Myogenesis: Methods and Protocols (Methods in Molecular Biology, v798)

METHODS

IN

MOLECULAR BIOLOGY™

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

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

Myogenesis Methods and Protocols Edited by

Joseph X. DiMario Department of Cell Biology and Anatomy, Chicago Medical School, Rosalind Franklin University of Medicine and Science, North Chicago, IL, USA

Editor Joseph X. DiMario, Ph.D. Department of Cell Biology and Anatomy Chicago Medical School Rosalind Franklin University of Medicine and Science North Chicago, IL, USA [email protected]

ISSN 1064-3745 e-ISSN 1940-6029 ISBN 978-1-61779-342-4 e-ISBN 978-1-61779-343-1 DOI 10.1007/978-1-61779-343-1 Springer New York Dordrecht Heidelberg London Library of Congress Control Number: 2011939748 © Springer Science+Business Media, LLC 2012 All rights reserved. This work may not be translated or copied in whole or in part without the written permission of the publisher (Humana Press, c/o Springer Science+Business Media, LLC, 233 Spring Street, New York, NY 10013, USA), except for brief excerpts in connection with reviews or scholarly analysis. Use in connection with any form of information storage and retrieval, electronic adaptation, computer software, or by similar or dissimilar methodology now known or hereafter developed is forbidden. The use in this publication of trade names, trademarks, service marks, and similar terms, even if they are not identified as such, is not to be taken as an expression of opinion as to whether or not they are subject to proprietary rights. Printed on acid-free paper Humana Press is part of Springer Science+Business Media (www.springer.com)

Preface Our understanding of the molecular and cellular mechanisms that control skeletal muscle development, regeneration, and adaptive responses to activity has increased dramatically in recent years. This expansion of knowledge has been fostered by innovative techniques and approaches that are either specifically designed or adapted for research in skeletal muscle biology. Myogenesis: Methods and Protocols presents detailed, step-by-step methods in the study of the molecular and cellular biology of skeletal muscle cells. Protocols from different model systems including mammalian, avian, zebrafish, and invertebrate skeletal muscle are included in this volume. Highlighted topics cover a wide range of interests and expertise including myogenic and stem cell isolation, investigation of models of exercise and disuse, viral vector delivery systems, calcium imaging, cell profiling, and protein–DNA and protein– protein interactions. The book presents model systems and state-of-the-art techniques developed and perfected by leading scientists in the field. The techniques are an invaluable resource for experienced and emerging scientists, including researchers such as molecular biologists, cell biologists, developmental biologists, skeletal muscle physiologists, and clinical scientists with interests in skeletal muscle stem cells and regenerative medicine. North Chicago, IL

Joseph X. DiMario

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

PART I

ISOLATION AND ANALYSIS OF SKELETAL MUSCLE PRECURSOR CELLS

1 Isolation and Characterization of Human Fetal Myoblasts . . . . . . . . . . . . . . . . Ariya D. Lapan and Emanuela Gussoni 2 Skeletal Muscle Satellite Cells: Background and Methods for Isolation and Analysis in a Primary Culture System . . . . . . . . . . . . . . . . . . . . . . . . . . . . Maria Elena Danoviz and Zipora Yablonka-Reuveni 3 Isolation of Muscle Stem Cells by Fluorescence Activated Cell Sorting Cytometry . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Alessandra Pasut, Paul Oleynik, and Michael A. Rudnicki 4 Mouse and Human Mesoangioblasts: Isolation and Characterization from Adult Skeletal Muscles . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Mattia Quattrocelli, Giacomo Palazzolo, Ilaria Perini, Stefania Crippa, Marco Cassano, and Maurilio Sampaolesi 5 Direct Electrical Stimulation of Myogenic Cultures for Analysis of Muscle Fiber Type Control . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Eric J. Cavanaugh, Jennifer R. Crew, and Joseph X. DiMario 6 Single Muscle-Fiber Isolation and Culture for Cellular, Molecular, Pharmacological, and Evolutionary Studies . . . . . . . . . . . . . . . . . . . . . . . . . . . Judy E. Anderson, Ashley C. Wozniak, and Wataru Mizunoya 7 Somite Unit Chronometry to Analyze Teratogen Phase Specificity in the Paraxial Mesoderm . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Sara J. Venters and Charles P. Ordahl

PART II

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NON-MAMMALIAN MODELS OF MYOGENESIS

8 Analysis of Skeletal Muscle Development in Drosophila. . . . . . . . . . . . . . . . . . Ginny R. Morriss, Anton L. Bryantsev, Maria Chechenova, Elisa M. LaBeau, TyAnna L. Lovato, Kathryn M. Ryan, and Richard M. Cripps 9 Immunocytochemistry to Study Myogenesis in Zebrafish . . . . . . . . . . . . . . . . Nathan C. Bird, Stefanie E. Windner, and Stephen H. Devoto 10 Immunofluorescent Localization of Proteins in Caenorhabditis elegans Muscle . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Kristy J. Wilson, Hiroshi Qadota, and Guy M. Benian

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PART III

EXPERIMENTAL MODELS AND ANALYSIS OF SKELETAL MUSCLE EXERCISE AND DISUSE

11 Resistance Loading and Signaling Assays for Oxidative Stress in Rodent Skeletal Muscle . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Stephen E. Alway and Robert G. Cutlip 12 Analysis of Skeletal Muscle Hypertrophy in Models of Increased Loading . . . . Sue C. Bodine and Keith Baar 13 Protein Overexpression in Skeletal Muscle Using Plasmid-Based Gene Transfer to Elucidate Mechanisms Controlling Fiber Size . . . . . . . . . . . . Chia-Ling Wu and Susan C. Kandarian 14 In Vivo Measurement of Muscle Protein Synthesis Rate Using the Flooding Dose Technique . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Marta L. Fiorotto, Horacio A. Sosa Jr., and Teresa A. Davis

PART IV

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MUSCLE PROFILING

18 Gene Profiling Studies in Skeletal Muscle by Quantitative Real-Time Polymerase Chain Reaction Assay . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Shephali Bhatnagar, Siva K. Panguluri, and Ashok Kumar 19 Analysis of Lipid Profiles in Skeletal Muscles . . . . . . . . . . . . . . . . . . . . . . . . . . Vassilis Mougios and Anatoli Petridou 20 Proteomic Analysis of Dystrophic Muscle . . . . . . . . . . . . . . . . . . . . . . . . . . . . Caroline Lewis, Philip Doran, and Kay Ohlendieck

PART VI

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GENERATION OF VIRAL VECTORS AND TRANSGENIC MICE

15 Recombinant Adeno-Associated Viral Vector Production and Purification . . . . Jin-Hong Shin, Yongping Yue, and Dongsheng Duan 16 Generation of Lentiviral Vectors for Use in Skeletal Muscle Research. . . . . . . . Christophe Pichavant and Jacques P. Tremblay 17 Generating Tamoxifen-Inducible Cre Alleles to Investigate Myogenesis in Mice . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Christoph Lepper and Chen-Ming Fan

PART V

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EXPERIMENTAL APPROACHES IN CALCIUM IMAGING OF SKELETAL MUSCLE

21 Detection of Calcium Release via Ryanodine Receptors . . . . . . . . . . . . . . . . . . Jerry P. Eu and Gerhard Meissner 22 Measurement of Calcium Release Due to Inositol Trisphosphate Receptors in Skeletal Muscle . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Mariana Casas, Francisco Altamirano, and Enrique Jaimovich 23 Detection of Calcium Sparks in Intact and Permeabilized Skeletal Muscle Fibers . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Noah Weisleder, Jingsong Zhou, and Jianjie Ma

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24 Analysis of Calcium Transients in Cardiac Myocytes and Assessment of the Sarcoplasmic Reticulum Ca2+-ATPase Contribution . . . . . . . . . . . . . . . . Anand Mohan Prasad and Giuseppe Inesi

PART VII

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RNA-MEDIATED GENE REGULATION

28 Determination of MiRNA Targets in Skeletal Muscle Cells . . . . . . . . . . . . . . . Zhan-Peng Huang, Ramón Espinoza-Lewis, and Da-Zhi Wang 29 shRNA-Mediated Gene Knockdown in Skeletal Muscle . . . . . . . . . . . . . . . . . . Muriel Golzio, Jean-Michel Escoffre, and Justin Teissié

PART IX

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ANALYSIS OF GENE PROMOTER TRANSCRIPTIONAL ACTIVITY

25 Analysis of Muscle Gene Transcription in Cultured Skeletal Muscle Cells. . . . . Charis L. Himeda, Phillip W.L. Tai, and Stephen D. Hauschka 26 Analysis of Fiber-Type Differences in Reporter Gene Expression of β-Gal Transgenic Muscle. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Phillip W.L. Tai, Catherine L. Smith, John C. Angello, and Stephen D. Hauschka 27 Determination of Gene Promoter Activity in Skeletal Muscles In Vivo. . . . . . . Sarah M. Senf and Andrew R. Judge

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ANALYSIS OF PROTEIN-DNA INTERACTIONS

30 Detection of NF-kB Activity in Skeletal Muscle Cells by Electrophoretic Mobility Shift Analysis. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Jason M. Dahlman and Denis C. Guttridge 31 Isolation of Nuclei from Skeletal Muscle Satellite Cells and Myofibers for Use in Chromatin Immunoprecipitation Assays . . . . . . . . . . . . . . . . . . . . . Yasuyuki Ohkawa, Chandrashekara Mallappa, Caroline S. Dacwag Vallaster, and Anthony N. Imbalzano 32 An Improved Restriction Enzyme Accessibility Assay for Analyzing Changes in Chromatin Structure in Samples of Limited Cell Number . . . . . . . . . . . . . . Yasuyuki Ohkawa, Chandrashekara Mallappa, Caroline S. Dacwag Vallaster, and Anthony N. Imbalzano 33 ChIP-Enriched in Silico Targets (ChEST), a ChIP-on-Chip Approach Applied to Analyzing Skeletal Muscle Genes . . . . . . . . . . . . . . . . . . . . . . . . . . Guillaume Junion and Krzysztof Jagla Index . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

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Contributors FRANCISCO ALTAMIRANO • Centro de Estudios Moleculares de la Célula, ICBM, Facultad de Medicina, Universidad de Chile, Santiago, Chile STEPHEN E. ALWAY • Laboratory of Muscle Biology and Sarcopenia, Department of Exercise Physiology and Center for Cardiovascular and Respiratory Sciences, West Virginia University School of Medicine, Morgantown, WV, USA JUDY E. ANDERSON • Department of Biological Sciences, Department of Human Anatomy and Cell Science, University of Manitoba, Winnipeg, MB, Canada JOHN C. ANGELLO • Department of Biochemistry, University of Washington, Seattle, WA, USA KEITH BAAR • Departments of Neurobiology, Physiology, and Behavior, and Physiology and Membrane Biology, University of California, Davis, CA, USA GUY M. BENIAN • Department of Pathology, Emory University, Atlanta, GA, USA SHEPHALI BHATNAGAR • Department of Anatomical Sciences and Neurobiology, University of Louisville School of Medicine, Louisville, KY, USA NATHAN C. BIRD • Department of Biology, Wesleyan University, Hall-Atwater Laboratories, Middletown, CT, USA SUE C. BODINE • Departments of Neurobiology, Physiology, and Behavior, and Physiology and Membrane Biology, University of California, Davis, CA, USA ANTON L. BRYANTSEV • Department of Biology, University of New Mexico, Albuquerque, NM, USA MARIANA CASAS • Centro de Estudios Moleculares de la Célula, ICBM, Facultad de Medicina, Universidad de Chile, Santiago, Chile MARCO CASSANO • Translational Cardiomyology, Stem Cell Research Institute, Catholic University of Leuven, Leuven, Belgium ERIC J. CAVANAUGH • Department of Cell Biology and Anatomy, Chicago Medical School, Rosalind Franklin University of Medicine and Science, North Chicago, IL, USA MARIA CHECHENOVA • Department of Biology, University of New Mexico, Albuquerque, NM, USA JENNIFER R. CREW • Department of Cell Biology and Anatomy, Chicago Medical School, Rosalind Franklin University of Medicine and Science, North Chicago, IL, USA STEFANIA CRIPPA • Translational Cardiomyology, Stem Cell Research Institute, Catholic University of Leuven, Leuven, Belgium RICHARD M. CRIPPS • Department of Biology, University of New Mexico, Albuquerque, NM, USA ROBERT G. CUTLIP • Health Effects Laboratory Division, National Institute for Occupational Safety and Health, West Virginia and Department of Exercise Physiology, West Virginia University School of Medicine, Morgantown, WV, USA CAROLINE S. DACWAG VALLASTER • Department of Cell Biology, University of Massachusetts Medical School, Worcester, MA, USA

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JASON M. DAHLMAN • Department of Molecular Virology, Immunology, and Medical Genetics, Human Cancer Genetics Program, Integrated Biomedical Sciences Graduate Program, The Ohio State University, Columbus, OH, USA MARIA ELENA DANOVIZ • Department of Biological Structure, School of Medicine, University of Washington, Seattle, WA, USA TERESA A. DAVIS • Department of Pediatrics, USDA/ARS Children’s Nutrition Research Center, Baylor College of Medicine, Houston, TX, USA STEPHEN H. DEVOTO • Department of Biology, Wesleyan University, Hall-Atwater Laboratories, Middletown, CT, USA JOSEPH X. DIMARIO • Department of Cell Biology and Anatomy, Chicago Medical School, Rosalind Franklin University of Medicine and Science, North Chicago, IL, USA PHILIP DORAN • Department of Biology, National University of Ireland Maynooth, Maynooth, Co. Kildare, Ireland DONGSHENG DUAN • Department of Molecular Microbiology and Immunology, School of Medicine, University of Missouri, Columbia, MO, USA JEAN-MICHEL ESCOFFRE • CNRS; IPBS (Institut de Pharmacologie et de Biologie Structurale), Université de Toulouse, Toulouse, France RAMÓN ESPINOZA-LEWIS • Cardiovascular Research Division, Department of Cardiology, Children’s Hospital Boston, Harvard Medical School, Boston, MA, USA JERRY P. EU • Department of Medicine, Division of Pulmonary, Allergy and Critical Care, Duke University, Durham, NC, USA CHEN-MING FAN • Department of Embryology, Carnegie Institution of Washington, Baltimore, MD, USA MARTA L. FIOROTTO • Department of Pediatrics, USDA/ARS Children’s Nutrition Research Center, Baylor College of Medicine, Houston, TX, USA MURIEL GOLZIO • CNRS; IPBS (Institut de Pharmacologie et de Biologie Structurale), Université de Toulouse, Toulouse, France EMANUELA GUSSONI • Program in Genomics and Division of Genetics, Children’s Hospital Boston, Boston, MA, USA DENIS C. GUTTRIDGE • Department of Molecular Virology, Immunology, and Medical Genetics, Human Cancer Genetics Program, Integrated Biomedical Sciences Graduate Program, Arthur G. James Comprehensive Cancer Center, The Ohio State University, Columbus, OH, USA STEPHEN D. HAUSCHKA • Department of Biochemistry, University of Washington, Seattle, WA, USA CHARIS L. HIMEDA • Department of Biochemistry, University of Washington, Seattle, WA, USA ZHAN-PENG HUANG • Cardiovascular Research Division, Department of Cardiology, Children’s Hospital Boston, Harvard Medical School, Boston, MA, USA ANTHONY N. IMBALZANO • Department of Cell Biology, University of Massachusetts Medical School, Worcester, MA, USA GIUSEPPE INESI • California Pacific Medical Center Research Institute, San Francisco, CA, USA KRZYSZTOF JAGLA • GReD, INSERM U931, CNRS UMR6247, Ferrand, France

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ENRIQUE JAIMOVICH • Centro de Estudios Moleculares de la Célula, ICBM, Facultad de Medicina, Universidad de Chile, Santiago, Chile ANDREW R. JUDGE • Department of Physical Therapy, University of Florida, Gainesville, FL, USA GUILLAUME JUNION • GReD, INSERM U931, CNRS UMR6247, Faculté de Medecine, Clermont University, Clermont-Ferrand, France; Gene Expression Unit, European Molecular Biology Laboratory – EMBL, Heidelberg, Germany SUSAN C. KANDARIAN • Department of Health Sciences, Boston University, Boston, MA, USA ASHOK KUMAR • Department of Anatomical Sciences and Neurobiology, University of Louisville School of Medicine, Louisville, KY, USA ELISA M. LABEAU • Department of Biology, University of New Mexico, Albuquerque, NM, USA ARIYA D. LAPAN • Biological and Biomedical Sciences, Harvard Medical School, Boston, MA, USA CHRISTOPH LEPPER • Department of Embryology, Carnegie Institution of Washington, Baltimore, MD, USA CAROLINE LEWIS • Department of Biology, National University of Ireland Maynooth, Maynooth, Co. Kildare, Ireland TYANNA L. LOVATO • Department of Biology, University of New Mexico, Albuquerque, NM, USA JIANJIE MA • Department of Physiology and Biophysics, UMDNJ-Robert Wood Johnson Medical School, Piscataway, NJ, USA CHANDRASHEKARA MALLAPPA • Department of Cell Biology, University of Massachusetts Medical School, Worcester, MA, USA GERHARD MEISSNER • Department of Biochemistry and Biophysics, School of Medicine, University of North Carolina, Chapel Hill, NC, USA WATARU MIZUNOYA • Department of Biological Sciences, Department of Human Anatomy and Cell Science, University of Manitoba, Winnipeg, MB, Canada; Department of Animal and Marine Bioresource Sciences, Kyushu University, Fukuoka, Japan GINNY R. MORRISS • Department of Biology, University of New Mexico, Albuquerque, NM, USA VASSILIS MOUGIOS • Department of Physical Education and Sport Science, Aristotle University of Thessaloniki, Thessaloniki, Greece YASUYUKI OHKAWA • Department of Epigenetics and SSP Stem Cell Unit, Kyushu University, Fukuoka, Japan KAY OHLENDIECK • Department of Biology, National University of Ireland Maynooth, Maynooth, Co. Kildare, Ireland PAUL OLEYNIK • Ottawa Hospital Research Institute, Ottawa, ON, Canada CHARLES P. ORDAHL • Department of Anatomy, School of Medicine, University of California, San Francisco, CA, USA GIACOMO PALAZZOLO • Translational Cardiomyology, Stem Cell Research Institute, Catholic University of Leuven, Leuven, Belgium SIVA K. PANGULURI • Department of Anatomical Sciences and Neurobiology, University of Louisville School of Medicine, Louisville, KY, USA

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ALESSANDRA PASUT • Department of Cellular and Molecular Medicine, University of Ottawa, Ottawa, ON, Canada ILARIA PERINI • Translational Cardiomyology, Stem Cell Research Institute, Catholic University of Leuven, Leuven, Belgium ANATOLI PETRIDOU • Department of Physical Education and Sport Science, Aristotle University of Thessaloniki, Thessaloniki, Greece CHRISTOPHE PICHAVANT • Department of Neurosciences, CHUL Research Center, Quebec City, QC, Canada ANAND MOHAN PRASAD • California Pacific Medical Center Research Institute, San Francisco, CA, USA HIROSHI QADOTA • Department of Pathology, Emory University, Atlanta, GA, USA MATTIA QUATTROCELLI • Translational Cardiomyology, Stem Cell Research Institute, Catholic University of Leuven, Leuven, Belgium MICHAEL A. RUDNICKI • Department of Cellular and Molecular Medicine, University of Ottawa, Ottawa, ON, Canada KATHRYN M. RYAN • Department of Biology, University of New Mexico, Albuquerque, NM, USA MAURILIO SAMPAOLESI • Translational Cardiomyology, Stem Cell Research Institute, Catholic University of Leuven, Leuven, Belgium SARAH M. SENF • Department of Applied Physiology and Kinesiology, University of Florida, Gainesville, FL, USA JIN-HONG SHIN • Department of Molecular Microbiology and Immunology, School of Medicine, University of Missouri, Columbia, MO, USA CATHERINE L. SMITH • Department of Biochemistry, University of Washington, Seattle, WA, USA HORACIO A. SOSA JR • Department of Pediatrics, USDA/ARS Children’s Nutrition Research Center, Baylor College of Medicine, Houston, TX, USA PHILLIP W.L. TAI • Department of Biochemistry, University of Washington, Seattle, WA, USA JUSTIN TEISSIÉ • CNRS; IPBS (Institut de Pharmacologie et de Biologie Structurale), Université de Toulouse, Toulouse, France JACQUES P. TREMBLAY • Department of Neurosciences, CHUL Research Center, Quebec City, QC, Canada SARA J. VENTERS • Department of Neurosurgery, School of Medicine, University of California, San Francisco, CA, USA DA-ZHI WANG • Cardiovascular Research Division, Department of Cardiology, Children’s Hospital Boston, Harvard Medical School, Boston, MA, USA NOAH WEISLEDER • Department of Physiology and Biophysics, UMDNJ-Robert Wood Johnson Medical School, Piscataway, NJ, USA KRISTY J. WILSON • Department of Pathology, Emory University, Atlanta, GA, USA STEFANIE E. WINDNER • Department of Biology, Wesleyan University, Hall-Atwater Laboratories, Middletown, CT, USA ASHLEY C. WOZNIAK • Department of Biological Sciences, Department of Human Anatomy and Cell Science, Faculty of Medicine, University of Manitoba, Winnipeg, MB, Canada

