cycler using the following protocol:

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Kohei Miyata et al. Denature

95  C 

2 min

*Denature

95 C

30 s

*Annealing

55–60  C

30 s

*Extension Elongation Hold



30 s (repeat * for 30–35 cycles)



5 min

72 C 72 C 

4 C

3. Check amplicon length by 2.5% agarose gel electrophoresis and estimate amplicon yield (see Note 6). 3.3.2 TA-Cloning

1. For the TA-cloning, using the pGEM-T Easy vector, the reaction conditions are as follows; Rapid ligation buffer

5 μL

pGEM-T easy vector

1 μL (50 ng)

PCR product

1–1.5 μL

T7 DNA ligase

1 μL

Water

1.5–2 μL

2. Incubate at 4  C overnight. 3. Add 2.5 μL of ligation product into 25 μL of DH5α competent cells and incubate on ice for 30 min. 4. Incubate at 42  C for 45 s followed by incubation on ice for 2 min. 5. Add 250 μL of SOC medium to competent cells; incubate at 37  C with shaking for 1 h. 6. Spread 50 μL of 2% X-gal on a dried LB agar plate containing ampicillin and dry plates again in the incubator for 30 min. 7. Plate 50 μL of cultured transformed cells on the LB-plates and incubate overnight. 3.4 Colony PCR and Direct Sequencing

The pGEM-T Easy vector cloning site is located between the T7 and SP6 promoter sites, with the M13 forward and reverse sequencing primer sites located outside of these. Colony PCR was therefore performed using the M13 forward and reverse primers, and the sequence reaction cycle was undertaken using T7 primers (see Note 7). 1. Prepare the colony PCR mixture (total volume 10 μL) as shown below:

Bisulfite Sequencing

11

GO Taq master mix

5 μL

5 μM M13 forward primer

0.5 μL

5 μM M13 reverse primer

0.5 μL

Water

4 μL

2. Perform the PCR on a thermal cycler using the following conditions: Denature *Denature

95  C

2 min



30 s



95 C

*Annealing

58 C

30 s

*Extension

72  C

30 s (repeat * for 25 cycles)

Elongation Hold



72 C

5 min



4 C

3. Load 2 μL of the PCR product to confirm amplicon length by 2.5% agarose gel electrophoresis. 4. Add 1 μL of ExoSAP-IT into the remaining PCR product. 5. Incubate at 37  C for 45 min, followed by an inactivation step at 85  C for 15 min. Thereafter store at 4  C. 6. Prepare the cycle sequencing reaction mixture as follows: ABI PRISM BigDye terminator

2 μL

5 BigDye terminator buffer

2 μL

T7 primer (1.6 pmol)

2 μL

PCR product

1 μL

Water

3 μL

7. Perform the cycle sequencing reaction using a thermal cycler set to the following protocol; Denature

96  C

3 min

*Denature

96  C

30 s

*Annealing



10 s



4 min (repeat * for 25 cycles)

50 C

*Extension

60 C

Hold

4 C

8. Add 2.5 μL of 125 mM EDTA and 30 μL of 100% ethanol to each well and seal the plate. 9. Vortex and incubate at room temperature for 15 min. 10. Centrifuge the plate at 4,000 rpm (1,650  g), 4  C for 30 min.

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11. Remove the seal, invert the plate, and centrifuge at 300 rpm for 1 min. 12. Add 30 μL of 70% ethanol to each well. 13. Reseal the plate and centrifuge at 4,000 rpm (1,650  g), 4  C for 5 min. 14. Remove the seal, invert the plate, and centrifuge at 300 rpm for 1 min. 15. Add 10 μL of Hi-Di Formamide to each well. 16. Reseal the plate, spin down, and incubate at 95  C for 2 min. 17. Place the reaction plate on ice in the dark until analysis. 18. Analyze with a Genetic analyzer (see Note 8).

4

Notes 1. In our previous study, muscle SCs were isolated according to a protocol based on extracting a single myofiber and culturing on collagen type I-coated dishes established by Dr. Naohiro Hashimoto [89]. We have also isolated the muscle SCs by a protocol based on cell sorting [10]. Muscle SCs were isolated as Ter119
/CD45
/CD31
/CD34þ/α7-integrinþ/Sca-1 cells. Isolated SCs were seeded on laminin-coated plates. For differentiation analysis, 26,000 cells/well were seeded on each well of a laminin-coated 8 well Nunc Lab-Teck Chamber slide, and cultured with Growth Medium (DMEM supplemented with 3% FCS þ 7% FBS). Cells were subsequently switched to Differentiation medium (DMEM supplemented with 2% horse serum) once the cells were grown up to 80% confluence [11]. 2. To generate the code of the bisulfite-converted sequence, Microsoft Word proved convenient. First, convert “CG” to “XX,” and then convert all “C” to “T.” This aids the design and visualization of the PCR primers as well as the position of the CpG sites. 3. A high-fidelity PCR enzyme is recommended for amplification in each step. 4. The PCR primer should contain as much of the thymine converted from non-CpG site cytosine as possible to avoid amplifying un-converted DNA. 5. If bisulfite PCR does not amplify sufficient PCR product, purification, and concentration (e.g., phenol extraction and ethanol precipitation) can be used to improve TA-cloning efficacy. 6. Purification of the PCR product may improve subcloning efficacy, but is not necessary.

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7. Sequencing analysis with over 20 colonies is recommended for statistical analysis. 8. QUMA (http://quma.cdb.riken.jp) or BiQ Analyzer (http:// biq-analyzer.bioinf.mpi-inf.mpg.de) was useful and convenient to quantify and visualize the bisulfite sequencing data. References 1. Zuo T, Tycko B, Liu TM et al (2009) Methods in DNA methylation profiling. Epigenomics 1:331–345 2. Bibikova M, Fan JB (2010) Genome-wide DNA methylation profiling. Wiley Interdiscip Rev Syst Biol Med 2:210–223 3. Frommer M, McDonald LE, Millar DS et al (1992) A genomic sequencing protocol that yields a positive display of 5-methylcytosine residues in individual DNA strands. Proc Natl Acad Sci U S A 89:1827–1831 4. Grunau C, Clark SJ, Rosenthal A (2001) Bisulfite genomic sequencing: systematic investigation of critical experimental parameters. Nucleic Acids Res 29:E65 5. Leontiou CA, Hadjidaniel MD, Mina P et al (2015) Bisulfite conversion of DNA: performance comparison of different kits and methylation quantitation of epigenetic biomarkers that have the potential to be used in non-invasive prenatal testing. PLoS One 10:e0135058 6. Yang AS, Este´cio MR, Doshi K et al (2004) A simple method for estimating global DNA

methylation using bisulfite PCR of repetitive DNA elements. Nucleic Acids Res 32:e38 7. Li LC, Dahiya R (2002) MethPrimer: designing primers for methylation PCRs. Bioinformatics 18:1427–1431 8. Wada MR, Inagawa-Ogashiwa M, Shimizu S et al (2002) Generation of different fates from multipotent muscle stem cells. Development 129:2987–2995 9. Hashimoto N, Murase T, Kondo S et al (2004) Muscle reconstitution by muscle satellite cell descendants with stem cell-like properties. Development 131:5481–5490 10. Mozzetta C, Consalvi S, Saccone V et al (2013) Fibroadipogenic progenitors mediate the ability of HDAC inhibitors to promote regeneration in dystrophic muscles of young, but not old Mdx mice. EMBO Mol Med 5:626–639 11. Albini S, Coutinho Toto P, Dall’Agnese A et al (2015) Brahma is required for cell cycle arrest and late muscle gene expression during skeletal myogenesis. EMBO Rep 16:1037–1050

Chapter 2 Whole Genome Chromatin IP-Sequencing (ChIP-Seq) in Skeletal Muscle Cells Karl Kamhei So, Xianlu Laura Peng, Hao Sun, and Huating Wang Abstract Transcriptional control of gene expression in skeletal muscle cell is involved in different processes ranging from muscle formation to regeneration. The identification of an increasing number of transcription factors, co-factors, and histone modifications has been greatly advanced by methods that allow studies of genomewide chromatin-protein interactions. Chromatin immunoprecipitation with massively parallel DNA sequencing, or ChIP-seq, is a powerful tool for identifying binding sites of TFs/co-factors and histone modifications. The major steps of this technique involve immunoprecipitation of fragmented chromatin, followed by high-throughput sequencing to identify the protein bound regions genome-wide. Here, in this protocol, we will illustrate how the entire ChIP-seq is performed using global H3K27ac profiling in myoblast cells as an example. Key words Transcription, ChIP-seq, Muscle cells

1

Introduction Transcriptional control through transcription factor/co-factor binding to DNA and subsequent induction of histone modifications is the premier mechanisms involved in regulation of skeletal muscle cell behaviors during muscle formation or regeneration [1, 2]. ChIP-seq has become a widely used method in studying proteinDNA interaction genome-wide [3]. In fact, it has become the new standard for transcriptional or epigenetic study in all types of cells or tissues. Adult muscle stem cells (also called muscle satellite cells) are responsible for muscle regeneration after acute or chronic muscle injury. Upon injury, quiescence muscle satellite cells quickly become activated and proliferate as myoblasts, which then differentiate and fuse into myofibers for repairing of damaged muscle [4]. Intensive efforts are being focused on dissecting the transcriptional and epigenetic regulation in the myogenic lineage progression [5, 6]. For example, MyoD, the master regulator of myoblast differentiation, is required to initiate the differentiation program in myoblast through

James G. Ryall (ed.), Skeletal Muscle Development, Methods in Molecular Biology, vol. 1668, DOI 10.1007/978-1-4939-7283-8_2, © Springer Science+Business Media LLC 2017

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inducing transcription of Myogenin [7]. MyoD and Myogenin then cooperate to induce expression of a vast number of muscle-specific genes such as Myosin heavy chain (MyHC). Myocyte Enhancer Factor 2D (Mef2D) was found to be associated with MyoD, inducing transcription of Histone deacetylase 9 (HDAC9), which causes transcriptional repression by histone deacetylation [8]. Interestingly, MyoD could also bind with histone acetyltransferase p300, causing histone acetylation on enhancers of myotube specific genes, consequently inducing transcription of these genes [7]. Emerging application of ChIP-seq in skeletal muscle cells has yielded a large number of ChIP-seq datasets for various TFs, cofactors, and histone modifications [7, 9]. Analyzing these ChIP-seq data has provided unprecedented global view of transcriptional/ epigenetic regulatory mechanisms in muscle cells. For example, our group has recently performed ChIP-seq for a ubiquitously expressed TF, Yin Yang1 (Yy1) in skeletal muscle myoblast and myotube cells; analyzing the data has yielded previously unknown insights into how Yy1 regulates myogenic differentiation by positively regulating Yam-1, novel muscle-associated large intergenic noncoding RNAs (lincRNAs) [10]. By combining multiple ChIPseq data, the genome-wide cooperativity between TFs can also be studied. For example, MyoD was found to be bound with Runtrelated transcription factor 1 (Runx1), Jun Proto-oncogene (c-Jun) and Forkhead box O3 (FoxO3) on enhancers of muscle-related genes during muscle differentiation [11, 12]. In addition, in satellite cells, ChIP-seq on H3K4me3 and H3K27me3 have allowed us to understand the genome-wide changes in the epigenomic landscapes in the progression of quiescent satellite cells (QSCs) to activated satellite cells (ASCs). Bivalent domains of these two histone marks indicate lineage-specific genes in QSCs [13]. During the transition process from QSCs to ASCs, the level of H3K4me3 remains fairly constant across the genome, while the level of H3K27me3 is low in QSCs but increases in ASCs, indicating a more repressed state [13]. The repressive mark H3K27me3 also increases in QSCs with aging, suggesting a loss of transcriptional potential with aging [13]. Global profiling of H3K27ac by ChIP-seq, on the other hand, will allow for the identification of active enhancer and promoter regions [14, 15]. ChIP involves many steps, including cell lysis, chromatin fragmentation, and immunoaffinity purification of chromatin that contain both desired protein and DNA [16]. The retrieved DNA will then be constructed into a library that can be sequenced on a highthroughput sequencer. The most popularly used sequencer on the market is from Illumina. Sequencing by Illumina sequencers involves binding of adapter ligated DNA library onto a flow cell,

ChIP-Seq in Muscle Cells

17

followed by massively parallel sequencing by synthesis (SBS). The downstream analysis of the sequencing data has been streamlined which includes alignment, peak calling, and result visualization [17, 18]. In this protocol we aim to illustrate the processes of ChIP, library construction and a simple version of bioinformatics analysis by performing H3K27ac ChIP-seq in C2C12 mouse myoblast cells.

2

Materials

2.1 ChromatinImmunoprecipitation Assay

1. 37% Formaldehyde Solution. 2. 2.5 M glycine. 3. Phosphate-buffered saline. 4. Block solution (0.5% bovine serum albumin in PBS). It is freshly made before use and keep cold. 5. Lysis Buffer 1 (150 mM NaCl, 50 mM HEPES-KOH, 1 mM EDTA, 10% glycerol, 0.2% NP-40, 0.25% Triton X-100). Add protease inhibitor cocktail just before use and keep cold. 6. Lysis Buffer 2 (200 mM NaCl, 10 mM Tris–HCl pH 8.0, 1 mM EDTA, 0.5 mM EGTA). Add protease inhibitor cocktail just before use and keep cold. 7. Lysis Buffer 3 (100 mM NaCl, 10 mM Tris–HCl pH 8.0, 1 mM EDTA, 0.5 mM EGTA, 0.1% Na-Deoxycholate, 0.5% N-lauroylsarcosine). Add protease inhibitor cocktail just before use and keep cold. 8. Wash buffer (500 mM LiCl, 50 mM HEPES-KOH, 1 mM EDTA, 0.5% NP-40, 0.5% Na-Deoxycholate). Keep cold. 9. Dynabeads Protein G (Life Technologies, cat. no. 1004D). 10. Magnetic particle concentrator (MPC) (Life Technologies, cat. no. A13346). 11. Cell scraper. 12. Centrifuge and microcentrifuge (Eppendorf, Hamburg, Germany, cat. no. 5810R and 5424R). 13. Sonicator (Covaris, Woburn, MA, model no. S220). 14. Rotator (Labnet, Edison, NJ, model no. H5500). 15. Temperature-controlled room.

2.2 Library Construction 2.2.1 End Repair

1. 200 ng ChIP-enriched and input DNA. 2. T4 DNA ligase buffer with 10 mM ATP (NEB, cat. no. B0202S). 3. dNTPs mix (10 mM each) (NEB, cat. no. N0447S).

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4. T4 DNA polymerase (3 U/μL) (NEB, cat. no. M0203L). 5. Klenow DNA polymerase (5 U/μL) (NEB, cat. no. M0210L) (Dilute to 1 U/μL with 1X NEBuffer 2 before use). 6. T4 polynucleotide kinase (10 U/μL) (NEB, cat. no. M0201L). 7. MinElute PCR Purification kit (Qiagen, cat. no. 28004). 2.2.2 A-Tailing

1. Buffer 2 (NEB, cat. no. B7002S). 2. dATP (1 mM) (NEB, cat. no. N0440S). 3. Klenow fragment (30 –50 exo minus) (5 U/μL) (NEB, cat. no. M0212L). 4. Digital dry bath (Labnet, model no. D1100). 5. MinElute PCR Purification kit.

2.2.3 Adapter Ligation

1. DNA ligase buffer (10) (NEB, cat. no. B0202S). 2. TruSeq DNA indexed adapter (Illumina, cat. no. 15026620). 3. DNA ligase (1 U/μL). 4. MinElute PCR Purification kit.

2.2.4 Enrichment of Adapter-Modified DNA Fragments by PCR

1. PCR grade water. 2. Library amplification primer (10) (KAPA, cat. no. KK2623). 3. Phusion® High-Fidelity DNA Polymerase (NEB, cat. no. M0530S). 4. Thermocycler.

2.2.5 Size Selection

1. Power supply. 2. Gel tank. 3. Agarose. 4. TAE (0.5). 5. Microwave oven.

2.2.6 Quantitation

1. KAPA Library Quantification Kit for Illumina platforms (KAPA KK4854). 2. Library (1000 diluted with water). 3. PCR grade Water.

2.3 Programs for Processing of Sequencing Data

1. SOAP2 (http://soap.genomics.org.cn/soapaligner.html) [19]. 2. MACS (http://liulab.dfci.harvard.edu/MACS/) [20]. 3. Integrated genome browser (IGB) (http://bioviz.org/igb/ index.html).

ChIP-Seq in Muscle Cells

3

19

Methods

3.1 Chromatin Immunoprecipitation

1. All procedures are done on ice or in a temperature-controlled room at 4  C.

3.1.1 Formaldehyde Crosslinking of Cells

2. Grow and collect 5  107 cells for each immunoprecipitation (see Note 1). 3. For adherent cells (for example, myoblast cell line C2C12 or activated satellite cells in culture): (a) Add 1% formaldehyde in PBS and incubate cells with formaldehyde solution for 10 min at room temperature. (b) Add 2.5 M glycine, making final concentration at 0.125 M and incubate for 10 min at room temperature to quench crosslinking reaction. (c) Rinse the cells with PBS twice and harvest cells using cell scraper. (d) Centrifuge the cells at 550  g for 5 min and store at 80  C. 4. For satellite cells freshly sorted by Fluorescence Activated Cell Sorting (FACS): (a) Centrifuge the cells at 550  g for 5 min, remove supernatant and resuspend in 1% formaldehyde in PBS and incubate cells with formaldehyde solution for 10 min at room temperature. (b) Add 2.5 M glycine, making final concentration at 0.125 M and incubate for 10 min at room temperature to quench crosslinking reaction. (c) Centrifuge at 550  g for 5 min, remove the supernatant, and add PBS to wash, repeat once. Discard supernatants and store at 80  C. 5. Crosslinked cells can be stored at 80  C for a few months if not immediately used.

3.1.2 Cell Sonication

1. Resuspend frozen cell pellets from previous steps in 10 mL of Lysis Buffer 1. Rotate on a rotator for 10 min at 4  C, then centrifuge at 1500  g for 5 min at 4  C. Discard the supernatant, repeat with Lysis Buffer 2. 2. Resuspend the pellet in 120 μL Lysis Buffer 3 and transfer to Covaris AFA microTUBE. 3. Sonicate at 4  C under optimized condition for individual cell type (see Note 2). 4. Transfer the sonicated chromatin to 1.5 mL Eppendorf tube and add 1/10 volume of 10% Triton X-100.

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5. Aliquot 1/10 volume of sonicated chromatin as input control and perform elution steps. Centrifuge at 20,000  g for 10 min at 4  C and collect supernatant. Supernatant can be stored at 80  C for a few weeks (see Note 3). 3.1.3 Preparation of Magnetic Beads

1. Add 100 μL of Dynabeads into 1 mL block solution in Eppendorf tube. 2. Place the tube in MPC. Allow beads to collect on the side of the Eppendorf tube. Remove supernatant with pipette. Add 1 mL block solution. 3. Repeat twice (three times in total). 4. Resuspend Dynabeads in 250 μL block solution with 5 μg of antibody. Place on a rotator at 4  C overnight (see Note 4). 5. Wash beads for three times in block solution, as in step 2. 6. Resuspend the antibody-coated beads in 100 μL block solution. 7. Add sonicated chromatin to antibody-coated beads and incubate on a rotator at 4  C overnight.

3.1.4 Wash

1. Collect Dynabeads using MPC. Invert the tube for twice to collect all beads. Remove supernatant and add 1 mL wash buffer to resuspend the beads. Repeat the wash for four more times. 2. Wash the beads once with 1 mL TE buffer þ 50 mM NaCl. 3. Collect the beads using MPC and remove supernatant.

3.1.5 Elution

1. Add 210 μL ChIP Elution Buffer and elute ChIP-enriched chromatin by incubating tubes at 65  C in a thermomixer for 45 min. Vortex or shake at 1500 rpm for 1 min in every 2 min interval. 2. Centrifuge the beads at 16,000  g for 1 min at room temperature. 3. Transfer 200 μL of supernatant to a clean Eppendorf tube. 4. Eluted chromatin can be stored at 20  C for a few days.

3.1.6 Reverse Crosslink

1. Thaw input chromatin and add elution buffer to 200 μL. 2. Reverse crosslink the ChIP-enriched DNA and input chromatin by incubating at 65  C in a thermomixer overnight.

3.1.7 Purification of ChIP-Enriched DNA

1. Add 200 μL of TE buffer to the tube from the previous step. 2. Add 4 μL of 20 mg mL1 RNaseA, mix and incubate at 37  C for 2 h. 3. Add 16 μL of 5 mg mL1 Proteinase K, mix and incubate at 55  C for 2 h.

ChIP-Seq in Muscle Cells

21

4. Add 400 μL Phenol:Chloroform:Isoamyl Alcohol (25:24:1), vortex and centrifuge at 10,000  g for 10 min. 5. Transfer the upper (aqueous) layer to a new Eppendorf tube with 16 μL of 5 M NaCl and 6 μL of 5 μg μL1 glycogen. 6. Add 800 μL of Ethanol. Incubate at 20  C for minimum 30 min. This step can be extended to overnight for higher yield. 7. Centrifuge at 20,000  g for 10 min at 4  C. Discard supernatant carefully and add 500 μL of 80% Ethanol. Centrifuge again at 20,000  g for 5 min at 4  C. 8. Remove supernatant and invert the tube to air dry for 30 min. Add 50 μL of 10 mM Tris–HCl to dissolve the pellet. 9. The DNA can now be used to perform qPCR or undergo library construction for high-throughput sequencing (see Note 5). 3.2 Library Construction 3.2.1 End-Repair

1. Dilute Klenow DNA polymerase with water to 1 U/μL. 2. Prepare the reaction mix as follows: ChIP-enriched DNA

40 μL

T4 DNA ligase buffer with 10 mM ATP

5 μL

dNTP mix

2 μL

T4 DNA polymerase

1 μL

Klenow DNA polymerase (1 U/μL)

1 μL

T4 polynucleotide kinase

1 μL

3. Incubate for 40 min at 20  C in a thermocycler. 4. Do the clean-up reaction with MinElute PCR Purification Kit using protocol as suggested by the manufacturer. Note that 250 μL buffer PB1 and 36 μL buffer EB are used here. 3.2.2 A-Tailing

1. Prepare the reaction mix as follows: End-repaired DNA from previous step

34 μL

Klenow buffer (NEBuffer 2)

5 μL

dATP (1 mM)

10 μL 0

0

Klenow fragment (3 –5 exo-)

1 μL

2. Incubate for 30 min at 37  C in a thermocycler. 3. Do the clean-up reaction with MinElute PCR Purification Kit using protocol as suggested by the manufacturer. Note that 250 μL buffer PB1 and 24 μL buffer EB are used here.

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3.2.3 Adapter Ligation

1. Prepare the reaction mix as follows (see Note 6): A-tailed DNA from previous step

22 μL

DNA ligase buffer

3 μL

TruSeq indexed DNA Adapter (1:10)

1 μL

DNA ligase

4 μL

2. Incubate for 25 min at 25  C in a thermocycler. 3. Do the clean-up reaction with MinElute PCR Purification Kit using protocol as suggested by the manufacturer. Note that 150 μL buffer PB1 and 21 μL buffer EB are used here. 3.2.4 Enrichment of Adapter-Ligated DNA Fragments by PCR

1. Prepare the reaction mix as follows:

2 Phusion Hi-fidelity PCR master mix

25 μL

10 KAPA Library amplification primer

5 μL

DNA from the previous step

20 μL

2. Amplify the adapter ligated DNA using the following PCR protocol: 30 s at 98  C. [10 s at 98  C, 30 s at 65  C, 30 s at 72  C] 12–14 cycles (see Note 7). 5 min at 72  C. Hold at 4  C. 3.3 Quality Control of ChIP-Seq Library

1. Run PCR products from the previous step on 2% agarose gel in TAE.

3.3.1 Size Selection

2. Excise 200–600 bp band and do gel recovery with MinElute Gel Extraction kit. Elute the library with 12 μL elution buffer.

3.3.2 Validation and Quantitation of DNA Library

1. 1 μL DNA library is needed for Bioanalyzer analysis to examine if adapter dimers are present (see Note 8). 2. 1 μL DNA library is needed for quantitation by library quantitation kit (KAPA). 3. Calculate the concentration and dilute it to 8 pM for cluster generation on an Illumina sequencer. 4. Sequencing with 50 bp, single read. Total 10 million reads are enough for standard ChIP-seq analysis.

ChIP-Seq in Muscle Cells

3.4 Bioinformatics Processing of the Sequencing Data

23

After sequencing is completed, select the sequenced reads with Phred score larger than 30 on the sequencer; store the raw sequences in a .fastq file.

3.4.1 Quality Control 3.4.2 Alignment

Map the sequenced reads (.fastq file) to the mouse reference genome (e.g., UCSC mm9 assembly) using SOAP2 [6], allowing maximum of two mismatches. Only uniquely mapped reads are retained for downstream analysis, stored in a .sam file.

3.4.3 Peak Calling

Identify the read enrichment peaks using MACS (Model-based Analysis for ChIP-seq; version 2.0.9, q-value < 105), with IgG ChIP-seq or input DNA as the control (see Note 9).

3.4.4 Visualization

Convert the mapped reads (.sam file) to .bedGrapgh, .wig or . BigWig file and upload to UCSC genome browser or submit to integrated genome browser (IGB) for the visualization of read intensity, as well as peak regions (Fig. 1).

4

Notes 1. 5  107 cells are sufficient for ChIP of histone marks, two times or more cells are needed for ChIP of transcription factors. 2. Sonication condition should be optimized for each cell type. The optimal size of sheared chromatin is 100–300 bp. The size of sheared chromatin should be examined before performing the ChIP. This can be done by running 200 ng of sheared input DNA on gel electrophoresis (Fig. 2). 3. Enrichment should be tested before library construction. Primers amplifying positive or negative control regions should be designed for enrichment test. The size of amplicon in qPCR should be between 100 and 150 bp. 4. Amount of antibody used should be optimized for each antibody. Typically, 5 μg of antibody for histone marks and 10 μg of antibody for transcription factors. Chr7 RefSeq

H3K27ac ChIP-seq

Fig. 1 Identification of Myod1 enhancer by H3K27ac ChIP-seq. By ChIPseq, enrichment of H3K27ac signal (bottom track) 22 kb upstream of Myod1 locus is identified in myoblast cells. The enhancer region is predicted by H3K27ac signal as indicated by the bar in genomic snapshot

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1 2 3 4 5 6 7

Lane

300bp 200bp 100bp

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Peak Power (W) /

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175

Duty %

/

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10

Cycle per Burst

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Fig. 2 Optimization of sonication condition on Covaris sonicator. Chromatins are sheared under various combinations of Peak Power, Duty%, Cycle per Burst, and Time (s). Left: 200 ng of chromatins are loaded in 1.5% agarose gel for visualization; right: various conditions of sonication in Covaris sonicator

5. In case of low enrichment or high background, preclear the lysate with 50 μL of magnetic beads overnight at 4  C with rotation. 6. In order to reduce the formation of adapter dimer, optimal amount of ChIP-ed DNA and adapter should be adjusted as follows: for 50–200 ng of ChIP-ed DNA, dilute the adapter with ddH2O at 1:10; for 201–1000 ng of ChIP-ed DNA, do not dilute the adapter. 7. In order to increase complexity of ChIP-seq library and reduce PCR artifact, a minimal number of PCR cycles should be used. For 50–200 ng of ChIP-ed DNA, perform 14 PCR cycles. For 201–1000 ng of ChIP-ed DNA, perform 12 PCR cycles. 8. It is critical to remove most of the adapter dimers before sequencing because they are short sequence (130–140 bp) which have higher preference to bind on the flow cell of Illumina sequencer, causing majority of reads to be adapter sequence in the output. For example, if the input library contains 1% of adapter dimer, 6.5% of reads will be adapter sequence; if the input library contains 5% of adapter dimer, 60.4% of reads will be adapter sequence. This will cause tremendous wastage of useful reads. 9. The number of identified peaks is associated with the sequencing depth, as well as the p- or q-value cutoff in the step of peak calling. Generally speaking, with the increasing number of sequenced reads, the number of peaks tends to gradually reach the plateau. However, considering the cost of sequencing, 10–20 million raw sequenced reads are usually generated for the downstream analysis. In addition, a looser p- or q-value cutoff in peak calling will lead to a larger number of resulting peaks. Therefore, one should take notion in criteria adjustment by manually checking the visualized read density and peak regions in order to decide the optimal cutoff for each experiment.

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Acknowledgment The work is substantially supported by General Research Funds (GRF) from the Research Grants Council (RGC) of the Hong Kong Special Administrative Region [14102315 and 14113514 to H.S.; 14133016, 14100415, 14116014, and 476113 to H. W.]; Focused Innovations Scheme: Scheme B to H.S. [Project Code: 1907307]; RGC Collaborative Research Fund (CRF) from RGC [Project Code: C6015- 14G to H.S. and H.W.]. References 1. Sincennes MC, Brun CE, Rudnicki MA (2016) Concise review: epigenetic regulation of myogenesis in health and disease. Stem Cells Transl Med 5(3):282–290 2. Brack AS, Rando TA (2012) Tissue-specific stem cells: lessons from the skeletal muscle satellite cell. Cell Stem Cell 10(5):504–514 3. Diehl AG, Boyle AP (2016) Deciphering ENCODE. Trends Genet 32(4):238–249 4. Tierney MT, Sacco A (2016) Satellite cell heterogeneity in skeletal muscle homeostasis. Trends Cell Biol 26(6):434–444 ˜ oz-Ca´noves P 5. Segale´s J, Perdiguero E, Mun (2015) Epigenetic control of adult skeletal muscle stem cell functions. FEBS J 282 (9):1571–1588 6. Fu X, Wang H, Hu P (2015) Stem cell activation in skeletal muscle regeneration. Cell Mol Life Sci 72(9):1633–1677 7. Blum R, Vethantham V, Bowman C, Rudnicki M, Dynlacht BD (2012) Genome-wide identification of enhancers in skeletal muscle: the role of MyoD1. Genes Dev 26(24):2763–2769 8. Sebastian S, Faralli H, Yao Z, Rakopoulos P, Palii C, Cao Y, Singh K, Liu QC, Chu A, Aziz A, Brand M, Tapscoot SJ, Dilworth FJ (2013) Tissue-specific splicing of a ubiquitously expressed transcription factor is essential for muscle differentiation. Genes Dev 27 (11):1247–1259 9. Asp P, Blum R, Vethantham V, Parisi F, Micsinai M, Cheng J, Bowman C, Kluger Y, Dynlacht BD (2011) Genome-wide remodeling of the epigenetic landscape during myogenic differentiation. Proc Natl Acad Sci U S A 108 (22):E149–E158 10. Lu L, Sun K, Chen X, Zhao Y, Wang L, Zhou L, Sun H, Wang H (2013) Genome-wide survey by ChIP-seq reveals YY1 regulation of lincRNAs in skeletal myogenesis. EMBO J 32 (19):2575–2588 11. Blum R, Dynlacht BD (2013) The role of MyoD1 and histone modifications in the

activation of muscle enhancers. Epigenetics 8 (8):778:784 12. Peng X, So K, He L, Zhao Y, Zhou J, Li Y, Yao M, Xu B, Zhang S, Yao H, Hu P, Sun H, Wang H (2017) MyoD- and FoxO3-mediated hotspot interaction orchestrates super-enhancer activity during myogenic differentiation. Nucleic Acids Res. doi:10.1093/nar/gkx488 13. Liu L, Cheung TH, Charville GW, Hurgo BMC, Leavitt T, Shih J, Brunet A, Rando TA (2013) Chromatin modifications as determinants of muscle stem cell quiescence and chronological aging. Cell Rep 4(1):189–204 14. Vernimmen D, Bickmore WA (2015) The hierarchy of transcriptional activation: from enhancer to promoter. Trends Genet 31(12):696–708 15. Shlyueva D, Stampfel G, Stark A (2014) Transcriptional enhancers: from properties to genome-wide predictions. Nat Rev Genet 15 (4):272–286 16. Arrigoni L, Richter AS, Betancourt E, Bruder K, Diehl S, Manke T, Bo¨nisch U (2016) Standardizing chromatin research: a simple and universal method for ChIP-seq. Nucleic Acids Res 44(7):e67 17. Thomas-Chollier M, Darbo E, Herrmann C, Defrance M, Thieffry D, van Helden J (2012) A complete workflow for the analysis of fullsize ChIP-seq (and similar) data sets using peak-motifs. Nat Protoc 7(8):1551–1568 18. Bailey T, Krajewski P, Ladunga I, Lefebvre C, Li Q, Liu T, Madrigal P, Taslim C, Zhang J (2013) Practical guidelines for the comprehensive analysis of ChIP-seq data. PLoS Comput Biol 9(11):e1003326 19. Li R, Yu C, Li Y, Lam TW, Yiu SM, Kristiansen K, Wang J (2009) SOAP2: an improved ultrafast tool for short read alignment. Bioinformatics 25(15):1966–1967 20. Zhang Y, Liu T, Meyer CA, Eeckhoute J, Johnson DS, Bernstein BE, Nusbaum C, Myers RM, Brown M, Li W, Liu XS (2008) Modelbased analysis of ChIP-Seq (MACS). Genome Biol 9(9):R137

Chapter 3 Analysis of RNA Expression in Adult Zebrafish Skeletal Muscle Tamar E. Sztal, Peter D. Currie, and Robert J. Bryson-Richardson Abstract The zebrafish is an excellent vertebrate model system to investigate skeletal muscle development and disease. During early muscle formation the small size of the developing zebrafish allows for the characterization of gene expression in whole embryos. However, as the zebrafish develops, access to the underlying skeletal muscle is limited, requiring the skeletal muscle to be sectioned for a more detailed examination. Here, we describe a straightforward and effective method to prepare adult zebrafish skeletal muscle sections, preserving muscle morphology, to characterize gene expression in the zebrafish adult skeletal muscle. Key words Zebrafish adult skeletal muscle, Pathology, Vibratome sectioning, RNA in situ hybridization, Skeletal muscle

1

Introduction Zebrafish are an excellent model system to study muscle development and disease. During early muscle formation the small size and optical clarity of the developing zebrafish allows for the characterization of gene expression in whole embryos. However, as the zebrafish grows in size a thick, pigmented, layer of skin encases the fish, preventing the direct visualization of the skeletal muscle beneath and thus a skeletal muscle section must be analyzed instead. Preliminary assessment of muscle pathology is usually based on a series of histological and immunological tests, performed on muscle samples, which have been indispensable in characterizing muscle development and in the identification of many neuromuscular disorders [1–4]. Most of these techniques involve preserving the skeletal muscle by either fixing the tissue in formaldehyde and then embedding them with paraffin wax or freezing the sample for cyrosectioning [5–8]. While both methods are sufficient to highlight important pathological features of the skeletal muscle, they

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either require the handling of toxic chemicals, which destroy the RNA, or can introduce freezing artifacts that severely disrupt muscle morphology [9, 10]. Furthermore, these techniques yield thin sections (approximately 10 μm), which can lead to a weak signal and many important details may be missed. We have developed a quick and easy technique to characterize the adult zebrafish skeletal muscle pathology using thick (100 μm) vibratome sections. Using the vibratome method to prepare skeletal muscle tissue, in addition to preserving RNA and protein integrity within the samples, we are able to maintain the morphology and 3D architecture of the muscle fiber. Staining of thicker sections also provides a clearer signal to identify the expression and localization of a gene of interest [11].

2

Materials Prepare all solutions using ultrapure water and analytical grade reagents. Prepare and store all reagents at room temperature unless otherwise indicated. Follow all waste disposal regulations when disposing waste materials.

2.1 Preparation of Muscle Sections

1. Embryo medium (E3): 5.0 mM NaCl (0.292 g), 0.17 mM KCl (0.013 g), 0.33 mM CaCl2 (0.044 g), 0.33 mM MgSO4 (0.081 g). Make up to 1 L with water and pH to 7.4. Store at room temperature for up to 1 week. 2. Tricaine (3-amino benzoic acidethylester) anesthetic solution: Make up in a glass bottle. Weigh 400 mg tricaine powder (Sigma) and add 97.9 ml of water and 2.1 ml of 1 M Tris (pH 9) then adjust pH to approximately 7. Store at 4
C. Add approximately 1 ml of tricaine solution per 25 ml of E3 (see Note 1). 3. 90 mm plastic Petri dishes. 4. Scalpel blade. 5. 50 ml plastic falcon tube. 6. 4% paraformaldehyde (see Note 2). 7. PBS solution: add 8 g of NaCl, 0.2 g of KCl, 1.44 g of Na2HPO4, 0.24 g of KH2PO4 to 800 ml of MQ water. Adjust the pH to 7.4 and make up to 1 L with MQ water. 8. For PBSTween (PBST): Add 5 ml of 20% Tween20 to 1 L of PBS solution. 9. 4% Low Melting Temperature Agarose (LMT): add 4 g of LMT powder to 100 ml of PBS in a conical flask on a heat block until dissolved (see Note 3). 10. TissueTek 25  20  5 mm plastic cryomold.

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11. Paintbrush. 12. Poly-L-lysine coated glass slides (see Note 4). 13. PAP pen. 2.2 RNA In Situ Hybridization

1. The following materials are needed to construct both the standard and overnight hybridization chambers: plastic airtight container, which is large enough to hold a microscope slide box with detachable lid, tissue paper, and Petri dishes. 2. Formamide (see Note 5). 3. 20 SSC stock solution: Add 175.3 g of NaCl and 88.2 g of sodium citrate to 800 ml of MilliQ water. Adjust the pH to 7.0 and make up to 1 L with MilliQ water. For 2 SSC, add 100 ml of 20 SSC to 900 ml of MilliQ water. 4. Proteinase K: add 10 ml of DEPC-treated water to 100 mg of Proteinase K. Store at 70
C (see Note 6). 5. Prehybridization solution: Add 25 ml of Formamide, 12.5 ml of 20 SSC, 0.025 ml of 100 mg/ml Heparin, 0.5 ml of 50 mg/ml yeast tRNA, 0.25 ml of 20% Tween20, 0.460 ml of 1 M citric acid and make up to 50 ml with MilliQ water (see Note 7). 6. Digoxygenin (DIG)-labeled RNA in situ hybridization probe (see Note 8 about synthesis). 7. Hybridization buffer: Add 60 ml of Formamide, 25 ml of 20 SSC, 0.5 ml of 20% Tween20, 0.96 ml of 1 M citric acid and make up to 100 ml with MillQ water (see Note 9). 8. Blocking solution: Add 0.4 ml of 100 mg/ml BSA and 0.4 ml of sheep serum to 20 ml of PBST (see Note 10). 9. Anti-DIG antibody solution. Dilute Anti-DIG antibody 1:5000 in blocking solution (see Note 11). 10. AP staining buffer: Add 5 ml of 1 M Tris pH 9.5, 2.5 ml of 1 M MgCl2, 1 ml of 5 M NaCl, 0.25 ml of 20% Tween 20 and make up to 50 ml with MilliQ water. 11. Staining solution: Add 0.02 ml of NBT/BCIP stain to 1 ml of AP staining buffer (see Note 12). 12. Aqueous mounting solution (see Note 13).

3

Methods Carry out all procedures at room temperature unless otherwise instructed.

3.1 Preparation of Muscle Sections

1. Anesthetize fish until they are not moving, place each in a Petri dish, and measure the total length using a ruler or calipers.

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Fig. 1 Preparing the zebrafish for sectioning. (a) Once the 15 mm zebrafish is anesthetized, the head and tail regions are cut off with a scalpel blade prior to fixation. To obtain optimal sections for RNA expression analyses, the region highlighted in yellow is the optimal region to use. (b) Schematic of the resultant section within the yellow region contains the myotome (colored in gray) with the muscle pioneer cells along the central midline (in red)

Once a fish of the desired length is identified, place the fish in the anesthetic for a further 5 min until dead. Once the fish is dead, place it in a Petri dish and cut off the head (between the gills and the pectoral fin, see Fig. 1a) and tail (between the anal and caudal fins; see Fig. 1a) using a scalpel blade. This region is chosen since it contains mostly myotome and avoids the intestine and anal regions, which provide hollow regions that are often difficult to section. Fix the fish by transferring it to a 50 ml Falcon tube containing 4% PFA and shake at 4
C overnight. Multiple fish can be fixed at a time but ensure the fish is completely covered by the PFA solution. 2. Pour off the 4% PFA and wash 4  15 min in PBST. 3. Pipette 4% LMT agarose into the plastic cyromold to form a layer covering the bottom and, using forceps, gently place the fish on the top of the agarose, with the anterior side to the left. 4. Fill the cryomold with the remaining agarose and allow it to set. 5. Once set, cut the agarose out of the cryomold and mount the agarose block in the correct orientation on the vibratome (with the anterior facing downward). Trim the vibratome block until the swim bladder and intestinal chambers are barely visible and the myotome encompasses the majority of the section. Cut 100 μm vibratome sections and place each section, using a soft paintbrush, in a plastic Petri dish filled PBS-Tween. 6. To mount the sections, carefully pick up the section using a soft paintbrush (size 6) and place in the center of a Poly-L-lysine slide. Use tweezers to spread the section out flat on the slide and use a pipette to remove any excess PBSTween from around the section. Multiple sections can be mounted on the slide but

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leave at least 1 cm between the sections and a border of at least 1.5 cm between the sections and the edge of the slide. 7. Allow the slides to dry for 2–3 h at room temperature and draw around the edge of the slide with a PAP pen. 8. Label the slide with a lead pencil and place them in a 37
C oven for 15 min. 3.2 In Situ Hybridization (on Mounted Sections)

Perform all steps with slides laying flat in a hybridization chamber at room temperature unless otherwise instructed. 1. To prepare the standard hybridization chamber, place a sheet of tissue paper at the bottom of a slide box, moisten with water, and rest slides laying flat across the slide box. To prepare the overnight hybridization chamber (see Fig. 2), place two sheets of tissue paper at the bottom of a slide box. 2. Prepare a solution of 100 ml formamide/100 ml 2 SSC pour 50 ml of this solution onto the tissue paper and rest slides laying flat across the slide box. Place a layer of blotting paper across the bottom of a large airtight plastic container and pour

Fig. 2 Assembly of the overnight hybridization chamber consists of (a) a sealable microscope slide box, with a layer of tissue paper on the bottom and microscope slides lay flat across the box. (b) Four Petri dishes are placed along the bottom of an airtight container, with a layer of tissue paper on the bottom, and the microscope slide box (a) is placed on the top of the Petri dishes. The tissue papers in both (a) and (b) are moistened with formamide/SSC solution

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the remainder of the formamide/SSC solution onto the blotting paper. Place four evenly spaced Petri dishes across the bottom of the plastic container on the top of the blotting paper and rest the slide box on the top of the Petri dishes. 3. Wash slides 4  15 min with PBSTween by pipetting the solution onto the slides ensuring that all of the sections are covered and then pouring the solution off the slides between each wash slide. 4. Incubate the sections for 5 min in 10 mM Proteinase K and then wash the sections twice with PBSTween and incubate for 30 min in 4% Paraformaldehyde (ensure this is performed in the fume hood). 5. Wash the slides for 5  5 min in PBSTween and transfer to the overnight hybridization chamber (work in the fume hood from hereon). 6. Pour off the PBSTween from the slides and incubate the sections for 90 min in Prehybridization Solution. 7. Replace the Prehybridization Solution with Prehybridization solution containing a DIG-labeled RNA probe (approximately 200 ng of RNA probe in 150 μl of Prehybridization solution) and incubate in the sealed hybridization chamber overnight at 65
C (see Note 14). 8. Pour off the probe and rinse slides with hybridization buffer to remove excess probe. 9. Wash slides for 15 min each with 75% hybridization buffer/ 25% 2 SSC, 50% hybridization buffer/50% 2 SSC, 25% hybridization buffer/75% 2 SSC, and 100% 2 SSC solutions at 65% in the sealed hybridization chamber. 10. Transfer slides to the standard hybridization chamber and wash for 2  30 min with 0.2 SSC at 65%. Wash slides for 15 min each with 75% 0.2 SSC/25% PBSTween, 50% 0.2 SSC/ 50% PBSTween, 25% 0.2 SSC/75% PBSTween, and 100% PBSTween. 11. Incubate the sections in Blocking solution for at least 2 h and then overnight in anti-Digoxygenin-antibody solution at 4
C. 12. Wash slides 6  15 min with PBSTween and then 3  5 min in staining buffer. Ensure that all of the PBSTween is removed from the slides before the stain is added. 13. Add 200 μl of NBT/BCIP stain to the slides and incubate in the dark until the stain develops (see Note 15). 14. Once the desired staining is obtained (Fig. 3), pour off the NBT/BCIP stain and rinse three times with PBSTween, then incubate for 30 min with 4% Paraformaldehyde. 15. Rinse with PBSTween and replace with an aqueous or glycerol mounting solution. Place a coverslip on the top of the slide and set aside to dry.

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Fig. 3 (a) In situ hybridization for myosin heavy chain (myhc) on skeletal muscle section, with increased magnification shown in (a0 ). (b, b0 ) In situ hybridization for laminin alpha 2 (lama2) and (c, c0 ) laminin alpha 4 (lama4). Expression of myhc and lama2 is expressed throughout the myotome, whereas lama4 is expressed in small cells between the muscle fibers (arrowheads), which are only evident due to the thicker nature of the sections. Full details of the probes used are available in [11]

4

Notes 1. Tricaine solution can be stored at 4
C; however, the solution may begin to precipitate, at which time it needs to be replaced. 2. Paraformaldehyde must be used in the fume hood. If paraformaldehyde is being made up from powder, wear a mask when weighing and dissolving the powder. Once the solution is dissolved, the mask may be removed. 3. The LMT agarose will take at least 1 h to dissolve on the heat block; however, it can also be heated in the microwave. Once

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the solution is dissolved, it will need to be kept on the hot plate to ensure that it does not set during the mounting steps. 4. As an alternative to Poly-L-lysine-coated slides, Superfrost microscope slides can also be used although these are slightly less adhesive. 5. For the overnight hybridization chamber only, commercial grade formamide can be used. 6. Monitor the slides under the microscope when the proteinase K is added to the sections. Different batches of proteinase K may vary in strength and may destroy the sections if not carefully monitored. 7. Prehybridization solution should be made up and dispensed in the fume hood. Once the solution is made up it can be stored for long term at 20
C. 8. Synthesis of Digoxygenin (DIG)-labeled RNA in situ probe [12]. 9. Hybridization solution should be made up and dispensed in the fume hood. Once the solution is made up it can be stored for long term at 20
C. 10. Once the blocking solution is made up it can be stored for long term at 20
C. 11. Anti-DIG antibody solution must be made up fresh and used immediately. In order to increase specificity, a pre-absorbed anti-DIG antibody can be diluted in the blocking solution instead. To make the pre-absorbed anti-DIG antibody homogenize zebrafish with a pestle in 0.1 ml of Blocking solution and make up to 1.0 ml with Blocking solution. The embryos do not need to be dechorionated and should include the same or older stages than will be used in the in situ hybridization. Add 0.01 ml of anti-DIG antibody and shake overnight at 4
C. Spin down the debris and collect the supernatant containing the pre-absorbed anti-DIG antibody, which can be stored at 4
C until needed. The pre-absorbed antibody can be used instead of the anti-DIG antibody stock at a final dilution of 1:2000 in Blocking solution. 12. Ensure that all of the PBST is removed before the staining solution is added. If the solution becomes cloudy, rinse the slides with AP staining buffer and add fresh staining solution to the sections. 13. A hard-set aqueous mounting medium such as VectaShield HardSet or Fluoromount is preferred. Alternatively, the sections can be mounted in an 80% Glycerol solution (80% Glycerol/20% PBS) with a coverslip applied over the top. The coverslip can be fixed using nail varnish and left to dry. Do not dehydrate the sections; otherwise, the PAP pen will

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dissolve over the slide and the morphology of the skeletal muscle will be disrupted. 14. Ensure that the overnight hybridization chamber can fit comfortably into the 65
C incubator without tipping on its side. 15. While the stain is developing, the sections will need to be closely monitored under a microscope. The staining may be easy to identify in regions throughout the muscle fibers (Fig. 3a, b) or in smaller cells within the myotome (Fig. 3c), which can be easily missed.

Acknowledgments The research was funded by an Australian National Health and Medical Research Council (NHMRC) Project Grant (APP1010110). TES is supported by MDA Developmental Grant (APP381325) and AFM Telethon Postdoctoral Fellowship (APP19853). PDC is supported by NHMRC Principal Research Fellowships (APP1002147, APP1041885). References 1. Nowak KJ, Ravenscroft G, Laing NG (2012) Skeletal muscle α-actin diseases (actinopathies): pathology and mechanisms. Acta Neuropathol 125:19–32 2. Joyce NC, Oskarsson B, Jin L-W (2012) Muscle biopsy evaluation in neuromuscular disorders. Phys Med Rehabil Clin N Am 23:609–631 3. Bo¨nnemann CG, Wang CH, Quijano-Roy S et al (2014) Diagnostic approach to the congenital muscular dystrophies. Neuromuscul Disord 24(4):289–311 4. Fetterman GH, Wratney MJ, Donaldson JS, Danowski TS (1956) Muscular dystrophy. I. History, clinical status, muscle strength, and biopsy findings. AMA J Dis Child 91:326–338 5. Dubowitz V, Sewry C, Oldfors A, Lane R (2007) Histological and histochemical changes. In: Muscle biopsy: a practical approach, 4th edn. Sauders Elsevier, pp 55–95 6. Kumar A, Accorsi A, Rhee Y, Girgenrath M (2015) Do’s and don’ts in the preparation of

muscle cryosections for histological analysis. J Vis Exp 99:e52793 7. Sheriffs IN, Rampling D, Smith VV (2001) Paraffin wax embedded muscle is suitable for the diagnosis of muscular dystrophy. J Clin Pathol 54:517–520 8. Shi S-R, Liu C, Pootrakul L et al (2008) Evaluation of the value of frozen tissue section used as “gold standard” for immunohistochemistry. Am J Clin Pathol 129:358–366 9. Meng H, Janssen PML, Grange RW et al (2014) Tissue triage and freezing for models of skeletal muscle disease. J Vis Exp 89:e51586 10. Chatterjee S (2014) Artefacts in histopathology. J Oral Maxillofac Pathol 18:S111–S116 11. Sztal T, Berger S, Currie PD et al (2011) Characterization of the laminin gene family and evolution in zebrafish. Dev Dyn 240:422–431 12. Broadbent J, Read EM (1999) Wholemount in situ hybridization of Xenopus and zebrafish embryos. Methods Mol Biol 127:57–67

Part II Assays for Proliferating Skeletal Muscle Cells

Chapter 4 Targeted Lipidomic Analysis of Myoblasts by GC-MS and LC-MS/MS Jordan Blondelle, Jean-Paul Pais de Barros, Fanny Pilot-Storck, and Laurent Tiret Abstract Lipids represent
10% of the cell dry mass and play essential roles in membrane composition and physical properties, energy storage, and signaling pathways. In the developing or the regenerating skeletal muscle, modifications in the content or the flipping between leaflets of membrane lipid components can modulate the fusion capacity of myoblasts, thus constituting one of the regulatory mechanisms underlying myofiber growth. Recently, few genes controlling these qualitative and quantitative modifications have started to be unraveled. The precise functional characterization of these genes requires both qualitative and quantitative evaluations of a global lipid profile. Here, we describe a lipidomic protocol using mass spectrometry, allowing assessing the content of fatty acids, glycerophospholipids, and cholesterol in the routinely used C2C12 mouse myoblast cell line, or in primary cultures of mouse myoblasts. Key words Myoblast fusion, Membrane lipid composition, LPC, VLCFA, MUFA, CL, Chromatography, Mass spectrometry

1

Introduction By promoting the movement of our eyes, mouth, tongue, lungs, trunk, and limbs, skeletal muscles constitute the largest vital and sociabilizing organ in our body, representing 31–38% of its mass [1]. Because of their sustained and sometimes intense mechanical activity also requested for thermoregulation, skeletal muscles are key actors of the bodily energetic metabolism. These pleiotropic roles of skeletal muscles rely on their early development, growth, and maintenance. Skeletal muscles are primarily composed of muscle fibers (myofibers) which are syncytia formed by the fusion of precursor cells, called myoblasts, derived from mesodermal progenitor cells [2]. Proliferation, myogenic differentiation, and fusion of myoblasts take place from embryonic and fetal to early postnatal stages [3]. A genetic cascade involving the activation of myogenic transcription factors regulates initial steps. This process is

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recapitulated throughout our life, upon exercise or injury; newly formed myofibers can thus be added to the existing or damaged muscle. Adult myoblasts emerge from a pool of quiescent progenitor stem cells located between the plasmalemma of the muscle fiber and the surrounding basal lamina, and called satellite cells [2]. Myoblast fusion relies on a three-step fusion mechanism between a competent myoblast and a muscle founder cell, regulated by a coordinated interplay between protein and lipid complexes [4]. Cell adhesion proteins of the Ig superfamily, which brings cells to within a close 20–50 nm distance from each other, mediate the first step of apposition between cell membranes. The second step is characterized by the activation of the actin-polymerization machinery and leads to the formation of transient actin-based structures including roundish F-actin-enriched focus at the site of fusion. The third step consists in the destabilization of the lipid bilayer which facilitates fusion between the two proximal outer leaflets of the bilayers (hemi-fusion stage), followed by fusion between the two distal inner leaflets (fusion pores) [5]. Then, cytoplasms of cells contact each other, leading to the mixture of intracellular components within the syncytium. Although numerous proteins have been identified as key components in the three steps, such as the actin-branching Arp2/3 complex, the GTPase Rac1, or the Myomaker fusogenic protein pathway [4, 6–10], there is increasing evidence that lipids may have promoting and sometimes complementary functions. Lipids play specific signaling roles. For example, they are able to form membrane microdomains, such as lipid rafts, which modulate trafficking and signaling pathways. Indeed during the second step of fusion, the phospholipid-bound Casein kinase 2 interacting protein-1 (CKIP-1) is dynamically targeted to the Arp2/3 complex [11], which is involved in building the F-actin focus through a transient local enrichment of phosphoinositides PI (4,5)P2, another profusion membrane lipid [12]. Lipid composition also regulates membrane dynamics through the modulation of its fluidity or its permissiveness to deformation and curvatures such as those observed during the hemi-fusion stage. An increase in membrane fluidity has been observed just before myoblast fusion [13, 14]. Moreover, myoblast differentiation and fusion are concomitant with modifications in lipids composition of membranes, which is likely, at least partially, responsible for the observed increase in their fluidity [14, 15]. In addition to the aforementioned transient local enrichment of the phosphoinositide PI(4,5) P2 [12], fusion-promoting modifications in membrane lipid composition also include the enrichment of unsaturated fatty acids [13, 15, 16] and the flipping of phosphatidylserine from the inner leaflet [17, 18]. During the myogenic differentiation, phosphatidylserine located in the outer leaflet can bind to Stabilin2, proved to promote fusion during myogenic differentiation [19]. Finally, lysophosphatidylcholine (LPC), a broad membrane fusion inhibitor [20],

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inhibits myoblast fusion when added to the culture medium [21] and a decrease in its content is observed during fusion [14]. Because fusion defects have been recently identified as a pathogenic mechanism per se in congenital centronuclear myopathies [14], in limb-girdle muscular dystrophy type 2B, 2L, in Miyoshi myopathy type 3 [22, 23] and in the Carey-Fineman-Ziter syndrome that includes a congenital myopathy with facial weakness [24], it is a therapeutic issue to identify regulatory pathways that could be targeted by pharmacological compounds. The genetic control of lipid composition modifications in competent myoblasts has just started to be identified in mammals, using both in vivo and in vitro models. In particular, the elongation of very long chain fatty acids, their unsaturation level, and the drop in LPC content during myoblast fusion are induced by the dynamic expression of an active, muscle-specific isoform of the Hacd1 gene [14], encoding an enzyme involved in the endoplasmic reticulum-resident fatty acid elongation machinery [25]. Determinants of the phosphatidylserine flipping remain to be deciphered, as well as other putative mechanisms involving lipids. A comprehensive nomenclature of lipids amenable to chemists, biologists, and the biomedical community was first proposed in 1976 and has since been regularly updated with a last version dated 2009 [26]. The accepted classification of eight categories is chemically based (Table 1). The present protocol allows quantifying lipids from four classes, which general chemical structure is reminded and an example is provided (Fig. 1). Table 1 Number of lipids identified in mammalians as of March 2017, in each of the eight categories established by the multi-institutional consortium LIPID Metabolites And Pathways Strategy (LIPID MAPS; www.lipidmaps. org and [26]). Standard abbreviations are indicated into brackets. Lipids quantified by the present protocol are from categories (a), (c), (d) and (e) Number of lipids per category [24] (a) Fatty acyls (FA)

7,010

(b) Glycerolipids (GL)

7,542

(c) Glycerophospholipids (GP)

9,620

(d) Sphingolipids (SP)

4,352

(e) Sterol lipids (ST)

2,835

(f) Prenol lipids (PR)

1,256

(g) Sacccharolipids (SL)

1,316

(h) Polyketides (PK)

6,742

Total

40,673

Fig. 1 Structure and abbreviated common name of lipids quantified by the present protocol, belonging to the FA, GP, SP, and ST categories

Targeted Lipidomic Analysis of Myoblasts pM

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mM

nM SM Cer GIc-Cer Lac-Cer

Minor lipids

SP

Major lipids

Cer-1P Sph Spa S1P Spa-1P

CHOL TAG DAG PE PC

GP

PS PG PA PI

Specific analysis profiling

Global analysis profiling

COX 5-LOX

FA

15-LOX

(this protocol)

CYP NAE

EC 10-12

10-11

FAG 10-10

10-9

10-8

10-7

10-6

10-5

10-4

10-3

Species concentration (M)

Fig. 2 Relative abundance of blood lipid (serum or plasma) contents and methodological strategies applied to quantify them. EC stands for endocannabinoids, which are included in the FA category. (Adapted from [27])

There is a great disparity in the range of endogenous concentrations from the most to least abundant member of each lipid within cells or bodily samples. This is illustrated by the examples of the serum and the plasma, reviewed in [27]. Consequently, there is no possibility of obtaining a comprehensive profile of minor lipids that, in global approaches, are masked by major classes. Methods targeting specific lipids can be used to overcome this sensitivity bottleneck (Fig. 2). Here, we describe a targeted mass spectrometry-based protocol that allows us to quantify fatty acyls, glycerophospholipids, sphingomyelins, and cholesterol in myoblasts, extracted either from the routinely used C2C12 mouse cell line or from primary cultures of myoblasts obtained from limb muscles of postnatal mouse pups. Combined with the fusion index of differentiating myoblasts, these analyses are mandatory to dissect the molecular pathways on which endogenous determinants or exogenous compounds act to promote fusion. The main sequential steps from cell culture to analysis of data are summarized (Fig. 3).