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CHIA-LING WU • Department of Health Sciences, Boston University, Boston, MA, USA ZIPORA YABLONKA-REUVENI • Department of Biological Structure, School of Medicine, University of Washington, Seattle, WA, USA YONGPING YUE • Department of Molecular Microbiology and Immunology, School of Medicine, University of Missouri, Columbia, MO, USA JINGSONG ZHOU • Department of Molecular Biophysics and Physiology, Rush University, School of Medicine, Chicago, IL, USA

Part I Isolation and Analysis of Skeletal Muscle Precursor Cells

Chapter 1 Isolation and Characterization of Human Fetal Myoblasts Ariya D. Lapan and Emanuela Gussoni Abstract Dissociated human fetal skeletal muscle contains myogenic cells, as well as non-myogenic cells such as adipocytes, fibroblasts, and lymphocytes. It is therefore important to determine an efficient and reliable isolation method to obtain a purer population of myoblasts. Toward this end, fluorescence-activated cell sorting in conjunction with robust myogenic cell surface markers can be utilized to enrich for myoblasts in dissociated muscle. In this chapter, we describe a method to significantly enrich for myoblasts using melanoma cell adhesion molecule (MCAM), which we have determined to be an excellent marker of human fetal myoblasts. The myoblasts resulting from this isolation method can then be expanded in vitro and still retain significant myogenic activity as shown by an in vitro fusion assay. The ability to isolate a highly myogenic population from dissociated muscle facilitates the in vitro study of skeletal muscle development and muscle diseases. Furthermore, robust expansion of these cells will lead to new insights in the development of cell-based therapies for human muscle disorders. Key words: Human muscle, Tissue dissociation, Fluorescence-activated cell sorter, Myoblast purification

1. Introduction Efficient isolation and maintenance of human myoblasts in vitro is an essential technique for the investigation of myogenic progenitor commitment and differentiation, the characterization of muscle development and human muscle disorders, and the development of cell-based therapies for muscle diseases. Initial studies involved the use of tissue explants or unpurified dissociated cells (1–7). However, due to the presence of non-myogenic mononuclear cells in human skeletal muscle, including adipocytes, fibroblasts, and lymphocytes, methods for the isolation of purer populations of myoblasts were developed. In 1974, Stephen Hauschka determined in vitro conditions for the clonal culture and differentiation

Joseph X. DiMario (ed.), Myogenesis: Methods and Protocols, Methods in Molecular Biology, vol. 798, DOI 10.1007/978-1-61779-343-1_1, © Springer Science+Business Media, LLC 2012

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of human muscle cells (8, 9). These culture conditions were further adapted by Blau and Webster, who introduced the use of preplating to remove fibroblasts followed by the generation of myogenic clones (10). For more specific and efficient isolation of myoblasts within a dissociated human muscle sample, Webster et al. utilized fluorescence-activated cell sorting (FACS) to positively select for cells expressing human neural cell adhesion molecule (NCAM) (11), a cell surface antigen shown to be expressed on myogenic cells (12). Alternatively, Baroffio et al. used FACS to enrich for human myoblasts on the basis of cell size; after expansion, the cells were tested for the expression of NCAM to confirm their myogenicity (13). The prospective isolation of pure populations of myogenic progenitors is highly desirable for translational research. Currently, many types of stem/progenitor cells are under study for their potential to repair diverse tissues, including skeletal muscle. As new technologies enable reliable propagation of multipotent or pluripotent cells, such as ES or induced pluripotent (iPS) cells, there is also an emerging need to optimize methods for the selection of lineage-specific progenitors obtained following the differentiation of these pluripotent cells. As a result, the identification of markers that prospectively enrich for cells with a specific potential is currently being researched. To identify such markers, gene expression studies on progenitor cells derived from embryonic or fetal tissues might be useful as these cells are likely still immature and retain long-term expansion potential, yet are developmentally specified toward a particular tissue. Recently, melanoma cell adhesion molecule (MCAM), also known as CD146, Mel-CAM, MUC18, A32 antigen, and S-Endo-1, was detected in human skeletal muscle as well as in other normal tissues such as smooth muscle, endothelium, and the nervous system (reviewed in refs. (14–16)). Interestingly, studies in chick embryos also showed that MCAM is expressed in somatic cells that specify the myotome during development (17). Following a microarray screening to identify genes regulated during human fetal myoblast fusion, MCAM was found highly expressed in proliferating myoblasts and significantly downregulated during fusion (18). Freshly isolated MCAM-positive cells were shown to undergo fusion in vitro whereas MCAM-negative cells did not (18). Furthermore, inhibition of MCAM expression in myoblasts by RNAi enhanced myoblast differentiation and fusion (18). Therefore, significant enrichment of myoblasts from dissociated muscle can be obtained by using MCAM as a positive selection marker in cell sorting. The present chapter describes the dissociation of human fetal skeletal muscle and the subsequent purification of myoblasts using cell sorting based upon MCAM expression. In vitro culture of these purified cells is then detailed as well as an assessment of myogenicity using an in vitro fusion assay.

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Isolation of Human Fetal Myoblasts

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2. Materials 2.1. Dissociation of Primary Human Fetal Skeletal Muscle Tissue (see Note 1)

1. Protected disposable scalpels with stainless steel blade size #10. 2. Sterile 10 cm tissue culture-treated plastic dishes. 3. Assorted sterile 5, 10, and 25 mL pipettes. 4. Sterile 0.22 μm PES (low protein binding) filters – 250 and 500 mL volumes. 5. 0.22 μm CN filter unit – 500 mL volume. 6. Sterile 15 and 50-mL conical centrifuge tubes. 7. BD Falcon sterile nylon cell strainers – 100 and 40-μm pore sizes. 8. 10× Hank’s balanced saline solution (HBSS), free of calcium chloride, magnesium chloride and magnesium sulfate, diluted to 1× with double distilled water and filter sterilized with a 0.22 μm CN filter. This solution can be stored at 4°C or room temperature. 9. Complete growth medium (500 mL): Mix 395 mL of high glucose Dulbecco’s modified Eagle’s medium (DMEM) with 100 mL of fetal bovine serum (FBS; see Note 2) and 5 mL of 100× penicillin–streptomycin–glutamine (PSG). Sterilize by filtering the solution through a 500 mL 0.22 μm PES filter unit. Store at 4°C and use within 1 month. 10. Sterile red blood cell lysis solution (Qiagen), stored at room temperature. 11. Sterile HEPES buffered saline solution, without phenol red. 12. 1 M calcium chloride solution (CaCl2·2H2O, FW 147). Dissolve 1.47 g powder in 10 mL of double distilled water. Store at 4°C. 13. Dispase stock solution: dissolve 1 g powder dispase II (Roche Applied Science) in 100 mL HEPES buffered saline. Add 316 mL of high glucose DMEM to generate a stock solution of 2.4 U/mL. Filter-sterilize the solution through a PES 500 mL filter; aliquot into 15-mL conical tubes (10 mL/tube) and store aliquots at −20°C. 14. Collagenase D stock solution: dissolve 2.5 g powder collagenase D (Roche Applied Science) in 250 mL solution of 1× HBSS supplemented with 1.25 mL of 1 M CaCl2. Sterilize by filtering through a PES filter unit. The filtered solution can be dispensed in 15-mL conical tubes (10 mL/tube) and stored at −20°C. 15. Sterile freezing medium: 90% FBS and 10% dimethyl sulfoxide (DMSO). Prepare freezing medium and immediately store

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on ice. Unused sterile freezing medium can be stored at 4°C for up to 4 weeks. 16. Sterile 1.8 mL CryoTube™ vials. 17. Bench top centrifuge. 18. Hemocytometer. 19. Sterile laminar flow hood. 20. −150°C freezer, liquid nitrogen storage tank. 21. Humidified 5% CO2 incubator set to 37°C. 2.2. Isolation of Myoblasts from Dissociated Human Fetal Skeletal Muscle

1. CryoTube™ vial containing dissociated human fetal skeletal muscle (Hu Fe SkM).

2.2.1. Thawing of Cryopreserved Sample Prior to FACS

4. Sterile tissue culture-treated plastic dishes – 10 or 15 cm size.

2. Sterile 0.22 μm PES filter – 500 mL volume. 3. Sterile 50-mL conical centrifuge tubes. 5. Sterile complete growth medium, as mentioned above. 6. Sterile laminar flow hood. 7. Water bath set to 37°C. 8. CO2 incubator, as above.

2.2.2. Preparation of Sample for FACS

1. Dissociated Hu Fe SkM sample thawed 1 day prior to FACS. 2. Sterile 0.22 μm PES filter – 500 mL volume. 3. Sterile 15 and 50-mL conical centrifuge tubes. 4. Sterile nylon cell strainers – 40-μm pore size. 5. Sterile 1× HBSS, as above. 6. Sterile cell dissociation buffer, enzyme free, PBS-based (Invitrogen) stored at room temperature. 7. Sterile 0.5% BSA/HBSS solution (500 mL): dissolve 2.5 g bovine serum albumin (BSA) in 1× HBSS. Sterilize by filtering the solution through a 500 mL 0.22 μm PES filter unit. Store at 4°C. 8. Antibodies (all stored at 4°C): (a) Anti-MCAM antibody, clone P1H12 (Millipore). (b) Mouse IgG1 monoclonal antibody (BD Pharmingen). (c) Alexa Fluor® 488 goat anti-mouse IgG (Invitrogen). Protect from light. 9. 10 mL syringe and 0.22 μm Acrodisc® Supor membrane low protein binding syringe filters (Pall Life Sciences). 10. 1 mg/mL Propidium iodide (PI): resuspend 10 mg powder in 10 mL of double distilled water, and then filter sterilize using a 0.22-μm syringe filter. Dispense in 1 mL aliquots and store at 4°C.

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11. Sterile FACS or 5 mL round-bottom tubes. 12. Sterile laminar flow hood. 13. Bench top centrifuge. 14. Inverted microscope. 15. Hemocytometer. 2.2.3. FluorescenceActivated Cell Sorting

1. FACS or 5-mL round-bottom tubes. 2. Cell sorting machine. 3. Cell sorting software.

2.3. In Vitro Culture and Analysis of Human Fetal Skeletal Myoblasts

1. Sterile 50-mL conical centrifuge tubes.

2.3.1. In Vitro Cell Culture

4. Sterile 0.22 μm PES (low protein binding) filter – 150 mL tube top volume.

2. Sterile tissue culture-treated plastic dishes. 3. BD Falcon black with clear bottom 96-well Microtest™ Optilux™ plates.

5. Sterile 1× HBSS. 6. Sterile complete growth medium, as above. 7. Differentiation medium (50 mL): Mix 48.5 mL of low glucose Dulbecco’s Modified Eagle’s Medium (DMEM) with 1 mL of horse serum (HS) and 0.5 mL of 100× PSG. Sterilize by filtering the solution through a 150 mL 0.22 μm PES filter unit. Store at 4°C and use within 1 month. 8. 0.15% Gelatin: add 0.75 g of gelatin to 500 mL of double distilled water. Do not shake. Sterilize the solution by autoclaving for 20 min and store at 4°C. 9. TrypLE™ Express Dissociation Enzyme with Phenol Red (Invitrogen). 10. Sterile laminar flow hood. 11. Water bath set to 37°C. 12. CO2 incubator. 13. Bench top centrifuge. 14. Inverted microscope. 15. Hemocytometer. 2.3.2. Immunocytochemistry for In Vitro Fusion Assay

1. 10× Phosphate buffered saline (PBS), diluted to 1× with double distilled water. Store at room temperature. 2. 4% Paraformaldehyde (4% PFA): Dilute 16% paraformaldehyde with 1× PBS. Use with caution as paraformaldehyde is extremely toxic; it is recommended that paraformaldehyde be used in a fume hood for safety. Aliquot and store at −20°C. Aliquots should not be repeatedly freezed and thawed; discard unused PFA after initial use.

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3. Permeabilization solution: Mix 50 μL of Triton ×-100 with 10 mL of 1× PBS. 4. Blocking solution: Mix together 1 mL of fetal bovine serum (FBS), 10 μL of Triton ×-100, and 9 mL of 1× PBS. 5. Antibodies (all stored at 4°C): (a) Anti-Human Desmin, clone D33 (Dako). (b) Alexa Fluor® 488 goat anti-mouse IgG (Invitrogen). Protect from light. 6. DAPI solution (a) DAPI stock solution (5 mg/mL): Dissolve 10 mg DAPI in 2 mL of double distilled water. Aliquot and store at −20°C. (b) DAPI working solution (100 ng/mL): Mix 2 μL of DAPI stock solution with 100 mL of PBS. Store at 4°C wrapped in aluminum foil to protect from light. 7. Inverted microscope with epi-fluorescence capabilities including ultraviolet/DAPI and FITC/GFP filter sets.

3. Methods 3.1. Dissociation of Primary Human Fetal Skeletal Muscle Tissue

All steps in this protocol should be performed in a sterile laminar flow hood using sterile tissue culture technique. 1. Preweigh one 10 cm tissue culture plate, and place the tissue sample to be dissociated in a second (non preweighed) 10 cm tissue culture plate. 2. Using sterile scalpels, remove and discard any remaining skin and bone from the muscle tissue. Tissue should be kept moist in sterile 1× HBSS. Add a few drops of sterile 1× HBSS to tissue as necessary, to prevent it from drying out. After skin is removed, place muscle tissue in the preweighed 10 cm tissue culture plate and weigh the plate again. Subtract from this number the tare of the empty plate to calculate the amount of muscle tissue to be dissociated. 3. Thaw frozen aliquots of dispase II and collagenase D in a 37°C water bath. Thawed collagenase and dispase stocks will be added at a volume of 3.5 mL each per gram of muscle tissue to be dissociated. Thaw only the amounts of collagenase D and dispase II necessary for dissociation. If an excess of enzymes is thawed, it can be refrozen once and reused. 4. Using sterile scalpels, mince muscle tissue until it resembles a fine paste. During mincing, add a few drops of sterile 1× HBSS to prevent exposed tissue from drying out. Tissue should always appear moist, but with no excess of liquid.

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5. After tissue is finely minced, add equal amounts of the thawed dispase II and collagenase D solutions. The final concentration will be 5 mg/mL for collagenase D and 1.2 U/mL for dispase II in this solution. Pipette minced tissue and enzyme solution up and down through a sterile 25 mL pipette a few times. 6. Incubate plate in a tissue culture incubator at 37°C with 5% CO2 for 15 min. Then pipette the digestion solution up and down through a sterile 25 mL pipette a few times and incubate again for 15 min. Repeat this step an additional 1–2 times, until the slurry easily passes though a sterile 5 mL pipette and all tissue chunks are dissolved. The total digestion time will range between 45 min and 1 h 15 min. 7. Add 2 volumes of complete growth medium to the digested slurry and filter the digestion solution through a 100-μm cell strainer over a 50-mL conical tube. Change cell strainer if it appears clogged. 8. Pellet cells for 10 min at 329 × g, room temperature. 9. Resuspend the pellet in 1 volume of complete growth medium (i.e., 3 mL) and add 7 volumes (i.e., 21 mL) of red blood cell lysis solution. Invert the tube a few times and then filter the solution through a 40-μm cell strainer over a 50-mL conical tube. 10. Count cells using a hemocytometer, and then pellet the cells for 10 min at 329 × g, room temperature. 11. Freeze cells at a concentration of 107 cells/mL in ice-cold freezing medium. Store cryovials at −80°C for ~2–3 days, then transfer them to −150°C where they can be permanently stored until necessary. 3.2. Isolation of Myoblasts from Dissociated Human Fetal Skeletal Muscle

All steps in this protocol except for cell sorting (see Subheading 3.2.3) should be performed in a sterile laminar flow hood using sterile tissue culture technique. Cell sorting should be performed in as clean an environment as possible.

3.2.1. Thawing of Cryopreserved Sample Prior to FACS

Cryopreserved cells should be carefully thawed and plated 1 day prior to cell sorting. This allows the cells to recover from the freezing process before undergoing FACS. 1. Prewarm complete growth medium in a water bath set to 37°C. Then, pipette 10 mL prewarmed medium into a sterile 50-mL conical tube. 2. Carefully and quickly thaw a vial of cryopreserved, dissociated Hu Fe SkM cells in a 37°C water bath and transfer the cells into the 50-mL conical tube with prewarmed proliferation medium using a 1-mL pipette. Rinse the inside of the cryovial with fresh complete growth medium to remove as many cells as possible. This step should be performed very quickly as the

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DMSO used during the cryopreservation process is toxic to the cells at room temperature. 3. Plate the cells in the prewarmed medium onto sterile, tissueculture treated plates at approximately 0.5–1 × 107 cells/10 cm plate or 1.5–3 × 107 cells/15 cm plate (see Note 3). If using a 15 cm plate, add 15 mL prewarmed medium to bring the total medium volume to 25 mL. 4. Incubate the cells in a CO2 incubator overnight at 37°C. 3.2.2. Preparation of Sample for FACS

1. Prewarm the following in a 37°C water bath: complete growth medium, 1× HBSS, and cell dissociation buffer. Place the 0.5% BSA/HBSS on ice. 2. Check your cells under a phase contrast microscope with 10× magnification (see Note 4). Ensure that there is no contamination and that the cells look healthy. 3. Set-up two 50-mL conical tubes per plate of cells thawed. 4. Save the old plate medium by carefully removing the medium from the plate and pipetting it into the first 50-mL conical tube. 5. Wash the cells with 5 mL (10 cm plate) or 10 mL (15 cm plate) 1× HBSS, and then save the wash by pipetting it into the second 50-mL conical tube (see Note 5). 6. Pipette 3 mL (10 cm plate) or 10 mL (15 cm plate) cell dissociation buffer onto the plate of washed cells and incubate in a humidified CO2 incubator for 10–12 min (see Note 6). 7. After incubation, remove the cells by gently lifting the cells off the plate. This is done by tilting the plate at an angle (~45°) and carefully pipetting the dissociation buffer currently in the plate onto the surface of the plate. Repeat this process 5–10 times (see Note 7). Pipette the dissociation buffer (containing the cells) into the 50-mL conical tube with the old medium (see Note 8). 8. Repeat step 7 twice more with 1× HBSS washes (5 mL for 10 cm plate and 10 mL for 15 cm plate). Save the washes in the second 50-mL conical tube. 9. Check the plate under a phase contrast microscope at 10× magnification for the presence of cells. There should be very few cells on the surface of the plate after this process. 10. Centrifuge the 50-mL conical tubes containing the cells and washes at 329 × g at 4°C for 10 min. 11. Resuspend the cells in 0.5–2 mL ice-cold 0.5% BSA/HBSS and combine the cells from both 50 mL conical tubes. 12. Filter the cells through a sterile 40 μm cell strainer over a 50 mL conical tube. Wash the cell strainer with an additional 0.5–1 mL of ice-cold 0.5% BSA/HBSS.

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13. Determine the cell concentration using a hemocytometer or other cell counting device. 14. Adjust the cell concentration to 2 × 107 cells/mL in ice-cold 0.5% BSA/HBSS. This may require an extra centrifugation step to spin down and resuspend the cells in a smaller volume. Cells should be centrifuged for 10 min at 329 × g at 4°C. 15. For FACS compensation controls, remove the following number of cells, pipette into a FACS tube (see Note 9), and adjust the volume to 200 μL by adding 175 μL ice-cold 0.5% BSA/ HBSS: (a) “No stain” compensation control → 0.5 × 106 cells. Store on ice in the dark. (b) “PI” compensation control → 0.5 × 106 cells. Add 0.4 μL of 1 mg/mL PI (final conc.: 2 μg/mL). Store on ice in the dark (see Note 10). 16. Primary antibody incubation: (a) For the primary antibody isotype control (mIgG), remove 0.5 × 106 cells (in 25 μL) and pipette into a 15 mL conical tube. Add 0.25 μL of unconjugated mIgG1 isotype control antibody (1:100 dilution). Place on ice. (b) For the remainder of the sample, add the unconjugated MCAM antibody to the sample at a final concentration of 0.5 μg/106 cells (1:100 dilution). Place on ice. Although this antibody is available in several conjugated formats, we have noted that the separation between the positive and negative MCAM populations is decreased by the conjugation. As a result, we recommend the use of the unconjugated antibody with a conjugated secondary antibody for maximum separation and sort purity (see Fig. 1). 17. Incubate the isotype control and MCAM samples on ice for 30 min. 18. After this incubation, wash the cells by adding ice-cold 0.5% BSA/HBSS (2–3 mL for isotype control and 10–20 mL for every 2 × 107 MCAM-labeled cells). Centrifuge the cells for 10 min at 329 × g at 4°C. 19. Resuspend the cells at 2 × 107 cells/mL in ice-cold 0.5% BSA/ HBSS. 20. Secondary antibody incubation (see Note 11): (a) For the primary antibody isotype control (mIgG), add 0.25 μL of Alexa Fluor® 488-conjugated anti-mouse antibody (1:100 dilution). Place on ice. (b) For the MCAM antibody-stained sample, add Alexa Fluor® 488-conjugated anti-mouse antibody at a 1:100 dilution. Place on ice.