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Fig. 3 Schematized methodological pipeline of the proposed protocol

2

Materials Prepare all solutions using ultrapure water (prepared by purifying deionized water, to attain a sensitivity of 18 MΩ·cm at 25  C) and analytical grade reagents. Prepare and store all reagents at room temperature (unless indicated otherwise). Chemicals are of the highest grade available.

Targeted Lipidomic Analysis of Myoblasts

2.1

C2C12 Cells

45

1. C2C12 cell vials (ATCC, Cat# CRL-1772TM) stored at 196  C in liquid nitrogen. 2. Proliferation medium (see Note 1). 3. Differentiation medium (see Notes 2 and 3). 4. Enzymatic dissociation medium (see Note 4). 5. Phosphate-Buffered Saline 1 (PBS). Store at 4  C. 6. Hemocytometer.

2.2 Myoblasts Isolation and Culture

1. M-proliferation medium (see Note 5). 2. M-complete proliferation medium (see Note 6). 3. Gelatin-coated dishes (see Note 7). 4. Collagenase/Dispase II solution (see Note 8). 5. 0.9% NaCl solution. 6. Scalpel blades n 10. 7. Mora fine scissors. 8. 100 μm cell strainers. 9. Centrifuge.

2.3

DNA Extraction

1. Automated Nucleic Acid Purification Maxwell 16 Research Instrument (Promega). Alternatively, use any other extraction instrument or method routinely used nearby. 2. Maxwell® 16 Buccal Swab LEV DNA Purification Kit (Promega, Cat# AS1295). 3. Assess concentration, purity, and yield using a spectrophotometric or a UV fluorescence tagging spectrometric quantitation method.

2.4 Tubes, Antioxidants, and Internal Standards

1. Glass tubes (see Note 9). 2. EDTA: ethylenediaminetetraacetic acid (see Note 10). 3. BHT: butylated hydroxytoluene (see Note 10). 4. Argon gas. 5. Lipid and deuterated fatty acids standards (Table 2).

2.5 Mass Spectrometry

1. GC MS: Agilent 7890A Gas Chromatograph equipped with a 7683 injector and a 5975C Mass Selective Detector (Agilent Technologies). 2. GC column: HP-5MS fused silica capillary column (30 m  0.25 mm inner diameter, 0.25 μm film thickness, Agilent Technologies). 3. HPLC: Agilent 1200 equipped with an autosampler, a binary pump, and a column oven (Agilent Technologies).

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Table 2 List of the internal standards used Name

Abbreviation

Company

Cat #

Lauric Acid (d3)

12:0 d3

CDN Isotopes

D-4027

Myristic Acid (d3)

14:0 d3

CDN Isotopes

D-3604

Palmitic Acid (d3)

16:0 d3

CDN Isotopes

D-1655

Stearic Acid (d3)

18:0 d3

CDN Isotopes

D-1825

Linoleic Acid (d4)

18:2 n-6 d4

Cayman

390150

Arachidic Acid (d3)

20:0 d3

CDN Isotopes

D-5254

Arachidonic Acid (d8)

20:4 n-6 d8

Cayman

69254-37-1

Behenic Acid (d3)

22:0 d3

CDN Isotopes

D-5708

Docosahexaenoic Acid (d5)

22:6 n-3 d5

Cayman

10005057

Lignoceric Acid (d4)

24:0 d3

CDN Isotopes

D-6167

Cerotic Acid (d4)

26:0 d3

CDN Isotopes

D-6145

Epicoprostanol

5β-Cholestan-3α-ol

Sigma

C2882

1,2-diheptadecanoyl-snglycero-3-phosphocholine

(17:0)2 PC

Avanti polar lipids

850360P

1-heptadecanoyl-2-hydroxy-snglycero-3-phosphocholine

17:0 LPC

Avanti polar lipids

855676P

1,2-diheptadecanoyl-sn-glycero3-phosphoethanolamine

(17:0)2 PE

Avanti polar lipids

830756P

1-myristoyl-2-hydroxy-sn-glycero3-phosphoethanolamine

14:0 LPE

Avanti polar lipids

856735P

N-heptadecanoyl-D-erythrosphingosylphosphorylcholine

d18:1/17:0 SM

Avanti polar lipids

860585P

1,2-dimyristoyl-sn-glycero3-phospho-L-serine

(14:0)2 PS

Avanti polar lipids

840033P

4. LC reverse phase chromatography column: ZorBAX EclipsePlus C18—2.1 mm  100 mm, 1.8 μm (Agilent Technologies). 5. MS/MS: Agilent 6460 QqQ triple quadrupole mass spectrometer equipped of a Jet Stream electrospray ionization source (ESI) (Agilent Technologies).

3

Methods All the procedures are carried out at room temperature, unless otherwise specified. Initial steps are performed under sterile cell culture conditions.

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3.1 C2C12 Cell Culture

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1. Warm the proliferation medium in a 37  C water bath for at least 20 min. 2. Fill up 10 cm culture dishes with 10 ml of pre-warmed proliferation medium. 3. Quickly thaw selected C2C12 cells (see Note 11) and plate them. 4. Let cells rest and attach for 6 h, then replace the medium with fresh proliferation medium. Cells should be maintained at 37  C in a humidity-saturated atmosphere containing 5% CO2. Keep cells at the optimal confluence (see Note 12). 5. If differentiation is required, count and seed the cells the afternoon before (see Note 13). Aspirate the proliferation medium and wash cells with pre-warmed PBS. Remove PBS and add 1 ml of 0.05% Trypsin-EDTA. Incubate cells at 37  C for 2–3 min (see Note 14). Apply mechanical stress to cells by pipetting up and down and transfer the 1 ml of dissociated cells in a 15 ml Falcon tube containing 4 ml of proliferation medium. Count cells and seed 1  105 cells/well in 3 cm plates. 6. Approximately 18 h later, control that cells are close to 70–80% confluency (see Note 15). Activate the differentiation of cells by replacing the proliferation medium with the differentiation medium.

3.2 Isolation and Culture of Primary Myoblasts

1. Prepare 0.2% Gelatin-coated 10 cm dishes before starting the muscle isolation (see Note 16). 2. Proceed with the euthanasia of the animal using the preferred method that has been validated by your Institution (see Note 17). Spray the animal fur with 70% ethanol to avoid the hair contamination of your samples. 3. Remove as much of non-muscle tissues as you can, including skin, fat, tendon, and nerve tissues around or attached to the muscle(s) (see Note 18). 4. In order to reduce cell death, quickly isolate muscles using fine scissors and place them in Petri dishes containing 10 ml of cold PBS (see Note 19). Move the dishes into a tissue culture hood. Proceed with the next steps with sterile tools and environment. This is critical to avoid contamination of your culture. 5. Place each muscle in a dry 6 cm plate and mince tissues with surgical scissors until disappearance of large pieces. 6. Add 0.5–1 ml of Collagenase/Dispase solution on the top of the muscle and further chop muscles (ten strockes) using a scalpel blade. Muscles should now have a slurry consistency.

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7. Transfer the mixture into a 1.7 ml Eppendorf tube and add Collagenase/Dispase solution up to 1.5 ml. 8. Incubate the tube on a Thermomixer (1000 rpm) at 37  C for 15 min. The mixture should be almost completely digested (see Note 20). 9. In the meantime, place a cell strainer on the top of a 50 ml Falcon tube and wet the filter with 1 ml of PBS. 10. Transfer the mixture containing cells to the cell strainer using a P1000 pipette. Use a cut end-tip (see Note 21). 11. Once the mixture has totally gone through the strainer, add 2–3 ml of PBS to rinse it. Without touching the membrane, pipet up and down and make sure that almost all the solution passed through the strainer. Finally, add another 5 ml of PBS and pipet up and down until all the solution passed through the filter. 12. Transfer filtered solution into a 15 ml Falcon tube and centrifuge for 5 min at 300  g. You should see a white pellet that contains cells. Aspirate the supernatant. 13. Resuspend cells in 10 ml of M-proliferation medium and plate in an uncoated 10 cm dish. Let them sit for 2 h in the incubator at 37  C (see Note 22). Collect the supernatant, spin for 5 min at 300  g (see Note 23). 14. Resuspend cells in 10 ml of M-complete proliferation medium and plate in a 10 cm-gelatin-coated dish. Incubate at 37  C with 5% CO2. 15. Refresh the medium with the M-complete proliferation medium 48 h later. At that stage, there will be a lot of debris and cell death and it is still possible to see a mix of myocyte and non-myocyte cells. Non-myocyte cells should disappear after the first passages; a differential pre-plating may be necessary (see step 13). Always perform passages before cells reach 70–80% confluency. 16. Refresh the medium every 48 h. Differentiation of cells may be triggered by replacing M-complete proliferation medium with differentiation medium. 3.3 Cell Extracts, Aliquots, and Storage

Culture can be stopped during proliferation of cells or at any specific time point of the differentiation process. 1. Aspirate the medium and wash cells with 0.9% NaCl solution. 2. Aspirate the 0.9% NaCl solution and add 1 ml of Trypsin-EDTA in the 10 cm dish. Incubate for 2–3 min (see Note 14). Resuspend cells in 5 ml of culture medium. Spin the cells at 300  g for 10 min. Wash the pellet with 0.9% NaCl. Aspirate the NaCl solution.

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49

3. Resuspend the cell pellet in 120 μl of PBS containing 10 nM of EDTA and 50 mg/ml BHT (see Note 23). 4. Separate the homogenized suspension into one aliquot of 20 μl (to quantitate DNA), stored at 20  C and one aliquot of 100 μl to quantitate lipids, stored at 80  C. 3.4 Preparation of Internal Standards

The internal standards used are lipids entities added in a constant amount to samples, blanks, and if necessary calibration standards (see Note 24). 1. The SI_Mix1 and SI_Mix2 prepared below are for 3–10 million cells (see Note 25). 2. Prepare fresh SI_Mix1 solution (cholesterol and phospholipids internal standards) using stock solutions (1 μg/μl) of standards. Ten microliters of SI_Mix1 solution contains 500 ng of epicoprostanol (5β-Cholestan-3α-ol), 50 ng of 17:0 LPC, 500 ng of (17:0)2 PC, 50 ng 14:0 LPE, 500 ng (17:0)2 PE, 500 ng d18:1/17:0 SM, 200 ng (14:0)2 PS in chloroform/ methanol/water 60/30/4.5 v/v/v. Keep on ice. 3. Prepare SI_Mix2 of deuterated fatty acid standard mix in ethanol using fatty acid stock solutions (1 μg/μl) kept at 80  C under argon. Five microliters of SI-Mix2 contains 260 ng of myristic acid-d3, 1128 ng of palmitic acid-d3, 840 ng of stearic acid-d3, 650 ng of linoleic acid d4, 1.04 ng of arachidic acidd3, 432 ng of arachidonic acid-d8, 1.08 ng of behenic acid-d3, 108 ng of DHA-d5, 0.52 ng of Lignoceric-d4, and 0.4 ng of cerotic acid-d4. Keep on ice (see Note 26).

3.5 Addition of Standards and Lipid Extraction

The following procedure is adapted from [28, 29]. 1. Thaw the aliquot of 100 μl prepared in Subheading 3.3 above on ice (see step 5). If the cell pellet corresponding to 3–10 million cells was stored dry at –80  C under argon, resuspend it in 100 μl of PBS containing 10 mM of EDTA and 50 mg/ml of butylated hydroxytoluene (PBS/EDTA/BHT solution). In both the cases, transfer the 100 μl in a new 2 ml microtube. Keep on ice. 2. Rinse the initial pellet tube with 120 μl of PBS/EDTA/BHT solution and transfer to the new 2 ml microtube (total volume: 220 μl). Sonicate the suspension and homogenize with a 20–200 μl micropipette. Transfer 50 μl of the homogenate in a 10 ml glass tube previously filled with argon for total fatty acids GC MS quantitation (see Note 15). Close tightly and keep at 20  C until the hydrolysis step of esterified fatty acids. 3. Add 50 μl 0.9% NaCl to 150 μl of the homogenate for total lipids extraction.

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4. Dispense 10 μl of SI_Mix1 solution using a digitally controlled analytical syringe. 5. Add 750 μl of chloroform/methanol 1/2 v/v. Mix vigorously for 10 min. 6. Add 250 μl of chloroform. Mix vigorously for 10 min. 7. Add 250 μl of ultrapure water. Mix vigorously for 10 min. 8. Centrifuge 5 min at 10,000  g. Transfer the lower organic phase into a new 1.5 ml microtube. 9. Add 8 μl of hydrochloric acid (3 mol/l) and 600 μl of chloroform to the remaining upper phase. Mix vigorously for 10 min. 10. Eliminate the upper aqueous phase and pool the lower organic phase to the previous one. 11. Evaporate the organic phase to dryness under vacuum. Solubilize the extract with 100 μl of a chloroform/methanol/water 60/30/4.5 v/v/v mix. Transfer to 300 μl glass inserts placed in 2 ml glass vials and store at 20  C until LC-MS/MS analysis. 3.6 Targeted GC NCIMS of Fatty Acids

1. Use the 50 μl aliquot from Subheading 3.5 (see step 2). 2. Dispense 5 μl of SI_Mix2 solution using a digitally controlled analytical syringe. 3. Dispense 1 ml of ethanol/BHT (50 mg/ml) and 60 μl of potassium hydroxide 10 mol/l. Fill the tube with argon and incubate 45 min at 56  C. 4. Add 1 ml of hydrochloric acid (1.2 mol/l) and extract the aqueous phase with 2 ml of hexane. 5. Mix vigorously for 10 min. Centrifuge for 10 min at 2000  g. 6. Collect the upper organic phase and evaporate to dryness under vacuum. 7. In a chemical hood, dispense sequentially 5 μl of pentafluorobenzyl bromide, 90 μl of acetonitrile, and 5 μl of diisopropylbenzylamine. Incubate at 37  C for 30 min. 8. Extract the resulting pentafluorobenzyl-fatty acid esters (PFBFAs) with 1 ml of water and 2 ml of hexane. Evaporate to dryness under vacuum. 9. Solubilize PFB-FAs with 100 μl of hexane. Transfer to 300 μl glass inserts placed in 2 ml glass vials. 10. Inject 1 μl of PFB-FAs in pulsed split mode on the 7890A Gas Chromatograph using the HP-5MS fused silica capillary column. The GC MS conditions are: carrier gas, helium at a flowrate of 1.1 ml/min; injector temperature setup at 250  C, pulsed split 10; oven temperature setup at 140  C, increased of 5  C/min to 300  C, and held for 10 min. The mass

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51

Fig. 4 GC-MS ion chromatograms from an analysis of a typical 1 million cells sample. (Upper panel) Total ion chromatogram profile of PBF-fatty acyl esters. Deuterated internal standards are displayed in red bold characters. (Lower panel) Extracted ion chromatograms of palmitoleic acids (n-9 and n-7 isomers), palmitic acid and tri-deuterated palmitic acid at m/z ¼ 253.2, 255.2 and 258.2, respectively

spectrometer operates under negative chemical ionization mode with methane as reactant gaz. Ion source and quadrupole temperatures were set up at 150  C. 11. Quantitate fatty acids by calculating their relative response ratios to their closest internal standard (Fig. 4). Calibration

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curves are obtained with fatty acid authentic standards (14:0, 15:0, 16:0, 17:0, 18:0, 19:0, 20:0, 21:0, 22:0, 24:0, 26:0, 16:1n-7, 18:1n-7, 18:1n-9, 18:2n-6, 18:3n-3, 18:3n-6, 20:2n-6, 20:3n-3, 20:4n-6, 20:5n-3, 22:3n-3, 22:5n-6, 22:6n-3) processed as cell pellets. 3.7 Targeted LC-MS/MS (QqQ) Identification of Glycerophospholipids and Sphingolipids in Single Reaction Monitoring Mode

1. Use the 300 μl aliquot from Subheading 3.5 (see step 11). 2. Inject 2–4 μl of sample. Lipids are separated by high performance liquid chromatography (HPLC) on the Agilent 1200. During analysis, the reverse phase chromatography column is maintained at 55  C. The mobile phases consist of (A) water/ methanol (60/40 v/v) 10 mM ammonium acetate, 1 mM acetic acid, and of (B) IPA/methanol (90/10), 10 mM ammonium acetate, 1 mM acetic acid. Lipids are separated by a linear gradient as follows: start at 40% of B for 1 min, reach 95% of B in 15 min, and then maintain at 100% for 1 min. Column is equilibrated with 40% B for 6 min before each injection. The flow rate is maintained at 0.25 ml/min. The autosampler temperature is set up at 5  C. 3. Set up the LC system coupled to the Agilent 6460 QqQ triple quadrupole mass spectrometer equipped of a Jet Stream electrospray ionization source (ESI) as follows: source temperature at 325  C, nebulizer nitrogen gas flow rate of 10 l/min at 20 psi, nitrogen sheath gas flow of 11 l/min, temperature at 300  C, capillary voltage of 3500 V, nozzle voltage of 1000 V. 4. Identify lipids of interest following the detailed fragmentation patterns of phospholipids by LC-MS/MS, recently reviewed [30].

3.7.1 LPC, PC, and SM

1. Perform analyses of LPC, PC, and SM by single reaction monitoring (SRM) positive mode using the MS/MS parameters listed (Table 3). The positive ion [MþH]+ represents the protonated form of the considered lipid. The product ion at m/z 184.1 represents the phosphocholine head group [C5H15O4NP]+ produced during the collision-induced dissociation process in the triple quadrupole mass spectrometer. An example of profile is shown (Fig. 5).

3.7.2 LPE and PE

1. Perform analyses of LPE and PE by single reaction monitoring (SRM) mode positive using the MS/MS parameters listed (Table 3). The positive ion [MþH]+ represents the protonated form of the considered lipid. The product ion at m/z [MþH–141]+ represents the ion resulting in a neutral loss of the phosphoethanolamine head group [C2H8O4NP] during the collision-induced dissociation process in the triple quadrupole mass spectrometer.

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53

Table 3 List of MS/MS spectrometric parameters used for single reaction monitoring quantitation of phospholipid and sphingomyelin species

Lipid class

Collision energy (V)

Polarity

Product ion

Fragmentor voltage (V)

[MþH]

+

184.1

138

20

Positive

PC

[MþH]

+

184.1

160

24

Positive

SM

[MþH]+

184.1

LPC

LPE PE

Precursor ion

[MþH]

+

[MþH]

+



[M–H]

(17:0)2PC

[MþCH3COO]

PS

[M–H]

20

Positive

[MþH–141]

110

12

Positive

[MþH–141]

+

136

12

Positive

172

50

Negative

172

50

Negative

150

19

Negative

241

PI



160 +

269.2 [M–H–87]



Fig. 5 LC-MS/MS single reaction monitoring elution profile of lysophosphatidylcholines (LPC) separated by reverse phase liquid chromatography. Each LPC molecule gives two peaks figuring 2-acyl (sn2) and 1-acyl (sn1) LPC isomer, respectively 3.7.3 PI and PS

1. Perform analyses of PI and PS by single reaction monitoring (SRM) negative mode using the MS/MS parameters listed (Table 3). The negative ion [M–H] represents the deprotonated form of the considered molecule. The product ion at m/z 241 results from the loss of the phosphoinositol head group

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[C6H10O8P] during the collision-induced dissociation process in the triple quadrupole mass spectrometer. As there is no PI internal standard commercially available, (17:0)2PC can be used for quantitation of PI. The negative ion [MþCH3COO] represents the (17:0)2PC with an acetyl adduct (m/z ¼ 820.6). The product ion at m/z 269.2 results from the loss of the C17:0 fatty acyl chain [C17H33O2]. The negative ion [M–H] represents the deprotonated form of the considered PS. The product ion at m/z [M–H–84]+ represents the ion resulting after neutral loss of the phosphoethanolamine head group [C3H5O2N] during the collision-induced dissociation process in the triple quadrupole mass spectrometer. 3.8 Targeted GC EIMS Identification of Cholesterol

1. Use the 300 μl aliquot from Subheading 3.5 (see step 11). 2. Transfer the lipid extract from the LC-MS/MS vial into a 10 ml glass tube. Rinse the insert with 100 μl of chloroform/methanol/water 60/30/4.5 v/v/v mix and transfer to the same 10 ml glass tube. Evaporate to dryness under vacuum. 3. Dispense 1 ml of ethanol/BHT (50 mg/ml) and 60 μl of potassium hydroxide 10 mol/l. Incubate for 45 min at 56  C. 4. Add 1 ml of ultrapure water and extract the aqueous phase with 5 ml of hexane. 5. Mix vigorously for 10 min. Centrifuge for 10 min at 2000  g. 6. Collect the upper organic phase and evaporate to dryness under vacuum. 7. Dispense 100 μl of a mixture of bis(trimethylsilyl)trifluoroacetamide/trimethylchlorosilane 4/1 v/v and incubate for 1 h at 80  C. 8. Evaporate to dryness under a gentle nitrogen stream in a chemical hood. 9. Solubilize the derivatized sterols as O-trimethylsilyl ether with 100 μl of hexane. Transfer to 300 μl glass inserts placed in 2 ml glass vials. 10. Inject 1 μl in pulsed split mode on the 7890A Gas Chromatograph coupled with the 5975C Mass Selective Detector. Separation is achieved on a HP-5MS 30 m  0.25 mm column using helium as carrier gas and the following GC conditions: injection at 250  C using the pulsed split mode (split 10), oven temperature program: initial temperature setup at 140  C, gradually increased until 280  C using a 15  C/min slope and then, until 300  C using a 2  C/min slope. The MSD was set up as follows: EI at 70 eV mode, source temperature at 230  C. Data is acquired in single ion monitoring (SIM) mode using m/z 368.3 and 370.3 as quantitation ions for cholesterol and epicoprostanol, respectively.

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55

3.9 SemiQuantitation of Lipids

1. Semi-quantify each lipid class species by calculating its response ratio, calculated as the ratio between the area of considered compound on the area of its respective internal standard. Quantitation of cholesterol is performed by calculating its relative response ratios to its closest internal standard. A standard calibration curve is prepared with cholesterol standards (0–2 μg) processed as sample extracts.

3.10 Normalization with DNA Quantitation

When myoblast fusion is experimentally impaired, their differentiation is delayed and their protein content putatively modified in comparison with newly formed syncytial myotubes. The use of protein quantitation to compare control and fusion-deficient batches of myoblasts (a between-samples calibrator) may thus bias the relative quantitation of lipids. One way to overcome this plausible limitation is to rather quantitate DNA, which content remains strictly dependent upon the number of nuclei, regardless of their inclusion in syncytia or their retention as singletons in fusiondeficient myoblasts. 1. Use the method you prefer to extract DNA from the 20 μl aliquot of cells. To avoid the precipitation and resuspension of DNA, we routinely use the automated Maxwell instrument (Promega) using the buccal swab kit and following the manufacturer’s recommendations. 2. Quantitate the total DNA content of the aliquot. 3. Normalize the lipid contents of your cell sample using DNA as a calibrator. 4. Perform relative quantitation of lipid content modifications between all the normalized samples.

3.11 Overview of Alternative and Complementary Analyses

This protocol represents a first step toward the targeted identification and semi-quantitation of some lipids known to contribute to myoblast fusion. A possible refinement of the method allowing a precise quantitation of individual glycerophospholipids and sphingomyelins consists in performing a LC-MS/MS profiling followed by a GC MS of fatty acids for each class of molecules. Complementary methods also include the non-targeted, comprehensive profiling of myoblasts, myofibers, muscles or essential organelles such as mitochondria using LC-High Resolution MS. These mass lipidomic analyses are highly sensitive and specific, allowing for example to identify minimal variations such as inter-breed metabolic differences in plasma [31]. Finally, it becomes feasible to map individual lipids on a tissue section by combining shotgun lipidomics with Liquid Extraction Surface Analysis (LESA), opening the opportunity to map the spatial distribution and molar abundance of hundreds of molecular lipid species within a tissue or across different anatomical regions [32].

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Notes 1. Supplement Dulbecco’s Modified Eagle’s medium (DMEM with 25 mM glucose) with 20% Fetal Bovine Serum and 100 UI/ml Penicilin—0.01 UI/ml Streptomycin (final concentrations). Filter. Store at 4  C. 2. Supplement Dulbecco’s Modified Eagle’s medium (DMEM with 25 mM glucose) with 2% Horse Serum and 100 UI/ml Penicilin—0.01 UI/ml Streptomycin (final concentrations). Filter. Store at 4  C. 3. The Horse serum should be chosen after assessing the fusion efficiency of batches from several companies. Use the same batch in all your comparative analyses. 4. 250 μg/ml Trypsin and 200 μg/ml EDTA solution in PBS. Store at 4  C. 5. Supplement Ham’s F-10 Media with 20% Fetal Bovine Serum and 100 UI/ml Penicilin—0.01 UI/ml Streptomycin (final concentrations). Filter. Store at 4  C. 6. Add 2.5 ng/ml of human recombinant bFGF to Mproliferation medium. The medium must be prepared extemporaneously; otherwise, bFGF will be inactivated. 7. Prepare 0.2% solution of Gelatin (Type A, Porcine skin) in ultrapure water, then autoclave. Spread 6–7 ml of 0.2% Gelatin in 10 cm plates in order to completely cover the plastic surface. Let it sit for a couple of hours under the hood and aspirate the gelatine. Gelatin-coated dishes may be used directly or stored at 4  C for weeks. 8. Dissolve 500 mg of Collagenase B in 50 ml of Dispase II. Add 250 μl of 0.5 M calcium chloride. 9. Plastic tubes release C14:0, C16:0 and to a lesser extent C18:0, added in plastic during the industrial process. This holds true for cones of micropipettes but there is no alternative. 10. EDTA, chelator in particular of the prooxidant copper(II) ion, Cu2+; BHT, a powerful synthetic antioxidant. 11. C2C12 differentiation is highly variable and depends on the batch and the passage. We highly recommend the use of low passages for a better differentiation. In order to test the differentiation of cells before further analyses, index fusion and myogenic factors expression should be assessed. 12. Do not allow cultures to become confluent. Cell density should never reach more than 70% confluency. High confluency will lead to activation of differentiation factors and will deplete the myoblastic population in the culture.

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13. Since the differentiation of C2C12 cells is highly dependent on their density, it is required to seed and trigger cell differentiation always at the same density for better reproducibility. 14. Do not incubate the cells with Trypsin-EDTA too long since it may cause a nonreversible destruction of cells’ anchorage proteins. Use brightfield microscope to assess that the cells are properly detached from the dish and quickly dilute in proliferation medium. 15. Fusion of C2C12 during the differentiation process requires cell-cell contact. By adding the differentiation medium at 70–80% confluence, cells will be allowed to perform one more cell cycle before entering in the myogenic program and reach the full confluence, leading to an efficient differentiation. 16. Gelatin may be replaced by 0.01% Rat Tail Collagen I diluted in 0.2% Acetic Acid. 17. While satellite cells are present throughout the life of the animal, their population represents around 30% of all nuclei contained in muscles during early development and their number decreases to 2–7% in adult muscles [33]. We recommend using 2–4-week-old mice to perform the experiment. 18. Satellite cells/myoblasts are found in all muscles. However, some muscles show higher content than others. For instance, oxidative muscles have more satellite cells than glycolytic muscles (i.e., soleus contains more satellite cells than the extensor digitorum longus (EDL) [33]. We recommend the use of muscles with high content of satellite cells. 19. PBS can be replaced by any other physiological medium. 20. If the mixture is not properly digested, pipette up and down several times with an end cut P1000 tip. That should avoid small pieces of muscles to get trapped inside the tip. Then reincubate for 15 min at 37  C. 21. A large proportion of non-muscle cells such as fibroblasts will attach to the plastic surface and myoblastic population will float. 22. While a single pre-plating may be sufficient, you can perform a second one to reduce even more the number of non-myocyte cells. 23. Alternatively, the cell pellet can be stored dry at – 80  C. In this case, vaporize argon on the pellet to remove air and firmly close your cryotube. Replacement of ambient air by the heavier argon eliminates dioxygen and prevents lipids of the pellet from oxidation.