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Fig. 1. Comparison of unconjugated and conjugated anti-MCAM, clone P1H12, antibodies. FACS plots illustrate a 17-week human fetal sample stained with (a) unconjugated anti-MCAM primary and PE-conjugated secondary antibodies, or (b) anti-MCAM antibody directly conjugated to PE, (c) FITC, or (d) AF488. Note the separation between the MCAM positive and negative populations. Populations positive for MCAM are marked by the gates.

21. Incubate the isotype control and MCAM samples with the secondary antibody on ice for 30 min. 22. After this incubation, wash the cells by adding ice-cold 0.5% BSA/HBSS (2–3 mL for isotype control and 10–20 mL for every 2 × 107 MCAM-labeled cells). Centrifuge the cells for 10 min at 329 × g at 4°C. 23. Resuspend the primary antibody isotype control in 200 μL icecold 0.5% BSA/HBSS and pipette into a new FACS tube. Add 0.4 μL of 1 mg/mL PI (final conc.: 2 μg/mL). Store on ice in the dark. 24. Resuspend the MCAM-labeled sample at 2 × 107 cells/mL in ice-cold 0.5% BSA/HBSS. 25. Transfer 0.5 × 106 cells (25 μL) of the MCAM-labeled sample to a new FACS tube. Adjust the volume to 200 μL by adding

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175 μL ice-cold 0.5% BSA/HBSS. Reserve these cells as the “Alexa Fluor® 488” compensation control. Store on ice in the dark. 26. Filter the remainder of the MCAM-labeled cells through a sterile 40 μm cell strainer over a 50 mL conical tube. Wash the cell strainer with ice cold 0.5% BSA/HBSS (approximately half the volume that the cells are currently in). 27. Add 1 mg/mL PI to the MCAM-labeled sample at a final concentration of 2 μg/mL. 28. Transfer the MCAM-labeled sample to a new FACS tube. Store on ice in the dark. 29. Prepare collection tube for sorted cells by pipetting 0.5 mL ice-cold 0.5% BSA/HBSS into a new FACS tube. Store on ice. 3.2.3. FluorescenceActivated Cell Sorting

It is beyond the scope of this chapter to review FACS or flow cytometry in detail. Please refer to literature specific to this technique, such as Current Protocols in Cytometry (Wiley and Sons) for further information. Here, gating specifications are briefly indicated. 1. Determine optimal excitation voltages and compensation values using the “no stain” and single color compensation controls. 2. Visualize the “PI” compensation control on a PI vs. forward or side scatter graph. Gate for live cells based upon PI exclusion (i.e., PI-negative cells) (see Fig. 2a). 3. Determine the positive and negative MCAM populations: visualize the “primary antibody isotype” control and MCAMlabeled sample on an Alexa Fluor® 488 vs. forward or side

Fig. 2. Example of FACS gating for human fetal cells immunostained with anti-MCAM antibody and PI: (a) Live cells selected by gating on the PI-negative population. (b) Cells labeled with mIgG isotype control. (c) Cells labeled with antiMCAM antibody. Comparison of the plots in (b, c) clearly defines a single MCAM-positive population in the anti-MCAM immunostained sample.

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scatter graph (see Note 12). There should be one MCAMnegative population in the “primary antibody isotype” control, while two populations, one MCAM-positive and one MCAMnegative, should be seen in the MCAM-labeled sample (see Fig. 2b, c). Gate and sort for the MCAM-positive cell population based upon this comparison. 3.3. In Vitro Culture and Analysis of Human Fetal Skeletal Myoblasts 3.3.1. In Vitro Cell Culture

All steps in this protocol should be performed in a sterile laminar flow hood using sterile tissue culture technique. 1. Coat sterile tissue culture-treated plates with 10 mL (10 cm plate) 0.15% gelatin for 1 h at 37°C. After incubation, remove the gelatin solution by aspiration. Then, slightly angle the plates for ~10 min to pool any excess liquid. Remove any excess liquid by aspiration. Coated plates can be stored at 4°C for up to 1 week; prewarm stored, coated plates in a CO2 incubator prior to use. 2. Prewarm complete growth medium in a water bath set to 37°C. 3. Resuspend sorted MCAM-positive cells at 0.5–1 × 106 cells/10 mL complete growth medium and plate on coated plates from step 1 (10 mL/plate). Gently rock plate(s) to evenly distribute cells, and then incubate in a CO2 incubator. Sorted cells will be small and have a bright, rounded appearance but will attach within 1–2 days postsorting. Figure 3a illustrates sorted MCAM-positive cells 1 day after sorting. 4. Propagate the cells to 60–75% confluency (see Fig. 3b; Note 13). This should take approximately 2–3 days; however, if necessary, replace the medium with fresh growth medium every 2 days until the plate is at 60–75% confluency.

Fig. 3. In vitro culture of FACS sorted MCAM-positive human fetal cells. (a) Sorted MCAM+ cells 1 day after sorting and plating. Most cells appear bright and rounded, while some cells have flattened and firmly adhered to the plate. (b) Sorted MCAM+ cells 7 days after sorting. Most cells are now elongated and firmly adhered to the plate.

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5. To passage cells: (a) Coat sterile tissue culture-treated plates as mentioned above. (b) Prewarm the following in a water bath set to 37°C: 1× HBSS, TrypLE™ Express dissociation enzyme, and complete growth medium. (c) Remove medium from plate by aspiration and wash the cells with 10 mL (10 cm plate) 1× HBSS. Remove HBSS by aspiration. (d) Pipette 1 mL TrypLE™ Express onto the plate and incubate in a humidified 37°C CO2 incubator for 2–3 min. Gently remove the cells from the plate using 9 mL complete growth medium and pipette into a sterile 50-mL conical tube. Wash any remaining cells from the surface of the plate with additional complete growth medium. (e) Centrifuge the cells at 329 × g at room temperature for 10 min. (f) Resuspend the cells in 10 mL fresh complete growth medium. (g) Determine the cell concentration using a hemocytometer and plate the cells at 0.5–1 × 106 cells in 10 mL complete growth medium/10 cm plate. (h) Cells should be passaged every 2–3 days and should not be grown past 75% confluency. 6. To freeze cells: (a) Trypsinize the cells as in step 5 and then resuspend in ice-cold freezing medium at desired cell concentration. (b) Store cryovials at −150°C where they can be permanently stored until necessary. 7. To perform an in vitro fusion assay: (a) Coat BD Falcon 96-well plates with 0.15% gelatin (50–100 μL/well) as above (see Note 14). (b) Trypsinize the cells and determine the cell concentration as in step 5 and then plate 7,500 cells in 100 μL complete growth medium/96-well. (c) Incubate the cells in a CO2 incubator overnight. (d) Carefully remove the growth medium from each well and replace with 100 μL prewarmed differentiation medium (see Note 15). Incubate the cells in a CO2 incubator overnight. (e) Replace the differentiation medium in each well daily during the course of the fusion assay.

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(f) Monitor the differentiation of the cells using a phase contrast microscope at 10 or 20× magnification. Fusion should occur within 1 week of exposure to differentiation medium. When mature myotubes of >10 nuclei/myotube are present, perform the following immunocytochemistry protocol to visualize the cells by fluorescence microscopy. 3.3.2. Immunocytochemistry for In Vitro fusion Assay (see Note 16)

1. Thaw 4% PFA at room temperature. 2. Carefully wash the cells with 50 μL of 1× PBS/96-well. Remove PBS using a 1–200 μL pipette (see Note 17). 3. Fix the cells with 50 μL 4% PFA for 20 min at room temperature; remove fixation liquid using a pipette, then permeabilize the cells with 50 μL permeabilization solution for 3 min at room temperature. 4. Remove permeabilization solution using a pipette, and then block the cells for 30 min at room temperature with 50 μL blocking solution. 5. Prepare the primary antibody solution by diluting the antihuman desmin antibody 1:100 in fresh blocking solution. Incubate the cells with primary antibody solution overnight at 4°C.

Fig. 4. In vitro fusion assay of sorted MCAM-positive and MCAM-negative cells. MCAM+ and MCAM− cells were isolated from a human fetal sample and cultured for 2 weeks prior to plating for a fusion assay. Cells were then differentiated for 3 days, fixed and stained for expression of desmin. MCAM+ cells fused as depicted in (a, b), while MCAM− cells were desmin-negative and not fused as shown in (c, d). (a, c) DAPI; (b, d) desmin of the same microscopic fields. White arrows point to typical myotubes observed in the MCAM+ fraction.

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6. Wash the cells 3 times with 1× PBS for 5 min at room temperature. The plate may be gently agitated on a rotating shaker. 7. Prepare the secondary antibody solution by diluting the Alexa Fluor® 488 anti-mouse antibody 1:1,000 in blocking solution. Incubate the cells in the dark with secondary antibody solution for 1 h at room temperature. 8. Wash the cells 3 times with 1× PBS for 5 min at room temperature in the dark. The plate may be gently agitated on a rotating shaker. 9. Store the cells in 100–200 μL DAPI working solution at 4°C and protect the cells from light with aluminum foil. 10. Visualize the cells by fluorescent microscopy using ultraviolet/ DAPI and FITC/GFP filter sets for DAPI and desmin, respectively. An example of fused MCAM-positive cells is shown in Fig. 4.

4. Notes 1. Institutional review and protocol approval are required prior to collection and processing of human fetal tissue. All personnel handling human tissue must receive appropriate safety and human subject education training. 2. FBS varies considerably between companies and even lot to lot from the same company. Therefore, several different FBS samples should be tested using the in vitro methods described in Subheading 3.3 to determine which lot/company works best for your myogenic cells. 3. No plate coating is required for this step as the cells are only plated for a day prior to use. 4. There will be many floating, live cells in your culture, which is normal for dissociated Hu Fe SkM. It is also likely that there will be small clumps of cells in the culture, and the number of clumps will vary. These clumps will be filtered out prior to cell sorting. Additionally, the dissociation process results in a large amount of debris in addition to cells. This will make the culture appear “dirty” (i.e., little black specks, etc.), but again, this is normal and should not be considered contamination. This debris will be removed during the FACS sample preparation process. 5. Given the large number of floating, live cells, it is important to save the old medium and the wash prior to removing the cells from the plate with cell dissociation buffer. This ensures that

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you maximize the number of cells available for FACS and subsequent analysis. 6. Cell dissociation buffer MUST be used to remove the cells from the plate when using the Chemicon anti-MCAM antibody. Trypsin removes/destroys the epitope for this antibody and therefore cannot be used for this protocol. 7. When the plate is tilted at an angle, the cells can be seen on the surface of the plate as a light opaque coating. Repeatedly rinse the cells off the plate until this coating is no longer visible. 8. The cell suspension containing cells and cell dissociation buffer must be pipetted into the old medium (or fresh medium) to deactivate the cell dissociation buffer. Prolonged exposure of the cells to the cell dissociation buffer may negatively affect the health of the cells. 9. Some FACS machines may require FACS tubes that are different in diameter/size from the tube specified in this protocol. Check that your tubes fit in your machine prior to use. 10. Propidium iodide (PI) is a membrane impermeant DNA dye that is used in this protocol to discriminate live from dead cells during the FACS. 11. For the best possible separation between antigen-expressing and nonexpressing populations, it is important to use a secondary antibody conjugated to a bright and photostable fluorophore. Invitrogen Alexa Fluor®-conjugated antibodies are brighter and more photostable than conventional conjugates such as FITC and Cy3, and they are recommended for applications requiring fluorescence detection in this protocol. 12. The Alexa Fluor® 488 dye is equivalent to FITC and GFP for fluorescence excitation and emission. Therefore, standard FITC and GFP filter sets for FACS and fluorescent microscopy can be used to visualize this dye. 13. Cells should never reach 100% confluency when proliferating as they will begin to differentiate and fuse on contact. The high serum growth medium will lower in serum concentration over time and will not be able to prevent fusion (see Note 15). 14. Black with optically clear bottom plates are recommended for optimal fluorescence detection. However, the anti-human desmin antibody used in this protocol is a robust antibody that can be detected in clear 96-well tissue culture plates if needed. 15. Low serum medium induces differentiation and fusion of myoblasts in culture (19–21). 16. This immunocytochemical protocol can also be utilized for the detection of other myogenic markers of proliferating or differentiating cells.

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17. A 1–200 μL pipette is recommended for removal of liquids in this immunocytochemical protocol due to the small surface area of a 96-well. Aspiration or large pipette tips (i.e., 1 mL pipette tip) may remove a significant number of cells from the well surface.

Acknowledgments This work was supported by grants from NIH/NINDS 2R01NS047727 and 5P50NS040828. References 1. Pogogeff IA, Murray MR (1946) Form and behavior of adult mammalian skeletal muscle in vitro. Anat Rec 95:321–335. 2. Geiger RS, Garvin JS (1957) Pattern of regeneration of muscle from progressive muscular dystrophy patients cultivated in vitro as compared to normal human skeletal muscle. J Neuropathol Exp Neurol 16:532–543. 3. Herrmann H, Konigsberg UR, Robinson G (1960) Observations on culture in vitro of normal and dystrophic muscle tissue. Proc Soc Exp Biol Med 105:217–221. 4. Goyle SS, Kalra SL, Singh B (1967) The growth of normal and dystrophic human skeletal muscle in tissue culture. Neurol India 15: 149–151. 5. Kakulas BA, Papadimitriou JM, Knight JO, Mastaglia FL (1968) Normal and abnormal human muscle in tissue culture. Proc Aust Assoc Neurol 5:79–85. 6. Skeate Y, Bishop A, Dubowitz V (1969) Differentiation of diseased human muscle in culture. Cell Tissue Kinet 2:307–310. 7. Bishop A, Gallup B, Skeate Y, Dubowitz V (1971) Morphological studies on normal and diseased human muscle in culture. J Neurol Sci 13:333–350. 8. Hauschka SD (1974) Clonal analysis of vertebrate myogenesis: II. Environmental influences upon human muscle differentiation. Dev Biol 37:329–344. 9. Hauschka SD (1982) Muscle cell culture: Future goals for facilitating the investigation of human muscle disease. In: Schotland DL (ed) Disorders of the motor unit. Wiley, New York. 10. Blau HM, Webster C (1981) Isolation and characterization of human muscle cells. Proc Natl Acad Sci 78:5623–5627. 11. Webster C, Pavlath GK, Parks DR et al (1988) Isolation of human myoblasts with the

fluorescence-activated cell sorter. Exp Cell Res 174:252–265. 12. Walsh FS, Ritter MA (1981) Surface antigen differentiation during human myogenesis in culture. Nature 289:60–64. 13. Baroffio A, Aubry JP, Kaelin A et al (1993) Purification of human muscle satellite cells by flow cytometry. Muscle Nerve 16:498–505. 14. Shih IeM (1999) The role of CD146 (Mel-CAM) in biology and pathology. J Pathol 189:4–11. 15. Ouhtit A, Gaur RL, Abd Elmageed ZY et al (2009) Towards understanding the mode of action of the multifaceted cell adhesion receptor CD146. Biochim Biophys Acta 1795:130–136. 16. Lecourt S, Marolleau JP, Fromigué O et al (2010) Characterization of distinct mesenchymal-like cell populations from human skeletal muscle in situ and in vitro. Exp Cell Res 316:2513–2526. 17. Pujades C, Guez-Guez B, Dunon D (2002) Melanoma cell adhesion molecule (MCAM) expression in the myogenic lineage during early chick embryonic development. Int J Dev Biol 46:263–266. 18. Cerletti M, Molloy MJ, Tomczak KK et al (2006) Melanoma cell adhesion molecule is a novel marker for human fetal myogenic cells and affects myoblast fusion. J Cell Sci 119: 3117–3127. 19. Simpson SB Jr, Cox PG (1967) Vertebrate regeneration system: culture in vitro. Science 157:1330–1332. 20. Yaffe D, Saxel O (1977) A myogenic cell line with altered serum requirements for differentiation. Differentiation 7:159–166. 21. Webster C, Filippi G, Rinaldi A et al (1986) The myoblast defect identified in Duchenne muscular dystrophy is not a primary expression of the DMD mutation. Hum Genet 74:74–80.

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Chapter 2 Skeletal Muscle Satellite Cells: Background and Methods for Isolation and Analysis in a Primary Culture System Maria Elena Danoviz and Zipora Yablonka-Reuveni Abstract Repair of adult skeletal muscle depends on satellite cells, myogenic stem cells located between the basal lamina and the plasmalemma of the myofiber. Standardized protocols for the isolation and culture of satellite cells are key tools for understanding cell autonomous and extrinsic factors that regulate their performance. Knowledge gained from such studies can contribute important insights to developing strategies for the improvement of muscle repair following trauma and in muscle wasting disorders. This chapter provides an introduction to satellite cell biology and further describes the basic protocol used in our laboratory to isolate and culture satellite cells from adult skeletal muscle. The cell culture conditions detailed herein support proliferation and differentiation of satellite cell progeny and the development of reserve cells, which are thought to reflect the in vivo self-renewal ability of satellite cells. Additionally, this chapter describes our standard immunostaining protocol that allows the characterization of satellite cell progeny by the temporal expression of characteristic transcription factors and structural proteins associated with different stages of myogenic progression. Although emphasis is given here to the isolation and characterization of satellite cells from mouse hindlimb muscles, the protocols are suitable for other muscle types (such as diaphragm and extraocular muscles) and for muscles from other species, including chicken and rat. Altogether, the basic protocols described are straightforward and facilitate the study of diverse aspects of skeletal muscle stem cells. Key words: Skeletal muscle, Satellite cell, Stem cell, Myogenesis, Pronase, Gelatin, Matrigel, Pax7, MyoD, Myogenin

1. Introduction This chapter aims to provide simple protocols for the isolation, culture, and analysis of satellite cells from adult skeletal muscle. We first detail background information about satellite cells (see Subheadings 1.1–1.3) and the range of cell isolation approaches developed over the years by us and others to analyze satellite cells (see Subheading 1.4). We then introduce an overview of our basic Joseph X. DiMario (ed.), Myogenesis: Methods and Protocols, Methods in Molecular Biology, vol. 798, DOI 10.1007/978-1-61779-343-1_2, © Springer Science+Business Media, LLC 2012

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satellite cell isolation and culture protocol (see Subheading 2) followed by practical details (starting with Subheading 3). We provide what we consider the simplest protocol that can be performed in any basic tissue culture laboratory, and in Subheading 1.4 we briefly discuss alternative approaches to purifying satellite cells. The basic approach provided in this chapter is an excellent means for analysis of satellite cells in culture when extreme purity is not needed. With careful attention to minimize connective tissue contribution, our standard protocol can yield cultures that are 80–95% pure based on staining for protein markers Pax7 and MyoD on culture day 4 (for additional details about these markers see Subheading 1.3). Collectively, our simple protocol for satellite cell isolation and culture has allowed detailed analyses of tissue-dissociated satellite cells. Standardized protocols for the isolation and culture of satellite cells are essential tools to enhance our understanding of cell autonomous and extrinsic factors that regulate their performance. 1.1. The Satellite Cell Is Defined by Its Niche

The functional units responsible for skeletal muscle contraction are cylindrical, multinucleated muscle fibers (myofibers). These contractile structures are established during embryogenesis, when mononuclear cells known as myoblasts fuse into immature muscle fibers or myotubes. Myonuclei (the myofiber nuclei) are postmitotic and under normal conditions cannot reenter a proliferative state to contribute additional nuclei. During postnatal life, myofiber growth, homeostasis, and repair rely on a population of mononuclear myogenic cells known as satellite cells (1–3). Satellite cells were initially described 50 years ago by their anatomical location on the surface of muscle fibers, between the myofiber plasmalemma and the basal lamina (4, 5) (for a schematic and electron microscope image see Fig. 1). However, the ultimate experimental proof that satellite cells are indeed myogenic progenitors has only been obtained by showing that cells derived from isolated myofibers

Fig. 1. A schematic (a) and EM micrograph (b) of satellite cell location. The myofiber basement and plasma membranes have been routinely detected by immunostaining with antibodies against laminin and dystrophin, respectively. (a) Myofiber nuclei depicted at the myofiber periphery represent the state of healthy adult myofibers; immature myofibers present in regenerating muscles display centralized myofiber nuclei (not shown). (b) Black arrows depict the basal lamina, white arrows depict apposing satellite cell and myofiber membranes; note the sarcomeric organization within the myofiber. A color version of this figure appeared in Yablonka-Reuveni and Day (2011).