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24. Such molecules must not be naturally present in samples, so to monitor this point it is highly recommended to first check the absence of these molecules within samples. 25. A 10 cm dish of C2C12 proliferating myoblasts at 70–80% confluence contains
2.5 millions cells. 26. SI_Mix2 vials can be stored at 80 for at least 1 year or at least 2 months at 20  C.

Acknowledgments The authors are grateful to Andrea A. Domenighetti (Rehabilitation Institute of Chicago, IL, USA) for her expert advice on cell culture; Victoria Bergas and He´le`ne Choubley for lipidomics technical assistance; Alexandre Prola for comments on the manuscript. This work was supported by the French National Research Agency (ANR; ANR-12-JSV1-0005), a French Government grant managed by the ANR under the program “Investissements d’Avenir” (Lipstic Labex ANR-11-LABX-0021), the Association Franc¸aise contre les Myopathies (AFM 16143 and Translamuscle), the CNM Project (www.labradorcnm.com), the University of Burgundy and the Institut National de la Sante´ et de la Recherche Me´dicale (INSERM). References 1. Janssen I, Heymsfield SB, Wang Z, Ross R (2000) Skeletal muscle mass and distribution in 468 men and women aged 18–88 yr. J Appl Physiol 89:81–88. doi: 10.1152/japplphysiol. zdg-1052-corr.2014 2. Randolph ME, Pavlath GK (2015) A muscle stem cell for every muscle: variability of satellite cell biology among different muscle groups. Front Aging Neurosci 7:190. doi:10.3389/ fnagi.2015.00190 3. Comai G, Tajbakhsh S (2014) Chapter one – molecular and cellular regulation of skeletal Myogenesis. In: Taneja R (ed) Current topics in developmental biology. Academic Press, Amsterdam, pp 1–73. doi:10.1016/B978-012-405943-6.00001-4 4. Kim JH, Jin P, Duan R, Chen EH (2015) Mechanisms of myoblast fusion during muscle development. Curr Opin Genet Dev 32:162–170. doi:10.1016/j.gde.2015.03.006 5. Zhao W-D, Hamid E, Shin W, Wen PJ, Krystofiak ES, Villarreal SA et al (2016) Hemifused structure mediates and controls fusion and fission in live cells. Nature 534:548–552. doi:10.1038/nature18598

6. Millay DP, O’Rourke JR, Sutherland LB, Bezprozvannaya S, Shelton JM, Bassel-Duby R et al (2013) Myomaker is a membrane activator of myoblast fusion and muscle formation. Nature 499:301–305. doi:10.1038/nature12343 7. Bi P, Ramirez-Martinez A, Li H, Cannavino J, McAnally JR, Shelton JM, Sa´nchez-Ortiz E, Bassel-Duby R, Olson EN (2017) Control of muscle formation by the fusogenic micropeptide myomixer. Science 356(6335):323–327. doi:10.1126/science.aam9361 8. Zhang Q, Vashisht AA, O’Rourke J, Corbel SY, Moran R, Romero A, Miraglia L, Zhang J, Durrant E, Schmedt C, Sampath SC, Sampath SC (2017) The microprotein Minion controls cell fusion and muscle formation. Nat Commun 8:15664. doi:10.1038/ncomms15664 9. Quinn ME, Goh Q, Kurosaka M, Gamage DG, Petrany MJ, Prasad V, Millay DP (2017) Myomerger induces fusion of non-fusogenic cells and is required for skeletal muscle development. Nat Commun 8:15665. doi:10.1038/ ncomms15665 10. He J, Wang F, Zhang P, Li W, Wang J, Li J, Liu H, Chen X (2017) miR-491 inhibits skeletal

Targeted Lipidomic Analysis of Myoblasts muscle differentiation through targeting myomaker. Arch Biochem Biophys 625–626:30–38. doi:10.1016/j.abb.2017.05.020 11. Baas D, Caussanel-Boude S, Guiraud A, Calhabeu F, Delaune E, Pilot F et al (2012) CKIP-1 regulates mammalian and zebrafish myoblast fusion. J Cell Sci 125:3790–3800. doi:10. 1242/jcs.101048 12. Bothe I, Deng S, Baylies M (2014) PI(4,5)P2 regulates myoblast fusion through Arp2/3 regulator localization at the fusion site. Development 141:2289–2301. doi:10.1242/dev. 100743 13. Prives J, Shinitzky M (1977) Increased membrane fluidity precedes fusion of muscle cells. Nature 268:761–763. doi:10.1038/ 268761a0 14. Blondelle J, Ohno Y, Gache V, Guyot S, Storck S, Blanchard-Gutton N et al (2015) HACD1, a regulator of membrane composition and fluidity, promotes myoblast fusion and skeletal muscle growth. J Mol Cell Biol 7:429–440. doi:10. 1093/jmcb/mjv049 15. Nakanishi M, Hirayama E, Kim J (2001) Characterisation of myogenic cell membrane: II. Dynamic changes in membrane lipids during the differentiation of mouse C2 myoblast cells. Cell Biol Int 25:971–979. doi:10.1006/ cbir.2001.0750 16. Briolay A, Jaafar R, Nemoz G, Bessueille L (2013) Myogenic differentiation and lipid-raft composition of L6 skeletal muscle cells are modulated by PUFAs. Biochim Biophys Acta 1828:602–613. doi:10.1016/j.bbamem. 2012.10.006 17. van den Eijnde SM, van den Hoff MJB, Reutelingsperger CPM, van Heerde WL, Henfling MER, Vermeij-Keers C et al (2001) Transient expression of phosphatidylserine at cell-cell contact areas is required for myotube formation. J Cell Sci 114:3631–3642 18. Jeong J, Conboy IM (2011) Phosphatidylserine directly and positively regulates fusion of myoblasts into myotubes. Biochem Biophys Res Commun 414:9–13. doi:10.1016/j.bbrc. 2011.08.128 19. Park S-Y, Yun Y, Lim J-S, Kim M-J, Kim S-Y, Kim J-E et al (2016) Stabilin-2 modulates the efficiency of myoblast fusion during myogenic differentiation and muscle regeneration. Nat Commun 7:10871. doi:10.1038/ ncomms10871 20. Yeagle PL, Smith FT, Young JE, Flanagan TD (1994) Inhibition of membrane fusion by lysophosphatidylcholine. Biochemistry (Mosc) 33:1820–1827

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21. Leikina E, Melikov K, Sanyal S, Verma SK, Eun B, Gebert C et al (2013) Extracellular annexins and dynamin are important for sequential steps in myoblast fusion. J Cell Biol 200:109–123. doi:10.1083/jcb.201207012 22. Griffin DA, Johnson RW, Whitlock JM, Pozsgai ER, Heller KN, Grose WE et al (2016) Defective membrane fusion and repair in Anoctamin5-deficient muscular dystrophy. Hum Mol Genet 25:1900–1911. doi:10.1093/ hmg/ddw063 23. Codding SJ, Marty N, Abdullah N, Johnson CP (2016) Dysferlin binds SNAREs (soluble N-Ethylmaleimide-sensitive factor (NSF) attachment protein receptors) and stimulates membrane fusion in a calcium-sensitive manner. J Biol Chem 291:14575–14584. doi:10. 1074/jbc.M116.727016 24. Di Gioia SA, Connors S, Matsunami N, Cannavino J, Rose MF, Gilette NM, Artoni P, de Macena Sobreira NL, Chan W-M, Webb BD, Robson CD, Cheng L, Van Ryzin C, RamirezMartinez A, Mohassel P, Leppert M, Scholand MB, Grunseich C, Ferreira CR, Hartman T, Hayes IM, Morgan T, Markie DM, Fagiolini M, Swift A, Chines PS, Speck-Martins CE, Collins FS, Jabs EW, Bo¨nnemann CG, Olson EN, Andrews CV, Barry BJ, Hunter DG, Mackinnon SE, Shaaban S, Erazo M, Frempong T, Hao K, Naidich TP, Rucker JC, Zhang Z, Biesecker BB, Bonnycastle LL, Brewer CC, Brooks BP, Butman JA, Chien WW, Farrell K, FitzGibbon EJ, Gropman AL, Hutchinson EB, Jain MS, King KA, Lehky TJ, Lee J, Liberton DK, Narisu N, Paul SM, Sadeghi N, Snow J, Solomon B, Summers A, Toro C, Thurm A, Zalewski CK, Carey JC, Robertson SP, Manoli I, Engle EC (2017) A defect in myoblast fusion underlies Carey-FinemanZiter syndrome. Nat Commun 8:16077. doi:10.1038/ncomms16077 25. Kihara A, Sakuraba H, Ikeda M, Denpoh A, Igarashi Y (2008) Membrane topology and essential amino acid residues of Phs1, a 3hydroxyacyl-CoA dehydratase involved in very long-chain fatty acid elongation. J Biol Chem 283:11199–11209. doi:10.1074/jbc. M708993200 26. Fahy E, Subramaniam S, Murphy RC, Nishijima M, Raetz CRH, Shimizu T et al (2009) Update of the LIPID MAPS comprehensive classification system for lipids. J Lipid Res 50: S9–S14. doi:10.1194/jlr.R800095-JLR200 27. Balgoma D, Checa A, Sar DG, Snowden S, Wheelock CE (2013) Quantitative metabolic profiling of lipid mediators. Mol Nutr Food

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31. Lloyd AJ, Beckmann M, Wilson T, Tailliart K, Allaway D, Draper J (2017) Ultra high performance liquid chromatography–high resolution mass spectrometry plasma lipidomics can distinguish between canine breeds despite uncontrolled environmental variability and nonstandardized diets. Metabolomics 13:15. doi:10.1007/s11306-016-1152-0 32. Almeida R, Berzina Z, Arnspang EC, Baumgart J, Vogt J, Nitsch R et al (2015) Quantitative spatial analysis of the mouse brain Lipidome by pressurized liquid extraction surface analysis. Anal Chem 87:1749–1756. doi:10.1021/ac503627z 33. Hawke TJ, Garry DJ (2001) Myogenic satellite cells: physiology to molecular biology. J Appl Physiol 91:534–551

Chapter 5 Measuring Mitochondrial Substrate Utilization in Skeletal Muscle Stem Cells C. Hai Ly and James G. Ryall Abstract Skeletal muscle stem cells (MuSCs) derived from the somatic mesoderm play a critical role in successful muscle regeneration following injury and trauma. MuSCs have been found to undergo rapid changes in metabolism following a change in cell state, such as that which occurs during the transition from quiescence to an actively proliferating state. There is mounting evidence that metabolism is critically important in the regulation of quiescence, activation, and differentiation and thus the development of new techniques that aim to further probe the metabolism of MuSCs is essential. The Seahorse XF Bioanalyzer is a powerful tool that simultaneously measures the extracellular rate of change in oxygen partial pressure and pH, providing a method to measure mitochondrial respiration and lactate production. In this chapter, we describe the use of key metabolic inhibitors that allow for the investigation of mitochondrial substrate utilization in primary MuSCs. Key words Metabolism, Muscle stem cells, Glycolysis, Glutaminolysis, Fatty acid oxidation

1

Introduction The precise developmental origin of skeletal muscle stem cells (MuSCs) has been the topic of intense research since their discovery in the 1960s. It has since been established that progenitor muscle cells of the trunk and limbs have a mesodermal somatic origin [1] while muscle progenitors in the head have multiple embryonic origins with substantial contributions from the cranial paraxial mesoderm [2]. Pax3+ and Pax7+ progenitor cells in the dermomyotome delaminate to form the myotome (the first skeletal muscles) and also migrate to more distant sites forming the limb bud mesenchyme. Both these processes are Pax3 dependent in mammals [3, 4 ]. In mice, these Pax3+ progenitors begin expressing Pax7 at E11.5 [5] and the expression of Pax7 is proposed to specify these cells to the myogenic lineage [6]. Pax3+/Pax7+ cells have been shown to persist into the latter stages of fetal and postnatal

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development, with many eventually occupying a sublaminal compartment and are presumed to be the postnatal MuSC [7, 8]. Traditional studies in cell biology initially suggested that cellular metabolism was a consequence of cell identity and lineage specification, playing a passive role in response to energy demand. However, recent studies in cancer biology and developmental and stem cell biology have identified a novel process termed “metabolic reprogramming” in which metabolites act as epigenetic cofactors to play an active role in determining cell and lineage specification [9–11]. Stem cell populations, including embryonic stem cells (ESCs) and inducible pluripotent stem (iPS) cells that exhibit high rates of proliferation, have been documented to preferentially utilize glycolysis in their proliferative phase and switch to oxidative phosphorylation (OXPHOS) during differentiation [9–12]. Utilizing whole transcriptome sequencing, it has been reported that quiescent MuSCs are highly enriched for the expression of genes involved in fatty acid oxidation but undergo metabolic remodeling following activation, with an increased expression of genes involved in glycolysis and glutamine metabolism [13]. These results have been supported at the functional level where the Seahorse XF Bioanalyzer has been used to document an increase in glycolysis following MuSC activation [13]. Studies remain ongoing to better understand the epigenetic consequences of this process of metabolic reprogramming [14]. The Seahorse XF Bioanalyzer is now a well-established and powerful tool for the real time analysis of cellular metabolism. The machine is designed so as to allow the user to introduce up to four compounds (or mix of compounds), which can provide a powerful analysis of metabolic pathways. The assay described in this chapter makes use of three compounds: 2-Cyano-3-(1-phenyl-1H-indol-3yl)-2-propenoic acid (UK5099, a mitochondrial pyruvate carrier inhibitor), 2-[6-(4-Chlorophenoxy)hexyl]-oxirane-2-carboxylic acid (etomoxir, an inhibitor of a mitochondrial fatty acid transporter), and Bis-2-(5-phenylacetamido-1,3,4-thiadiazol-2-yl)ethyl sulfide (BPTES, an inhibitor of mitochondrial glutamine utilization). Using these three compounds in different combinations allows for the determination of dependence, capacity, and flexibility of a cell with respect to the utilization of a particular mitochondrial fuel, in this case; glucose, fatty acids, or glutamine (see Fig. 1). Dependence is defined as the reliance on a particular fuel to maintain baseline respiration, the capacity is the ability to increase the utilization of a particular fuel when other metabolic resources are limited or inhibited and the flexibility is the difference between capacity and dependence. Mounting evidence shows that changes in metabolism underlie the extensive growth and proliferation observed in cancer [15]. The observation that non-tumorigenic cells also display a preference for glycolysis during rapid division would suggest that the metabolic rewiring in cancer may also be important in the regulation of

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Glucose

Glycolysis

Palmitate

BPTES

Pyruvate

Etomoxir

UK5099

Acetyl CoA

α-Ketoglutarate 4H+ + O2

TCA Cycle 2H2O

Fig. 1 A schematic showing the site of actions of the three metabolic inhibitors utilized in these assays. UK5099 inhibits the mitochondrial pyruvate transporter (MPC), Etomoxir blocks the action of carnitine palmitoyl-transferase 1a (CPT1a) and BPTES inhibits glutaminolysis by inhibiting glutaminase 1 (GLS1)

proliferation and differentiation in other cell types, including MuSCs [13, 16, 17]. This assay will help researchers better understand the metabolic shifts that occur in MuSCs in response to a change in cell state.

2

Materials

2.1 Cells and Growth Media

1. MuSCs isolated via fluorescence activated cell sorting (see Note 1). 2. Dulbecco’s modified Eagle’s medium (DMEM; 25 mM glucose, 1 mM sodium pyruvate and 2 mM L-glutamine). 3. Horse serum. 4. Fetal bovine serum. 5. Penicillin/Streptomycin (1000 U/mL and 100 μg/mL). 6. Basic fibroblast growth factor.

2.2

XF Assay Media

1. Seahorse base medium. 2. 100 mM Sodium pyruvate. 3. 200 mM L-glutamine. 4. 45% D-glucose solution. 5. 5 M hydrochloric acid (HCl) and 5 M sodium hydroxide (NaOH).

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2.3 Analysis of Metabolic Fuel Dependency and Flexibility

1. XF24 cell culture microplates (see Note 2). 2. XF24 sensor cartridge. 3. Seahorse XF24 Bioanalyzer. 4. Seahorse calibration fluid. 5. Cell counter (optional). 6. UK5099 stock solution (25 mM in DMSO, store at
20  C). 7. Etomoxir stock solution (25 mM in DMSO, store at
20  C). 8. BPTES stock solution (25 mM in DMSO, store at
20  C). 9. Trypsin EDTA (0.25%). 10. Phosphate-buffered saline.

3

Methods

3.1 Cell Seeding and XF Sensor Hydration (Day Prior to Assay)

1. All experiments must be performed at 37  C, so the Seahorse XF Bioanalyzer should be switched on and allowed to equilibrate to 37  C on the day prior to the assay. 2. Make growth media for cell seeding by aliquoting 39.5 mL of DMEM into a 50 mL falcon tube and adding 5 mL of horse serum, 5 mL of fetal bovine serum, 0.5 mL penicillinstreptomycin (1000 U/mL and 100 μg/mL, respectively), and 2.5 ng/mL of basic fibroblast growth factor. 3. Seed cells into the XF cell culture microplate at the predetermined seeding density (see Note 2). Cells are seeded in 200 μL of growth media and should be ejected in a forceful manner to ensure proper mixing of the cells (see Fig. 2) (see Note 3). It is recommended that users include at least n ¼ 4/ group (see Note 4), and at least three wells must contain 200 μL of media alone for background normalization. 4. Place the XF cell culture microplate into a humidified incubator at 37  C with 5% CO2 for 16–24 h to allow attachment of cells (see Note 5).

Fig. 2 Aiming the ejection of the cell suspension at the bottom corner and ejecting forcefully creates a vortex motion of the fluid, allowing mixture of the suspension during ejection and even distribution of cells in the well

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5. The XF sensor cartridge should be hydrated by placing 500 μL of Seahorse calibration fluid into each well of the XF sensor cartridge (see Note 6). 6. Wrap the XF sensor cartridge plate with parafilm to prevent evaporation and then place into a 37  C CO2 free incubator overnight (see Note 7). 3.2 Preparation of Seahorse Assay Medium (Day of Assay)

1. Prepare assay medium by adding 500 μL of glucose solution (200 g/L), 500 μL of glutamine solution (200 mM), and 500 μL of sodium pyruvate solution (100 mM) to 48.5 mL of Seahorse base medium in a 50 mL falcon tube. The final concentrations are 25 mM glucose, 2 mM glutamine, and 1 mM sodium pyruvate. 2. Allow assay medium to equilibrate to 37  C in a water bath before proceeding. 3. Adjust the pH of the Seahorse assay media with 5 M NaOH or 5 M HCl to 7.40  0.02 (see Note 8). Very small increments of acid and base (0.5–1 μL) should be added since this is an unbuffered medium. 4. Return the assay medium to the water bath to allow the temperature to return to 37  C.

3.3 Washing and Equilibrating Cells

1. Cells must be washed with Seahorse assay medium prior to beginning the assay (see Note 9). 2. Assuming cells were seeded in 200 μL remove 180 μL from each well (see Notes 10 and 11). 3. Add 500 μL of Seahorse assay medium to each well. 4. Remove 500 μL from each well. 5. Add 955 μL of Seahorse assay medium to each well. 6. Remove 300 μL from all wells, this should leave 675 μL of media per well. Place the cell microplate into a CO2 free incubator at 37  C for 1 h to allow degassing of the cell culture microplate.

3.4 Preparation of Drug Solutions

1. Remove aliquots of stock drug solutions from freezer and allow to thaw and equilibrate to room temperature. 2. Decant 10 mL of Seahorse assay media into 2  5 mL aliquots in 15 mL falcon tubes and label these as Glucose A and Glucose B to test glucose utilization (or Glutamine A and B to test glutamine utilization, or FA A and B to test fatty acid utilization). 3. Prepare Glucose A by adding 4 μL of 25 mM UK5099 stock solution to 5 mL of Seahorse assay media to a 10 concentration of 20 μM (see Notes 12 and 13 for the preparation of Glutamine A and FA A).

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4. Prepare Glucose B by adding 6 μL of 25 mM BPTES stock solution and 8 μL of 25 mM Etomoxir stock solution to 5 mL of Seahorse assay media to a 10 concentration of 30 μM and 40 μM respectively (see Notes 14 and 15 for the preparation of Glutamine B and FA B). 5. Glucose A and Glucose B should be warmed to 37  C and the pH corrected to 7.40  0.02 before proceeding (see Note 16). 6. Obtain the XF sensor cartridge that has been hydrating in calibration fluid from the CO2 free incubator. 7. Remove parafilm and plastic lid. Ensure the XF sensor cartridge is correctly oriented before proceeding with the following steps. An easy way to do so is to make sure that the side with the “Seahorse” logo is on your right and the “blue line corner” is on the bottom left-hand side (see Fig. 3a, b). 8. To test the dependency of a particular fuel add 75 μL of 10 solution A into port A and 83 μL of 10 solution B into port B (see Note 17). To additionally test the capacity and flexibility of the cells with respect to a particular fuel add 75 μL of 10 solution B to port A followed by 83 μL of 10 solution A to port B (see Note 4). Ensure that the correct solutions are going into the right injection ports (see Fig. 4a, b). 9. Replace the lid on the XF sensor cartridge and place it back into the CO2 free incubator while setting up the protocol on the Seahorse XF Bioanalyzer occurs. 3.5 Seahorse Bioanalyzer Assay Protocol

1. Prior to starting the assay it is necessary to design the assay protocol and configure the plate layout. While the plate layout will be user dependent, an example assay protocol is presented in Fig. 5 (see Note 18).

Fig. 3 (a) Note the cut edge of the sensor cartridge, the plate should be oriented with this edge at the bottom left hand corner. (b) Lifting up the sensor cartridge component will reveal the plate containing the calibrant. There are three landmarks to be aware of; “the blue line corner” which should be at the bottom left corner, the “Utility Plate” logo which should be on the left and the “Seahorse” logo which should be on the right

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Fig. 4 A schematic showing the layout and orientation of the drug ports for each well should the plates have been correctly oriented in the previous steps. The hole in the middle is where the metal probe in the Seahorse Bioanalyzer will be lowered into. A schematic showing the locations to load solutions A and B for (a) the capacity assay and (b) the dependence assay is illustrated

Fig. 5 A sample assay protocol involving a 3 min mix step, a 2 min wait step and a 3 min measure step. The duration of these steps can be altered by individual users

2. The Seahorse assay wizard should be used to enter the assay protocol and plate layout. 3. Begin the assay by selecting “Start Assay” on the assay wizard. 4. Begin the equilibration and calibration step by placing the XF sensor cartridge in the calibration solution into the Seahorse XF Bioanalyzer (see Note 19).

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5. Following the calibration of the probes, remove the plate containing the calibration fluid and replace with the cell culture microplate containing the cells, ensure that the lid of the microplate is removed as the probes in the sensor cartridge will be lowered into the cell culture microplate. 6. At the completion of the assay, remove the sensor cartridge and cell culture microplate from the Seahorse XF Bioanalyzer. The user may now choose to isolate the cells and measure total protein or perform a cell count to use as a control reference for cell numbers (see Note 20). 3.6 Analysis of the Data

At the completion of the assay the Seahorse XF software will generate two files saved at a predetermined directory; an .xfd file that can be opened with the Seahorse XF reader software or the Seahorse Wave software and a .xls file that can be opened with Microsoft excel or a spreadsheet reader (see Note 21). 1. Open either the .xfd file or the .xls file and locate the “Special Operations” tab (see Fig. 6) (see Notes 22 and 23). 2. Select either AUC Analysis or AUC Anova Analysis, this will generate a spread sheet with a summary of the data (see Fig. 6). 3. The glucose dependence can be calculated by taking the area under the curve (AUC) of the basal metabolism and by

Fig. 6 Analyses of the data generated can be performed on the XF reader or on the spreadsheet generated. Users can perform Area under the curve analyses by going into the special operations tab

Measuring Mitochondrial Substrate Utilization A

Injection of UK5099

Injection of Etomoxir/BPTES

B

Injection of Etomoxir/BPTES

69

Injection of UK5099

Glucose Dependence Basal Respiration

Basal Respiration

Non-mito Respiration C

Glucose Capacity Non-mito Respiration

D

Glucose Dependence

Basal Respiration

Flexibility Glucose Capacity Dependence

Non-mito Respiration

Fig. 7 (a) Schematic representation of a seahorse curve. The glucose dependence is the area above the curve following the administration of the MPC inhibitor but below the continuation of basal respiration. This can be calculated by taking the area under the curve of basal respiration and subtracting the area under the curve following administration of the inhibitor of the glucose pathway. (b) A schematic showing glucose capacity, this is the area under the curve following the addition of Etomoxir and BPTES which should force the cells to solely rely on glucose. We then subtract any oxygen consumption following the addition of UK5099 as this is attributed to non-mitochondrial processes. (c, d) The combination of the dependence assay and the capacity assay allows an in-depth look at the fuel pathway of interest. By subtracting the dependence from the capacity, we arrive at the measurement of fuel flexibility, the capacity of cells to increase the utilization of a particular fuel as a compensatory mechanism to losing the use of the other two pathways for mitochondrial metabolism. By combining the fuel dependency assay and the fuel capacity assay, we are able to calculate the flexibility of a cell with respect to a certain fuel. By subtracting the glucose dependence from the glucose capacity we have the glucose flexibility, a measurement of the cells ability to upregulate glucose utilization to compensate for the loss of fatty acid oxidation and glutaminolysis

subtracting the AUC following the introduction of the inhibitor UK5099 (see Note 24) (see Fig. 7a). 4. The calculation of the glucose pathway capacity is performed by calculating the AUC following the addition of Etomoxir and BPTES and then by subtracting the AUC following the injection of the glucose pathway inhibitor, which is the nonmitochondrial respiration (see Fig. 7b). 5. When both the fuel dependency and fuel capacity are performed in conjunction, it is possible to characterize a further property of the cells, the fuel flexibility. The cells flexibility with respect to a certain fuel can be obtained by taking the fuel capacity of the cell and subtracting the fuel dependency of the cell (see Fig. 7c, d).

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Notes 1. MuSCs in this study have been isolated following an adapted protocol by Liu and colleges [18]. It should be noted that C2C12 cells, an immortalized murine myoblast line, can also be used to study the metabolism of myogenic cells. 2. The protocol described has been optimized for the Seahorse XF24, users should adapt cell numbers for use on the Seahorse XF24e, XF96/96e, or XFp. Also, it is important to note that cell size and metabolism can vary significantly depending on whether quiescent or activated MuSCs (or C2C12 cells) are utilized and whether experiments will involve fetal or adult MuSCs. Cell numbers should be optimized by individual users with this in mind. 3. When seeding cells the tip of the pipette should be pointed at the bottom edge of the wells and ejected forcefully to create a vortex, this helps ensure even distribution of the cells in the well as the solution is mixed during ejection. 4. If only the dependence on a particular fuel is to be measured then one group of cells is required for each condition. If capacity and flexibility details are to be determined, then two groups of cells are required for each condition. Therefore, depending on the number of conditions it may be necessary to use one plate for each substrate (glucose, FA, glutamine) analyzed. 5. Immediately following isolation, quiescent MuSCs will begin to activate. As such, any experiments where the quiescent state is of interest, cells cannot be cultured for a prolonged period of time. Quiescent MuSCs should be isolated, seeded at the appropriate density, and the assay should be commenced immediately. 6. The volume of calibration fluid may vary but it is recommended between 500 and 750 μL to be utilized to properly hydrate the fluorometric probes, the tips of the probes themselves must be immersed in the calibration fluid. 7. Ideally, the XF sensor cartridge is hydrated overnight, but if necessary calibration can be completed by hydrating the XF sensor cartridge for 4 h with no discernible difference to readings. 8. The Seahorse XF Bioanalyzer relies on changes to pH to make inferences regarding glycolytic flux; thus, it is important that the pH of the media is finely tuned and consistent across experiments. 9. Most commercially available cell culture media contains a pH buffering agent that will interfere with the reading and

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accuracy of the Seahorse XF Bioanalyzer. It is therefore essential to wash the cells with the Seahorse assay medium to remove any buffering capacity. 10. Care should be taken to not touch the pipette tips to the bottom of the wells, as this can disturb the cell layer at the bottom. Assay media being added to the wells should also be done gently for the same reason. 11. It is recommended that washing steps be performed one row at a time so as to avoid cells drying out. 12. Prepare Glutamine A by adding 6 μL of 25 mM BPTES stock solution to 5 mL of Seahorse assay media to a 10 concentration of 30 μM. 13. Prepare FA A by adding 8 μL of 25 mM Etomoxir stock solution to 5 mL of Seahorse assay media to a 10 concentration of 40 μM. 14. Prepare Glutamine B by adding 4 μL of 25 mM UK5099 stock solution and 8 μL of 25 mM Etomoxir stock solution to 5 mL of Seahorse assay media to a 10 concentration of 20 μM and 40 μM respectively. 15. Prepare FA B by adding 4 μL of 25 mM UK5099 stock solution and 6 μL of 25 mM BPTES stock solution to 5 mL of Seahorse assay media to a 10 concentration of 20 μM and 30 μM respectively. 16. Correcting the pH of the drug solutions is important so as to not cause any spikes in pH readings during measurements that may interfere with data analysis. 17. Note that these injection volumes have been calculated such that the injection volume is 1:10 of the final volume in the well following the injections to achieve the desired concentration of drugs. Should you start with an alternative volume new calculations will need to be made (e.g., 75 μL injected in port A and 675 μL of volume in the well for a total volume of 750 μL). The final volume in each well following all injections in an XF24 cell microplate should not exceed 1000 μL and in the XF96 cell microplate should not exceed 450 μL. Be careful to not load the solutions into the ports too forcefully, as it is possible to break the surface tension of water and also break the membrane at the bottom of the port causing the solutions to prematurely enter the wells. 18. Following any injections or prior measurements the Seahorse XF Bioanalyzer moves through three key steps: a mixing step in which the probes on the sensor cartridges are raised and lowered to thoroughly mix any drugs introduced, a wait step to allow equilibration, and then a measurement step in which the oxygen consumption rate and the extracellular acidification

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rates are concurrently measured. There are three measurement periods in this protocol; baseline respiration, respiration following injection of port A and respiration following the injection of port B. The mix, wait, and measurement steps are looped three times for each period. We use a protocol with a 3 min mix step, 2 min wait step, and a 3 min measurement step. 19. It is essential that the lid of the sensor cartridge be removed prior to being placed into the Seahorse XF Bioanalyzer, as failure to do so may cause damage to the metal probes inside the Seahorse XF Bioanalyzer. 20. It is not advised to use these cells in further culturing experiments as the concentrations of drugs used for these experiments are well beyond the LD50 of these and most cell types. 21. The OCR and raw O2 partial pressures of each well should be examined individually before proceeding with any data analysis to ensure consistent and stable baseline OCR/O2 values. 22. Data can also be analyzed utilizing an excel module that automatically calculates the different parameters of the assay; the Seahorse XF Mito Fuel Report Generator. 23. The AUC can be calculated by utilizing the inbuilt software in the XF reader or by opening the excel file generated. At the time of writing, the Wave software does not currently support the calculation of AUC. 24. Here, an examination of the mitochondrial fuel flexibility assay with respect to glucose is presented, but the same principles apply for the analysis of glutamine and fatty acid utilization. References 1. Scaal M, Christ B (2004) Formation and differentiation of the avian dermomyotome. Anat Embryol 208(6):411–424 2. Harel I, Nathan E, Tirosh-Finkel L, Zigdon H, Guimaraes-Camboa N, Evans SM et al (2009) Distinct origins and genetic programs of head muscle satellite cells. Dev Cell 16(6):822–832 3. Kassar-Duchossoy L, Giacone E, GayraudMorel B, Jory A, Gomes D, Tajbakhsh S (2005) Pax3/Pax7 mark a novel population of primitive myogenic cells during development. Genes Dev 19(12):1426–1431 4. Tajbakhsh S, Rocancourt D, Cossu G, Buckingham M (1997) Redefining the genetic hierarchies controlling skeletal myogenesis: Pax-3 and Myf-5 act upstream of MyoD. Cell 89 (1):127–138 5. Relaix F, Rocancourt D, Mansouri A, Buckingham M (2004) Divergent functions of murine

Pax3 and Pax7 in limb muscle development. Genes Dev 18(9):1088–1105 6. Hutcheson DA, Zhao J, Merrell A, Haldar M, Kardon G (2009) Embryonic and fetal limb myogenic cells are derived from developmentally distinct progenitors and have different requirements for beta-catenin. Genes Dev 23 (8):997–1013 7. Gros J, Manceau M, Thome V, Marcelle C (2005) A common somitic origin for embryonic muscle progenitors and satellite cells. Nature 435(7044):954–958 8. Relaix F, Rocancourt D, Mansouri A, Buckingham M (2005) A Pax3/Pax7-dependent population of skeletal muscle progenitor cells. Nature 435(7044):948–953 9. Folmes CD, Nelson TJ, Martinez-Fernandez A, Arrell DK, Lindor JZ, Dzeja PP et al (2011) Somatic oxidative bioenergetics

Measuring Mitochondrial Substrate Utilization transitions into pluripotency-dependent glycolysis to facilitate nuclear reprogramming. Cell Metab 14(2):264–271 10. Folmes CD, Nelson TJ, Terzic A (2011) Energy metabolism in nuclear reprogramming. Biomark Med 5(6):715–729 11. Panopoulos AD, Yanes O, Ruiz S, Kida YS, Diep D, Tautenhahn R et al (2012) The metabolome of induced pluripotent stem cells reveals metabolic changes occurring in somatic cell reprogramming. Cell Res 22(1):168–177 12. Koopman R, Ly CH, Ryall JG (2014) A metabolic link to skeletal muscle wasting and regeneration. Front Physiol 5:32 13. Ryall JG, Dell’Orso S, Derfoul A, Juan A, Zare H, Feng X et al (2015) The NAD+-dependent SIRT1 deacetylase translates a metabolic switch into regulatory epigenetics in skeletal muscle stem cells. Cell Stem Cell 16(2):171–183

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14. Ryall JG, Cliff T, Dalton S, Sartorelli V (2015) Metabolic reprogramming of stem cell epigenetics. Cell Stem Cell 17(6):651–662 15. Agathocleous M, Harris WA (2013) Metabolism in physiological cell proliferation and differentiation. Trends Cell Biol 23 (10):484–492 16. Wang T, Marquardt C, Foker J (1976) Aerobic glycolysis during lymphocyte proliferation. Nature 261(5562):702–705 17. Brand KA, Hermfisse U (1997) Aerobic glycolysis by proliferating cells: a protective strategy against reactive oxygen species. FASEB J 11 (5):388–395 18. Liu L, Cheung TH, Charville GW, Rando TA (2015) Isolation of skeletal muscle stem cells by fluorescence-activated cell sorting. Nat Protoc 10(10):1612–1624

Chapter 6 Microcontact-Printed Hydrogel Microwell Arrays for Clonal Muscle Stem Cell Cultures Victor M. Aguilar and Benjamin D. Cosgrove Abstract Adult muscle stem cells (also called satellite cells) are an anatomically defined population of cells that are essential for muscle regeneration. In aging and dystrophic diseases, muscle stem cells (MuSCs) exhibit functional and molecular heterogeneity; therefore, single-cell assay technologies are critical to illuminate the mechanisms of pathological stem cell dysfunction. Here, we describe the process of generating mechanically tunable hydrogels with micro-scale well features (“microwells”) using micro-contact printing for clonal muscle stem cell culture assays. Microcontact printing (μCP) is a simple and versatile method for generating cell culture substrates for micro-scale features for spatially restricting the cultures of single cells and their progeny. We explain how to use photolithography and polydimethylsiloxane casting to generate stamps capable of printing purified extracellular matrix proteins onto soft, hydrated poly(ethylene glycol) hydrogels to generate arrayed microwells in a defined pattern. We summarize methods to analyze the viability, migration, and differentiation of individual MuSC clones within hydrogel microwell cultures. Key words Microcontact printing, Photolithography, Hydrogel, Stem cell expansion, Live-cell imaging, Migration, Self-renewal, Laminin

1

Introduction Muscle stem cells (MuSCs; also called satellite cells) contribute to muscle regeneration through self-renewal divisions, which generate both myoblast progenitors capable of myogenic differentiation and additional stem cells [1]. The self-renewal function and molecular identity of MuSCs become increasingly variable and heterogeneous in a number of muscle diseases [2]. Therefore, clonal assessment of individual MuSCs provides a useful method to dissect their functional variability with the goals of refining their functional hierarchy and how heterogeneity contributes to disease pathogenesis [3, 4]. Microcontact printing (μCP) is a versatile approach for patterning culture substrates to provide defined geometries for adhering cells to surface-tethered chemicals and bioactive molecules [5]. μCP entails the transfer of biological molecules–most prominently

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proteins, peptides, and lipids–from an inert patterned stamp to a variety of solid substrates [6]. The resultant geometric pattern on the cell-culture substrate generates an “array” of adherent islands capable of restricting the migration of cells and isolating them to enable long-term clonal analysis [7]. Here, we detail a protocol for using microcontact printing to fabricate poly(ethylene glycol) (PEG) hydrogels with a geometrically defined array of “microwells” with surface-tethered adhesion proteins for the purpose of culturing individual MuSCs and monitoring their clonal progeny. This protocol focuses on fabricating PEG hydrogel discs sized to fit in a multiwell plate with an array of evenly spaced “microwells” with the extracellular matrix protein laminin conjugated to their bottom surface. Proteins are transferred from an inert silicone stamp made from polydimethylsiloxane (PDMS) into PEG hydrogels as they are polymerizing. The PDMS stamps are designed to have an array of positive features (“posts”) that will generate complimentary negative features (“microwells”) on the hydrogel disc after polymerization through μCP [8]. The μCP procedure involves the use of buffer-exchanged and dehydrated sodium dodecyl sulfate-polyacrylamide gel electrophoresis (SDS-PAGE) gels rehydrated with a laminin protein solution. PDMS stamps are first generated from a micro-fabricated silicon wafer, engineered with negative well features by standard photolithography methods. Photolithography is a method that uses high intensity light (typically ultraviolet) to generate a physical pattern on a silicon wafer [9]. While methods have been developed to generate clean patterns across specialized bare wafers, most techniques rely on the use of photochemical compounds known as photoresists. Photoresists vary in chemistry and dissolve (positive photoresists) or harden (negative photoresists) under high intensity light, which is exposed in patterns across the wafer surface through the use of a photomask. Photomasks are designed with a geometric pattern by standard computer-aided drafting software, and a mask pattern imparted on the wafer is mirrored on the final hydrogel substrate. Array designs can be achieved with many types of photoresists, with differing feature-size and depth limits. SU-8, a common negative photoresist, is often employed for microelectromechanical systems (MEMS) given its broad range for feature heights, spanning from a few nanometers to over 400 μm [10]. In this protocol, we describe the use of SUEX, a dry negative photoresist custom-made to specific heights through a layering protocol [11]. SUEX yields more even surfaces and thicker resist layers than SU-8. The silicon wafer is finally etched and/or washed with solvents to remove residues and yield a pattern. Once patterns are finished and treated, PDMS is mixed and cured over the patterned silicon wafer to create stamps. We synthesize PEG hydrogels from two monomer precursors, a 8-arm PEG-vinylsulfone (VS) and a 4-arm PEG-thiol (SH), both at

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10 kDa MW, resuspended in separate storage buffers. The PEG is synthesized through a Michael-type addition reaction and can readily incorporate primary amines and thiols in proteins and peptides [12]. We recommend generating PEG hydrogel microwells with a 150 μm depth (to prevent cells leaving microwells through detachment) and with a 300–600 μm diameter. Smaller diameters limit how long clonal cultures can be maintained without resulting in high (>50%) cell confluency. In 300 μm microwells, we recommend culturing MuSCs for only 4–5 days (resulting in
16-cell clones). In 600 μm microwells, we recommend culturing for 7 days (
128-cell clones). We typically fabricate the microwells onto 15 mm diameter, 1.6 mm thick PEG hydrogel discs, suitable for fitting into 24-well-size tissue culture plates. Using a staggered pattern, approximately 500 microwells at 600 μm diameter can fit onto these discs, resulting in ~393 wells/cm2 culture surface (Fig. 1). To achieve clonal density, we typically seed 500

Fig. 1 Generation of PEG hydrogel microwells by micro-contact printing. (a) Photolithography using SUEX to form silicon wafer with repeating well pattern. Chrome mask patters are designed to establish an etching of SUEX photoresist and determine wafer, PDMS, and hydrogel features. (b) 8 cm silicon wafer with 600 μm well features. (c) Profilometry using a Tencor P10 instrument at a horizontal scanning speed of 1000 μm/ s demonstrates well features are ~600 μm wide and ~170 μm deep. (d) The setup for microcontact printing in hydrogels involves the use of a slider system (PTFE sliders þ PDMS stamp) and a set of binder clip clamps. PDMS stamps, coated with proteins such as laminin, are used to transfer the protein to the PEG hydrogel during its polymerization. The Michael addition reaction incorporates the protein onto the surface of PEG gel, specifically at the bottoms of each microwell feature. The polymerization is considered completed when there is a noticeable separation of the gel from the stamp (bottom far left gel). These show three stamps per slide sandwich but the protocol describes volumes for four stamps

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FACS-sorted α7-integrin+ CD34+ mouse MuSCs per well (see Notes 1 and 2). We typically use long-term time-lapse microscopy to track the migration and cell divisions of all cells in each microwelldefined MuSC clone. Time-lapse sequences are tracked, segmented, and analyzed to generate clonal lineage-trees for the assessment of self-renewal and differentiation at the clonal level [4, 3, 13].

2

Materials This section describes starting reagents and instruments required for the assembly of microwell hydrogels. Prepare and store all reagents at room temperature (unless indicated otherwise). Please use appropriate personal protective equipment (PPE) throughout each protocol and discard waste (including sharps, chemicals, and biohazardous materials) following institutional guidelines.

2.1 Silicon Wafer and PDMS Microfabrication

1. Computer-aided design software (e.g., L-edit). 2. Photolithography contact aligner (e.g., ABM Contact Aligner). 3. Molecular vapor deposition instrument (e.g., Applied Microstructures MVD100). 4. Profilometer (e.g., Tencor P1). 5. SUEX photoresist. 6. Silicon wafers, 10 cm diameter. 7. Hot plates (2). 8. AZ EBR solvent (e.g., Microchemicals). 9. Perfluorooctyltrichlorosilane (FOTS) solution. 10. Isopropyl alcohol. 11. PDMS solution (e.g., Dow Corning Sylgard 184 Silcone Elastomer Kit). 12. Digital scale and weigh dishes. 13. Vacuum desiccator chamber. 14. Hybridization oven. 15. Razor blades. 16. Hollow metal stainless steel cutting tube, 14 mm diameter (These can be designed and fabricated in most machine shops. Alternatively, a commercially available 12 or 14 mm biopsy punch could be used).

2.2 Hydrogel Microwell Fabrication

1. Biosafety cabinet. 2. 0.45 μm filter units. 3. 0.45 μm luer-lock syringe filter.

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4. PEG-VS monomer 8-arm 10 kDa (e.g., JenKem USA catalog # 8ARM(TP)-VS-10K). Store at 20  C. 5. PEG-SH monomer 4-arm 10 kDa (e.g., JenKem USA catalog # 4ARM-SH-10K). Store at 20  C. 6. Triethanolamine (TEA) buffer, 0.2 M pH 8.0 (e.g., SigmaAldrich catalog # T0449). 7. Ultrapure ddH2O. 8. 70% ethanol. 9. Sonicator. 10. Laminin, 0.5 mg/mL (e.g., Roche Diagnostics catalog # 11243217001). Stored at 20  C. 11. 1 PBS, pH 7.4. 12. Dialysis cassettes, 3 mL volume and >7 kDa molecular-weight cutoff (e.g., Thermo Fisher # 66370, 3 mL Slide-A-Lyzer dialysis cassette 10 kDa MWCO). 13. Magnetic stir plate. 14. 10% Bis-Tris gels, 1 mm thick (e.g., Invitrogen catalog # NP0301). 15. Antibiotic/antimycotic solution (e.g., Corning # 30-004-CI). 16. Hot plate. 17. Plain glass (not frosted) micro-slides, 2 in.  3 in.  1 mm (VWR catalog # 48382-180). 18. PTFE sheets, 0.8 or 1.6 mm (Alta Aesar catalog # AA45151HB or 45194-HB). 19. PTFE-coated spatula (Ted Pella catalog # 13523). 20. PTFE-coated tweezers (Ted Pella catalog # 5002-35 and 5002-37). 21. Plastic-ware: 35 mm dishes, 60 mm dishes, 10 cm dishes, 15 cm dishes, 1.7 mL Eppendorf tubes, 15 mL tubes. 22. Syringes. 2 mL and 3 mL. 23. Needle, 18G. 24. Aluminum foil. 2.3 Muscle Stem Cell Culture and Immunostaining

1. Humidified 37  C cell-culture incubator. 2. Tissue-culture-treated 24-well plates. 3. Myogenic growth medium: 42% DMEM low-glucose, 42% Ham’s F10 Nutrient Mix, 15% FBS, 1% Penicillin/Streptomycin, and 2.5 ng/mL FGF2. 4. p38 MAP kinase inhibitor SB202190. 5. Time-lapse microscope with motorized stage, environmental chamber, and appropriate acquisition software. Recommend

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phase-contrast or Hoffman contrast objectives at 10–20 magnification. 6. Cell counting, segmentation, and tracking software (e.g., ImageJ, CellProfiler, or Baxter Algorithm for Matlab). 7. Paraformaldehyde. 8. Blocking buffer: 20% goat serum and 0.5% Triton X-100 in PBS. 9. Staining buffer: 0.5% BSA and 2.5 mM EDTA in PBS. 10. Laminin antibody (e.g., Millipore # 05-206). 11. BSA conjugated with AlexaFluor 555 (e.g., Thermo Fisher # A34786). 12. Myogenin antibody (e.g., Santa Cruz # sc576). 13. Pax7 antibody (e.g., Santa Cruz # sc81648). 14. AlexaFluor 488-conjugated donkey anti-rabbit antibody (e.g., Jackson Immuno Research Laboratories # 711-545-152). 15. AlexaFluor 647-conjugated donkey anti-mouse IgG1 antibody (e.g., Jackson Immuno Research Laboratories # 715-605-150). 16. Topro-3 (e.g., Thermo Fisher # T3605).

3

Methods This protocol can be split into two stages, with the wafer and PDMS preparation steps (Subheadings 3.1 and 3.2) preceding the PEG hydrogel fabrication and MuSC cultures Subheadings 3.3–3.9.

3.1 Fabricating Patterned Silicon Wafers

The photolithography required to this protocol can be conducted in an academic or commercial micro/nanofabrication facility with the help of trained staff. Given the small scale of these features, a high-grade clean room facility is required for this work. Photolithography requires the use of key equipment such as a contact aligner, an etching machine, and a photoresist stripping machine (etching and stripping instruments are needed if using SU-8 instead of SUEX) [6]. Photolithographic patterns are created through the use of a chrome mask, a glass pane that has the pattern etched on it to suit the type of photoresist being used (positive or negative). This is a simplified tutorial based on the use of negative photoresist SUEX. Please consult with an expert in microfabrication practices for understanding how to use all the tools and materials needed for this process. 1. Design a chrome mask to imprint a pattern into silicon wafers (see Note 3). Generate a simple staggered pattern of dots using

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a computer-aided design software such as L-Edit (Fig. 1a). Chrome mask writers can invert designs to accommodate for the use of the opposite type of photoresist. 2. Dehydrate silicon wafers in an oven overnight at 95  C. 3. Coat silicon wafers with photoresist. Laminate SUEX (available at defined thicknesses) over wafer using hot rollers at 65  C with spinning speeds adjusted to SUEX thickness per manufacturer’s instructions. Use a 200 μm thick coating of SUEX to achieve ~150 μm deep features. 4. Heat wafer at 65  C for at least 15 min. 5. Develop wafers using a contact aligner. Exposure strength and duration will vary depending on photoresist thickness. For 200 μm thick SUEX, develop for 10 min at 1700 mJ/cm2 exposure with the use of a “365L” filter. Expose in 20 s intervals punctuated by 10 s rest periods. 6. Place wafer on a hot plate at room temperature and then set to 65  C. Once the plate reaches 65  C, bake for a total of 5 min. During this step, set another hot plate to 95  C. 7. Transfer the wafer from the 65  C plate to the 95  C plate and bake it for 15 min. Then, turn off the hot plate and allow the wafer to gradually cool to room temperature. 8. After a sufficient cooling time (usually 1 h), remove the unpolymerized photoresist from the wafer. Submerge the wafer, patterned-side down, in the photoresist solvent EBR until features appear. For 200 μm thick SUEX, this usually takes 15–30 min. Then, transfer the wafer into a fresh EBR solution bath. 9. Remove the excess EBR solution by submerging the wafer in a bath of isopropanol for 10 min. 10. Treat silicon wafers with a passivation coating to protect them during the PDMS casting process. Use a molecular vapor deposition instrument to coat the wafer with a layer of perfluorooctyltrichlorosilane (FOTS) (see Note 4). Wafer should appear as in Fig. 1b. 11. Characterize the “microwell” features on the coated silicon wafer using a profilometer, set to a horizontal scanning speed of 1000 μm/s, to ensure desired feature pattern and depth were achieved (Fig. 1c). 3.2 Making PDMS Stamps

1. Secure the silicon wafer, patterned-side-up, to a 15 cm cellculture dish by taping the perimeter to the dish bottom. Clean the silicon wafer surface with compressed air to remove debris from the features. 2. Prepare the PDMS solution. Mix the PDMS pre-polymer with the curing agent at a 10:1 weight ratio in a disposable weighing

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dish. Use a clean metal spatula to vigorously mix for 10 min. If the silicon wafer is mounted in a 15 cm dish, 30–35 g of PDMS solution is required per wafer (see Note 5). 3. For best results, place the mixed PDMS in a vacuum desiccator chamber for 10 min to remove excess air bubbles. 4. After mixing and degassing, carefully pour the PDMS solution to at least 2 mm thickness over the wafer, avoiding any sudden movement. 5. After pouring, degas the PDMS again in the vacuum desiccator chamber for 40–60 min. 6. Cure the PDMS on the wafer at 70  C overnight in an oven (see Note 6). Then, allow the PDMS and wafer to cool to room temperature. 7. Remove the cured PDMS from the wafer very carefully to reduce wear on the wafer and minimize the possibility of wafer cracking. Use a razor blade to cut the PDMS safely around the perimeter of the silicon wafer and then pry the PDMS off the wafer with a metal spatula. Removed PDMS should be stored feature-side down in another 15 cm dish and can be kept covered at room temperature for up to 3 months. 8. Cut PDMS stamps using a 14 mm diameter hollow metal stainless steel cutting tube. It is recommended that 4 stamps and 2 “spacer strips” (typically 5 mm  30 mm rectangles) are cut from the same area of PDMS (see Note 6). Spacer strips can be manually cut using a razor blade. 3.3 Preparing PEG Monomers

Fabricating microwell hydrogel substrates for cell culture requires the use of a biosafety cabinet and sterile materials and procedures. All solutions should be prepared in 0.45 μm-filtered buffers to minimize particulate debris and preserve sterility. Reagents should be stored at the recommended temperatures. Frozen solutions should be gradually thawed and store on ice. This is especially critical for the PEG monomer solutions before the synthesis reaction. All material transfer into and out of the biosafety cabinet should be in covered, sterile containers or preceded by thorough cleaning and sterilization with 70% ethanol solutions. After use, excess reagents should be discarded in chemical or biohazardous waste as appropriate. We recommend synthesizing hydrogel microwell substrates in sets of four 24-well-size hydrogel discs, and the following instructions (Subheadings 3.3–3.8) describe fabricating one set of 4 discs. 1. Dilute 8-arm 10 kDa PEG-VS monomer to 100 mg/mL in sterile-filtered 0.2 M triethanolamine (TEA) buffer (see Note 7). Weigh 100 mg PEG-VS in a sterile 1.7 mL tube (see Note 8). Transfer to a biosafety cabinet. Add 920 μL sterile

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TEA buffer to bring mixture to a total volume of 1000 μL. Tap and invert the tube to mix and vortex for 10 s until dissolved (see Note 9). After dissolved, keep on ice. 2. Transfer to a 3 mL syringe, then attach a 0.45 μm sterile filter and pass the solution through a filter to a new tube. Aliquot at 130 μL in 1 mL tubes. Store aliquots at 20  C, protected from light (see Note 10). 3. Dilute 4-arm 10 kDa PEG-SH monomer to 100 mg/mL in sterile-filtered ddH2O. Weigh 100 mg PEG-SH in a sterile 1.7 mL tube (see Note 8). Transfer to a biosafety cabinet. Add 920 μL sterile ddH2O to bring mixture to a total volume of 1000 μL. Tap and invert the tube to mix and vortex for 10 s until dissolved (see Note 9). After dissolved, keep on ice. 4. Transfer to a 3 mL syringe, then attach a 0.45 μm sterile filter and pass the solution through filter to a new tube. Then, aliquot at 260 μL in 1 mL tubes. Store aliquots at 20  C, protected from light (see Note 10). 3.4 Preparing Proteins for MicroContact Printing

All the following procedures using PEG solutions should be conducted in a biosafety cabinet. Protein solutions for micro-contact printing need to be prepared without Tris or any other buffer reagents containing primary amines (which will compete with the Michael addition) and at pH 7.4–8.0. Therefore, some protein solutions will require dialysis prior to use. We typically use laminin, a protein found in the basal lamina of the skeletal muscle stem cell niche [14]. 1. Thaw undialyzed laminin at 4  C overnight prior to dialysis. 2. Place 6 L of sterile PBS in a 4  C cold room. 3. Prepare a dialysis cassette and buffers. Wet the dialysis cartridge in 1–1.5 L of PBS.in a 2 L beaker with stir bar on a magnetic stir plate in the cold room. 4. Transfer laminin using a 2 mL syringe fitted with a 18G needle into the 10 kDa MWCO dialysis cassette. After injecting laminin, use the syringe to remove excess air from the cassette. Seal the used entry port with tape. Float the dialysis cassette in 1.5 L PBS for 1 h with gentle stirring (approximately 600 RPM) and cover with aluminum foil. 5. After 1 h, discard the used PBS and replace with 1.5 L fresh sterile PBS, re-cover with foil and dialyze overnight. The following day, replace the PBS with fresh PBS once again for 1 h. 6. Remove the dialyzed laminin using a sterile 2 mL syringe fitted with a 18G needle and aliquot into sterile 1.7 mL tubes (see Note 11). Store at 4  C for up to 2 months.

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3.5 Preparing Acrylamide Gels

For the micro-contact printing procedure, acrylamide gels are used to transfer proteins to PDMS stamps. We use prefabricated 10% Bis-Tris gels at 1 mm thickness but custom recipes can be used. 1. Remove 10% Bis-Tris gel from its casting cartridge. Cut off comb, cut into 4 equal “quarters,” and transfer into a sterile 15 cm dish. 2. Remove the Tris buffer through three washes with sterile PBS, each with at least 1 h incubation time. 3. Store buffed-exchange acrylamide gel “quarters” in sterile PBS with 1% antibiotic/antimycotic at 4  C for up to 2 months.

3.6 Preparing PDMS Stamps

1. Clean the feature-side (posts) of four circular PDMS stamps (cut to 13 mm diameter) with clear adhesive tape to remove debris and dust. Gently push an unused portion of the tape onto the post surface twice. 2. Vigorously spray four PDMS stamps and two spacers with 70% ethanol two times and shake out excess ethanol (see Note 12). Place stamps and spacers in an open 60 mm tissue culture dish feature-side up. Fill dish with enough 70% ethanol to cover all stamps and let them sit for 15 min to sterilize. 3. Remove all the ethanol and let stamps to dry to feature-size up and then cover (see Note 13).

3.7 Protein Transfer to PDMS Stamps

1. Set a hot plate to 65  C. 2. Collect one pre-washed Bis-Tris gel “quarter” per four PDMS stamps. Transfer from antibiotic storage solution and place into a 60 mm tissue culture dish. 3. Wash Bis-Tris gel with sterile PBS twice and then aspirate all PBS. Dry uncovered in biosafety cabinet for 5 min. 4. Prepare protein coating solution (see Note 14). Dilute 33 μL dialyzed laminin in 67 μL sterile PBS to ~100 mg/mL final concentration in 100 μL per gel-quarter. Store at room temperature for 15 min while drying Bis-Tris gel. 5. Place the 60 mm dish containing a Bis-Tris gel quarter on the 65  C hot plate with the dish lid slightly ajar. Let the gel dry until it begins to slightly curl at the sides (usually within 10–15 min) (see Note 15). Remove the dish, place in the biosafety cabinet, and let it cool to room temperature for 15 min (see Note 16). 6. Pipet 70 μL of the laminin solution onto the top of the partially dried Bis-Tris gel-quarter drop wise. Make sure to distribute the drops evenly over the gel-quarter, and then gently spread the protein solution over the entire gel-quarter with a pipette tip or a clean spatula. 7. Let the gel-quarter dry 15–25 min until no liquid remains visible on its surface (see Notes 17 and 18).