2

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23

produce myogenic progeny, able to proliferate, differentiate, and self-renew in vitro and in vivo (6–13). Satellite cells were initially described using electron microscopy (4, 5, 14, 15). More recent methods facilitate monitoring these cells by light microscopy based on expression of a range of specific markers that can be detected by immunostaining (16, 17). In particular, specific expression of the paired box transcription factor Pax7 and availability of an excellent antibody for immunodetection of this protein provides a uniform means to identify satellite cells in their native position in a range of species including mouse (10, 12, 13, 18, 19), rat (20), chicken (21, 22), and human (23, 24). Additionally, genetically manipulated reporter mice permit direct detection of satellite cells based on specific expression of a fluorophore or b-galactosidase (b-gal) (13, 17, 19, 25, 26). We demonstrated that transgenic expression of GFP under the control of nestin regulatory elements (NES-GFP) allows detection of satellite cells in freshly isolated myofibers. NES-GFP mice also facilitate isolation of satellite cells using fluorescent-activated cell sorting (FACS) and subsequent studies of purified populations (13, 19). The Myf5nLacZ/+ mouse has also provided a means to identify satellite cells in intact muscle and isolated myofibers (2, 11, 19, 26, 27). In this mouse, one of the Myf5 alleles was modified to direct lacZ expression, resulting in b-gal expression in satellite cells as originally reported by Beauchamp et al. (26). We frequently use crosses of NES-GFP with Myf5nLacZ/+ mice, allowing the detection of satellite cells by means of direct fluorescence and X-gal staining (19). Satellite cells are considered the major, if not only, source of myogenic progeny in adult muscle (2, 3). Other cell types isolated from skeletal muscle, such as mesoangioblasts, pericytes, and myoendothelial cells also seem to have some myogenic potency (28–30), but whether these cell types participate in normal muscle maintenance and repair remains unclear. The isolation of the latter cell types requires special enrichment approaches and these cells do not appear to contribute to our myogenic preparations. The majority of cells in our standard preparations of freshly isolated myogenic progenitors display the satellite cell phenotype; i.e., preparations from Myf5nLacZ/+/NES-GFP mice are enriched with Pax7+/b-gal+/ GFP+ cells (shown by cytospin and mRNA expression analyses of freshly isolated cells). Hence, we refer to our freshly isolated cells prepared by the basic approach detailed herein as preparations of satellite cells or myogenic progenitors. Once satellite cells are cultured and proliferate, the resulting cells are referred to as myogenic progeny. 1.2. Functional Satellite Cells Are Required Throughout Life

In the juvenile growth phase, when muscles enlarge, satellite cells are proliferative and add nuclei to growing myofibers (21, 31–34). In most adult muscles, satellite cells are typically quiescent until their activation is invoked by muscle injury (1, 35–37). Subtle injuries may lead to minimal proliferation of activated satellite

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cells whereas major trauma can recruit greater numbers of satellite cells and promote prolonged proliferation prior to differentiation. As small myofiber injuries can occur routinely during daily activity, a mechanism for repair is essential for muscle maintenance throughout life. Activation of myogenic precursors is controlled by proximal signals from the muscle niche, microvasculature and from inflammatory cells (38–41). Systemic factors may also regulate satellite cell activation (42–44). Following their activation, satellite cells may contribute to repair of damaged myofibers and also generate new myofibers following cell division and fusion of myoblast progeny. Satellite cell behavior is under stringent regulatory control to balance various actively maintained states, including quiescence, entry into proliferation and continuity of the cell cycle, and terminal differentiation (45, 46). Furthermore, apart from their ability to fortify myofibers and contribute to muscle regeneration, satellite cells have the capacity to replenish a reserve pool and self-renew, qualifying them as tissue-specific stem cells (11, 47). It is not known, however, to what extent individual satellite cells differ with regard to their amplification and renewal potential (19, 47). During early growth, muscle satellite cells may represent about 30% of the nuclei, whereas in the healthy adult satellite cells represent approximately 2–7% of nuclei within skeletal muscle (1, 21). The number of satellite cells per myofiber or per cross-sectional area may vary immensely between muscles. For example, the fast twitch extensor digitorum longus (EDL) contains fewer satellite cells compared with the slow twitch soleus (1, 12, 48). Additionally, myofiber ends may have a higher concentration of satellite cells than the rest of the myofiber (22). There are also reports of an age-associated decline in satellite cell number, where the presence and extent of decline may vary by muscle (12, 19, 20, 49). Satellite cell performance may also decline in the aging environment, a possible contributory factor to age-associated muscle deterioration, also known as sarcopenia (20, 50). However, additional studies suggest that initial performance of skeletal muscle progenitors is delayed, but not necessarily impaired in old age and that factors beyond satellite cell activity alone may play a role in reducing muscle repair in old age (44, 51). Indeed, satellite cell activity can be rejuvenated upon exposure of old muscle to a juvenile environment by cross-transplantation or by parabiosis of young and old mice (42, 52). Muscle wasting associated with muscular dystrophy is also thought to lead to exhaustion of satellite cells due to the continuous demand for reparative myogenic cells (53–55). Overall, satellite cells are vital to skeletal muscle homeostasis and regeneration throughout life, and understanding the regulation of myogenic stem cells will likely provide valuable insights into muscle wasting in aging and disease.

2

1.3. Detection of Satellite Cell Progeny by Temporal Expression Patterns of Myogenic-Related Transcription Factors

Myogenic Stem Cells

25

At the molecular level, myogenesis of satellite cells is highly orchestrated to ensure that specific genes are regulated in a temporally organized manner according to genetic blueprints, cell cycle requirements, and environmental factors. The resulting pattern of gene expression yields terminally differentiated myoblasts, capable of adding myonuclei to existing myofibers in addition to fusing together to form new myofibers during muscle growth and repair (3, 45, 56, 57). To monitor various stages of satellite cell myogenesis in culture, we focus primarily on the expression patterns of Pax7 and the myogenic regulatory factors MyoD, myogenin, and Myf5. As demonstrated in our published studies, the temporal expression patterns of these genes do not vary for mouse, rat, or chicken satellite cell progeny. For additional background information about the functional roles of Pax7 and the myogenic regulatory factors in myogenesis, the reader should refer to additional publications (e.g., (58–60); for a comprehensive review see (3)). Satellite cell progeny can be distinguished from their quiescent progenitors based on distinctive gene expression patterns (2, 3, 57). In particular, expressions of MyoD and myogenin have been used extensively in conjunction with Pax7 (8, 10, 12, 46) (see Fig. 2). Proliferating progeny (myoblasts) continue to express Pax7, but distinctly from their quiescent progenitors, also express MyoD. A decline in Pax7 along with the induction of the muscle-specific transcription factor myogenin marks myoblasts that have entered the differentiation phase and initiated cell cycle withdrawal. Coinciding with or soon after the upregulation of myogenin, differentiating myoblasts initiate expression of various genes encoding structural proteins, such as sarcomeric myosin, and fuse into myotubes (12, 21, 39, 61). During myoblast differentiation,

Fig. 2. The molecular signature of satellite cell progeny in a primary cell culture: proliferation, differentiation, and self-renewal. A color version of this figure appeared in YablonkaReuveni and Day (2011).

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a subpopulation of mononucleated cells downregulate MyoD expression and exit the cell cycle, but maintain Pax7 expression. These cells define a reserve population that presumably reflects satellite cell self-renewal (10–12, 19, 46, 47, 57). Both quiescent and proliferating satellite cells also express the myogenic regulatory factor Myf5 as determined by mRNA analysis (13, 19, 62). Myf5 promoter activity can also be observed through b-gal detection in satellite cells and their proliferating progeny in myogenic cultures from the aforementioned Myf5nlacZ/+ mice (19, 26). However, detection of the Myf5 protein has not been reported in quiescent satellite cells, though proliferating progeny do express Myf5 protein (46, 63). Thus, it is possible that while the Myf5 promoter is active in quiescent satellite cells, Myf5 protein is not produced until cells begin to proliferate. Ultimately, Myf5 expression declines when myoblasts enter differentiation, while MyoD expression persists well into the differentiation stage when satellite cells are maintained in our standard culture conditions (3, 12, 46). 1.4. Classic and Contemporary Approaches for Satellite Cell Isolation

Much of our understanding of satellite cell biology has arisen from cell culture studies. The information provided in this section focuses on primary cultures of bona fide satellite cells. Studies with myogenic cell lines (including rat L6 and L8, and mouse C2, C2C12 and MM14) have also permitted extensive biochemical and molecular analyses of aspects of myogenesis, though these models do not always fully adhere to the biology of satellite cells (64–68). A comprehensive description of myogenic cells lines from the American Tissue Culture Collection (ATCC) and other sources can be found in our recent review (3). Two main cell culture approaches have been employed by us and other investigators in the study of bona fide satellite cells: 1. Cultures of isolated myofibers where the satellite cells remain in their native position underneath the myofiber basal lamina (8, 12, 69). This approach allows the study of satellite cells and their progeny in their in situ position and after they migrate out from the parent myofiber. We have described protocols for single myofiber isolation and culture as a means to study satellite cells at great details in other book chapters in this Methods in Molecular Biology Series (70, 71). 2. Primary myogenic cultures prepared from mononucleated cells dissociated from whole muscle. Protocols for obtaining primary myogenic cultures involve releasing satellite cells from their niche. Steps of mincing, enzymatic digestion, and repetitive triturations of the muscle are required for breaking both the connective tissue network and the myofibers to release the satellite cells from the muscle bulk. Depending on the enzymatic procedure and the purpose for cell isolation, enrichment

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for satellite cells beyond the basic isolation protocol is often unnecessary. Indeed, the basic isolation protocol that is detailed next in this chapter has been used by us in many cell culture studies of satellite cells (12, 21, 46, 72). Alternatively, satellite cells can be enriched from whole muscle cell suspensions by various approaches that reduce the presence of fibroblastic cells, typically present to some degree in the preparation, and remove myofibril debris present in the initial cell suspension. The following three different approaches have been described for myogenic preparation cleanup: (a) initial plating on uncoated tissue cultures dishes that results in separation of cells based on adhesion characteristics, where cells that remain in suspension after a short period are collected for culturing (i.e., differential plating) (73–75); (b) fractionation on Percoll density gradients (62, 76–78); (c) cell sorting by forward and side scatter (79, 80). In studies where further enrichment of satellite cells is warranted, cells can be isolated by FACS using antibodies that react with satellite cell surface antigens (47). First, cells are released from the muscle tissue using collagenase or collagenase-dispase, enzyme preparations that preserve cell surface antigens compared to Pronase or trypsin digestion methods. Studies from various laboratories (performed mainly with mouse tissue) have established that satellite cells can be isolated based on negative selection for CD45, CD31, and Sca1, and positive selection for CD34 and a7 integrin (25, 47, 81). Additional cell surface antigens, including CXCR4, b1 integrin, and syndecan-4 have also been used for isolation from adult muscle (82–84). A range of fluorescence-based reporter systems in genetically manipulated mouse strains have also permitted reliable isolation of purified populations of satellite cells. For example, we have isolated satellite cells from different muscle groups of transgenic NES-GFP mice (13, 19), and Pax3-/Pax7-driven GFP reporter expression has also been used for isolation by FACS (25, 85), with the limitation that the Pax3 reporter is only expressed in satellite cells from selective muscles (25). Mice with a GFP reporter gene inserted into the Myf5 locus also permit isolation of myogenic cells by FACS (86–88); however GFP expression is below detection level in many of the satellite cells, which reduces the usefulness of these Myf5GFP mice for satellite cell isolation by FACS. Additionally, Cre-Lox mouse models are useful for isolating satellite cells and identifying their progeny. Fluorescent reporters can be permanently turned on in cells derived from myogenic progenitors upon expression of Cre-recombinase driven by promoters of myogenic genes such as Pax3, Myf5, and MyoD (89–92). When using such Cre-Lox mouse models to sort satellite cells, one should be careful to ensure that the reporter is not expressed in additional cell types during embryogenesis. For example, Myf5-Cre expression has been reported in nonmyogenic regions

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(93–95). It is also important to note that some head muscles (e.g., extraocular muscles) develop via Pax3-independent pathways and satellite cells in these muscles do not express the Pax3-Credriven reporter ((17, 96) and our unpublished studies).

2. About Our Basic Protocols for Satellite Cell Isolation and Analysis

2.1. Strain and Age of Animals

In this chapter, we describe the basic methodologies regularly used in our laboratory for the isolation, culture, and characterization of myogenic progenitors from adult mouse skeletal muscle. As detailed in the previous section, we also use contemporary approaches for satellite cell isolation that are based on fluorescence reporter expression and/or based on expression of cell surface antigens. However, such approaches require the availability of special resources and reagents. Here, we describe a basic and straightforward method that we frequently use to isolate and characterize satellite cell performance in culture. This procedure can be performed in any tissue culture facility, using wildtype and mutant mouse muscles of various ages (12), and is suitable for satellite cell isolation from rat (62) and chicken (46) muscles. Figure 3 shows representative micrographs of myogenic cultures emanating from satellite cells isolated using our basic procedure from adult mouse hindlimb muscles. Our standard protocol for immuncytochemical analysis of satellite cell cultures provides quantitative insight into the “myogenicity” of the cell preparation (i.e., the presence and frequency of myogenic cells) and progression of satellite cell progeny from proliferation to differentiation and production of reserve cells. Table 1 summarizes the source and characteristics of a set of monoclonal antibodies used in our laboratory for the analysis of myogenesis in primary cultures of mouse satellite cells, which are also applicable to rat satellite cells (8, 12, 13, 19, 20, 62). For analysis of chicken satellite cells, we rely on the same Pax7 and MF20 antibodies as in Table 1, but for the detection of myogenic regulatory factors we use rabbit polyclonal antibodies developed against the chicken proteins (21, 46, 97). In the following subheadings, we discuss some important considerations that should be taken in mind when establishing satellite cell primary cultures. The protocols in this chapter focus on the isolation and culture of myogenic progenitors from adult (3–6 month-old) C57BL/6 mice. Aged mice and other mouse strains have also been used in our studies following the same procedures (12, 19). However, muscles from younger mice may contribute more cells due to age-associated decline in satellite cells in some muscles (12, 19).

Fig. 3. Phase micrographs depicting the morphology of mouse myogenic cultures seeded on gelatin-coated (a–c) and Matrigelcoated (d) dishes. Cells were isolated by Pronase digestion and cultures were maintained in rich growth medium according to protocols detailed in this chapter. (a–d) Show the cultures on days 3, 5, 7, and 7, respectively. Round cells observed during early culture days (a, b) are proliferating myoblasts. Multinucleated myotubes can already be observed on day 5 (b) and enlarge on subsequent days (c, d). Residual debris resulting from tissue dissociation, which is present in early culture days and can be mistakenly considered a contamination (see step 30, Subheading 4.1), is noticeable at same focal level as the proliferating cells (a). The identity of myoblasts and myotubes can be further confirmed by their characteristic protein expression (see Fig. 2) using immunostaining with antibodies detailed in Table 1. Images were taken with a 20× objective.

Table 1 Mouse monoclonal antibodies frequently used in our studies for analyzing progeny of mouse satellite cells as they transit through proliferation, differentiation, and renewal

a

Antibodya

Clone

Isotypeb

Sourcec

References

Anti-Pax7

Pax7

IgG1

DSHBd

(12, 13, 98)

Anti-MyoD

5.8A

IgG1

BD Biosciences

(12, 63, 99)

Anti-myogenin

F5D

IgG1

DSHB

(12, 63, 100, 101)

Anti-sarcomeric myosin

MF20

IgG2b

DSHB

(12, 102)

The antibodies against Pax7 and sarcomeric myosin were prepared originally against chicken proteins (98, 102). The antibody against sarcomeric myosin recognizes an epitope shared by all isoforms of sarcomeric myosin heavy chain in skeletal and cardiac muscle in a wide range of species b The isotype of each antibody is provided to help in designing double-immunostaining studies. We routinely perform such studies using the anti-sarcomeric myosin in combination with the antibodies against MyoD, myogenin, and Pax7 (12). Isotype-specific secondary antibodies are available from a variety of commercial sources. We obtain such antibodies (Alexa Fluor conjugated) from Invitrogen c The same monoclonal antibodies against Pax7, MyoD, and myogenin are available from additional sources d The Developmental Studies Hybridoma Bank (DSHB) is under the auspices of the National Institute of Child Health and Human Development and maintained by The University of Iowa, Department of Biology, Iowa City, IA. http:// dshb.biology.uiowa.edu

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Also, the contribution of nonmyogenic cells in the preparation may increase with age or in different mouse strains, and consequently, some conditions may need modification (i.e., duration of enzymatic digestion, extent of tissue trituration, cell straining conditions to remove debris, centrifugation speed of harvested suspension, etc.) to minimize the proportion of undesired cell types. 2.2. Muscles

Herein we detail our standard procedure for the isolation of satellite cells from hindlimb muscles of adult mice. For this preparation, we typically pool the fast twitch muscles tibialis anterior (TA) and gastrocnemius from both hindlimbs, using one mouse per preparation. For additional details about TA and gastrocnemius anatomy and isolation procedures see Notes 1 and 2. This approach can also be used for isolating myogenic progenitors from limb, body, and head muscles. However, the contribution of connective tissue and vasculature may vary between muscles, and the tissue isolation procedure should be modified accordingly to minimize cells derived from such structures. The purity of the resultant preparation of isolated satellite cells (and cultures emanating from this preparation) is directly dependent on the amount of effort spent meticulously cleaning the muscle of these additional structures.

2.3. Digestive Enzyme for Muscle Dissociation

Our procedure is based on cell dissociation from whole muscle using Pronase digestion (see item 7, Subheading 3.4 and steps 4 and 13, Subheading 4.1). Pronase (available from Calbiochem) consists of a mixture of proteases isolated from the extracellular fluid of Streptomyces griseus. Because of its particular protease content, which includes several types of endo and exopeptidases, Pronase has a broad activity (103, 104). Pronase digestion may not be optimal for prospective satellite cell enrichment by antigen-based cell sorting because of extended digestion of surface antigens. However, myogenic cell preparations isolated by Pronase digestions show a lower level of nonmyogenic cells compared to that observed when collagenase or collagenase/ dispase enzyme solutions are used. It is possible that certain nonmyogenic populations do not survive well after Pronase digestion and this may lead to the increased purity of these cultures.

2.4. Cell Yield, Choice of Culture Dish and Cell Seeding Density

Cell yields can vary depending on the age of the animal. Muscles from neonatal and young mice (1-month old or less) yield considerably more myogenic progenitors than muscles from adult mice. As mentioned earlier, variations are also observed when working with different muscles. For the mouse strain (C57BL/6) and hindlimb muscles (TA and gastrocnemius) used for the protocol described herein, each preparation typically yields 2–5 × 105 cells. We commonly use 24-well or 35-mm culture dishes. We generally use 35-mm dishes for training or when performing single

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comparisons. In such cases, we initiate the cultures at 5–10 × 104 cells per plate. For multiple replicates across multiple time points, we use 24-well plates where starting cell density can be proportionally matched with that of the 35-mm plates based on surface area. Alternatively, seeding densities can be further reduced, and depending on experimental goals may range from 5 × 104 to 1 × 103 for primary cultures. Although not further detailed below in the protocol section, it is noteworthy that in some of our studies we also use 48-well trays where we seed 2–10 cells per well; in such studies we aim to achieve clonal growth for monitoring progeny of individual satellite cells. 2.5. Culture Medium

The standard growth medium used for our mouse satellite cell cultures consists of high glucose Dulbecco’s Modified Eagle Medium (DMEM) supplemented with 20% fetal bovine serum, 10% horse serum, and 1% chicken embryo extract (CEE). This serumrich growth medium supports both proliferation and differentiation of myogenic cells (12). See Subheading 3.4 and Notes 3–6 for details about recommended cell culture reagents, our protocol for preselection of optimal sera lots and preparation of CEE, and final medium preparation. Some variations can be found from laboratory to laboratory with regard to the basic culture media (e.g., Ham’s F10 instead of DMEM, or a mixture of the two), serum type and concentration, and source of growth factors (e.g., purified growth factors, especially fibroblast growth factor, instead of CEE). Differences in culture conditions may explain some divergences in satellite cell behavior among different laboratories. For example, some published protocols rely on first using serum-rich growth medium that supports proliferation followed by a switch to serum-poor medium to support differentiation. There are also reported variations in medium composition when preparing cultures from other species. For example, for primary cultures of chicken satellite cells we typically use medium containing 10% horse serum and 5% chicken embryo extract (21, 46, 76, 97). To study the effects of specific growth factors on myogenic cell performance, we typically maintain the cells for 3 days in our standard rich growth medium to allow for optimal cell adherence, then switch the cells into serum-low (e.g., DMEM containing 2% horse serum) or serum-deprived media. Prior to switching to serum-low medium, the cultures are rinsed extensively with DMEM to remove traces of the rich medium that otherwise adhere to the cell layer and reduce the observed effect of the additives being examined.