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8. Place the four PDMS stamps feature-side down onto the justdry Bis-Tris gel quarter. Use toothed tweezers to grab and place the stamp with the features facing down and do not disturb. The stamp posts should become visibly darker when they have contacted the gel surface but the inter-post areas should remain more clear. If the posts do not appear darker, softly push the stamp down with the tweezers until a change is observed. 9. Leave stamps on the protein-coated gel for 30 min to allow protein transfer. The hydrogel microwell polymerization system consists of a “sandwich” system comprised of: two 2 in.  3 in. glass micro-slides; two “spacers” each consisting of PTFE and PDMS pieces; four proteincoated PDMS stamps with PEG gel solutions added to the top; and two binder clip clamps (see Fig. 1d). We typically prepare a solution containing 3.35 wt%/vol final PEG concentration to achieve at 12 kPa Young’s modulus hydrogel, which is optimal for MuSC self-renewal [3, 4, 15]. Here, we describe PEG synthesis for use on 13 mm stamps with a 1.6 mm PTFE spacer. The PEG volume should be changed if the stamp or spacer size is changed. Tuning the hydrogel to a softer or stiffer modulus is possible by changing the total PEG concentration (see Note 20). See Table 1 with recipes for other Young’s moduli.

3.8 Microcontact Printing of Proteins into PEG Microwells

1. After the protein transfer to PDMS starts, thaw PEG-VS and PEG-SH aliquots on ice. 2. After the protein transfer to PDMS completes, set up the PEG hydrogel polymerization sandwich. Place one glass slide on a closed 35 mm dish and put one PDMS spacer on each narrow side. Spray two PTFE spacers to make them “sticky” to the PDMS and set them gently place on the top of each of the two stamps (see Note 19). Table 1 Formulations for PEG hydrogels with varying weight percentages for tuning hydrogel Young’s modulus. These formulations are sufficient for four 220-μL hydrogels made at 24-well-size (12.7 mm diameter and 1.6 mm thickness) Young’s modulus (kPa)

Total PEG (wt%/vol)

Total volume for 4 gels (μL)

TEA (μL)

100 mg/mL PEG-SH (μL)

100 mg/mL PEG-VS (μL)

10

3.19

891.8

607.3

189.7

94.8

12

3.35

891.8

593.1

199.2

99.6

20

3.88

891.8

545.8

230.7

115.3

30

4.35

891.8

503.9

258.6

129.3

60

5.29

891.8

420.0

314.5

157.2

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3. Carefully transfer four protein-coated PDMS stamps with tweezers and place them on the glass side feature-side up. Gently push the edges of the PDMS stamps down to secure their flat sides to the glass. Quickly proceed to prepare PEG hydrogel synthesis solution sufficient for four stamps. 4. Prepare gel solutions by adding 100 mg/mL PEG-SH and 100 mg/mL PEG-VS solutions into TEA buffer. To make enough PEG solution for four stamps, add 593 μL TEA buffer, 199 μL PEG-SH, and then 100 μL PEG-VS to make a 892 μL final PEG solution. Return the mixture and unused reagents to ice as quickly as possible (see Note 20). 5. To mix the gelling solution, lightly vortex the tube for 5 s, then return it to ice for 5 s and repeat 4. 6. Carefully pipet 200–220 μL of the mixed PEG solution to the top of each PDMS stamp. The gel should form a bead on the top of the stamps. 7. Clean the top glass slide quickly with 70% ethanol and place gently on the top of the gel setup. Carefully secure both the narrow ends of the slide sandwich using binder clips, centering the clip in the middle of each spacer (see Note 21). 8. Close the binder clip handles and place the gel system within a 15 cm tissue-culture dish (see Fig. 1d). Close the lid and place inside a humidified 37  C cell-culture incubator. 9. Incubate for ~18–36 h to complete the PEG polymerization process. Stop when the PEG hydrogels have mostly receded from the PDMS stamps and an air gap can be seen. 10. Transfer back to the biosafety cabinet, dissemble the sandwich setup, and remove each gel will a spatula. 11. Use PBS to help the PEG hydrogel disc dissociate from the PDMS stamp. Transfer the PEG hydrogel discs to separate wells of a 6-well plate containing 2 mL of sterile PBS per well for 1 h to rehydrate (see Note 22). 12. Adhere hydrogel discs to a 24-well tissue-culture plate by adding a small amount of PEG solution under each gel. Remove all PBS from hydrogel disc storage plate. 13. Prepare a PEG hydrogel solution for gluing 4 hydrogel discs as in Subheading 3.9, steps 1–4 above. Add 39.9 μL TEA buffer, 13.4 μL PEG-SH, and then 6.7 μL PEG-VS to make a 60 μL final PEG “gluing” solution. Vortex as in Subheading 3.9, step 6. 14. Pipet 12 μL of the “gluing” solution to the bottom surface of each culture well and smear evenly with a bent plastic pipette tip. Move quickly to complete this for 4 wells in 37  C) Bis-Tris gel will cause protein denaturing and/or degradation. 17. This is a critical step. If the stamp is placed on the gel-quarter with excess liquid remaining, protein will coat the bottom and sides of the PDMS posts and thus the hydrogel will have protein on all surface features, not only bottom of the microwells. 18. Test the dryness of the gel-quarter by pushing a 200 μL pipette tip onto its surface to make a halo mark. If the mark does not fill in with remaining protein solution, the gel-quarter is sufficiently dry. 19. The thickness of the PTFE spacer determines the thickness of the gel. Switching the spacer thickness requires adjusting the PEG gel solution volume per stamp to achieve the desired diameter. 20. Given that the 8-arm PEG-VS contains twice as many reactive groups as the 4-arm PEG-SH per μL aliquot volume, twice as much PEG-SH monomer solution is added to the PEG reaction. 21. Make sure the sandwich system remains flat as tipping will allow the gel to flow off the stamps. 22. Optionally: Inspect the quality of each microwell hydrogel using a phase-contract microscope and select hydrogel discs with the most consistent microwell pattern for gluing and use in a culture experiment. 23. Normoxia and hypoxia studies could use other oxygen concentrations. 24. It can be helpful to record cell cultures from 24 to 36 h postseeding to identify motile, viable cells and select ~100 viable cells for long-term time-lapse acquisition.

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Acknowledgments This work was supported by NIH grant R00AG042491 (to B.D. C.). We would like to acknowledge helpful discussions with Helen Blau, Penney Gilbert, Karen Havenstrite, and Matthias Lutolf, who were instrumental in developing this technology, and the advice of the staff technicians at the Cornell NanoScale Science and Technology Facility (CNF), a shared national user facility funded in part by the NSF-NNCI program and New York State. References 1. Cosgrove BD, Sacco A, Gilbert PM, Blau HM (2009) A home away from home: challenges and opportunities in engineering in vitro muscle satellite cell niches. Differentiation 78 (2–3):185–194. doi:10.1016/j.diff.2009.08. 004 2. Tierney MT, Sacco A (2016) Satellite cell heterogeneity in skeletal muscle homeostasis. Trends Cell Biol 26(6):434–444. doi:10. 1016/j.tcb.2016.02.004 3. Gilbert PM, Havenstrite KL, Magnusson KE, Sacco A, Leonardi NA, Kraft P, Nguyen NK, Thrun S, Lutolf MP, Blau HM (2010) Substrate elasticity regulates skeletal muscle stem cell self-renewal in culture. Science 329 (5995):1078–1081. doi:10.1126/science. 1191035 4. Cosgrove BD, Gilbert PM, Porpiglia E, Mourkioti F, Lee SP, Corbel SY, Llewellyn ME, Delp SL, Blau HM (2014) Rejuvenation of the muscle stem cell population restores strength to injured aged muscles. Nat Med 20 (3):255–264. doi:10.1038/nm.3464 5. Khademhosseini A, Langer R, Borenstein J, Vacanti JP (2006) Microscale technologies for tissue engineering and biology. Proc Natl Acad Sci U S A 103(8):2480–2487. doi:10.1073/ pnas.0507681102 6. Bernard A, Renault JP, Michel B, Bosshard HR, Delamarche E (2000) Microcontact printing of proteins. Adv Mater 12(14):1067–1070. doi:10.1002/1521-4095(200007) 12:143.0.Co;2-M 7. Wang Y, Shah P, Phillips C, Sims CE, Allbritton NL (2012) Trapping cells on a stretchable microwell array for single-cell analysis. Anal Bioanal Chem 402(3):1065–1072. doi:10. 1007/s00216-011-5535-9 8. Lutolf MP, Blau HM (2009) Artificial stem cell niches. Adv Mater 4(21):32–33. doi:10.1002/ adma.200802582

9. Berkowski KL, Plunkett KN, Yu Q, Moore JS (2005) Introduction to photolithography: preparation of microscale polymer silhouettes. J Chem Educ 82(9):1365–1369 10. Lorenz H, Despont M, Fahrni N, LaBianca N, Renaud P, Vettiger P (1997) SU-8: a low-cost negative resist for MEMS. J Micromech Microeng 7(3):121–124. doi:10.1088/0960-1317/ 7/3/010 11. Johnson D, Voigt A, Ahrens G, Dai W (2010) Thick epoxy resist sheets for MEMS manufactuing and packaging. Proc IEEE Micr Elect: pp. 412–415 12. Lutolf MP, Hubbell JA (2003) Synthesis and physicochemical characterization of end-linked poly(ethylene glycol)-co-peptide hydrogels formed by Michael-type addition. Biomacromolecules 4(3):713–722. doi:10.1021/ bm025744e 13. Magnusson KE, Jalden J, Gilbert PM, Blau HM (2015) Global linking of cell tracks using the Viterbi algorithm. IEEE Trans Med Imaging 34(4):911–929. doi:10.1109/TMI.2014. 2370951 14. Yin H, Price F, Rudnicki MA (2013) Satellite cells and the muscle stem cell niche. Physiol Rev 93 (1):23–67. doi:10.1152/physrev.00043.2011 15. Blau HM, Cosgrove BD, Ho AT (2015) The central role of muscle stem cells in regenerative failure with aging. Nat Med 21(8):854–862. doi:10.1038/nm.3918 16. Sacco A, Doyonnas R, Kraft P, Vitorovic S, Blau HM (2008) Self-renewal and expansion of single transplanted muscle stem cells. Nature 456(7221):502–506. doi:10.1038/ nature07384. nature07384 [pii] 17. Gobaa S, Hoehnel S, Roccio M, Negro A, Kobel S, Lutolf MP (2011) Artificial niche microarrays for probing single stem cell fate in high throughput. Nat Methods 8 (11):949–955. doi:10.1038/nmeth.1732

Chapter 7 Isolation, Culture, and Differentiation of Fibro/Adipogenic Progenitors (FAPs) from Skeletal Muscle Robert N. Judson, Marcela Low, Christine Eisner, and Fabio M. Rossi Abstract Fibro/Adipogenic Progenitors (FAPs) are a multipotent progenitor population resident in skeletal muscle. During development and regeneration, FAPs provide trophic support to myogenic progenitors that is required for muscle fiber maturation and specification. FAPs also represent a major cellular source of fibrosis in degenerative disease states, highlighting them as a potential cellular target for anti-fibrotic muscle therapies. Effective and reproducible methods to isolate and culture highly purified FAP populations are therefore critical to further understand their biology. Here, we describe a fluorescent activated cell sorting (FACS) based protocol to isolate CD31 /CD45 /Integrin-α7 /Sca1+ FAPs from murine skeletal muscle including details of tissue collection and enzymatic muscle digestion. We also incorporate optimized methods of expanding and differentiated FAPs in vitro. Together, this protocol provides a complete workflow to study skeletal muscle derived FAPs and compliments downstream analytical, drug screening, and disease modeling applications. Key words FAPs, Fibro-adipogenic progenitors, Skeletal muscle, MSCs, Mesenchymal progenitors, Myogenesis, Skeletal muscle regeneration, Satellite cells, Fibrosis

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Introduction Fibro-Adipogenic Progenitors (FAPs) are a multipotent progenitor population resident in skeletal muscle. Originally defined by their bi-potent adipogenic and fibrogenic differentiation potential [1–3], FAPs share functional and phenotypic characteristics with mesenchymal stem/stromal cells (MSCs) found in almost all mammalian tissues [4]. Unlike somite-derived myogenic progenitors that are the principal precursors of mature skeletal muscle myofibers, FAPs play a critical supportive role in myogenesis. During development, FAPs provide a source of extracellular matrix (ECM) proteins needed for the anatomical organization of muscle groups and are critical for myofiber maturation and specification [5]. In the

Robert N. Judson and Marcela Low contributed equally to this work. James G. Ryall (ed.), Skeletal Muscle Development, Methods in Molecular Biology, vol. 1668, DOI 10.1007/978-1-4939-7283-8_7, © Springer Science+Business Media LLC 2017

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adult, FAPs are largely quiescent but become activated upon skeletal muscle damage and provide trophic support for muscle stem cells (satellite cells) during regeneration [1, 6]. Evidence suggests FAPs also play a significant role in tissue degeneration, representing the major cellular source of fibrosis, fibrofatty infiltrate, and ossification [1, 3, 7] in a number of diseases. As a result, FAPs have become an obvious potential therapeutic target for reparative disorders, including muscular dystrophies. Successful isolation and culture of highly purified FAPs from skeletal muscle therefore represents an important method for developmental biology, drug screening, and disease modeling involving these cells. Prospective isolation by fluorescent activated cell sorting (FACS) has become the gold standard method of purifying distinct cellular subsets from heterogenous populations. FACS allows cells to be separated based on size, granularity, and most importantly, fluorescence emission using fluorophore-conjugated antibodies. In this chapter, we provide our group’s latest protocol to isolate FAPs from murine skeletal muscle. We define FAPs based on a cell surface antigen profile of CD31 /CD45 /Integrin-α7 /Sca1+. This multipotent progenitor population is distinct from satellite cells and is capable of fibroblastic, adipogenic, chondrogenic, and osteogenic differentiation. We also provide a detailed protocol of how to expand FAPs in vitro following isolation and induce tri-lineage differentiation. Together, this protocol provides a complete workflow to study skeletal muscle derived FAPs and compliments downstream analytical applications to further understand their biology and therapeutic potential.

2 2.1

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1. Mice: 8–12 week old mice (see Note 1). 2. 70% (v/v) Ethanol. 3. Forceps. 4. Scissors. 5. Sterile Ca2+/Mg2+ free phosphate-buffered saline (PBS). 6. Sterile tissue culture dishes (6-well plates, 35 mm dishes, or 60 mm dishes depending on the amount of muscle being collected). 7. Collagenase II (250 CDU/mL). 8. Calcium Chloride (250 mM). 9. 37  C warm room, incubator, or water bath (see Note 2). 10. Sterile 1 mL syringes. 11. Sterile 15 and 50 mL conical tubes.

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12. Transfer pipettes. 13. Benchtop centrifuge. 14. Collagenase D (1.5 U/mL)/Dispase II (2.4 U/mL). 15. Benchtop vortex. 16. 40 μm cell strainers. 17. FACS Buffer: Ca2+/Mg2+-free PBS supplemented with 2% fetal bovine serum (FBS) and 2 mM EDTA (pH 7.9). 2.2 Antibody Staining

1. FACS Buffer, as above. 2. Sterile 15 and 50 mL conical tubes. 3. 1.5 mL microcentrifuge tubes. 4. Benchtop centrifuge. 5. Microcentrifuge. 6. Experimental antibody solution: FITC-conjugated rat antimouse CD31 (eBioscience, clone: 390)—1:500, Alexa Fluor 488-conjugated rat anti-mouse CD45 (AbLab, clone: I3/2)— 1:1000, PeCy7-conjugated rat anti-mouse Sca-1 (eBioscience, clone: D7)—1:5000, Alexa Fluor 647-conjugated rat antimouse integrin alpha7 (AbLab, clone: R2F2)—1:1500. 7. Propidium iodide (PI) solution. 8. Hoechst 33342 (H) solution. 9. Sterile 5 mL FACS tubes with 40 μm cell strainer cap. 10. Sterile 5 mL polystyrene FACS tubes. 11. FAP expansion media (described below). 12. Cell sorter.

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1. FAP expansion media: Dulbecco’s modified eagle medium (DMEM) supplemented with 10% FBS, 2.5 ng/mL Basic fibroblast growth factor (bFGF) and Penicillin 10,000 U/ mL: Streptomycin 10,000 μg/mL. 2. Adipogenic media: Dulbecco’s modified eagle medium (DMEM), supplemented with 10% FBS, 0.25 μM dexamethasone, 0.5 mM isobutylmethylxanthine, 1 μg/mL insulin, 5 μM troglitazone and Penicillin 10,000 U/mL and Streptomycin 10,000 μg/mL. 3. Osteogenic media: Dulbecco’s modified eagle medium (DMEM) (Invitrogen) supplemented with 10% FBS, 10 nM dexamethasone, 5 mM β-glycerophosphate, 50 μg/mL ascorbic acid and Penicillin 10,000 U/mL and Streptomycin 10,000 μg/mL.

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4. Chondrogenic media: Dulbecco’s modified eagle medium (DMEM), supplemented with 10% FBS, 1 ng/mL TGFβ-1 and 50 μg/mL ascorbic acid. 5. Benchtop centrifuge. 6. Sterile culture plates (see Note 3). 7. Sterile 15 mL conical tubes. 8. Ca2+/Mg2+-free PBS. 9. Hemocytometer. 10. Class II biosafety cabinet. 11. Tissue culture incubator (humidified, 37  C, 5% CO2).

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Methods All the steps should be performed on ice unless otherwise specified.

3.1 Muscle Dissection

1. Complying with an approved institutional protocol, euthanize mice at 8–12 weeks of age. Place mice on paper towel and spray liberally with 70% (v/v) ethanol to disinfect and reduce fur contamination. 2. Using forceps and scissors make a small abdominal incision through the skin, being careful not to breach the peritoneum. 3. Remove skin from the hind legs using forceps to pull down the skin from the abdominal incision, exposing the hind limb muscles. 4. Using scissors and forceps, use blunt dissection technique to remove whole muscle groups (quadriceps, tibialis anterior, posterior tibial muscles, etc.) from around the tibial and femoral bones. Repeat the process for both legs, liberating muscles as closely as possible to their tendinous connections (see Note 4). Separate muscle groups into 6-well plates, 35 mm dishes, or 60 mm dishes depending on the sample size and number of mice harvested. Keep the plates on ice. 5. For single, unstained and fluorescence minus one (FMO) controls prepare additional tissue as a “control sample.” Use one muscle group from a separate mouse.

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3.2.1 Two-Step Digestion

Muscle digestion can be performed using either a two-step or onestep protocol (see Note 5). 1. Using two pairs of forceps, tear the tissue along the muscle fibers until the muscle is in small pieces. Gentle tearing of the muscle is required for maintaining sample quality and ensuring cell survival.

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2. Prepare activated Collagenase II and Collagenase D/Dispase II by adding 10 μL/mL of 250 mM calcium chloride solution. Keep prepared solutions on ice until use. 3. First digestion: Add previously prepared Collagenase II. The volume of enzyme necessary will depend on the number of samples and size of plate used (see Note 6). 4. Incubate the plate at 37  C for 30 min. 5. After incubation, use a syringe plunger to gently yet thoroughly mash the tissue until there are no longer any large chunks of muscle (see Note 7). 6. Add ~1–5 mL of sterile ice-cold PBS to each dish and using transfer pipettes, transfer the tissue sludge into either a 15 mL conical tube for separate muscle groups, or a 50 mL conical tube for the whole leg. Rinse the dish as necessary to recover all the tissue. Keep the tubes on ice. 7. Top up the tubes with sterile ice-cold PBS and centrifuge at 130  g for 5 min at 4  C. 8. Discard the supernatant and proceed immediately to the second digestion. 9. Second digestion: Resuspend the tissue pellet in previously prepared Collagenase D/Dispase II solution (see Note 6). 10. Incubate the tubes at 37  C for 1 h with gentle rotation. To help ensure even digestion, it is recommended that at 15 min intervals, samples are vortexed at low speed for 8–10 s. 11. Add ~5–10 mL ice-cold FACS buffer (15 mL tube) or 20–30 mL (50 mL tube) and titurate using Pasteur pipettes. Transfer the tubes to ice. 12. Place a 40 μm cell strainer on the top of a 50 mL conical tube and filter the suspension, keeping the tubes on ice (see Note 8). 13. Fill the tubes with FACS buffer and centrifuge at 515  g for 5 min at 4  C. For large samples, such as a whole leg, divide the filtrate into three separate 50 mL conical tubes and top each tube up to 50 mL to wash the samples more efficiently. 14. Discard the supernatant and immediately proceed to antibody staining. 3.2.2 One-Step Digestion

Alternatively, you can use the following protocol for digestion (see Note 5). 1. Transfer dissected muscles into a culture dish of appropriate size. 2. Using scissors, cut the samples into 2–4 mm pieces.

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3. Transfer the pieces into either a 15 mL conical tube for individual muscles groups or a 50 mL conical tube for the whole leg. 4. Prepare activated Collagenase D/Dispase II by adding 10 μL/ mL of 250 mM calcium chloride solution. Keep prepared solution on ice until use. 5. Add previously prepared Collagenase D/Dispase II solution using 350–600 μL for individual muscle groups or 3 mL for both legs of a mouse. 6. Perform a quick spin of the tubes (a few seconds at low speed) to draw tissue into the enzyme solution. 7. Incubate the tubes at 37  C for 1 h with gentle rotation. To help ensure even digestion, it is recommended that at 15 min intervals, samples are vortexed at low speed for 8–10 s. 8. Top up the tubes with ice-cold PBS and vortex or titurate using a transfer pipette to thoroughly mix the sample. 9. Using a transfer pipette, filter cells through 40 μm cell strainer placed on the top of a 50 mL conical tube (see Note 9). 10. Top up each tube with ice-cold PBS and centrifuge at 515  g for 7 min at 4  C. 11. Discard the supernatant and proceed immediately to antibody staining. 3.3 Antibody Staining and Cell Sorting

1. For the controls: resuspend pellet from the “control sample” using 1 mL of FACS buffer. Label 9 sterile 1.5 mL microcentrifuge tubes for unstained, single color (Hoechst, PI, FITC, PeCy7, and APC), and fluorescence minus one (FMO for FITC, PeCy7, and APC) controls. Aliquot 100 μL of cells into each tube. These samples will be used for compensation and as gating controls. 2. Add appropriate antibodies to each tube. Cover samples from light and incubate for 30 min on ice. 3. For the experimental sample, prepare the experimental antibody solution in FACS buffer. 4. Resuspend and combine pellets from multiple muscle groups in 1 mL of experimental antibody solution (see Note 10). Mix thoroughly, cover samples from light, and incubate for 30 min on ice. 5. Wash control samples by adding 500 μL FACS buffer to each tube and centrifuging at 515  g for 10 min at 4 ˚C. 6. Wash experimental samples by adding 20–40 mL of FACS buffer to the tube and centrifuging at 515  g for 10 min at 4 ˚C.

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7. Discard the supernatant and resuspend pellets from control samples in 500 μL of FACS buffer containing PI (1:1000) and Hoechst (1:500). 8. Resuspend pellets from experimental sample in at least 1–3 mL of FACS buffer containing PI (1:1000) and Hoechst (1:500). 9. To ensure the sample does not contain cell clumps or debris, perform a final filtration of cells using a 5 mL FACS tube with 40 μm cell strainer lid. Proceed immediately to cell sorting. 10. For cell sorting, set up the cell sorter according to the manufacturer’s instructions. The unstained sample should be used to set correct voltages for each channel. Use single and FMO control samples to set compensation values. 11. Establish the gating strategy to sort FAPs. A typical gating strategy is shown in Fig. 1. Briefly, first, gate for live cells (PI , Hint) followed by CD31/CD45 and Integrin α7 lineage negative cells. From here, gating for the Sca1+ population captures FAPs. 12. Collect FAPs in 2 mL of FAP expansion medium in precooled 5 mL polystyrene tubes. Typical FAP yields from a single mouse are approximately 70,000–100,000 viable cells (see Note 11). 3.4 In Vitro Culture and Differentiation of FAPs

1. After cell sorting, pellet FAPs by centrifugation at 515  g for 10 min at 4 ˚C. 2. Working in a biosafety cabinet, use the sterile technique to carefully remove the supernatant and resuspend cells in a small volume of FAP expansion media (see Note 12). 3. Plate cells onto tissue culture treated dishes at a density of 10,000 cells/cm2. The number of cells can be obtained directly from the cell sorting machine or alternatively, counted using a hemocytometer (see Note 13). 4. Incubate cells in a humidified tissue culture incubator under standard growth conditions at 37 ˚C and 5% CO2. Change media every 48 h to maintain expansion (see Note 14). 5. For mesenchymal differentiation assays, allow cells to grow to 85–95% confluence and remove expansion media. Wash once with PBS and add adipogenic or osteogenic media (see Note 15). For chondrogenic differentiation, pellet 3–5  105 cultured FAPs in a 15 mL falcon tube by centrifugation at 510  g for 5 min and incubate in 1 mL chondrogenic media for 21 days under standard conditions at 37 ˚C and 5% CO2 (see Note 16).

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Fig. 1 Gating strategy for cell sorting of FAPs. FAPs from murine skeletal muscle are gated based on size and shape (FSC, SSC), viability (PI , H+), and cell surface markers (CD45 /CD31 /integrinɑ7 /Sca1+)

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Notes 1. We recommend 8–12 week old mice for maximum cell yield and ease of digestion. Older mice yield less cells and may have more fibrotic tissue in the muscle requiring additional digestion time. All protocols involving the use of rodents must adhere to all relevant institutional and regional guidelines. 2. A 37  C warm room, incubator or water bath should be equipped with some method to gently rotate or agitate the samples to ensure even tissue digestion. 3. Size of sterile tissue culture plates will vary depending on the number of cells collected and plating density. 4. Muscle groups should be dissected and separated into anterior tibial muscles: Tibialis anterior (TA), extensor digitorum longus (EDL). Posterior tibial muscles: Plantaris, gastrocnemius and soleus. Anterior femoral muscles: Quadriceps. 5. One-step digestion, used for interstitial cell isolation, has some advantages such as decreased processing time and increased cell yield. However, this method can also lead to increased debris and possible contamination of desired populations by other cell types. In this case, additional wash steps after enzymatic digestion help to remove debris and use of a viability dye is highly recommended to help gate out any remaining debris during cell sorting. When choosing which protocol to use (one step versus two step), consider the downstream applications; while cell yield may be important for certain assays, population purity may be much more crucial for other applications such as cell culture or RNAseq. 6. For single muscle groups in separate wells of a 6-well plate, use the following volumes: one TA (250 μL) or one Gastrocnemius (500 μL), for Collagenase II and Collagenase D/Dispase II. For 60 mm dishes containing both legs from one mouse, use 2 mL of Collagenase II and 1 mL of Collagenase D/Dispase II. Some modifications in volume of enzymes or digestion time also are allowed, e.g., fibrotic samples or aging muscles. 7. If after the first step digestion you still see some undigested muscles, try to gently dissociate them; however, do not expect full digestion at this step. 8. For a faster filtration, let the tubes sediment and wait until the pieces of muscles, still remaining in the tubes, go to the bottom. Filter just the top part. 9. If large chunks of muscle are observed at the end of this digestion, the sample can be filtered through a 70 μm cell strainer before being passed through the 40 μm filter.

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10. The volume of antibody cocktail should be adjusted to match the number of cells for optimal staining. When staining all hindlimb muscles use 1 mL of antibody cocktail per mouse and perform the staining in a 50 mL falcon tube. 11. Cell sorting can have negative effects on cell viability which may influence the quality of FAP cultures. Ensure cells are sorted into cooled media and remain on ice following the sort. Cells should be processed and plated in vitro as quickly as possible following sorting. 12. The volume of FAP expansion media used will depend on the size of the pellet and the desire plate density. 500 μL is typical. 13. Cells are initially seeded at high density to boost cell survival and improve the expansion. We have found seeding at lower cell densities can lead to cell death and poor initiation of FAP cultures. Also, cell density will depend on the source of your sample, damage muscle usually requires less cell density than undamaged muscle. 14. Passaging of FAPs is not recommended as extended culturing leads to loss of phenotypic characteristics and multipotency of cells. However, if large numbers of cells are required (e.g., for the analysis by Western Blot or IP), FAPs can be expanded for 1–2 passages. 15. For adipogenic differentiation, change media every 3–4 days until vacuoles are observed (7–14 days). Visualization of adipogenic differentiation can be detected by Oil Red O or Perilipin staining. For osteogenic differentiation, change media every 3–4 days until bone matrix formation is observed (up to 21 days). Visualization of osteogenic differentiation can be detected by Alizarin Red S or silver nitrate (von Kossa) staining. 16. For chondrogenic differentiation, perform half volume (500 μL) media changes every 3–4 days for at least 21 days. For the visualization of chondrogenic differentiation, chondrocytic pellets can be fixed, paraffin embedded and then sections stained with Alcian Blue and Nuclear Fast Red. References 1. Joe AWB, Yi L, Natarajan A, Le Grand F, So L, Wang J, Rudnicki MA, Rossi FMV (2010) Muscle injury activates resident fibro/adipogenic progenitors that facilitate myogenesis. Nat Cell Biol 12:153–163 2. Uezumi A, Fukada S-I, Yamamoto N, ‘ichi TS, Tsuchida K (2010) Mesenchymal progenitors distinct from satellite cells contribute to ectopic

fat cell formation in skeletal muscle. Nat Cell Biol 12:143–152 3. Uezumi A, Ito T, Morikawa D, Shimizu N, Yoneda T, Segawa M, Yamaguchi M, Ogawa R, Matev MM, Miyagoe-Suzuki Y, Takeda S’i, Tsujikawa K, Tsuchida K, Yamamoto H, Fukada S-I (2011) Fibrosis and adipogenesis originate from a common mesenchymal progenitor in skeletal muscle. J Cell Sci 124:3654–3664

FAP Isolation and Culture 4. da Silva Meirelles L, Chagastelles PC, Nardi NB (2006) Mesenchymal stem cells reside in virtually all post-natal organs and tissues. J Cell Sci 119:2204–2213 5. Mathew SJ, Hansen JM, Merrell AJ, Murphy MM, Lawson JA, Hutcheson DA, Hansen MS, Angus-Hill M, Kardon G (2010) Connective tissue fibroblasts and Tcf4 regulate myogenesis. Development 138:371–384 6. Fiore D, Judson RN, Low M, Lee S, Zhang E, Hopkins C, Xu P, Lenzi A, Rossi FMV, Lemos

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DR (2016) Pharmacological blockage of fibro/ adipogenic progenitor expansion and suppression of regenerative fibrogenesis is associated with impaired skeletal muscle regeneration. Stem Cell Res 17:161–169 7. Lounev VY, Ramachandran R, Wosczyna MN, Yamamoto M, Maidment ADA, Shore EM, Glaser DL, Goldhamer DJ, Kaplan FS (2009) Identification of progenitor cells that contribute to heterotopic skeletogenesis. J Bone Joint Surg Am 91:652–663

Chapter 8 Human Satellite Cell Isolation and Xenotransplantation Steven M. Garcia, Stanley Tamaki, Xiaoti Xu, and Jason H. Pomerantz Abstract Satellite cells are mononucleated cells of the skeletal muscle lineage that exist beneath the basal lamina juxtaposed to the sarcolemma of skeletal muscle fibers. It is widely accepted that satellite cells mediate skeletal muscle regeneration. Within the satellite cell pool of adult muscle are skeletal muscle stem cells (MuSCs), also called satellite stem cells, which fulfill criteria of tissue stem cells: They proliferate and their progeny either occupies the adult MuSC niche during self-renewal or differentiates to regenerate mature muscle fibers. Here, we describe robust methods for the isolation of enriched populations of human satellite cells containing MuSCs from fresh human muscle, utilizing mechanical and enzymatic dissociation and purification by fluorescence-activated cell sorting. We also describe a process for xenotransplantation of human satellite cells into mouse muscle by injection into irradiated, immunodeficient, mouse leg muscle with concurrent notexin or bupivacaine muscle injury to increase engraftment efficiency. The engraftment of human MuSCs and the formation of human muscle can then be analyzed by histological and immunofluorescence staining, or subjected to in vivo experimentation. Key words Human, Skeletal muscle, Stem cell, Satellite cell, Xenotransplantation

1

Introduction

1.1 Human Satellite Cells

Satellite cells and their subpopulation of skeletal muscle stem cells (MuSCs) are the principal mediators of adult mammalian skeletal muscle regeneration [1–5]. MuSCs repair muscle after injury via activation from a quiescent state, proliferation, and differentiation or undergo self-renewal to either repair damaged muscle or maintain the stem cell pool (reviewed in [6, 7]). Isolation of purified satellite cells was necessary to prove the identity of the populations of cells within skeletal muscle that have myogenic regenerative potential. Furthermore, transplantation of isolated cells is a standard experimental approach that leads to fundamental insights into muscle stem cell biology. Satellite cells were initially isolated from mice and eventually purified at the single-cell level [3, 4]. However, technical challenges of working with and difficulty in obtaining human tissue initially hampered progress of human muscle stem cell biology [8–20]. Thus, until recently, it was not feasible to study

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defined populations of endogenous human muscle cells to prove muscle stem cell function without passage in culture, which alters cellular properties and does not maintain the same representative populations of cells found in the body. Significant progress has been made in the last 2 years that has led to extensive characterization of human satellite cells and definitive isolation and transplantation of human satellite cells with stem cell function [21, 22]. Our laboratory has published the isolation of highly purified human satellite cells by flow cytometry, and has shown they fulfill criteria of stem cells [22]. Development of xenotransplantation protocols that support the production of human muscle fibers required sufficient immunodeficiency and crippling of endogenous host muscle stem cells to accept human xenografts, while preserving an environment capable of supporting regeneration by transplanted cells. In this chapter, we describe detailed protocols for the isolation of highly purified human satellite cells that express the canonical transcription factor paired box protein 7 (PAX7) using fluorescence-activated cell sorting (FACS) with surface markers CD56 and CD29 in addition to negative selection. We also describe methods for xenotransplantation of satellite cells into NOD scid gamma (NSG) immunodeficient mice, using adjuvant radiation and mild toxin injury. We demonstrate how xenografted muscles can then be secondarily injured to assess the regeneration capacity of human MuSCs in vivo. Finally, we provide protocols for the analysis of engrafted human MuSCs and the formation of human muscle by histological and immunofluorescence staining. 1.2 Experimental Design 1.2.1 Tissue Dissociation

1.2.2 FACS Preparation and Isolation

Human muscle tissue obtained from operating room biopsies range in composition of muscle, fat, tendon, connective tissue, and fascia. Muscle tissue itself is composed of myofiber bundles that contain large amounts of connective tissue. Efficient satellite cell isolation begins with muscle trimmed of contaminating tissues so it can then be processed with meticulous mechanical and enzymatic digestion to produce single-cell suspensions. Mastery of these steps is paramount if one hopes to achieve a high yield of isolated satellite cells from whole muscle. Magnetic column separation with metal microbead-conjugated antibodies greatly helps to remove the majority of endothelial and hematopoietic cells and fiber debris from bulk digests. This drastically improves the efficiency and speed of satellite cell isolation in the subsequent steps. Once single cells have been dissociated from the bulk tissue, satellite cells can be selectively isolated via FACS. Cell suspensions are stained with an antibody cocktail consisting of sytox blue, antiCD31, anti-CD34, anti-CD45, anti-CD29, and anti-CD56. CD31
/CD34
/CD45
/sytox
/CD29þ/CD56þ satellite cells can be readily distinguished and separated from other cell

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types including endothelial, hematopoietic, and myofiber debris. The satellite cell-enriched population can subsequently utilized in experiments tailored to the users’ specific interests such as phenotypic characterization, mRNA purification, culture, and transplantation. 1.2.3 Xenotransplantation of Human Satellite Cells

Freshly isolated human satellite cells can be engrafted into mouse muscle. NSG immunodeficient mice, irradiation, and limited injury with either notexin or bupivacaine allows successful transplantation of human satellite cells into the tibialis anterior muscle. 18 gray (Gy) irradiation to the hind limb efficiently compromises the native mouse skeletal muscle stem cell population, significantly improving the engraftment of human cells. Sorted satellite cells are suspended directly in the muscle toxin of choice and both cells and toxin are co-injected into the same location within the recipient tibialis anterior muscle after irradiation. Mice are housed for a period of 5 weeks while human MuSCs contribute to the regeneration of the injured muscle and recapitulate the niche-associated stem cell population. Additionally, this time allows for the maturation of regenerated human muscle. If warranted, expansion of human muscle and the human satellite cell population is facilitated by muscle reinjury; otherwise, mouse sacrifice and muscle analysis can proceed.

1.2.4 Characterization and Analysis of Human Satellite Cell Engraftment

Once engrafted and matured, both human muscle and human MuSCs can be readily distinguished from host mouse muscle and cells in situ. Techniques described will provide researchers with the tools to characterize and analyze both these components in vivo in a systematic fashion via immunofluorescence staining and histology. Utilization of this model will enable researches to analyze human muscle and satellite cell differentiation, response to injury, and selfrenewal.

1.2.5 Expected Results

l

Protocol is efficacious on all cranial and somite mesoderm derived muscles tested.

l

Ten-to-twenty thousand satellite cells isolated per gram of muscle tissue.

l

Isolation of a pure population of satellite cells >95% Pax7 positive.

l

Can perform further assays in vitro or in vivo after transplantation.

l

Efficient xenotransplantation with as little as 1000 cells.

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Materials Reagents

1. 3 Gill’s Hematoxylin. 2. Alexa Fluor 488 goat anti-mouse IgG1 (Thermo). 3. Alexa Fluor 594 goat anti-mouse IgG2b (Thermo). 4. Antibody Diluent (DAKO). 5. Anti-CD29-488, Clone T52/16 (Ebioscience). 6. Anti-CD31-450, Clone WM-59 (Ebioscience). 7. Anti-CD31-microbeads (Miltenyi Biotec). 8. Anti-CD34-450, Clone4H11 (Ebioscience). 9. Anti-CD45-450, Clone H130 (Ebioscience). 10. Anti-CD45-microbeads (Miltenyi Biotec). 11. Anti-CD56-APC-vio-770 (Miltenyi Biotec). 12. Betadine. 13. 0.5% bupivacaine. 14. Collagenase XI. 15. Cy5 donkey anti-rabbit (Jackson Immunology). 16. Eosin. 17. Ethanol. 18. Fc blocking reagent (Miltenyi Biotec). 19. Heat inactivated fetal bovine serum (FBS). 20. Isopentane. 21. Ketamine. 22. Liquid nitrogen. 23. Mouse monoclonal IgG1 anti-PAX7 (DHSB). 24. Mouse monoclonal IgG2b anti-human LAMIN A/C (Vector Laboratories). 25. Mouse monoclonal IgG2b anti-human SPECTRIN (Leica Microsystems). 26. NSG mice older than 8 weeks of age. 27. Paraformaldehyde (PFA). 28. Permount. 29. Protein Block, Serum-Free (DAKO). 30. Rabbit polyclonal anti-laminin (Sigma-Aldrich). 31. Scott’s Water. 32. Sytox blue. 33. Optimal Cutting Temperature compound (OCT). 34. 0.25% trypsin.

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35. Mounting medium with DAPI. 36. Xylazine. 37. Xylene. 2.2 Media, Buffers, and Solutions

1. ACK Lysing Buffer. 2. Digestion media: Dulbecco’s Modified Eagle Medium (DMEM) with high glucose, 10% FBS, and 1% Penicillin/ Streptomycin. 3. FACS buffer: phosphate-buffered saline without calcium or magnesium, 2% FBS, 2 mM EDTA, and 10 mM HEPES. 4. MACS buffer: phosphate-buffered saline without calcium or magnesium, 0.5% BSA, 2 mM EDTA, and 10 mM HEPES. 5. Muscle collection media: Dulbecco’s Modified Eagle Medium (D-MEM) with high glucose, 30% FBS. 6. Phosphate-buffered saline with Tween (PBST): PBS, supplemented with 0.1% Tween-20. 7. Phosphate-buffered saline without magnesium or calcium (PBS w/o Caþþ or Mgþ). 8. Satellite cell collection media: Dulbecco’s Modified Eagle Medium (D-MEM) with high glucose, 20% FBS, 1% Penicillin/Streptomycin, and 10 μM rho associated protein kinase inhibitor. 9. Sulfamethoxazole/Trimethoprim, 200 mg/40 mg per 5 ml solution.

2.3

Equipment

1. 10 cm glass petri dishes. 2. 10 ml syringe vials. 3. 1000 μl, 200 μl, 20 μl, and 10 μl pipets and corresponding tips. 4. 15 ml conical tubes with screw caps. 5. 15 mm  15 mm  5 mm cryomolds. 6. 18-gauge needles. 7. 37  C mixing incubator. 8. 37  C water bath. 9. 5–0 chromic gut absorbable suture. 10. 50 ml conical tubes with screw caps. 11. 70 μm nylon mesh strainers. 12. Automatic serological pipet with 5 ml, 10 ml, and 25 ml pipettes. 13. BD FACSAriaII (BD Sciences). 14. Box cutting blades. 15. Cryostat.

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16. Disposable scalpels size 15. 17. 1.5 ml microcentrifuge tubes. 18. Hair removal cream. 19. Hamilton syringe needles, 32-gauge (Hamilton Company, Cat# 7803-04). 20. Hamilton syringe, 50 μl Model 1705 RN SYR, (Hamilton Company Cat# 7655-01). 21. Heating pad. 22. Irradiator. 23. Lead block. 24. MACS MultiStand (Miltenyi Biotec). 25. Microscope slide coverslips. 26. Microscope slides. 27. MidiMACS Separator (Miltenyi Biotec). 28. Miltenyi LD depletion columns (Miltenyi Biotec). 29. Paper tape. 30. Petrolatum ophthalmic ointment. 31. Polystyrene round-bottom tube with cell-strainer cap, 5 ml. 32. Q-tips. 33. Refrigerated centrifuge. 34. Rodent isoflurane anesthesia machine (or another approved means of obtaining surgical anesthesia). 35. Skin glue. 36. Surgical set: forceps, scissors, and needle driver. 37. Upright fluorescence microscope.

3

Methods

3.1 Tissue Dissociation and FACS Preparation

1. Collect muscle in muscle collection media, store at 4  C until processing (see Note 1). 2. Resuspend collagenase XI at 1 mg/ml in digestion media (see Note 2). 3. Trim excess fat, tendon, connective tissue, and fascia from the muscle tissue (see Fig. 1a and Note 3). 4. Use a box cutting blade and size 15 disposable scalpel to chop the muscle tissue in a 10 cm glass petri dish into 1 mm2 pieces (see Fig. 1b and Note 4). 5. When thoroughly minced, transfer to a 50 ml conical tube.

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Fig. 1 Tissue Dissociation Steps. (a) Whole muscle tissue before beginning digestion. The muscle has been extensively trimmed of fat and connective tissue. (b) Muscle in (a) after mechanical mincing with scalpel and box cutting blade. Note the size of the minced tissue pieces (c). Tissue digestion after 35 min in collagenase XI. (d) Trituration procedure using an 18-gauge needle attached to a 10 ml syringe. (e) Tissue clumping during (d). Clumps are placed into a glass petri dish and mechanically minced until they are able to pass through the needle. (f) Tissue digest after trituration and complete digestion in collagenase XI

6. Add collagenase XI at 12.5 mg per gram of muscle to the tube (see Fig. 1c). 7. Place on mixing platform in a 37  C incubator for 35 min (see Note 5). 8. Break up the tissue by trituration with an 18-gauge needle; put clumps aside in original petri dish and chop up with box cutting blade, then pass through needle (see Fig. 1d, e and Note 6). 9. Return everything to the original 50 ml conical tube and place it back in the 37  C mixing incubator.

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10. Incubate tissue in collagenase XI for a TOTAL time of 1 h 10 min (see Fig. 1f). 11. Begin warming 3 ml of 0.25% trypsin for every gram of tissue to 37  C. 12. After 1 h and 10 min TOTAL time of tissue digestion, dilute and rinse the suspension with PBS w/o Caþþ or Mgþ by filling up the 50 ml conical tube (see Note 7). 13. Centrifuge the sample at 2000 RPM (750  g) for 4 min at room temperature, then aspirate the supernatant (see Note 8) 14. Resuspend the pellet completely and wash again with PBS w/o Caþþ or Mgþ (see Note 9). 15. Centrifuge the sample again at 2000 RPM for 4 min at room temperature, and then aspirate the supernatant. 16. Resuspend the pellet completely in the residual liquid. 17. Add 3 ml of 0.25% trypsin for every gram of tissue (starting weight), mix gently and place in a 37  C warmer for 10–12 min (see Note 10). 18. Add 1.5 ml of FBS per gram of tissue (starting weight) to quench trypsin and mix gently. 19. Dilute with equal volume of 2% FBS in PBS and pass the suspension through a 70 μm nylon mesh strainer directly into a new 50 ml conical tube (see Note 11). 20. Wash the filter with 2% FBS in PBS. 21. Centrifuge the sample at 2000 RPM for 4 min at 4  C, and then aspirate the supernatant. 22. Resuspend the pellet and perform red blood cell lysis by adding 1 ml ACK per gram of tissue (starting weight) and incubate on ice for 5–7 min (see Note 12). 23. Wash the sample by filling the conical with 2% FBS in PBS. 24. Centrifuge the sample at 2000 RPM for 4 min at 4  C, and then aspirate the supernatant. 25. Resuspend pellet in remaining liquid and transfer to a 15 ml conical tube. 26. Wash the original 50 ml conical tube with up to 0.5 ml of MACS buffer and add to the new 15 ml conical tube. 27. Take a 50 μl aliquot of cells for compensation controls, place 10 μl into five 1.5 ml microcentrifuge tubes, and keep on ice. 28. Add 5 μl Fc blocking reagent for every gram of tissue (starting weight) to the 15 ml conical sample tube, mix gently and incubate for 5 min on ice (see Note 13).

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29. Add 5 μl of anti-CD45 magnetic beads and 2 μl of anti-CD31 magnetic beads for every gram of tissue (starting weight) to the sample tube, mix gently, and incubate for 20 min on ice. 30. Wash the sample by filling the 15 ml conical tube with 10 ml of MACs buffer. 31. Centrifuge the sample at 2000 RPM for 4 min at 4  C, and then aspirate the supernatant. 32. Resuspend the pellet completely in residual liquid and run the sample on a MACS LD column (connected to a MidiMACS separator magnet attached to a MACS MultiStand) primed with MACs buffer (see Note 14). 33. Collect 5 ml of pass-through in a new 15 ml conical on ice and then elute the bound fraction (optional) in a separate 15 ml conical by adding 3 ml MACS buffer to the column and plunging after the removal of the column from the magnet (see Note 15). 34. Wash the samples with MACS buffer by filling the conical tube to 15 ml. 35. Centrifuge the samples at 2000 RPM for 4 min at 4  C, then aspirate the supernatant. 36. Resuspend the pellets gently in 100 μl FACS buffer and transfer to 1.5 ml microcentrifuge tubes. 3.2

FACS Isolation

The following FACS Staining Protocols are Adapted From [22] 1. Create the antibody staining cocktail: 1X antibody for every 2.5 g of tissue (starting weight); 5 μl anti-CD31-450, 5 μl antiCD34-450, 5 μl anti-CD45-450, 20 μl anti-CD560APCVio770, and 7 μl anti-CD29-488. 2. Add the appropriate volume of antibody cocktail each sample and incubate for 30 min on ice (stain pass-through and bound fractions separately each with 1 antibody cocktail). 3. Wash each sample with 1 ml FACS buffer. 4. Centrifuge the samples at 2000 RPM for 4 min at 4  C, and then aspirate the supernatant. 5. Resuspend the pellets in 300 μl sytox blue diluted 1:1000 in FACS buffer. 6. Make single color isotype compensation controls for all colors in addition to control cells without any stain and cells with only sytox blue staining (use the five 10 μl aliquots set aside in Subheading 3.1, step 27). 7. Place all samples and controls into separate 5 ml polystyrene round-bottom tubes with cell-strainer caps by running the cells through the strainer cap.

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8. Keep the tubes ice and cover to protect from light. 9. Make satellite cell collection tubes; add 500 μl satellite cell collection media to 1.5 ml microcentrifuge tubes. 10. Run single-color isotype compensation controls on the flow cytometer to determine gating (see Note 16). 11. Run samples on the flow cytometer and sort satellite cells; sort scheme: FSC/SSC -> viable singlets -> negative depletion of CD31/CD34/CD45/sytox positive cells -> CD29/CD56 gate (see Fig. 2). 12. Collect CD31
/CD34
/CD45
/sytox
/CD29þ/ CD56þ cells directly into the satellite cell collection media tubes. 3.3 Mouse Irradiation (Day Prior to or the Day of Transplants) (See Note 17) 3.4 Xenotransplantation of Human Satellite Cells

1. Anesthetize NSG mice according to an approved protocol (we use ketamine/xylazine). 2. Shield the bodies of the mice with lead blocks, leaving the left hind leg exposed (see Note 18). 3. Irradiate with 18 Gy. 1. Follow all guidelines and protocols set forth by your intuition’s animal care and use regulatory board. 2. Prepare satellite cells for transplantation: centrifuge collection tubes at 2000 RPM for 10 min at 4  C, and then aspirate the supernatant (see Note 19). 3. Resuspend the pellets in chilled 0.50% bupivacaine (50 μl per mouse) on ice or chilled 0.002 μg/ml notexin (50 μl per mouse) on ice. 4. Keep cell/toxin suspensions on ice until injection. 5. Prepare mouse surgery area and surgical equipment. 6. Anesthetize mice according to your approved protocol (we use isoflurane sedation). 7. Weigh and place anesthetized mouse on sterile mat over heating pad on its back. 8. Protect eyes with sterile petrolatum ophthalmic ointment. 9. Remove hair from entire left leg below the knee with chemical or physical means (we use hair removal cream). 10. Use a Q-tip to clean the leg with 70% ethanol. 11. Use a Q-tip to apply betadine to entire anterior hind leg. 12. Use Q-tip to clean the leg again with 70% ethanol. 13. Using surgical forceps, grab the skin at the mid lower leg and make a 5 mm transverse incision over the tibialis anterior with surgical scissors (see Note 20).

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%Parent %Total #### 100.0 57.7 57.7 51.2 29.5 93.9 27.7 21.5 77.4 2.0 9.4

Fig. 2 Representative FACS profiles for the isolation of human satellite cells. Satellite cells are in the P5 population. The population hierarchy is shown on the bottom right. All cells are gated on cell size and granularity using forward and side scatter. The P1 population is gated for viable singlets using forward and side scattering. The P3 population is then gated for live CD34/CD45/CD31/sytox negative cells. This P4 population is gated on CD29 and CD56 to isolate satellite cells. The CD29/CD56 gate is shown as both dot and contour plots for ease of identification. We typically isolate the uppermost 75% of the CD56 positive population

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14. Using a 50 μl Hamilton syringe with an attached 32-gauge needle, inject 50 μl of cell/toxin suspension directly into the tibialis anterior muscle belly (see Note 21). 15. Close the incision with 5–0 chromic gut absorbable suture. 16. Cover the incision with thin layer of skin glue. 17. Administer analgesic medications and tag mic according to protocol (see Note 22). 18. Place mouse in clean recovery cage over heating pad until fully recovered. 19. We recommend administering antibiotics (sulfamethoxazole/ trimethoprim supplemented water) for a period of 7 days. 20. House mice for desired experimental length and monitor per protocol (see Note 23). 3.5 Tibialis Anterior Dissection and Preparation (See Note 24)

1. Sacrifice mice according to protocol. 2. Generously spray down the left leg with 70% ethanol. 3. Using scissors and forceps, remove skin from entire leg taking care to not damage underlying muscle. 4. Remove tibialis anterior by cutting distally at the tendon and proximally at the knee. 5. Set in OCT within a 15 mm  15 mm  5 mm mold. 6. Freeze in liquid nitrogen chilled isopentane (see Note 25). 7. Store frozen tissue blocks at
80  C.

3.6 Characterization and Analysis of Human Satellite Cell Engraftment

The following staining protocols are adapted from [22] and references therein including [9, 15, 20, 23]. 1. Using a cryostat, cut tissue sections at 6–10 μm depending on desired thickness (we use 6 μm for immunofluorescence staining and 10 μm for H&E histology) (see Note 26). 2. Store microscope slides at
80  C. 3. For immunofluorescence staining: warm slides at room temperature for 10 min. 4. Fix sections in 4% PFA at room temperature for 10 min. 5. Wash in PBST for 1 min. 6. Block with DAKO Protein Block for 10 min (see Note 27). 7. Incubate with the following primary antibodies diluted in DAKO Antibody Diluent: mouse monoclonal IgG2b antihuman SPECTRIN (1:200), mouse monoclonal IgG2b antihuman LAMIN A/C (1:500), mouse monoclonal IgG1 antiPAX7 (1:20), and rabbit polyclonal anti-laminin (1:500) for 5 h at room temperature or overnight at 4  C. 8. Wash in PBST for 1 min.

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9. Incubate secondary antibodies for 1 h at room temperature diluted in DAKO Antibody Diluent: Alexa Fluor 594 goat antimouse IgG2b (1:500), Alexa Fluor 488 goat anti-mouse IgG1 (1:500), Cy5 donkey anti-rabbit (1:500). 10. Wash in PBST for 1 min. 11. Mount sections with mounting medium with DAPI using a coverslip. 12. Slides are ready for examination using an upright fluorescence microscope (see Fig. 3a and Note 28). 13. For hematoxylin and eosin (H&E) histology: bring slides to room temperature for 10 min. 14. Rehydrate as follows; 5 min in xylene, twice; 5 min in 100% ethanol, twice; 5 min in 95% ethanol; 5 min in 80% ethanol and then wash in water for 30 s. 15. Place slides in 3 Gill’s Hematoxylin for 4 min and then wash with water for 30 s. 16. Place slides Scott’s water for 3 min and then wash in water for 30 s. 17. Place in Eosin for 2 min and then wash with water for 30 s (see Note 29). 18. Dehydrate as follows: 1 min in 80% ethanol; 2 min in 95% ethanol, twice; 3 min in 100% ethanol, twice; 2 min in xylene, twice. 19. Mount slides with Permount using coverslips and air dry overnight before imaging (see Fig. 3b).

4

Notes 1. Muscle tissue can be stored in collection media for up to 12 h at 4  C until processing. 2. Collagenase batches vary in their intrinsic enzymatic activity. Vials are labeled as >1200 Collagenase Digestive Units per mg. Optimization of enzyme concentration may be necessary based on a given enzyme batch. We recommend a starting concentration of 1–2 mg/ml and optimizing as the user becomes more familiar with the process of isolating satellite cells. The collagenase is resuspended in 10% FBS to inhibit contaminating proteases in the collagenase XI preparations. If collagenase XI is used without FBS then extensive titration of enzyme solution is required. 3. We recommend removing non-muscle tissue from collected muscles before storage; however, users can trim at any time prior to processing. It is important to start Subheading 3.1,

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Fig. 3 Analysis of engrafted human muscle and satellite cells after xenotransplantation into mouse tibialis anterior muscle. (a) Representative immunofluorescence images for the identification of human myofibers (SPECTRIN) and sublaminar human MuSCs (LAMIN A/C and PAX7 costaining—arrows) engrafted in mouse muscle. Both human and mouse myofibers are stained gray (LAMININ). (b) Representative H&E sections of transplanted muscle at two different magnifications

step 4 with a piece of muscle trimmed of fat, tendon, connective tissue, and fascia, as much as possible. This will reduce clumping during subsequent digestion steps and improve the dissociation of single cells resulting in increased satellite cell yield. 4. Perform the mincing in a small amount (about 3 ml) of collagenase XI digest solution to enhance the mechanical digestion process. Take care to mince the muscle tissue with the box cutting blade and scalpel to as small (50 Hz).

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Methods

3.1 Cardiomyocyte Differentiation from hPSCs

This protocol describes the generation of cardiomyocytes and stromal cells from hPSCs; it involves the culture of hPSCs (Subheading 3.1.1) and the subsequent differentiation toward the cardiac lineage (Subheading 3.1.2). It is important to follow these steps quickly and with precision to maintain health of the cells and also ensure good differentiation outcomes.

3.1.1 Passaging, Seeding, and Culture of hPSCs

Day–4 (see Note 21). 1. Prepare a 1% (v/v) hESC qualified Matrigel solution by adding 0.5 mL of hESC qualified Matrigel to 49.5 mL of PBS. 2. Coat a new T-25 with 1% (v/v) hESC qualified Matrigel solution for 1 h at room temperature. 3. Prepare mTeSR-1 culture medium by combining 40 mL of mTeSR-1 base medium containing 0.5% P/S (0.2 mL), with 10 mL of mTeSR-1 5 supplement. 4. 10 min before enzymatic passaging of cells; aspirate Matrigel solution, add 10 mL of mTeSR-1 medium and place into cell culture incubator until required (see Note 22). 5. To passage hPSCs, take existing cell cultures and aspirate medium. Wash culture with 1 mL of TrypLE, aspirate and then add 2 mL of TrypLE. Incubate for 3 min at 37
C (see Note 23). 6. After 3 min, add and mix 3 mL of mTeSR-1 medium to the T25 and remove all 5 mL of cell suspension and place into a 15 mL graduated polypropylene tube. Take a small aliquot of the solution (~20–200 μL) for counting, while cell suspension is being centrifuged (see the next step). 7. Centrifuge cell suspension at 300  g for 3 min. After centrifugation remove the supernatant, and add 1 mL of fresh mTeSR1 medium. 8. Add 500,000 cells to the newly coated T-25 (based on the cell count) and place in an incubator for cells to culture. Day–1 9. Change hPSC culture medium. Warm 5 mL of fresh mTeSR-1 medium, aspirate old medium, and add fresh mTeSR-1 medium (see Fig. 1b).