2.6. Plate Coating Matrices

Adhesion of myogenic progenitors to cell culture dishes can be significantly improved by coating the plastic substrate with a variety of extracellular matrix constituents or derivatives. In addition to cell adhesion, matrix components can influence the extent

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of myogenic cell proliferation, differentiation, and renewal (12, 13, 105, 106). In our laboratory, the main matrices used for coating tissue culture plates for satellite cell cultures are Matrigel and gelatin. Matrigel is a solubilized basement membrane preparation extracted from the Engelbreth-Holm-Swarm mouse sarcoma, a tumor rich in extracellular matrix proteins. Its major component is laminin, followed by collagen IV, entactin, and heparan sulfate proteoglycan (107). Matrigel is available from BD Biosciences and can be obtained in its standard format or in its growth factor reduced format. In our studies we have typically used the growth factor reduced format. Matrigel must be carefully handled on ice when aliquoting and coating tissue culture dishes with dilutions. For additional details on Matrigel source and handling, see item 8, Subheading 3.4 and Notes 7 and 8. Gelatin is produced by partial hydrolysis of type I collagen extracted from connective tissues. It can be purchased in a tissue culture grade powder form and easily reconstituted in water to the desired concentration. For specific details about gelatin source and our preparation of gelatin solution, see item 8, Subheading 3.4 and Notes 9 and 10. Gelatin is readily available, inexpensive, and easy to use, which makes it an ideal product for training new team members and for use in standard cultures. However, long-term high-density myogenic cultures may spontaneously detach from plates coated with gelatin. In addition, satellite cell progeny typically demonstrate a more limited proliferative period, earlier differentiation, smaller myotubes and more meager development of reserve cells when grown on this substrate compared to Matrigel-coated dishes. Matrigel also allows a more even cell distribution upon initial cell plating compared to that observed when cells are seeded on gelatin-coated dishes. Additionally, when plated on Matrigel-coated dishes, myogenic progenitors can reach high cell densities and form complex myotube networks, typically without detaching from the substrate. The latter features have prompted us to use Matrigel especially when seeding cells at low density or when aiming to obtain single cell clones. Disadvantages of Matrigel include higher cost and the requirement for more careful handling. Other commercially available matrices that we have tested in pilot experiments that may provide reasonable alternatives include (a) GelTrex (a Matrigel-like product from Invitrogen); and (b) Attachment Factor (Invitrogen), a ready made gelatin-based product. 2.7. Fixation and Immunostaining

For immunostaining analyses using the antibodies listed in Table 1, we typically fix the cultures with a paraformaldehydesucrose solution that is prepared in our laboratory. For further details about fixation approach and fixative composition, see

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Subheading 4.2, and Notes 11 and 12. It should be noted that fixatives should be optimized for preservation of both the cells and the antigens being analyzed. We perform all immunostaining steps in a manner that maintains sterility; handling antibodies strictly in the tissue culture hood minimizes possible bacterial contamination and helps maintain antibody stocks for years.

3. Materials 3.1. General Comments

1. The quantities of glassware, media, and reagents as well as the time intervals for enzymatic digestion described in this chapter are appropriate for the isolation of satellite cells from TA and gastrocnemius muscles of both hindlimbs of one adult (3–6 month-old) C57BL/6 mouse. We typically do not pool muscles from multiple mice into a single preparation as cell yields are not necessarily increased linearly when using more muscle bulk. 2. All procedures are performed using sterile materials, supplies, and techniques. Before transferring solutions/media into the tissue culture hood, spray the glass/plastic containers with 70% ethanol and wipe dry.

3.2. General Equipment

The following facilities are required for the cultures described in this chapter: 1. Standard humidified tissue culture incubator (37°C, 5% CO2 in air). 2. Tissue-culture laminar flow hood. 3. Water bath (37°C). 4. Hair trimmer (optional, for shaving hair from the hindlimbs prior to muscle dissection). 5. Stereo dissecting microscope with transmitted light base (microscope is either placed inside a tissue culture hood or in an isolation box/clean area). 6. Surgical tools for harvesting the muscles. Two types of forceps with extra fine-tips are recommended in particular to clean the muscles: (a) straight 110-mm (41/4″), and (b) curved 115-mm (41/2″). We typically sterilize dissection tools with a glass bead sterilizer, which is useful for quick sterilizing of tools as needed. 7. Table-top centrifuge. 8. Inverted phase contrast microscope for monitoring cell culture. 9. Inverted fluorescence microscope for analysis of immunolabeled culture dishes.

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10. Hemacytometer and cover glass. Cover glasses can be purchased separately if replacement is needed. 11. Pipette controller (motorized pipette filler), essential for triturating the tissue after enzymatic digestion. 3.3. Plastic and Glassware Supplies

1. Standard (9″) Pasteur pipettes. 2. Wide-bore pipettes prepared from the standard (9″) Pasteur pipettes. Using a file or a diamond knife cut the narrow end of these pipettes to prepare a set of them with a bore diameter of approximately 3 mm. Shake the pipette to remove any glass fragments, fire polish sharp ends, and autoclave. These pipettes are used to transfer muscle fragments. 3. Pasteur pipettes (9″) with cotton plug. 4. Serological glass pipettes (1-mL). 5. Serological glass pipettes (10-mL). 6. Syringe filters, 0.22-mm PVDF low protein binding filters and 1- or 3-cc disposable plastic syringes. Bottle top filters, 0.22 mm. 7. Cell strainer, 40-mm nylon mesh. 8. Polypropylene conical centrifuge tubes, sterile, 15 and 50 mL. 9. Plastic Petri dishes, 100-mm. 10. Tissue culture dishes, 35-mm. 11. Twenty four-well multiwell tissue culture dishes.

3.4. Cell Isolation and Culture Reagents

1. DMEM (Dulbecco’s Modified Eagle Medium; high glucose, with 4,500 mg/L glucose, 4 mM L-glutamine, 110 mg/L sodium pyruvate), supplemented with 100 U/mL penicillin and 100 mg/mL streptomycin. The term DMEM used from here on in this chapter refers to DMEM with antibiotics. 2. Fetal bovine serum (FBS; standard, not heat inactivated; Invitrogen/Gibco; see Note 3). Original bottles are stored at −80°C for long term (years); once thawed and aliquoted, stored at −20°C. 3. Chicken embryo extract (CEE) (available commercially from several sources); or, as in our studies, prepared by the investigator (see Notes 4 and 5); stored at −80°C for long term (years) or at −20°C when aliquoted. 4. Horse serum (HS; standard, not heat inactivated; Hyclone; see Note 6). Original bottles are stored at −80°C for long term (years); once thawed and aliquoted, stored at −20°C. 5. Standard growth medium for satellite cell cultures is made up of DMEM, 20% fetal bovine serum, 10% HS and 1% CEE. Culture medium is stored at 4°C and used within 3 weeks from preparation. 6. DMEM containing 10% HS to resuspend cells after enzymatic digestion.

2

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7. Pronase (Calbiochem, reconstituted in DMEM) used for muscle digestion as described in Subheading 4.1, step 4. 8. Matrigel (BD Biosciences) for coating tissue culture dishes (see Note 8 for instructions). Matrigel can be purchased in its standard format. We usually dispense Matrigel into aliquots of 0.1–0.2 mL and freeze them at −20°C. See Note 7 for handling details. 9. Gelatin (Type A, Sigma-Aldrich) can be used as an alternative for coating tissue culture dishes (see Note 10 for procedure). Prepare and store 5 mL aliquots of 2% gelatin solution as indicated in Note 9. 3.5. Reagents and Solutions for Fixing and Immunostaining

Unless otherwise stated, the following solutions are stored at 4°C and prewarmed at room temperature before use. 1. Prefixation rinse solution: DMEM as in item 1, Subheading 3.4. 2. Fixative: 4% paraformaldehyde in a sodium phosphate buffer containing 0.03 M sucrose (for further hazardous material details and composition/preparation of the fixative solution see Notes 11 and 12). To maintain quality and effectiveness of fixative, only prewarm the amount that is required for immediate use. 3. Postfix rinse solution: Tris-buffered saline (TBS); 0.05 M Tris, 0.15 M NaCl, pH 7.4 (for preparing this solution, see Note 13). 4. Detergents: Triton X–100; Tween 20. 5. Detergent solutions: TBS containing 0.5% Triton X-100 (TBSTRX100); TBS containing 0.05% Tween 20 (TBS-TW20). 6. Blocking reagent: Normal goat serum (standard serum, does not need to be a product that is sold specifically for immunostaining). Can be stored at −80°C for long term (years); once thawed and aliquoted, store at −20°C. 7. Blocking Solution: TBS containing 2% normal goat serum (TBS-NGS). 8. Mounting media: Vectashield (Vector Laboratories) and (1) sterile 25% glycerol solution in TBS for 24-well plates; or (2) cover glass, 22 mm2, for 35-mm dishes.

4. Methods 4.1. Cell Isolation and Culture

1. Prewarm 30 mL of DMEM to 37°C and then keep at room temperature throughout the procedure. 2. Coat the tissue culture dishes with gelatin or Matrigel following the instructions described in Notes 8 and 10, respectively.

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3. Add 5 mL of DMEM to three 60-mm Petri dishes and place the dishes in the tissue culture incubator until muscle dissection begins. 4. Prepare 1 mL of 1% Pronase solution; we prepare this solution fresh for each experiment, dissolving 0.01-g Pronase in 1-mL DMEM. Use a 0.22-mm syringe filter attached to a 1- or 3-cc syringe to filter the Pronase solution into a 15-mL conical centrifuge tube. At the time of muscle digestion (see steps 13 and 14, Subheading 4.1), this solution will be diluted tenfold in the DMEM containing the muscle fragments to make 3 mL of a final digest in 0.1% solution. 5. Euthanize one mouse according to institutional regulations, shave (optional) the hindlimbs and spray them (regardless if shaving or not) lightly with 70% ethanol. 6. Harvest the TA and gastrocnemius muscles from both hindlimbs and place them in a 60-mm Petri dish with DMEM (see Notes 1 and 2 for further description of these muscles and how to isolate them). 7. Rinse the muscles by gently swirling the plate and transfer them to the second 60-mm Petri dish. 8. Under the dissecting microscope, using the straight and curved fine point forceps (described in Subheading 3.2, item 6), carefully remove from each muscle the tendons, fat, vessels, and bits of connective tissue as much as possible. 9. Transfer the cleaned muscles to the third 60-mm Petri dish with DMEM and cut into small fragments (about 3 mm3) but do not mince (if fragments are too small, the mechanical trituration that follows the enzymatic digestion step is less effective in releasing cells). Further inspect the muscle fragments to eliminate, as much as possible, any remaining connective tissue. 10. Using a sterilized wide-bore Pasteur pipette, transfer the suspension of muscle fragments to a 15-mL conical tube and allow the fragments to settle down. Alternatively, muscle fragments can be collected by low speed centrifugation (~200 × g) for 4 min. 11. Aspirate and discard the supernatant. Add DMEM to the settled muscle fragments up to a final volume of 2 mL, including the muscle bulk. Shake the tube gently to loosen the pelleted tissue and transfer tube contents to a 35-mm dish using a widebore Pasteur pipette. 12. Use 700 mL of DMEM to rinse the 15-mL tube of any remaining muscle bits and add this volume to the 35-mm dish. 13. Add 300 mL of 1% Pronase to the plate, generating a final volume of 3 mL (including muscle bits) and a final concentration of 0.1% Pronase.

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14. Place the 35-mm dish inside the tissue culture incubator for 60 min. Gently swirl the dish every 15–20 min during digestion (alternatively, one can use a low speed agitator placed inside the tissue culture incubator). 15. At the end of the digestion period, transfer the muscle fragments and Pronase solution to a 15-mL conical tube using a wide-bore Pasteur pipette. 16. Spin down the suspension by low-speed centrifugation at 400 × g for 5 min. 17. Aspirate the supernatant carefully, without disturbing the loose pellet of digested muscle pieces. Resuspend the muscle bulk in 5 mL of 10% HS in DMEM (prewarmed at 37°C, then kept at room temperature until used). At this stage the still attached satellite cells can be released by mechanical trituration in a manner that avoids damaging the desired cells. We perform two cycles of muscle trituration (detailed below, see steps 18–22) so that cells released early in the process can be harvested and set aside, after which further trituration releases the remaining cells. It is critical that the enzymatic digestion does not only fully dissociate the tissue, but only loosens the cells; without the mechanical trituration steps, cell yields are poor. 18. First muscle trituration: Vigorously triturate muscle fragments by passing them repetitively (about 20 times) through a 10-mL glass pipette until the tissue bits pass easily through the tip of the pipette. Shearing of the tissue with the mechanical trituration is critical to efficient cell release. Allow the suspension to settle in the 15-mL conical tube so that the remaining larger bits separate from the supernatant that contains the released cells. 19. Without disturbing the precipitated material, collect the supernatant and transfer it to a 15-mL conical tube. 20. Second muscle trituration: Add 5 mL of 10% HS into the 15-mL conical tube containing the remaining muscle pieces and repeat the muscle trituration process, now using a 9″ cottonplugged glass Pasteur pipette until all the muscle pieces easily passes through it. 21. Allow the suspension to settle as in step 19 and collect the supernatant in the same 15-mL conical tube as in step 20. 22. Place a 40-mm cell strainer onto a 50-mL conical tube. 23. Using a 10-mL glass pipette, transfer the pooled supernatants from the two triturations to the 40-mm cell strainer. Make sure the suspension passes through the strainer by carefully tapping the side. This step eliminates residual large debris from the cell suspension. 24. For maximal cell recovery, allow an additional 1-mL DMEM to drip through the cell strainer to recover the residual cells trapped by debris in the unit.

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25. Centrifuge the strained cell suspension at ~1,000 × g for 10 min (see Note 14) to recover the cells released during the trituration steps. 26. Carefully aspirate supernatant (which is discarded) and resuspend the cell pellet in 1 mL of standard growth medium (prewarmed at 37°C and held at room temperature until needed) using a 1-mL glass pipette. We recommend removal of the supernatant manually with a Pasteur pipette, and not by mechanical aspiration to minimize the risk of aspirating the delicate cell pellet as well. 27. Using a micropipette, collect 10 mL of the cell suspension (ensure the suspension is mixed gently just before removing the aliquot for cell counting as cells settle very fast when held in the tube for processing) and transfer it to the edge of one of the hemacytometer chambers, previously cleaned with 70% ethanol, dried and covered with the cover glass (the cell suspension should run under the cover glass by capillarity). Count only the small round cells while avoiding red blood cells. For increased accuracy, we recommend counting another 10-mL sample of the cell suspension in the second hemacytometer’s chamber. 28. Plate cells in the Matrigel- or gelatin-coated culture dishes. When using 24-well trays, plate cells at a density of 1–2 × 104/ well (standard density) or 1–2 × 103/well (low density). When using 35-mm dishes, plate 1–2 × 105 cells (standard density) or less, depending on the experimental goal. 29. Culture the cells undisturbed in the incubator for 3 days. 30. Rinse the cultures 1–2 times with 1 mL of prewarmed DMEM before adding fresh medium at the first medium change. This helps to remove debris that is apparent in the primary cultures and can be easily mistaken for contamination to an inexperienced observer. Cultures should be rinsed very gently to minimize cell detachment. If warranted, the level of debris in the cell suspension can be further reduced before culturing the cells (see Note 15), but the debris also disappears with time as cultures get more dense. 31. Replace the culture medium with fresh medium every 3 days. Note, however, that medium may need to be changed more frequently at late time points depending on the density of the cells. 4.2. Cell Culture Fixation and Immunostaining

1. Warm DMEM and fixative solution to room temperature. DMEM can be first warmed in a water bath set at 37°C then held at room temperature until needed. The 4% paraformaldehyde fixative solution should be allowed time to equilibrate to room temperature prior to its use. For both items, warm only

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the required volume for the experiment. Extensive warming of the fixative solution multiple times results in deterioration of the paraformaldehyde. 2. Rinse cultures with DMEM three times. Following the final rinse add 250 mL of DMEM to each well in the 24-well plate or 500 mL to each 35-mm dish. 3. Add an equal volume of the 4% paraformaldehyde fixative solution to the culture medium in each well or dish (250 or 500 mL as above). Allow 10 min at room temperature for the fixation, then carefully remove the culture medium-paraformaldehyde fixative mixture and rinse each well three times with TBS. 4. Add 500 mL of TBS-TRX100 for 5 min at room temperature to permeabilize the cells. Alternatively, the permeabilization step can be omitted if considering using antibodies different from those listed in Table 1, as some antigens might be sensitive to this detergent. Also, cultures can be treated with TBSTRX100 later (but then blocking solution detailed below needs to be reapplied prior to antibody staining). Note that from this step on, unless otherwise stated, the volumes of each reagent are the same for either 24-well or 35-mm dishes. 5. Add 500 mL of blocking solution (TBS-NGS) to each well or dish to block nonspecific antibody binding. 6. Cultures are then kept at 4°C overnight or longer (see Note 16). 7. Allow plates to warm up to room temperature for at least 10 min before starting the antibody staining procedure. 8. Dilute the appropriate primary antibody in blocking solution. For antibodies listed in Table 1, we typically use antibody formulations as previously published (12, 19). 9. Rinse the cultures three times with TBS-TW20. 10. Aspirate the final TBS-TW20 rinse and add 150 mL of the primary antibody solution for 1 h at room temperature, followed by an overnight incubation at 4°C in a humidified chamber. Primary (and secondary – see step 11) antibodies are applied at the center of the dish. When using 24-well plates, a light and continuous swirling on a flat surface is required to ensure optimal spreading of the antibody across the well; otherwise, antibody solution rapidly accumulates at the periphery (see Note 17). When working with 35-mm dishes, plates are manually swirled only upon applying the antibody then maintained without any disturbance during the labeling period, allowing the antibody solution to spread throughout the plate by capillarity. 11. Bring the plate to room temperature as in step 7 and dilute the appropriate secondary antibody in the blocking solution.

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For antibodies listed in Table 1, we typically use secondary antibodies diluted as previously published (12, 19). 12. Rinse cultures three times with TBS-TW20 warmed up at room temperature. 13. Aspirate the final TBS-TW20 rinse and add the diluted secondary antibody (same volume and swirling conditions as for the primary antibody, see step 10) for 1–2 h at room temperature. 14. Aspirate the secondary antibody and wash three times with TBS-TW20. 15. For nuclear visualization, add 100 mL of DAPI solution (4¢,6-diamidino-2-phenylindole, dihydrochloride; stock concentration 10 mg/mL, working concentration 1 mg/mL diluted in TBS-NGS prior to use) for 30 min at room temperature (see Note 18). 16. Rinse the cultures twice with TBS-TW20 followed by a final rinse with TBS. 17. Aspirate the TBS and mount cultures in Vectashield mounting medium. The mounting medium prevents the stained cultures from drying and retards fading of the immunofluorescent signal. Add 1 drop at the center of each well of the tray or each 35-mm culture dish. If working with a 35-mm culture dish, complete the mounting process by covering with a cover slip. We prefer not to use cover slips when working with 24-well trays. Instead, we add 300 mL of glycerol mounting solution (25% glycerol in TBS) following the initial drop of Vectashield to allow mounting medium coverage of individual wells in 24-multiwell trays.

5. Notes 1. The information provided here is to assist in the identification and isolation of the tibialis anterior and gastrocnemius muscles. We recommend the following literature and links for anatomical descriptions and schematic images of mouse muscles, though they refer to rat and human muscles. (a) Tibialis anterior (TA): The TA is a superficial muscle of the anterior compartment of the lower hindlimb, located in a medial position (108, 109). It arises from the lateral condyle and the upper lateral surface of the tibia. Its tendon passes across the medial surface of the dorsum of the foot and inserts on the medial cuneiform bone and the first metatarsal. The TA muscle is responsible for the dorsiflexion and inversion

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of the foot. For a schematic image of this muscle see: http://www.bartleby.com/107/illus437.html. (b) Gastrocnemius: The gastrocnemius muscle is the most superficial muscle in the posterior part of the lower hindlimb (103, 104). It consists of two heads that arise from the lateral and medial condyles of the femur. The distal end of the gastrocnemius muscle is the Achilles’ or calcaneal tendon, which is attached to the posterior surface of the calcaneus. Located deep to the gastrocnemius and closely connected to it is the soleus muscle. These two muscles are collectively called triceps surae and together, they are responsible for the plantarflexion of the foot. For a schematic image of the gastrocnemius location see: http://www.bartleby.com/107/illus438.html. For additional information about TA and gastrocnemius muscles, refer to: http://www.bartleby.com/107/129.html. 2. Harvesting hindlimb TA and gastrocnemius muscles: (a) To begin with the TA extraction, secure the mouse in a supine position to the dissecting board by pinning down the hindlimb to be dissected and the diagonal forelimb. (b) Use straight operating scissors to cut through the skin, opening a small incision above the knee. (c) Holding the skin with fine forceps, insert rounded-tip scissors beneath the incision and carefully open the scissors to loosen the skin from the underlying muscles. (d) Extend the incision to a point just in front of the digits. (e) Loosen the skin as you go, being careful not to cut the underlying muscles or blood vessels. (f) Cut and remove the skin from the knee to the paw. (g) Identify the tendon at the insertion of the TA. (h) Place one arm of the very fine point forceps underneath the tendon and carefully pull proximally, with the forceps under the TA muscle, to drag the fascia that covers the muscle. Then use the forceps to pull the fascia upward toward the knee and discard it. (i) Use micro scissors to cut the tendon at the insertion of the TA, as far as possible from the muscle itself. (j) Using very fine point forceps grasp the tendon and carefully pull it in order to lift the TA muscle gently away and upward. (k) With the TA lifted, cut the proximal attachment against the knee with micro scissors and place the removed muscle in a 60-mm Petri dish with DMEM.