3.1.2 Cardiac Differentiation of hPSCs

Days 0, 1, and 2 10. Prepare RPMI base media by adding 500 mL RPMI 1640 Medium with 5 mL of P/S and 500 μL of 200mM AA2P.

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11. The initial differentiation steps require RPMI w/B27 Ins medium. To prepare RPMI w/B27 Ins medium take 49 mL of RPMI base media and add 1 mL of B27 supplement, minus insulin. 12. On day 0, remove mTeSR-1 culture medium and wash with 2.5 mL of RPMI base medium. 13. On days 0, 1, and 2, aspirate medium and add 5 mL of RPMI w/B27 Ins medium with 5 ng/mL BMP-4 (1: 4000), 9 ng/mL Activin A (1: 4000), 5 ng/mL bFGF (1: 4000), and 1 μM CHIR99021 (1: 10,000). Add growth factors and small molecules fresh daily and warm to 37
C before addition (see Note 24). Day 3 14. Aspirate medium and wash with 2.5 mL of RPMI base medium. 15. Aspirate wash medium and add 10 mL of RPMI w/B27 Ins medium with 5 μM IWP-4 (1: 1000). Add IWP-4 fresh and warm to 37
C before addition. Days 6, 8, and 10 16. The final differentiation steps require RPMI w/B27þ Ins medium. To prepare RPMI w/B27þ Ins medium take 49 mL of RPMI base media and add 1 mL of B27 supplement, containing insulin. 17. Aspirate medium and add 5 mL of RPMI w/B27þ Ins medium with 5 μM IWP-4 (1: 1000). Add IWP-4 fresh daily and warm to 37
C before addition. Day 13 18. Aspirate medium and add 5 mL of RPMI w/B27þ Ins (no additional factors) medium. Warm to 37
C before addition. 19. Cells are cultured until Day 15 and subsequently harvested for tissue generation (Fig. 1b) (see Note 25). 3.2 Manufacture of 3D Casting Well and Mechanical Loading Poles

Before manufacturing the polymer casting well and mechanical loading poles, you will require the respective templates. The casting well is made from a Teflon ring template, while the mechanical loading poles template was machined from polycarbonate (see Note 26). Template designs and dimensions can be found in Fig. 1c, d and can be readily fabricated at workshop facilities. 1. Preheat hotplate to a low/medium setting of approximately 50–60
C. 2. Mix the PDMS elastomer with the curing agent. The base and curing agent are mixed in a 10 to 1 ratio (w/w). First measure

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out the desired weight of base into a clean graduated tube and wait for the elastomer to settle to the bottom (the liquid will be highly viscous). Once weighed, add the appropriate amount of curing agent using a bulb pipette, to the elastomer and stir thoroughly for several minutes. 3. To remove bubbles from the elastomeric solution, place tube with the lid off and upright in the vacuum desiccator and vacuum for 30 min. Once bubbles have been removed, the PDMS is ready to be used for molding. 4. To fabricate the casting wells, pour the PDMS into a well of the 12-well plate to a height of approximately 0.5 cm. Place the Teflon ring template into the well, allowing a portion of the ring to remain above the surface level of the PDMS. Two Teflon rings can be placed in each well of a 12-well plate (Fig. 1e, f) (see Note 27). 5. To fabricate the mechanical loading poles, pour the PDMS into the template, wiping away any excess to ensure PDMS is level with the template (Fig. 1j). 6. Place both templates onto a hotplate for 1 h to cure PDMS (see Note 28). 7. Once cured the templates can be removed. For the casting well, use a pair of forceps to remove the Teflon rings and cut around the outside of the well, using a scalpel, to remove the pair of casting well molds from the 12-well plate (Fig. 1g). For the mechanical loading poles, again peel them off the template and cut them into individual pairs of poles using a scalpel (Fig. 1k, l). 8. To ensure sterility, autoclave the casting well and mechanical loading poles ready to be used in tissue fabrication. 3.3 Engineered Heart Tissue Fabrication and Culture

To fabricate the bioengineered cardiac tissues, the day 15 hPSCderived cardiac cells (Subheading 3.3.1) first need to be dissociated and then incorporated into a 3D collagen gel (Subheading 3.3.2).

3.3.1 Cardiac Differentiation Dissociation

1. Aspirate the differentiation medium and add the collagenase type I solution (2.5 mL per T-25) and incubate at 37
C for 1 h. Collagenase solution should be preheated to 37
C before use. 2. After 1 h, add PBS (7.5 mL per T-25) and gently pipette up and down ensuring all the cells have been collected. Cells should be in suspension in large cell aggregates. 3. Place cell suspension into a graduated tube and centrifuge for 3 min at 300  g. 4. Remove the supernatant and resuspend in Trypsin-EDTA (5 mL per T-25) and incubate for 10 min at 37
C, constantly

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agitating the cell suspension. Trypsin-EDTA solution should be preheated to 37
C before use. 5. Prepare MEMα medium by adding 450 mL of MEMα, GlutaMAX Supplement, no nucleosides with 5 mL of P/S, 500 μL of 200 mM AA2P and 50 mL of FBS. 6. After 10 min, add MEMα medium (5 mL per T-25) to neutralize the Trypsin-EDTA and filter cell suspension through a 100 μm cell strainer. MEMα medium should be preheated to 37
C before use. 7. Count cell suspension and then centrifuge for 3 min at 300  g. 8. Remove the supernatant and resuspend in MEMα medium at the required cell density (see Subheading 3.3.2). 3.3.2 Engineered Heart Tissue Fabrication

The first four steps of fabricating the cardiac tissues must be done on ice, using “ice cold” reagents, this is critical to ensure mixtures will not gel until desired. All steps must be performed aseptically. Day 15-EHT Fabrication 1. Using forceps, place a PDMS casting well into a well of a 6-well plate (one casting well per well) and ensure each casting well is firmly stuck down. Place the whole plate on ice to cool (see Note 29). 2. After the cardiac differentiation has been dissociated, resuspend the cells at a density of 4.55  106 cells/mL in cold MEMα medium (see Note 30). 3. To make up the bioengineered cardiac tissue mixture, add the following precooled reagents together in the same order as listed, ensuring each reagent is thoroughly mixed at each step (see Note 31): (a) 36 μL of acid soluble Collagen I (7.3 mg/mL) (see Note 30) (b) 4.3 μL of 10 DMEM (c) 4.0 μL of 0.1 M NaOH (d) 121 μL cell suspension (0.55  106 cells at 4.55  106 cells/mL) (ensure the cell suspension is gently mixed to avoid cell death). The mixture describes the amount of reagents per EHT (plus 10% excess). To make up the mixture for multiple EHTs, simply multiply the above mixture by the number of EHTs required. If the acid soluble Collagen I is at a different concentration than that listed, the amount of each reagent needs to be adjusted (see Note 30). 4. Pipette 150 μL of cell/gel mixture into each casting well using a 1 mL pipette, avoiding the generation of bubbles within the

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mixture. Run the pipette tip around the entire base of the casting well to break the surface tension of the solution and spread the mixture around the entire circumference (Fig. 1h). 5. Centrifuge the 6-well plate for 10 s at 100  g to ensure the mixture is sitting on the base of the casting mold. 6. Place the 6-well plate into the incubator at 37
C for 30 min. This will cause the mixture to gel and turn an opaque pink color. 7. After the mixture has gelled, gently add 7 mL of pre-warmed MEMα medium and place the 6-well plate into the incubator. Day 17 8. Aspirate the medium, being careful not to disturb the EHT, and gently add 7 mL of pre-warmed MEMα medium. Day 20 5 days after EHTs have formed, they are transferred from their casting wells to mechanical loading poles. These mechanical loading poles are designed to impart 10% strain upon the tissues, allowing formation of aligned cardiomyocyte bundles (Fig. 1n). 9. By day 5 EHTs will have condensed around the center of the casting well and have enough mechanical integrity to be handled. Using a 200 μL pipette tip gently move around the base of the casting well and move the EHT up the edge and off the top of the PDMS mold, being careful not to damage the tissue. 10. Once the tissue has been removed, remove the PDMS casting well and place a set of mechanical loading poles into the well using sterile forceps. Clamp the ends of the poles closed using sterile forceps, and use a 200 μL pipette tip to gently place the tissue over each pole (see Note 32). Release the forceps allowing the tissue to take the mechanical load (Fig. 1m). 11. Place the EHT, on the mechanical poles, into a new well of a 12-well plate with 2 mL of fresh pre-warmed MEMα medium. Then place the 12-well plate into a cell culture incubator, changing the medium (2 mL of fresh, pre-warmed MEMα medium) every 2–3 days until required. 3.4 EHT Cryoinjury Model

1. Using a pair of sterile forceps, shape the end of the dry ice pellet into a sharp edge (Fig. 1I). 2. Aspirate the medium from well, and then press the piece of dry ice lightly into the exposed edge of the tissue for 3 s (see Note 33). 3. Replace the medium with 2 mL of fresh pre-warmed MEMα medium and place back into a cell culture incubator. Change the medium every 2–3 days until required (see Notes 34 and 35).

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3.5 EHT Electrical Stimulation and Force Measurement

EHT force of contraction can be measured using an organ bath or calculated by the deflection of the mechanical loading posts (see Note 36). Herein this section, we detail EHT electrical pacing (Subheading 3.5.1) and approximation of contraction force via post deflection (Subheading 3.5.2).

3.5.1 Electrical Stimulation

EHTs will contract spontaneously, and as such they do not require electrical stimulation to access their force of contraction. However following cryoinjury, more accurate results will be obtained if EHTs are “paced” at a consistent rate using electrical stimulation. 1. Using a scalpel, strip (remove) the PVC protecting coating from the both ends of a piece of wire (approximately 30 cm long, removing 0.5 cm from either end). Bend each of the stripped ends into an “L-shape.” This is performed for two pieces of wire. 2. To establish the electrical stimulation device, insert the connection cable, via the banana plug, into the digital stimulator output ports (positive and negative). Then attach a piece of wire to each of the alligator clips, creating a negative and positive terminal. 3. Remove well-plate lid and place the positive and negative wires into the culture medium, ensuring the wires are either side of the EHT (see Notes 37 and 38). 4. Set the digital stimulator to a frequency of 1 Hz, 10 ms squarewave pulse and a voltage of 100 mV (see Notes 39 and 40). 5. Stimulate the EHT and record the resultant EHT contractions for a 10 second period on a phase contrast microscope at a high-capture rate (i.e., every 20 ms).

3.5.2 Force Measurement

Using the recorded videos, EHT contraction force is calculated by the formulae for the general deflection of a cantilever:   3EI F ¼ δ L3 F ¼ Contraction force

I ¼ Area moment of inertia

E ¼ Young’s modulus

L ¼ Length of pole

δ ¼ Post deflection

The area moment of inertia of a “filled rectangle” is calculated by bh 3 b ¼ base width h ¼ height: I ¼ 12 As a result, the EHT contraction force within in our system is calculated by F ¼

3Ebh 3 δ 12L 3

F ¼ 6μN =μm δ:

E PDMS ¼ 1:5 N=mm2

b ¼ 2 mm

h ¼ 2 mm

L ¼ 10 mm

:

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Therefore, for every μm of pole deflection the tissue is producing 6 μN of force (see Note 41). The force measurement of each EHT can be automated using the aforementioned equation in conjunction with software that tracks the movement of the pole (i.e., Matlab). This allows visualization and calculation of EHT contraction parameters (force, beating rate, etc.) (Fig. 1p) (see Note 42).

4

Notes 1. For protocols to generate bioengineered skeletal muscle tissues, please refer to [12, 16, 17]. The main discrepancy between fabricating engineered skeletal muscle and cardiac muscle is the starting cell population. In brief, engineered skeletal muscle is usually generated from a starting population of myoblasts (satellite cells). Myoblasts are incorporated into the 3D collagen mixture (as per Subheading 3.3.2) and cultured in standard myoblast growth medium (i.e., base medium plus 20% FBS). After tissue formation (2 days), growth medium is replaced with differentiation medium (i.e., base medium plus 2% horse serum) and cultured as per protocol (Subheadings 3.3.2 and 3.4). 2. Human pluripotent stem cells (hPSCs) can be embryonic stem cells or induced pluripotent stem cells in origin. The hPSCs must be adapted to feeder-free conditions, in a chemically defined medium with enzymatic single cell passaging. We recommend culturing hPSCs in mTeSR on a Matrigel substratum, passaged using TrypLE enzymatic digestion. For more information regarding hPSCs adaptation, refer to Stover and Schwartz [18] or “Maintenance of Human Pluripotent Stem Cells in mTeSR-1 Technical Manual” [19]. 3. MTeSR-1 is used for the feeder-free culture of hPSCs. The mTeSR-1 culture medium kit comes with mTeSR-1 basal medium (400 mL) and mTeSR-1 5X supplement (100 mL). 5 supplement should be aliquoted in 10 mL lots and stored at 20
C until required. 4. Upon arrival P/S should be aliquoted (2.5/5 mL). Concentrated stocks should be stored at 20
C. 5. Upon arrival Matrigel should be aliquoted (500 μL). This should be performed upon ice, as the solution will irreversibly form a gel if left at room temperature. Concentrated stocks should be stored at 20
C. 6. PBS is made as per the manufacturer’s instructions and contains: Potassium Chloride (KCl)—200 mg/L, Potassium

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Phosphate monobasic (KH2PO4)—200 mg/L, Sodium Chloride (NaCl)—8000 mg/L and Sodium Phosphate dibasic (Na2HPO4) anhydrous—1150 mg/L. 7. AA2P is reconstituted to 200 mM in milli-Q water. The solution should be sterile filtered and aliquoted. Concentrated stocks should be stored at 20
C. 8. BMP-4 is first reconstituted to 500 μg/mL in 4 mM hydrochloric acid, and then diluted to 20 μg/mL in milli-Q water containing 0.1% (w/v) bovine serum albumin (BSA). Concentrated stocks should be stored at 20
C in aliquots. 9. Activin A is reconstituted at 36 μg/mL in PBS containing 0.1% BSA (w/v). Concentrated stocks should be stored at 20
C in aliquots. 10. bFGF is reconstituted at 20 μg/mL in PBS containing 0.1% BSA (w/v). Concentrated stocks should be stored at 20
C in aliquots. 11. CHIR99021 should be made up to 10 mM in Dimethyl sulfoxide (DMSO). Concentrated stocks should be stored at 20
C in aliquots. 12. IWP-4 should be made up to 5 mM in DMSO. Concentrated stocks should be stored at 20
C in aliquots. 13. Upon arrival B-27 supplement minus insulin should be aliquoted into 1 mL aliquots and stored at 20
C. 14. Upon arrival B-27 supplement should be aliquoted into 1 mL aliquots and stored at 20
C. 15. Sylgard 184 silicone elastomer kit contains two components, a polydimethylsiloxane (PDMS) elastomer base and curing agent. These components can be purchased separately if desired. 16. Collagenase Type I should be reconstituted to a 0.2% (w/v) solution in PBS with calcium and magnesium (PBSþ) containing 20% FBS. This should be dissolved, filtered, aliquoted, and stored at 20
C. 17. PBSþ is made as per manufacturer’s instructions and contains: Calcium Chloride (CaCl2) (anhyd.)—100 mg/L, Magnesium Chloride (anhydrous)—47 mg/L, Potassium Chloride (KCl)—200 mg/L, Potassium Phosphate monobasic (KH2PO4)—200 mg/L, Sodium Chloride (NaCl)— 8000 mg/L and Sodium Phosphate dibasic (Na2HPO4) anhydrous—1150 mg/L. 18. Upon arrival Trypsin-EDTA should be aliquoted and stored at 20
C.

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19. Upon arrival FBS should be aliquoted and stored at 20
C. 20. 10 DMEM is generated as per manufacturer’s instructions to give a 10 concentrated DMEM solution. 21. hPSCs are designed to be passaged and seeded upon a Friday, so the differentiation protocol can be started the following Tuesday. 22. Refrain from warming mTeSR multiple times before use in cell culture, as this will degrade hPSCs maintenance growth factors and result in a reduced capacity to maintain pluripotency. 23. Incubate for only 3 min (or close to as possible), as incubating for too long will result in significant cell death. 24. Avoid freeze-thaw cycles for the growth factors, thawed aliquots may be stored in the fridge and used for up to 2 weeks. 25. After 15 days of differentiation, cells should consist of approximately 60–80% cardiomyocytes (α-Actinin and Cardiac troponin T positive) and 20–40% of CD90+ stromal cells (by flow cytometry). The stromal cells are critical for the integrity and function of the EHT. If the ratio of cardiomyocytes to stromal cells is not within this approximate range, the tissue will not form or function adequately. 26. Other high density polymers, with the capacity to be heated to 60
C without deformation or warping, could also be used. 27. Ensure that there are no bubbles present in the PDMS prior to curing, if bubbles are present, vacuum PDMS again in the vacuum desiccator (20–30 min). 28. To check if the PDMS is cured, press a pipette tip against the polymer. The PDMS should be elastic but firm. If the pipette tip penetrates the PDMS surface, even slightly, and is still a liquid, place back onto the hotplate and leave to cure for longer. 29. Ensure the casting well is stuck to the 6-well plate prior to being cooled on ice, otherwise PDMS will not adhere. 30. If Collagen I is at a different concentration than that listed (7.3 mg/mL) the amount of each reagent will need to be adjusted. The amount of Collagen I (mg of Collagen/EHT) must be kept constant, as well as the Collagen ratio to 10 DMEM and 0.1 M NaOH, to ensure the pH is neutralized and the mixture will gel. The cell suspension density and volume per EHT must also be adjusted to ensure a constant 550,000 cells per EHT and a mixture total of 150 μL. Three examples are listed below:

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Current protocol

Example 1 Example 2 Example 3

Collagen I per EHT (mg)

0.263

0.263

0.263

0.263

Collagen I density (mg/mL)

7.3

3.65

6.4

5.5

Collagen I volume (μL)

36

72

41.1

47.8

10 DMEM (0.119 μL per μL of Collagen I)

4.3

8.6

4.9

5.7

0.1 M NaOH (0.111 μL per μL of Collagen I)

4

8

4.6

5.3

Cells per EHT

550,000

550,000

550,000

550,000

Cells suspension volume (μL)

121

77

115

107

Cell suspension density (cells/mL)

4.55  106 7.17  106 4.79  106 5.16  106

Total mixture volume (μL)a

165

165

165

165

150 μL per EHT þ10%

a

31. Mix each newly added reagent thoroughly, ensuring no bubbles are formed in the process. 32. Using forceps, position the pole so the base is held against the edge of the well. When handling the EHT, place the pipette tip through the center of the EHT and avoid directly poking any area of the tissue. 33. Apply the dry ice probe to an area of the EHT that lies on the top of the pole to ensure contact has been made. 34. Following dry ice application there should be a localized injury on one side of the EHT (Fig. 1o). 35. Cryoinjury is usually performed 7 days after EHTs are placed on mechanical loading poles (Day 27); however, this could be performed at any time point after the EHT has formed. 36. Organ baths allow the study of muscle contractions; however, it consists of several expensive pieces of equipment and requires highly trained operators. In our experience, force calculations, via pole deflection, are more amenable to our EHT studies and force approximation using these mechanical engineering equations closely reflects values obtained using force transducers/sensors.

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37. Replace wires every 2–3 EHTs recorded, as wire terminals will corrode. 38. As this is not performed aseptically, EHTs are not further cultured after electrical stimulation. 39. EHTs can be stimulated up to a frequency of 3–4 Hz. 40. Bioengineered skeletal muscle can be stimulated to produce “twitch” contractions (e.g., 1 Hz stimulation) or “tetanus” contractions (>20 Hz stimulation) [12]. 41. In the absence of electrical stimulation, force measurements can be taken at any time point after the EHT has been placed on the mechanical loading poles. 42. The cryoinjury model described should yield an approximate 50% decrease in force at 6 h post-injury. References 1. Lee RT, Walsh K (2016) The future of cardiovascular regenerative medicine. Circulation 133(25):2618 2. Blau HM, Cosgrove BD, Ho ATV (2015) The central role of muscle stem cells in regenerative failure with aging. Nat Med 21(8):854–862 3. Tabebordbar M, Wang ET, Wagers AJ (2013) Skeletal muscle degenerative diseases and strategies for therapeutic muscle repair. Ann Rev Pathol 8(1):441–475 4. Porrello ER et al (2011) Transient regenerative potential of the neonatal mouse heart. Science 331(6020):1078 5. Porrello ER et al (2013) Regulation of neonatal and adult mammalian heart regeneration by the miR-15 family. Proc Natl Acad Sci 110 (1):187–192 6. Ranga A, Gjorevski N, Lutolf MP (2014) Drug discovery through stem cell-based organoid models. Adv Drug Deliv Rev 69-70:19–28 7. Huh D, Hamilton GA, Ingber DE (2011) From 3D cell culture to organs-on-chips. Trends Cell Biol 21(12):745–754 8. Sato T et al (2009) Single Lgr5 stem cells build crypt-villus structures in vitro without a mesenchymal niche. Nature 459(7244):262–265 9. Takebe T et al (2013) Vascularized and functional human liver from an iPSC-derived organ bud transplant. Nature 499(7459):481–484 10. Lancaster MA et al (2013) Cerebral organoids model human brain development and microcephaly. Nature 501(7467):373–379

11. Takasato M et al (2014) Directing human embryonic stem cell differentiation towards a renal lineage generates a self-organizing kidney. Nat Cell Biol 16(1):118–126 12. Madden L et al (2015) Bioengineered human myobundles mimic clinical responses of skeletal muscle to drugs. eLife 4:e04885 13. Tiburcy M et al (2017) Defined engineered human myocardium with advanced maturation for applications in heart failure modelling and repair. Circulation 135(19):1832–1847 14. Eder A et al (2016) Human engineered heart tissue as a model system for drug testing. Adv Drug Deliv Rev 96:214–224 15. Eschenhagen T et al (2012) Physiological aspects of cardiac tissue engineering. Am J Physiol Heart Circ Physiol 303(2):H133 16. Juhas M et al (2014) Biomimetic engineered muscle with capacity for vascular integration and functional maturation in vivo. Proc Natl Acad Sci 111(15):5508–5513 17. Cheng CS et al (2016) Cell density and joint microRNA-133a and microRNA-696 inhibition enhance differentiation and contractile function of engineered human skeletal muscle tissues. Tissue Eng A 22(7–8):573–583 18. Stover AE, Schwartz PH (2011) Adaptation of human pluripotent stem cells to feeder-free conditions in chemically defined medium with enzymatic single-cell passaging. Methods Mol Biol 767:137–146 19. Maintenance of human pluripotent stem cells in mTeSR™1 (2015). Stem Cell Technologies

INDEX A

G

Adipogenic............................................... 93–99, 101, 102

Green fluorescence protein (GFP) ...................... 127–133

B

H

Bioanalyzer ............................... 22, 62, 64, 66–68, 70–72 Bisulfite sequencing .................................................... 3–12

Histone ...................................................... 15, 16, 23, 174 Human......................................... 56, 105–121, 135, 148, 149, 195–200, 203, 204, 209, 210, 212–224 Hydrogel................................. 75–91, 147–152, 155–162

C C2C12 cells ............................... 45, 47, 57, 70, 128–131, 136, 138, 142 Calcium............ 56, 94, 97, 98, 109, 150, 197, 213, 221 Cardiomyocyte .................. 210–212, 214, 215, 218, 222 ChIPseq ........................................................................... 23 Cholesterol .................................................. 43, 49, 54, 55 Chondrogenic ........................................... 94, 96, 99, 102 Chromatin .................................................................15–24 Chromatography ................................................ 46, 51, 53 Contraction ....................... 180, 205, 212, 219, 220, 223 Cryoinjury ............................................................ 209–224

I

D

L

Damage.................. 15, 72, 94, 102, 105, 116, 152, 161, 162, 171, 190, 196, 203–205, 209, 212, 218 Degeneration..................................................94, 147, 148 Diaphragm............................................................ 177–191 DNA ............... 3–12, 15–18, 20–22, 24, 45, 49, 55, 119 Dysferlin ............................................................... 195, 201

Laminin.............................33, 76, 77, 79, 80, 83, 84, 87, 91, 108, 116, 121, 147, 148 Lipidomics .......................... 39–41, 43–45, 47–51, 54–58 Lysophosphatidylcholine (LPC) ..............................40, 53

E

Mass spectrometry ............................................. 43, 45, 46 Membrane .............................. 7, 9, 40, 48, 71, 136, 180, 195–200, 203–206 Mesenchymal stem cell ................................................... 93 Metabolism fatty-acid oxidation ...................................... 55, 62, 69 glutaminolysis......................................................63, 69 glycolysis .................................................................... 62 mitochondria .......................................................63, 69 Methylation ................................................................. 3–12 Microcontact-printing .................................75–87, 89–91 Microwell ...................................................................77, 88 Mouse .............................................. 7, 23, 43, 78, 80, 86, 89, 96, 99, 101, 102, 107, 108, 114, 116, 118, 121, 128, 148, 150, 156–158, 167–172, 179, 182, 184, 186, 190, 201

Electrophoresis ........................................ 5, 10, 11, 23, 76 Embryo .........................27, 28, 34, 39, 61, 62, 167–176, 178, 182, 184–186, 188, 190, 220 Endocytosis ................................................................... 196 Epigenetic ........................................................... 15, 16, 62 Extracellular.....................71, 76, 93, 135, 139, 141, 180

F Fibro-adipogenic progenitors (FAPs) ....................93–99, 101, 102 Fibroblast.................. 63, 64, 94, 95, 180, 182, 197, 212 Fibrosis............................................................................. 94 Fluorescence activated cell sorting (FACS) ........... 63, 78, 86, 94, 95, 97, 98, 106, 107, 109–115

Immunofluorescence ................. 107, 116, 118, 167–176 Immunoprecipitation.....................................3, 17, 19–21 Injury .................................................... 15, 105–107, 196, 204, 206, 209, 210, 212–224 Innervation ................................. 167, 177, 178, 180–191 In-situ hybridization ....................................29, 30, 32–34 Intracellular ......................... 40, 136, 180, 196, 200, 202 Irradiation.........................107, 114, 120, 138, 140, 142, 196, 200, 201, 203–205

M

James G. Ryall (ed.), Skeletal Muscle Development, Methods in Molecular Biology, vol. 1668, DOI 10.1007/978-1-4939-7283-8, © Springer Science+Business Media LLC 2017

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226 Index

Muscle MuSCs ...............................61–63, 70, 75, 78, 86, 88, 105–107, 118 myoblasts .............................15, 39, 55, 57, 128, 148, 167, 168, 174, 180, 220 myofiber...............................15, 39, 55, 93, 106, 118, 167, 169, 171, 174, 178, 180, 182 myotubes .......................................128, 136, 148, 196 satellite cells ................................................... 5, 15, 16, 19, 40, 57, 75, 94, 105–107, 114, 118, 120, 197, 201, 209, 220 Myogenesis differentiation.................................15, 16, 39, 40, 75, 136, 138–140, 142 elongation.................................................................. 41 fusion ................................................................ 57, 182 maturation ....................................... 93, 135–143, 182 migration .......................................................... 76, 182 proliferation .....................................39, 105, 168, 174 quiescence.................................................................. 15 regeneration................................................15, 75, 135 self-renewal ....................................................... 75, 105 Myopathy................................................................ 41, 195 Myosin ..................16, 33, 129, 132, 133, 168, 170, 201

O Optogenetic.......................................................... 136–138 Osteogenic................................................. 94, 95, 99, 102

Phospholipids ..................................................... 40, 49, 51 Photolithography ...................................... 76–78, 80, 150 Polyethylene glycol (PEG) ........................ 76, 77, 79, 80, 82, 83, 85–87, 90 Progenitor .................................5, 39, 40, 61, 75, 89, 93, 94, 167, 179, 180, 182 Protein .................................................... 3, 15–17, 28, 40, 55, 68, 75–77, 83–85, 87, 91, 106, 108, 109, 116, 121, 127, 135, 136, 147, 170, 178, 180, 181, 195, 201, 212

R RNA .............................................. 5–7, 16, 27–32, 34, 35

S Sarcolemma ....................... 195, 196, 199, 200, 205, 206 Sequencing ......................................................... 15–24, 62 Somite .............................................................93, 107, 167

T Transcription ............ 15, 16, 23, 39, 106, 168, 178, 179 Transfection................................................. 138, 142, 143 Transplantation ............................................105–107, 120 Trophoblasts.................................................................. 201

X Xenotransplantation ............................ 105–114, 116–121

P

Z

Perivascular .................................................................... 201 Phosphatidylserine ....................................................40, 41

Zebrafish .......................................................27–32, 34, 35