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(l) Do the same (see steps b–k) for the other hindlimb before proceeding to the next steps (gastrocnemius isolation). (m) Turn the mouse over in a prone position, pin it appropriately, and identify the gastrocnemius. (n) Using dressing forceps, pull away the upper hindlimb muscles that cover the proximal portion of the gastrocnemius. (o) Place very fine point forceps under the Achilles tendon and move it proximally underneath the gastrocnemius. (p) Cut the Achilles tendon and lift the gastrocnemius as done for the TA. (q) With the gastrocnemius lifted, cut its proximal side as close as possible to its origin. Remove the soleus muscle, which is intertwined with the main tendon of the gastrocnemius, and place the gastrocnemius in the same 60-mm Petri dish containing the isolated TA muscles. Repeat steps p and q for the other hindlimb. 3. Fetal bovine serum (FBS) should be precharacterized by comparing sera from several suppliers. We select FBS based on the capacity of the serum to support proliferation and differentiation of mouse primary myoblasts cultured when seeding cultures at a wide range of cell concentrations. Only sera able to support good growth and differentiation at both high and low cell density are employed in our studies. Primary myogenic cultures for these tests are prepared as described here. The vendor listed for this product in item 2, Subheading 3.4 is provided as an example for what we found to be optimal when the serum selection was performed. 4. We prepare chicken embryo extract (CEE) in our laboratory using 10-day-old White Leghorn embryos (70, 110). The procedure is similar to a previously described method (111) but uses the entire embryo. We recommend this approach over purchasing CEE if the investigator can obtain embryonated chicken eggs, as the quality is higher and the cost lower than that of purchased CEE. 5. Preparation of chicken embryo extract: (a) Embryonated chicken eggs (8 dozen, White Leghorn; from Charles River or local sources with a good egg fertility index) are maintained in a standard egg incubator (incubation conditions: a dry temperature of 38°C, a wet temperature of 30°C, and relative humidity of 56%). (b) After 10 days, batches of 15–30 eggs are removed from the incubator and transferred into the tissue culture hood. All steps from here on are performed in a sterile manner.

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(c) Place the eggs lengthwise in the rack and spray with 70% ethanol to sterilize. Wait for several minutes until the ethanol evaporates. (d) Crack open one egg at a time into a 100-mm Petri dish. (e) Remove the embryo from surrounding membranes by piercing it with fine forceps. Rinse the embryo by transferring it through three 100-mm Petri dishes containing DMEM supplemented with antibiotics (see item 1, Subheading 3.4 for DMEM-antibiotics formulation). Swirl embryo a few times in each dish for a good rinse. (f) Empty the egg remains from the initial 100-mm dish (described in step d) into a waste beaker and repeat steps d–f until the final rinse dish contains about 30 embryos. (g) Transfer the embryos with fine forceps into a 60-mL disposable syringe, force through the opening with the syringe plunger, and collect the suspension into a 500-mL sterile glass bottle. (h) The extract is diluted with approximately an equal volume of DMEM (supplemented with antibiotics) and gently agitated for 2 h at room temperature. To ensure good agitation, keep maximum volume to one-half bottle capacity and place the bottle at 45° angle during the agitation. (i) The extract is frozen at −80°C for a minimum of 48 h. It is then thawed, dispensed into 50-mL conical tubes, and centrifuged at approximately 500 × g for 10 min to remove residual tissue. (j) The supernatant is pooled, divided into 40-mL aliquots, and kept frozen at −80oC for long-term storage. For shortterm storage, we typically prepare aliquots of 2.5 mL that are kept frozen at −20°C. (k) Prior to use, the CEE-thawed aliquot should again be centrifuged at about 800–1,000 × g for 10 min to remove aggregates. The supernatant is then collected and added to the DMEM-based medium to prepare the rich growth medium for myogenic stem cell cultures. The growth medium is then passed through a sterile 0.22-mm filter to clear remaining particles and sterilize. All details of supplies for generating the medium are in Subheading 3.4. To ensure optimal cell growth conditions, we typically prepare only 250-mL medium each time, and use it within a few weeks. 6. Horse serum (HS) should be precharacterized by comparing sera from various suppliers. We select HS based on its capacity to support proliferation and differentiation of primary chicken myoblasts cultured at standard and clonal densities (21, 46). The vendor product listed in item 4, Subheading 3.4 for HS

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source is provided as an example for what we found to be optimal when the serum selection was performed. 7. Matrigel (BD Biosciences) is shipped on dry ice and stored at −20°C until aliquoted. Matrigel should be thawed on ice; never use a warmer temperature, as it will prematurely gel. To ensure Matrigel stability, we follow the manufacturer’s handling instructions, when thawing the product on ice (overnight in an ice bucket placed at 4°C). Once liquefied, Matrigel is aliquoted with prechilled 1-mL serological glass pipettes into tubes chilled on ice. Typically, we aliquot 0.1 and 0.2 mL each into 2-mL cryogenic vials sealed with O-rings. These aliquots are stored at −20°C. 8. Coating tissue culture dishes with Matrigel: All steps are done on ice, unless otherwise noted. Matrigel stock is first diluted to create a working mixture used to coat plates (see Note 7). We here describe the coating of 24-well plates only. (a) Thaw the required amount of Matrigel by placing frozen aliquot(s) on ice for at least 30 min and as much as 1.5 h to allow the Matrigel stock to completely liquefy for subsequent dilution to the working solution. We observed some batch-to-batch variation in the time it takes to thaw the aliquots; therefore, for consistency, we typically allow Matrigel aliquots to thaw for 1.5 h. (b) Prechill a 50-mL conical tube on ice and transfer the thawed Matrigel into the tube. Add ice-cold DMEM to dilute the Matrigel to a final concentration of 1 mg/mL. Gently mix the Matrigel and DMEM by several repetitive drawings through a 1-mL glass pipette. An optimal Matrigel stock is at ~10 mg/mL protein concentration, further diluted at 1:10 for the working Matrigel solution. Stock protein concentration can vary greatly from lot to lot and should be monitored. Allow the diluted Matrigel solution to cool on ice for 15 min. (c) After 15 min, use a chilled 1 mL glass pipette to draw up the diluted Matrigel solution and coat the dishes with an appropriate volume (150 mL per well for a 24-well plate). In our experience, 2 mL of working Matrigel solution can be used to coat an entire 24-well plate; we typically coat 6–8 wells at a time as detailed next. (d) Per each series of 6–8 wells, leave the culture plate/dish coated with the Matrigel working solution on ice for 7 min, then use the same pipette as before (held cooled in a tube on ice) to remove the Matrigel solution and place it back in the 50-mL conical tube that is kept on ice. This will leave a thin coat of Matrigel at the bottom of the wells/dishes.

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(e) Once all of Matrigel solution has been placed back in the tube, use the same pipette to coat the next set of wells in the tray. Always be sure to leave the aliquot of diluted Matrigel in each well for 7 min. (f) Having coated all the desired wells per 1 tray, tilt the dishes and use a 20-mL pipette tip to carefully remove residual Matrigel and place it back in the 50-mL conical tube kept on ice. (g) Incubate the Matrigel-coated dishes in the tissue culture incubator for at least 1 h. (h) About 10 min before plating the isolated cells, take the Matrigel-coated dishes out of the incubator to the tissue culture hood and open the lid. This will allow evaporation of water that otherwise will condense on the underside of the lid when moving the dish from the warm incubator to room temperature. If allowed to form, the condensation will drip into the well, disturbing the Matrigel coating. (i) The working Matrigel solution can be used to coat additional dishes after completing one tray coating. Matrigel that has been used to coat too many dishes, however, is less effective in supporting cell adhesion. We typically limit reuse of diluted Matrigel to three rounds of coating and work with a larger volume of diluted Matrigel if coating more than 1 tray. Also, we only use Matrigel that has been diluted the day of the fiber isolation to maintain consistency. 9. Preparation of 2% gelatin solution: (a) Weigh 2 g of gelatin powder and transfer it to a 250-mL glass bottle containing 100 mL of deionized water. (b) Autoclave (only at this stage will gelatin powder completely dissolve). (c) Allow the solution to cool to room temperature. (d) Aliquot 5 mL into 15 mL conical tubes and store at 4°C. Gelatin solution will solidify upon refrigeration. Aliquots stored at 4°C are good for years. 10. Coating tissue culture dishes with gelatin: (a) Place 2% gelatin aliquot in a 37°C water bath until completely liquefied; then keep in the tissue culture hood until used. (b) Distribute 150–200 mL of gelatin solution into each well of the 24-well plate, or 300–500 mL into 35-mm culture dishes. (c) Swirl gently the 24-well plate or the 35-mm dish to allow even coating of the plating surface. Inspect plates to ensure even spreading of the gelatin solution as some regions may remain uncoated initially.

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(d) Allow the gelatin-coated dish to sit at room temperature for at least 1 h. (e) Using a Pasteur pipette remove the entire volume of gelatin solution from the wells. That will leave a thin coat of gelatin at the bottom of the wells/dishes. Gelatin solution can be reused several times (at least 10 times) without affecting cell adhesion and growth. (f) Let the gelatin-coated dishes sit in the tissue culture hood for at least 30 min before plating the isolated cells. 11. Paraformaldehyde is a white powder with a formaldehyde-like odor. It is a rapid fixative and a potential carcinogen. When handling paraformaldehyde, wear gloves, a mask, and goggles. It is important to refer to the MSDS instructions and institutional regulations for further information regarding storage, handling, and first-aid. 12. Preparation of 100 mL of 4% paraformaldehyde with 0.03 M sucrose, in a fume hood: (a) Mix 4 g of paraformaldehyde powder and 80 mL of deionized water in a glass beaker; cover with parafilm. (b) Warm the solution to 60°C with continuous stirring to dissolve the powder. (c) Allow the solution to cool to room temperature. (d) Add about 1–4 drops of 1 N NaOH, until the opaque color of the solution clears. (e) Add 10 mL of 1 M sodium phosphate. (f) Adjust the pH to 7.2–7.4 using color pH strips. (g) Add 1.026 g of sucrose. (h) Bring volume to 100 mL. (i) Filter through a 0.22-mm disposable filter unit into a bottle. (j) Store at 4°C in an aluminum foil-wrapped bottle for no more than 1 month. 13. Preparation of Tris-buffered saline (TBS): To make one liter of 10× TB: (a) Weigh 60.5 g of Tris-Base into a beaker. (b) Add 700 mL deionized water to the beaker. (c) Place the beaker on top of a magnetic stirrer. (d) When the powder has dissolved, adjust the pH to 7.4. (e) Add deionized water to bring the volume up to 1 L, mix well, autoclave or sterilize through filter, and store at 4°C. To make one liter of TBS: (a) Weigh 8.766 g NaCl in a beaker. (b) Add 100 mL of 10× TB to the beaker and mix vigorously.

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(c) When the powder has dissolved, add deionized water to bring the volume up to 1 L; mix well, sterilize through filter and store at 4°C. 14. The optimal speed to centrifuge the cell suspension should be further “fine tuned” by the investigator. High centrifugation speed results not only in preparations with better cell yields, but also with higher debris content. Debris may represent an obstacle not only for further analysis and/or treatments, such as flow cytometry and cell sorting, but also for the survival and adhesion of isolated myogenic stem cells. 15. To minimize debris resulting from muscle digestion, cell suspensions of freshly isolated satellite cells can be further purified by Percoll density centrifugation. Although this approach aims mostly to remove debris (76), a modification of this procedure that includes a multi-step Percoll gradient can also fractionate cell subpopulations (62). 16. For some antibodies the cultures may be blocked for just 2–4 h at room temperature if overnight blocking is not desired. 17. For even and continuous distribution of the antibodies (both primary and secondary), it is recommended to place 24-well plates on a gyrating platform rotator. This is important since, without agitation, the antibody solution tends to rapidly accumulate at the well periphery, leading to uneven staining across the culture. 18. DAPI is potentially harmful. Avoid prolonged or repeated exposure; we typically dissolve the entire powder in its original container and generate a concentrated stock solution. A readymade DAPI reagent is available from Molecular Probes. It is important to refer to the MSDS instructions and institutional regulations for further information regarding storage, handling, and first aid.

Acknowledgments We thank Lindsey Muir for reviewing this manuscript and providing valuable comments. We are also grateful to the granting agencies that funded this study. Our current research is supported by grants to Z.Y.R. from the National Institutes of Health (AG021566; AG035377; AR057794) and the Muscular Dystrophy Association (135908). M.E.D is supported by the Genetic Approaches to Aging Training Program (T32 AG000057). The development of the protocols described here could not be possible without early support to Z.Y.R from the American Heart Association, the USDA Cooperative State Research, Education and Extension Service, and the National Institutes of Health (AG013798).

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References 1. Hawke, TJ., and Garry, DJ. (2001) Myogenic satellite cells: physiology to molecular biology. J Appl Physiol 91, 534–551. 2. Zammit, PS., Partridge, TA., and YablonkaReuveni, Z. (2006) The skeletal muscle satellite cell: the stem cell that came in from the cold. J Histochem Cytochem 54, 1177–1191. 3. Yablonka-Reuveni, Z., and Day, K. (2011) Skeletal muscle stem cells in the spotlight: the satellite cell., in Regenerating the Heart: Stem Cells and the Cardiovascular System (Stem Cell Biology and Regenerative Medicine Series) (Cohen, I., and Gaudette, G., Eds.) Springer, Humana Press, Chapter 11, pp. 173–200. 4. Mauro, A. (1961) Satellite cell of skeletal muscle fibers. J Biophys Biochem Cytol 9, 493–495. 5. Katz, B. (1961) The terminations of the afferent nerve fibre in the muscle spindle of the frog. Philos Trans Royal Soc Lond 243, 221–240. 6. Bischoff, R. (1975) Regeneration of single skeletal muscle fibers in vitro. Anat Rec 182, 215–235. 7. Konigsberg, U.R., Lipton, B.H., and Konigsberg, I.R. (1975) The regenerative response of single mature muscle fibers isolated in vitro. Dev Biol 45, 260–275. 8. Yablonka-Reuveni, Z., and Rivera, A.J. (1994) Temporal expression of regulatory and structural muscle proteins during myogenesis of satellite cells on isolated adult rat fibers. Dev Biol 164, 588–603. 9. Rosenblatt, J.D., Lunt, A.I., Parry, D.J., and Partridge, T.A. (1995) Culturing satellite cells from living single muscle fiber explants. In Vitro Cell Dev Biol Anim 31, 773–779. 10. Zammit, P.S., Golding, J.P., Nagata, Y., Hudon, V., Partridge, T.A., and Beauchamp, J.R. (2004) Muscle satellite cells adopt divergent fates: a mechanism for self-renewal? J Cell Biol 166, 347–357. 11. Collins, C.A., Olsen, I., Zammit, P.S., Heslop, L., Petrie, A., Partridge, T.A., and Morgan, J.E. (2005) Stem cell function, self-renewal, and behavioral heterogeneity of cells from the adult muscle satellite cell niche. Cell 122, 289–301. 12. Shefer, G., Van de Mark, D.P., Richardson, J.B., and Yablonka-Reuveni, Z. (2006) Satellite-cell pool size does matter: defining the myogenic potency of aging skeletal muscle. Dev Biol 294, 50–66. 13. Day, K., Shefer, G., Richardson, J.B., Enikolopov, G., and Yablonka-Reuveni, Z. (2007) Nestin-GFP reporter expression

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2 25. Montarras, D., Morgan, J., Collins, C., Relaix, F., Zaffran, S., Cumano, A., Partridge, T., and Buckingham, M. (2005) Direct isolation of satellite cells for skeletal muscle regeneration. Science 309, 2064–2067. 26. Beauchamp, J.R., Heslop, L., Yu, D.S., Tajbakhsh, S., Kelly, R.G., Wernig, A., Buckingham, M.E., Partridge, T.A., and Zammit, P.S. (2000) Expression of CD34 and Myf5 defines the majority of quiescent adult skeletal muscle satellite cells. J Cell Biol 151, 1221–1234. 27. Ono, Y., Boldrin, L., Knopp, P., Morgan, J.E., and Zammit, P.S. (2010) Muscle satellite cells are a functionally heterogeneous population in both somite-derived and branchiomeric muscles. Dev Biol 337, 29–41. 28. Dellavalle, A., Sampaolesi, M., Tonlorenzi, R., Tagliafico, E., Sacchetti, B., Perani, L., Innocenzi, A., Galvez, B.G., Messina, G., Morosetti, R., Li, S., Belicchi, M., Peretti, G., Chamberlain, J.S., Wright, W.E., Torrente, Y., Ferrari, S., Bianco, P., and Cossu, G. (2007) Pericytes of human skeletal muscle are myogenic precursors distinct from satellite cells. Nat Cell Biol 9, 255–267. 29. Zheng, B., Cao, B., Crisan, M., Sun, B., Li, G., Logar, A., Yap, S., Pollett, J.B., Drowley, L., Cassino, T., Gharaibeh, B., Deasy, B.M., Huard, J., and Peault, B. (2007) Prospective identification of myogenic endothelial cells in human skeletal muscle. Nat Biotechnol 25, 1025–1034. 30. Tedesco, F.S., Dellavalle, A., Diaz-Manera, J., Messina, G., and Cossu, G. (2010) Repairing skeletal muscle: regenerative potential of skeletal muscle stem cells. J Clin Invest 120, 11–19. 31. Moss, F.P., and Leblond, C.P. (1971) Satellite cells as the source of nuclei in muscles of growing rats. Anat Rec 170, 421–435. 32. Campion, D.R. (1984) The muscle satellite cell: a review. Int Rev Cytol 87, 225–251. 33. Schultz, E. (1996) Satellite cell proliferative compartments in growing skeletal muscles. Dev Biol 175, 84–94. 34. White, R.B., Bierinx, A.S., Gnocchi, V.F., and Zammit, P.S. (2010) Dynamics of muscle fibre growth during postnatal mouse development. BMC Dev Biol 10, 21. 35. Schultz, E., Gibson, M.C., and Champion, T. (1978) Satellite cells are mitotically quiescent in mature mouse muscle: an EM and radioautographic study. J Exp Zool 206, 451–456. 36. Snow, M.H. (1978) An autoradiographic study of satellite cell differentiation into regenerating myotubes following transplanta-

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Chapter 3 Isolation of Muscle Stem Cells by Fluorescence Activated Cell Sorting Cytometry Alessandra Pasut, Paul Oleynik, and Michael A. Rudnicki Abstract Satellite cells are a heterogeneous population of muscle progenitors with stem cell properties responsible for the regeneration of adult skeletal muscle. Increasing interest in the therapeutic potential of satellite cells has challenged researchers with the need to purify a homogenous population of muscle progenitors. Here we provide a detailed protocol for the isolation of a pure population of satellite cells using fluorescence activated cell sorting. We give specific guidelines to ameliorate the reproducibility of the satellite cell isolation protocol with the goal to standardize procedures across labs. This protocol identifies satellite cells within adult skeletal muscle as an enriched population of Integrin A7+/CD34+ double positive cells and CD45, CD31, CD11b, and Sca1 negative (Lin−) cells (Integrin A7+/CD34+/Lin−).. Functional assay shows that Integrin A7+/CD34+/Lin− satellite cells possess high myogenic potential and ability to regenerate muscle depleted satellite cells upon transplantation. Key words: Satellite cells, Markers, FACS, Cytometry, Fluorochromes

1. Introduction By providing a lifelong reservoir of muscle progenitors, satellite cells are the main contributors of muscle regeneration (1). Although accounting for only a small percentage of total myonuclei, satellite cells are able to repopulate damaged muscles by activating and differentiating into mature myofibers and at the same time self renew the original pool (2–5). Heterogeneity is a well-established and most likely unique feature of satellite cells (1, 4). Thus appropriate tools that guarantee the isolation of homogenous isolation of homogenous populations are central to the manipulation of satellite cells and to effectively establish the ability of different satellite cell subpopulations to regenerate injured muscles.

Joseph X. DiMario (ed.), Myogenesis: Methods and Protocols, Methods in Molecular Biology, vol. 798, DOI 10.1007/978-1-61779-343-1_3, © Springer Science+Business Media, LLC 2012

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Fluorescence activated cell sorting (FACS) is an assay in which single cell properties either physical or chemical are simultaneously analyzed in a fluid stream system and used to separate a heterogeneous sample into distinct groups of cells (6, 7). In a modern flow cytometer, a laser of a selected wavelength is directed toward a flowing stream of single cells and the properties of light scattering is used to infer parameters such as cell size, cell granularity, or DNA content (6, 7). The availability of fluorescently tagged antibodies that specifically recognize and bind to cell surface antigens is an efficient tool to further discriminate stem cells from more differentiated cells within adult tissues or to separate a subpopulation of stem cells within the same pool. Satellite cell isolation by FACS is an incredibly resourceful but often challenging tool. When starting from a heterogeneous sample such as muscle tissue, several steps need to be performed to obtain a highly pure population of muscle progenitors. In this chapter we offer guidelines and suggestions to improve tissue digestion, single cell preparation, sample labeling, and sample sorting. Satellite cells do not express a unique stem cell marker; rather they can be distinguished from other muscle cells by using a combination of both negative and positive markers. The cell surface markers CXC motif receptor R-4 (CXCR4), the vascular cell adhesion molecule 1 (V-CAM-1), Integrins A7B1 or CD34 are used by different labs to identify satellite cells by FACS (2, 3, 5, 8–12). In the protocol herein described satellite cells are identified as an enriched population of Integrin A7/CD34 double positive cells and CD45, CD31, CD11b, and Sca1 negative (Lin−) cells (Integrin A7+/CD34+/Lin−). More importantly, transplanted Integrin A7+/ CD34+/Lin− cells can efficiently regenerate muscles by both repairing and fusing with damaged fibers and contributing to a self-renewal pool (3, 5). FACS coupled with high throughput gene expression studies allowed the elucidation of novel signaling pathways and molecules involved in satellite cell self-renewal and activation (5, 9). It is thus clear that the development of reproducible and standardized methods for the isolation of stem cells is of paramount importance especially in the context of translational research.

2. Materials 2.1. Reagents

1. Sterile surgical tools. 2. Netwell Mesh Filters, 74 Mm (Costar). 3. CellTrics disposable filters, 50 Mm (Partec). 4. Characterized Hyclone Fetal Bovine Serum (FBS; Thermo Scientific).

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5. 2.5 U/mL Dispase II (Roche). 6. 2.5 U/mL Collagenase B (Roche). 7. Monoclonal antibody anti-mouse Integrin A7 clone 3C12 (MBL). 8. Alexa647 anti-mouse IgG1 (Molecular Probes, Invitrogen). 9. Phycoerythrin (PE) anti-mouse CD11b (eBioscience). 10. Phycoerythrin (PE) anti-mouse CD45 (eBioscience). 11. Phycoerythrin (PE) anti-mouse Ly-6A-E (Sca1; BD Bioscience). 12. Phycoerythrin (PE) anti-rat CD31 (BD Bioscience). 13. Biotin anti-mouse CD34 (eBioscience). 14. Streptavidin-APC-Cy7 (BD Bioscience). 15. Hoechst 33342 (Sigma). 16. Phosphate Buffer Saline (PBS). 2.2. Equipment

1. Cell sorter equipped with three lasers: 488 nm laser for the excitation of PE fluorochrome, 633 nm laser for the excitation of APC and APC-Cy7 fluorochrome, and UV for the excitation of Hoechst dye.

3. Methods Mononuclear cells from a digested muscle preparation are processed using the Beckman-Coulter MoFlo cytometer (DakoCytomation) equipped with 488, 633 nm, and UV lasers. The Summit V4.3 software suite was used to analyze all results. The protocol is divided into four sections: preparation of single cell suspension, cell staining, performing FACS, and downstream applications. All procedures are to be performed at room temperature unless otherwise specified. 3.1. Single Cell Suspension

1. Dissect hind limb muscles from 6 to 8 week old mice for optimal satellite cell yield. Use a razor blade or other appropriate tools to first remove any hairs. Expose the muscle completely by cutting through the thick membrane surrounding the tissue. Dissect the muscles following their lengths and anatomy. Collect muscles in cold PBS until ready to proceed with all the samples. 2. If necessary, wash muscles in cold PBS a couple of times. 3. Using small scissors or other appropriate tools, carefully mince the muscles. If any bones are left, separate the bones from the muscles. Remove intramuscular fat pads (white and soft tissue).

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4. Proceed with enzymatic digestion by adding 5 mL of collagenasedispase solution (2.5 U/mL) per each muscle preparation (two hind limbs) and incubate for 10–12 min at 37°C. 5. Coat a plastic pipette with FBS to prevent cells from sticking to the pipette walls and resuspend minced muscles up and down a few times to help tissue digestion. 6. Repeat steps 4 and 5 until the tissue is almost entirely digested and appears as a homogenous solution with few or no tissue chunks. Depending on the enzyme activity, the incubation time can vary between 30 and 60 min. 7. Add 2 volumes of 10% FBS in PBS to inactivate enzyme activity and filter the solution through a 74 Mm mesh filter. Centrifuge at 239 × g for 5 min. Keep the cell pellet and transfer the supernatant to a clean tube. 8. Centrifuge the supernatant one more time. Combine the two cell pellets in a new tube. 3.2. Cell Staining

1. Resuspend the cell pellet in 1 mL of 2% FBS in PBS. Take an aliquot (10 ML) and count the number of cells. It is recommended to dilute the aliquot at least 2 times to obtain an accurate measurement of total cell number. The staining protocol and corresponding volumes listed below are suitable for 10 million cells. Optional: if erythrocytes are presents in the cell pellet, before proceeding to count, add Red Blood Cell Lysis buffer (Sigma) as per manufacturer instruction and then proceed to count. Erythrocytes will anyway be excluded during the sorting strategy (see Subheading 3.3, step 2). 2. Prepare the following controls: (a) Set aside an aliquot of 50,000 cells to determine the threshold for cell autofluorescence (unstained control) (see Subheading 3.3, step 3). (b) Set aside three aliquots of 50,000 cells to be used as single color controls. In this specific case, the single color controls are Alexa647, APC-Cy7, and PE. The ratio of labeled antibody to cell number is the same used for the actual sample. The purpose of this control will be discussed in Subheading 3.3, steps 4 and 5. Dilute each control up to 500 ML in 2% FBS in PBS and filter the cell suspension using a 50 Mm disposable filter to ensure a single cell suspension. Keep solutions on ice, protected from light until ready to process. 3. Adjust the final volume of the actual sample to 1 mL before proceeding further. 4. Add to the cell suspension 5 ML of monoclonal antibody antimouse Integrin A7 (1 mg/mL) and 10 ML of biotin anti-mouse

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CD34 (0.5 mg/mL) per 10 million cells. Because different batches of antibodies might have slightly different concentrations, it is always recommended to titrate each new batch of antibody (see Note 1). 5. Incubate cells on ice for 15 min and gently shake cell suspension every 5 min to prevent cell clumps. 6. Add 9 mL of cold 2% FBS in PBS and centrifuge for 5 min at 239 × g to wash unbound primary antibody. 7. Resuspend the pellet in 1 mL of cold 2% FBS in PBS. Add 5 ML of Alexa647 anti-mouse IgG (2 mg/mL) and 2.5 ML of Streptavidin APC-Cy7 (0.2 mg/mL). Add 2.5 ML of PE antimouse CD45 (0.2 mg/mL), 2.5 ML of PE anti-mouse CD11b (0.2 mg/mL), 2.5 ML of PE anti-mouse Sca1 (0.2 mg/mL), and 2.5 ML of PE anti-rat CD31 (0.2 mg/mL), also referred to as Lin−. If a UV laser is available, add 5 ML of Hoechst dye (1 mg/mL). Incubate cells on ice for 15 min protected from light. If a UV laser is not available, use another cell viability dye such as propidium iodide (PI) to discriminate dead-live cells (see Note 2). 8. Add 9 mL of cold 2% FBS in PBS and centrifuge for 5 min at 239 × g to wash unbound secondary antibody. 9. Resuspend cells in 500 ML of cold 2% FBS in PBS and filter cell suspension through a 50 Mm disposable filter to ensure a single cell suspension. Keep cells on ice until ready to sort. 3.3. Performing FACS

These are general rules that should be followed when sorting satellite cells. Instrument calibration, cleaning procedure, or other routine operations are here omitted due to space limitations. 1. The following dot plots are used to create a satellite cell sorting profile on Summit or any equivalent flow cytometry/FACS software: (a) Side scatter (SSC) in logarithmic scale (log) vs. forward scatter (FSC) in linear scale (lin). SSC represents the light scattered and collected at a 90° angle. It is an indication of intrinsic cellular granularity. FSC represents the light scattered and collected at a 180° angle. It is an indication of cell size. (b) FSC linear vs. pulse width is also known as doublet discrimination. It distinguishes between singlets (single cells) and doublets (two cells joined together). This plot is needed to guarantee that a purified population of single cells is sorted. (c) SSC log vs. PE log distinguishes between Lin− and Lin+ cells. (d) Alexa647 log vs. APC-Cy7 log for compensation purposes.

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(e) SSC log vs. Hoechst log distinguishes between dead (Hoechst−) and live (Hoechst+) cells. (f) SSC log vs. Alexa647 log distinguishes between Integrin A7− and Integrin A7+ cells. (g) SSC log vs. APC-Cy7 log distinguishes between CD34− and CD34+ cells. 2. Analyze the autofluorescence control by running the sample at a rate of roughly 1,000 events (cells)/s. Since these cells do not contain any label, they will appear as negative events. While analyzing the sample on cycle (noncumulatively), adjust the sensitivities of each detector until the events are displayed in the first decade of the X-axis in the corresponding fluorescence plots (PE, Alexa647, Alexa647-Cy7, and Hoechst). Collect and save the data for 50,000 events by switching to cumulatively data collection (turn cycle off). In the SSC-log vs. FSC lin plot (see Fig. 1a) draw a region (R1) being careful to exclude the line of events that have extremely low FSC and variable SSC. These events represent dead cells, debris, and erythrocytes. Satellite cells have low FSC and low-medium SSC. 3. In the FSC lin vs. pulse width plot (see Fig. 1b), draw a region (R2) that encompasses the majority of the events. These are the singlets. Doublets are characterized by large pulse width and should not be included in R2. Doublets account for a maximum of 10% of a properly prepared sample. Any result larger than this indicates the lack of a single cell suspension and the sample should be refiltered. Inclusion of doublets could also result in the collection of false positives. 4. Analyze the single color controls separately and save 50,000 events each. These cells will be viewed as negative events in the corresponding fluorescence plot. In the SSC log vs. PE log plot (see Fig. 1c) draw a region around the negative population. This region (R3) represents Lin− (PE−) cells and accounts for 70% of total events. 5. Analyze the Alexa647 single color control. Compensate out any Alexa647 signal detected on the APC-Cy7 plot (see Note 3). Analyze the sample and save 50,000 events. 6. In the SSC log vs. Hoechst log plot (see Fig. 1d), draw a region (R4) around the Hoechst+ cells. Hoechst+ cells should account for 70–90% of total events. 7. In the SSC log vs. Alexa647 log plot (see Fig. 1e), construct and apply a gate (G1) that includes R1, R2, R3, and R4. Satellite cells should appear as a small, tight population of events with low-medium SSC and high Alexa647 signal. Draw a region (R5) around this population.

a

d

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Hoechst log

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Pulse Width

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R5:Integrin ŷ7+/Lin–

R2

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R6:Integrin ŷ7+/CD34 +/Lin –

R3

Fig. 1. Isolation of satellite cells from a heterogeneous muscle preparation by FACS. Dot plots representing the sequential gating strategy used to identify satellite cells from a heterogonous muscle sample. (a) Satellite cells appear as low FSC and low to medium SSC events. FSC is shown in linear scale while SSC is shown in logarithmic scale. (b) Doublets discrimination ensures that only a single cell suspension is analyzed and sorted. (c) Hoechst is used to discriminate between live (Hoechst+) and dead (Hoechst−) events. (d) CD45 and CD11b blood lineage cells, Sca1 mesenchymal progenitors, and CD31 endothelial cells are excluded by gating on PE-events. PE-events account for ~70% of the total events in a standard muscle preparation (e) The round gate represents a distinct population of Integrin A7 and Lin− cells in the Alexa647 log vs. SC log plot. (f) The majority of satellite cells are double positive for the cell surface markers Integrin A7 and CD34. Integrin A7+/CD34+/Lin− satellite cells account for 1–4% of total events. The Summit softwareV4.3 is used to derive FACS analysis.

SSC log

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8. In the SSC log vs. APC-Cy7 log plot (see Fig. 1f), construct and apply a new gate (G2) that includes R1, R2, R3, R4, and R5. Satellite cells appear in a similar location as found in Fig. 1e. Draw a region (R6) around these cells. 9. The sort logic applied to collect satellite cells is R1 + R2 + R3 + R4 + R5 + R6. This sorting strategy identifies satellite cells as Integrin A7+/CD34+/Lin− cells. Sort into collection tubes containing 2% FBS in PBS kept on ice. To facilitate and ameliorate cell recovery, the sort is performed under low pressure with a maximum flow rate of 3,000 events/s while using a 100 Mm nozzle. Higher concentration serum or alternative media ( i.e.: primary myoblasts medium) can be used to collect cells during the sort. After the first sort is complete, it is possible to run the sample another time for highest purity; however, the final yield of satellite cells might be heavily depleted because of either cell death or decreased fluorescence intensity of the antibodies resulting in higher rates of false negatives. 3.4. Downstream Applications

Each cell sorting must be followed by a careful characterization of the phenotype which should include immunostaining on freshly cytospunned cells and real time PCR (Q-PCR) for satellite cell specific markers to confirm the nature of the sorted population. This protocol allows for the isolation of an extremely pure population of satellite cells with the ability to both participate in new fiber formation as well as replenish satellite cell pools in impaired muscles (3, 5, 8). If performing immunostaining on freshly sorted cells, resuspend cells in low volume (max 200 ML) and cytospin at 700 rpm (Cytospin 4, ThermoShendon) for 5 min using appropriate coated slides (Shandon Double Cytoslide coated). Increasing the speed might result in damaging cell integrity and poor quality staining. Care should also be taken during the immunostaining. Gentle washing is recommended to avoid losing cells. Integrin A7+/ CD34+/Lin− satellite cells contain more than 90% Pax7+ satellite cells and express satellite cell specific markers by Q-PCR (5). When sorting satellite cells for gene expression analysis (microarray), it is recommended to decontaminate the FACS sorter and minimize the presence of RNAse by using specific RNAse-free products and cleaning solutions. After the sorting, briefly centrifuge the cells to remove serum. Sorted cells do not pellet very easily, so extreme care should be taken to avoid disturbing the pellet. Resuspend the pellet in an appropriate volume of Trizol or other suitable RNA extraction buffer and immediately proceed with the isolation. Addition of Glycol Blue or other RNA carriers during the RNA isolation protocol might be useful to identify the pellet. When combining multiple sortings, it is best to isolate RNA after each sorting and then concentrate RNA at the desired volume using a speed-vacuum centrifuge system. Freezingthawing Trizol-cell pellets decreases the final RNA yield.

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Flow cytometry and FACS are undoubtedly valuable tools to obtain homogenous populations of stem cells. Continuous advances in technology made FACS suitable for different downstream applications such as gene expression studies, transplantation studies or single cell analysis. However, the increasing number of publications using FACS data requires unifying and standardizing the procedures to obtain and properly interpret FACS data. Here we summarize and emphasize the importance of the following information: (1) Machine settings: model of cytometer, number of lasers and sorting parameters (nozzle diameter and pressure); (2) Sample handling: if starting from heterogeneous sample, the type of tissue dissociation (sonication, homogenization, enzymatic, mechanical), antibodies list and dilutions, number of labeled cells, type of live-dead assay. (3) Flow analysis: software used to perform the sorting, sorting scheme, description of compensation, and type of controls used (i.e., fluorescence minus one (FMO) or single control). A detailed list of minimum accompanying information is also provided by Lee et al. (13). Flow data must always be accompanied by a complete functional assay. The use of standard procedures to isolate and identify satellite cells from muscle tissues and a careful description of the sorting strategy will allow interpreting and comparing flow data across labs and impact the reproducibility of FACS derived data.

4. Notes 1. Antibody labeling optimization (Titration). Flow cytometry is informative under saturating labeling condition (5). The goal with an antibody titration is to optimize the ratio of labeled antibody to cell number. To perform an antibody titration, prepare aliquots containing the same number of cells. Save one aliquot as an unstained (autofluorescence) control. To the rest, add an increasing amount of the antibody of interest. Analyze each aliquot on a flow cytometer/FACS instrument from the lowest antibody concentration to the highest. The fluorescence intensity should increase in strength until it reaches a maximum above which the signal does not increase anymore. The corresponding antibody concentration is called saturating condition and represents the amount of labeled antibody that should be used. When titrating an antibody for the first time, it is also useful to determine whether the antibody of choice can nonspecifically bind to antigen or other receptors on target cells thus contributing to either increased cell autofluorescence or increased background signal. To do so, it is recommended to run isotype controls for each antibody used for FACS. Isotype controls must match the species, the isotype and the

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fluorochrome of the antibody of interest and be used at the same saturating condition to obtain meaningful information. Isotype controls do not substitute for single color controls. 2. Viability staining. The goal of viability staining is to distinguish between live and dead cells in a FACS profile and to exclude dead cells from the sorted cells. We have employed Hoechst (where Hoechst+ cells are live cells) because its fluorescence emission is significantly separated from the other fluorochromes here employed. Other viability stains include propidium iodide (PI) or costained with PI and Hoechst. 3. Compensation. Compensation is a procedure by which it is possible to remove artifacts (i.e., inclusion of false positives) when fluorochromes with spectral overlap (spillover) are used in the same sample (14). In the present sort, Alexa647 and the tandem dye APC-Cy7 are the two fluorochromes with spectral overlap. When performing FACS using multiple fluorochromes it is best to follow these simple rules: (1) Low abundant or weak antigens should be labeled with the brightest fluorochromes available. (2) If compensation due to spectral overlap needs to be performed, it is preferable to choose dyes with sufficiently distinct spectral overlap to clearly distinguish negative from positive cells and minimize the inclusion of contaminants. In this case, the two antigens used to identify satellite cells (Integrin A7 and CD34) are conjugated with APC and APC-Cy7, respectively, which do not show spectral overlap with PE thus decreasing the inclusion of contaminants in the final sort. Here compensation is applied only to remove any Alexa647 signal from the APC-Cy7 plot. To do this, load the autofluorescence control and draw quadrant regions on SSC log vs. APC-Cy7 log (see Fig. 2a). Then, load the Alexa647 single color control on the same plot (see Fig. 2b). As shown in Fig. 2b, Alexa647 spillover into the APC-Cy7 channel results in the appearance of APC-Cy7 positive cells. Apply compensation until no APC-Cy7 positive events are seen (see Fig. 2c). For accurate description of compensation using Summit Software refer to Summit Software training guide (15). Alongside single color controls, FMO controls should also be analyzed. To prepare FMO controls, label the sample with all the antibodies except for one (i.e., Alexa647-FMO: cells are stained with all antibodies except Alexa647).

Alexa647 log comp

APC-Cy7 log comp

c

APC-Cy7 log comp

Fig. 2. Compensation strategy. Alexa647 and APC-Cy7 tandem dye have similar emission spectra. For this reason, any spillover of the Alexa647 in the APC-Cy7 channel must be corrected. (a) All events in the autofluorescence control sample should appear as double negative on the APC-Cy7 vs. Alexa647 plot. (b) Alexa647 spillover into the APC-Cy7 channel results in the appearance of false positive events. (c) After compensation is applied no Alexa647 cells are detected into the APC-Cy7 channel. The Summit softwareV4.3 is used to derive compensation analysis.

APC-Cy7 log comp

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References 1. Rudnicki MA, Le Grand F, McKinnell I, Kuang S (2008) The molecular regulation of muscle stem cell function. Cold Spring Harb Symp Quant Biol. 73:323–31 2. Kuang S, Kuroda K, Le Grand F, Rudnicki MA (2007) Asymmetric self-renewal and commitment of satellite stem cells in muscle. Cell 129(5):999–1010 3. Sacco A, Doyonnas R, Kraft P, Vitorovic S, Blau HM (2008) Self-renewal and expansion of single transplanted muscle stem cells. Nature 456(7221):502–6 4. Collins CA, Olsen I, Zammit PS, Heslop L, Petrie A, Partridge TA, Morgan JE (2005) Stem cell function, self-renewal, and behavioral heterogeneity of cells from the adult muscle satellite cell niche. Cell 122(2):289–301 5. Le Grand F, Jones AE, Seale V, Scimè A, Rudnicki MA (2009) Wnt7a activates the planar cell polarity pathway to drive the symmetric expansion of satellite stem cells. Cell Stem Cells 5;4(6):535–47 6. HM Shapiro (2003) Practical Flow Cytometry. Fourth edition, Wiley-Liss Inc 7. Hoffman RA (2007) Current Protocols in Cytometry. Wiley Interscience 8. Blanco-Bose WE, Yao C-C, Kramer RH, Blau HM (2001) Purification of mouse primary myoblasts based on A7 Integrin expression. Exp Cell Res 265:212–220 9. Fukada S, Uezumi A, Ikemoto M, Masuda S, Segawa M, Tanimura N, Yamamoto H, Miyagoe-Suzuki Y, Takeda S (2007) Molecular signature of quiescent satellite cells in adult skeletal muscle. Stem Cells 25(10):2448–59

10. Cerletti M, Jurga S, Witczak CA, Hirshman MF, Shadrach JL, Goodyear LJ, Wagers AJ (2008) Highly efficient, functional engraftment of skeletal muscle stem cells in dystrophic muscles. Cell 134(1):37–47 11. Fukada S, Higuchi S, Segawa M, Koda K, Yamamoto Y, Tsujikawa K, Kohama Y, Uezumi A, Imamura M, Miyagoe-Suzuki Y, Takeda S, Yamamoto H (2004) Purification and cellsurface marker characterization of quiescent satellite cells from murine skeletal muscle by a novel monoclonal antibody. Exp Cell Res 296(2):245–55 12. Beauchamp JR, Heslop L, Yu DS, Tajbakhsh S, Kelly RG, Wernig A, Buckingham ME, Partridge TA, Zammit PS (2000) Expression of CD34 and Myf5 defines the majority of quiescent adult skeletal muscle satellite cells. J Cell Biol 151(6):1221–34 13. Lee JA, Spidlen J, Boyce K, Cai J, Crosbie N, Dalphin M, Furlong J, Gasparetto M, Goldberg M, Goralczyk EM, Hyun B, Jansen K, Kollmann T, Kong M, Leif R, McWeeney S, Moloshok TD, Moore W, Nolan G, Nolan J, NikolichZugich J, Parrish D, Purcell B, Qian Y, Selvaraj B, Smith C, Tchuvatkina O, Wertheimer A, Wilkinson P, Wilson C, Wood J, Zigon R; Scheuermann RH, Brinkman RR (2008) MIFlowCyt: the minimum information about a flow cytometry experiment. Cytometry A 73(10):926–30 14. Roederer M (2001) Spectral Compensation for Flow Cytometry: Visualization Artifacts, Limitations, and Caveats. Cytometry 45:194–205 15. Prursley S (2007) Summit Software Training Guide, Beckman Coulter Inc

Chapter 4 Mouse and Human Mesoangioblasts: Isolation and Characterization from Adult Skeletal Muscles Mattia Quattrocelli, Giacomo Palazzolo, Ilaria Perini, Stefania Crippa, Marco Cassano, and Maurilio Sampaolesi Abstract Mesoangioblasts (MABs) are mesoderm-derived stem cells, associated with small vessels and originally described in the mouse embryonic dorsal aorta. Similar though not identical cells have been later identified and characterized from postnatal small vessels of skeletal muscle and heart. They have in common the expression of pericyte markers, the anatomical location, the ability to self-renew in culture, and to differentiate into various types of mesodermal lineages upon proper culture conditions. Currently, the developmental origin of MABs and the relationship with other muscle stem cells are not understood in detail and are the subject of active research. This chapter provides an outline of the latest techniques for isolation and characterization of adult MABs from human and mouse skeletal muscles. Key words: Mesoangioblasts, Pericytes, Muscle stem cells, FACS, Surface antigens, Cocultures, Mesodermal lineages

1. Introduction Skeletal muscle regeneration is mainly sustained by satellite cells (1), local myogenic progenitors localized underneath the basal lamina of muscle fibers. Differently, cardiac muscle is less efficient to regenerate and tends to develop scar tissue after injuries (2). In the last years, several groups have reported the presence of local stem/progenitor cells able to differentiate into cardiac (3–6) and skeletal muscle lineages (7–13). In this chapter, we provide protocols for isolation, cloning, expansion, and characterization of mesoangioblasts (MABs) derived from murine and human adult skeletal muscles (see Subheading 3.1 and 3.2). However, it could be possible to apply slightly modified

Joseph X. DiMario (ed.), Myogenesis: Methods and Protocols, Methods in Molecular Biology, vol. 798, DOI 10.1007/978-1-61779-343-1_4, © Springer Science+Business Media, LLC 2012

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techniques to isolate cardiac MABs, not considered in this chapter due to constrain of space. The corresponding GLP/GMP human cells have been isolated and characterized (Molmed, Italy), ready to use for clinical trials. We will also describe various cell differentiation methods, i.e., coculture with C2C12 myoblasts (see Subheading 3.3), spontaneous differentiation (see Subheading 3.4), induction of smooth muscle (see Subheading 3.5), osteoblasts (see Subheading 3.6), and adipocytes (see Subheading 3.7). In addition, we will present procedures for collagen-based coating of tissue culture surfaces (see Subheading 3.8) and freezing procedures for MABs (see Subheading 3.9). MABs must be cultured under physiological O2 conditions (5% O2, 5% CO2, 90% N2). Basic animal handling, dissection, and tissue culture skills are necessary for successful isolation and propagation of MABs. Expertise in histochemistry, biochemistry, and molecular biology is required for MABs characterization. Importantly, sterile conditions in either Class II biohazard flow hoods (recommended for human material) or laminar flow hoods are required. Institutional Animal Care and Use Committee (IACUC) should approve the protocols. Muscle biopsies should be performed under general or local anesthesia with the minimum degree of pain. Approval of Institutional Ethics Committee and patients’ informed consent are necessary in the case of human samples.

2. Materials Researchers consider cell culture as black art, due to the enormous variables that require much effort to find solutions to upcoming problems. However, a systematic approach for each step in the process will help to solve critical issues. Diligently follow the protocols and refer to the notes for troubleshooting. Primary culture from adult skeletal muscle results in a mixed population of cells that includes MABs. Because of the inability of other cell types to rapidly proliferate under these conditions, MABs increase their number in proportion and take over the mixed population, making it easier for their isolation and cloning. All reagents are provided by Gibco, unless otherwise stated. 2.1. Basic Materials

1. Skeletal muscle fragments from murine or human samples (see Subheading 3). 2. Ca2+/Mg2+-free phosphate-buffered saline (PBS), sterile. 3. DMEM-20 medium (see Subheading 2.3), sterile. 4. MEGA-5 medium (see Subheading 2.3), sterile. 5. TrypLE Express Trypsin, sterile.

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6. 3.5-, 6-, 10-, and 15-cm Petri dishes. 7. 3.5 cm Collagen-coated dishes (see Subheading 3.8). 8. Rounded-edge disposable scalpels, sterile. 9. Curved forceps, sterile. 10. Sharp-edged straight forceps, sterile. 11. 5% CO2, 5% O2, 90% N2 incubator. 2.2. FluorescenceActivated Cell Sorter Isolation Supplementary Materials

1. Fluorescence-activated cell sorter (FACS)-suitable polystyrene and polypropylene tubes.

2.3. Medium Recipes

1. DMEM-20 medium (250 mL): 200 mL DMEM high glucose, supplemented with 50 mL of heat-inactivated fetal bovine serum (FBS), 1% penicillin/streptomycin solution (100 units), 2 mM glutamine, 1 mM sodium pyruvate, and 1× nonessential amino acid solution. Filter through a 0.22 Mm membrane to ensure sterility. Store at 4°C for up to 2 weeks.

2. Phycoerythrin-conjugated monoclonal anti-human/mouse alkaline-phosphatase (AP), clone B4-78 (R&D, USA). 3. Phycoerythrin-conjugated mouse IgG1 Isotype Control (R&D, USA).

2. MEGA-5 medium (250 mL): 237.5 mL DMEM Megacell (Sigma), supplemented with 12.5 mL of heat-inactivated FBS, 1% penicillin/streptomycin solution, 2 mM glutamine, 1× nonessential amino acid solution, 0.1 mM 2-mercaptoethanol, and 1.25 Mg human bFGF (Peprotech). Filter through a 0.22 Mm membrane to ensure sterility. Store at 4°C for up to 2 weeks. 3. DMEM-10 medium (250 mL): 225 mL DMEM high glucose, supplemented with 25 mL of heat-inactivated FBS, 1% penicillin/ streptomycin solution, 2 mM glutamine, 1 mM sodium pyruvate. Filter through a 0.22 Mm membrane to ensure sterility. Store at 4°C for up to 2 weeks. 4. Freezing medium (FM; 50 mL): 45 mL heat-inactivated FBS supplemented with 5 mL Hybri-MAX® DMSO (Sigma). Filter through a 0.22 Mm membrane to ensure sterility. Store at 4°C for up to 4 weeks. 5. Differentiation medium (DM; 250 mL): 245 mL DMEM high glucose, supplemented with 5 mL of heat-inactivated Horse Serum (HS), 1% penicillin/streptomycin solution, 2 mM glutamine, and 1 mM sodium pyruvate. Filter through a 0.22 Mm membrane to ensure sterility. Store at 4°C for up to 2 weeks. 6. Smooth muscle induction medium (SMM; 250 mL): 245 mL DMEM high glucose, supplemented with 5 mL of heat-inactivated horse serum (HS), 1% penicillin/streptomycin solution, 2 mM

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glutamine, 1 mM sodium pyruvate, and 1.25 Mg TGFB (Peprotech). Filter through a 0.22 Mm membrane to ensure sterility. Store at 4°C for up to 2 weeks. 7. Osteogenic induction medium (OM; 170 mL): 170 mL human mesenchymal stem cell (hMSC) Osteogenic Basal Medium, supplemented with manufacturer’s single aliquots of dexamethasone, glutamine, ascorbic acid, 2-glycerophosphate, mesenchymal cell growth supplement (MCGS), and penicillin/ streptomycin (all reagents by Lonza). Store at 4°C in the dark for 4 weeks. 8. Adipogenic induction medium (AM; 170 mL): 170 mL hMSC Adipogenic Induction Medium, supplemented with manufacturer’s single aliquots of dexamethasone, indomethacin, recombinant insulin, glutamine, 3-isobutyl-1-methyl-xanthine (IBMX), MCGS and gentamicin sulfate and amphotericin B (GA1000), (all reagents by Lonza). Store at 4°C in the dark for 4 weeks. 9. Adipogenic maintenance medium (AMM; 170 mL): 170 mL hMSC AMM, supplemented with manufacturer’s single aliquots of recombinant insulin, glutamine, MCGS, and GA1000 (all reagents by Lonza). Store at 4°C in the dark for 4 weeks.

3. Methods Murine adult MABs, differently from their embryonic counterparts, express pericyte markers (such as NG2, CD140a, CD140b, and alkaline phosphatase) and lack endothelial markers, such as CD31 (see Table 1). Regarding differences in isolation and cloning of MABs, primary cultures from adult tissues show a slower

Table 1 MABs markers under proliferation conditions (by FACS/WB/ RT-PCR/qPCR) Murine MABs

Positive

Negative Human MABs

Positive

Negative

AP(+), NG2(+), Sca1(++), CD34(+), CD44(++), CD117(+), CD140a(++), CD140b(++) CD31, CD45, CD56, CD133 AP(+), NG2(+), SSEA4(+), CD13(++), CD44(++), CD49f(+), CD56(+), CD90(++), CD140a(+), Cd140b(++) CD31, CD34, CD45, CD133

(+) = Slightly positive; (++) = highly positive

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growth rate and lower cloning efficiency compared with embryonic counterparts. Mouse adult muscle fragments can be stored in DMEM-20 medium up to 24 h at 4°C prior to processing. We have also defined “pericyte-derived cells,” the human MABs isolated from adult skeletal muscle, since they express a range of pericyte markers and do not express endothelial markers, such as CD31 and CD34 (14). Although the origin of adult MABs is not completely clarified, it is likely that adult pericyte-derived cells originate from embryonic MABs (11). 3.1. FACS-Isolation of Adult Murine MABs

1. Dissect skeletal muscles from juvenile (see Note 1) murine hind limbs in sterile conditions and remove carefully any trace of fur, skin, and fat. 2. Rinse immediately the muscles in 5 mL of PBS in a 10 cm dish to remove blood cells. In case of separated isolations from different muscles, it is recommended to use a Petri dish per muscle. 3. Transfer muscles in a new 10 cm dish and dissect them in ~2 mm fragments with a sterile round-shaped scalpel. Discard eventual traces of fibrous tissue. 4. Transfer muscle fragments onto a 3.5 cm collagen-coated Petri dish (see Subheading 3.8 and Note 2) with a sterile curved forceps. Tissue fragments (usually six to seven per dish) should be positioned at 8–9 mm distance from each other to ensure optimal yields. 5. Carefully drop 100 ML of prewarmed DMEM-20 on top of each piece and incubate at 37°C (see Note 3) for 18–24 h in a 5% CO2, 5% O2 humidified incubator (see Note 4). 6. Cover the fragments with 1.8 mL of DMEM-20/dish. Slowly pipet the medium on the side of each dish to avoid fragment detachment. Remove eventual nonattached fragments with a sterile curved forceps (see Note 5). Incubate at 37°C in a 5% CO2, 5% O2 humidified incubator generally for at least 72 h. 7. Starting from this time point, check the fragments daily to monitor the extent of cell spread from the biopsies. In case of medium acidification and toning, gently remove the medium and rinse with fresh DMEM-20. As soon as the spreading cell layer reaches approximately 5 mm (see Fig. 1 and Note 6) from each fragment, proceed immediately to next step. 8. Carefully remove the fragments with a sterile sharp-edged forceps, gently remove the medium and wash with PBS. Detach the cells with 600 ML of prewarmed trypsin, incubating the dish for 2–3 min at room temperature. Once detached, add 1 mL of DMEM-20, mix and collect by gentle pipetting in a 15 mL tube and centrifuge for 5 min at 300 × g at room temperature.

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Fig. 1. Schematic representation of fluorescence-activated cell sorter (FACS)-isolation of mesoangioblasts (MABs). The scheme represents muscle fragments on collagen-coated dishes, cells spreading from the fragment, sorting of AP+/AP− fractions by FACS-analysis and a typical AP staining of the sorted populations. White bar: 100 Mm.

9. Resuspend homogeneously the pellet in 2 mL of DMEM-20, avoiding any clump. Plate the cells onto a new 3.5 cm collagen-coated Petri dish and incubate at 37°C in a 5% CO2, 5% O2 humidified incubator. 10. Upon 70–75% confluence (generally after 48–72 h), detach the cells as described above, remove the supernatant and resuspend in 1 mL of PBS and prepare three sterile FACS-suitable capped polystyrene tubes as follows: (1) 105 cells – Blank sample; (2) 105 cells – Isotype sample; (3) 2 × 105 cells – sorting sample. 11. Spin down for 5 min at 300 × g at room temperature, remove the supernatant and resuspend the pellets by pipetting or gentle vortexing as follows: (1) Blank sample – 200 ML of PBS; (2) Isotype sample – 50 ML of PBS supplemented with 0.2 Mg of the appropriate isotype; (3) Sorting sample – 50 ML of PBS supplemented with 0.25 Mg of the appropriate FACS-antibody targeting AP (see Subheading 2.2 and Note 7). 12. Incubate for 30 min at room temperature in the dark. 13. Spin down for 5 min at 300 × g at room temperature, remove the supernatant, and wash the cells in 200 ML of PBS. 14. Repeat the previous step and proceed to sort MABs as AP-positive cell fraction, according to the fluorescent dye and the FACS machine used. Collect the MABs in sterile FACSsuitable polypropylene tubes, containing 500 ML of DMEM20 (see Note 8). 15. Once the sorting process is completed, transfer the sorted cell suspension into a new 15 mL tube, spin down for 5 min at 300 × g at room temperature, remove the supernatant, gently resuspend

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the cells in 2 mL of DMEM-20, and plate onto a new 3.5 cm collagen-coated dish. 16. Culture and expand MABs on collagen-coated plastic in DMEM-20 at 37°C in a 5% CO2, 5% O2 humidified incubator. Cell passages are recommended upon 80–85% confluence at a 1:4 ratio. Cell batches can be stored in liquid nitrogen according to long-term storage procedures (see Subheading 3.9). MABs retain their proliferation/differentiation properties for about 20–30 passages (see Table 1). During later passages, MABs generally undergo senescence or loss-of-potency effects. 3.2. FACS-Isolation of Adult Human MABs

1. Follow the same procedure as for FACS-isolation of murine MABs (see above), replacing DMEM-20 medium with MEGA-5 medium. 2. Collagen-coated plastic surfaces are required during the isolation, but are not necessary during subsequent expansion steps. 3. Expand human MABs in MEGA-5 medium in a 5% CO2, 5% O2 humidified incubator, splitting 1:2 upon 80–85% confluence. Human MABs generally retain their proliferation/differentiation features for ~20 passages (see Fig. 2 and Table 1). During later passages, human MABs usually undergo extensive senescence and apoptosis.

3.3. Coculture with C2C12

1. Expand murine C2C12 myoblasts (ATCC) in DMEM-10 medium at 37°C in a 5% CO2, 5% O2 humidified incubator, splitting 1:5 upon 70% confluence. Change DMEM-10 medium daily and avoid myotube formation. 2. At day 0 of differentiation, start the cocultures seeding together 2 × 104 C2C12 myoblasts and 104 murine or 4 × 104 human MABs per 3.5 cm collagen-coated Petri dish. Incubate at 37°C with DMEM-20 medium in case of murine MABs or with MEGA-5 medium in case of human MABs. 3. After 24 h, remove medium, wash with PBS, add DM medium, and incubate. 4. Refresh DM medium every 2–3 days, until appearance of myotubes (usually after approximately 5–10 days) and proceed to analyses (see Note 9).

3.4. Spontaneous Differentiation

1. Expand murine or human MABs at 37°C in a 5% CO2, 5% O2 humidified incubator. 2. At day 0 of differentiation, seed 105 murine MABs or 2 × 105 human MABs per 3.5 cm collagen-coated Petri dish. Incubate at 37°C with DMEM-20 medium in case of murine MABs or with MEGA-5 medium in case of human MABs.

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Fig. 2. Main proliferation/differentiation features of human MABs. During proliferation, human MABs appear like spindleshape cells, expand for 20 passages in approximately 60 days and express AP, PdgfrA, PdgfrB, and NG2 at a detectable protein level (top). In case of spontaneous differentiation, human MABs start to express Myogenin (Myog, center, left ) and generate myosin heavy chain (MyHC) positive myotubes (center, right). Human MABs, upon application of appropriate protocols and media, can differentiate toward other mesodermal lineages, such as smooth muscle cells, osteocytes, and adipocytes (bottom). Bars: 100 Mm.

3. After 24 h, remove medium, wash with PBS, add DM medium and incubate. 4. Refresh DM medium every 2–3 days, until appearance of myotubes (only in case of human MABs, usually after approximately 10–12 days) and proceed to analyses (see Note 10). To enhance myotube formation, MABs can be cultured on Matrigel coated Petri dishes (15).

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1. Expand murine or human MABs at 37°C in a 5% CO2, 5% O2 humidified incubator. 2. At day 0 of differentiation, seed 2 × 105 murine MABs or 3 × 105 human MABs per 3.5 cm Petri dish. Incubate at 37°C with DMEM-20 medium in case of murine MABs or with MEGA-5 medium in case of human MABs. 3. After 24 h, remove medium, wash with PBS, add SMM medium and incubate for 5–7 days, refreshing the medium every 2 days. Proceed to analyses.

3.6. Osteogenic Differentiation

1. Expand murine or human MABs in 3.5 cm Petri dishes at 37°C in a 5% CO2, 5% O2 humidified incubator. 2. Upon 100% confluence, remove medium, wash with PBS, add OM medium and incubate. 3. Refresh OM medium every 4 days for 2–3 weeks and proceed to analyses (see Note 11).

3.7. Adipogenic Differentiation

1. Expand murine or human MABs in 3.5 cm Petri dishes at 37°C in a 5% CO2, 5% O2 humidified incubator. 2. Upon 100% confluence, remove medium, wash with PBS, add AM medium and incubate. 3. After 72 h, remove AM medium, wash with PBS, and add AMM medium. Incubate for 72 h. Alternate AM with AMM (3 days/each) two times more. 4. Refresh the AMM medium and incubate for additional 6 days. Proceed to analyses (see Note 12).

3.8. Collagen-Coating

1. Dissolve 100 mg calf skin collagen in 20 mL glacial acetic acid overnight at room temperature while stirring. 2. Carefully mix and add the acid collagen solution to 80 mL culture-grade water. Filter through a 0.22 Mm membrane to ensure sterility. Store at 4°C for up to 16 weeks. 3. To coat a dish, add collagen solution until the bottom is homogeneously covered. Incubate 5 min at room temperature, remove the collagen solution, and dry the dish out. Incubate the dish at 37°C overnight in a sterile oven. 4. After 24 h, wash the surface at least three times with PBS. Before seeding cells, ensure the correct pH by covering the bottom with a RedPhenol-containing medium. If the medium tones, wash again with PBS.

3.9. Long-Term Storage of Cell Batches

1. After expansion, remove medium, wash with PBS, cover the cell layer with a proper amount of trypsin. 2. Incubate for 5 min at 37°C in a 5% CO2, 5% O2 humidified incubator. Add the same amount of DMEM-20 for murine

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MABs or MEGA-5 for human MABs and carefully collect the cells in a 15 mL tube. Count the viable cells and spin down for 5 min at 300 × g at room temperature. 3. Resuspend in a suitable amount of FM medium (1 mL/2 × 106 cells) and pipet 1 mL of cell suspension/cryovial. 4. Incubate the cryovials in isopropanol-containing cryobox overnight at −80°C. After 24 h, transfer the vials into −150°C freezers or liquid N2 tanks for long-term storage.

4. Notes 1. Adult MABs can be isolated starting from 7-day-old mice. Before this age, it is possible to eventually isolate fetal progenitors. In juvenile mice, such as 2- or 4-week-old, MABs yields are slightly poorer and older mice are expected to yield fewer MABs. 2. Prior to use, ensure a correct removal of any acid traces by means of several PBS washes, in case of acid collagen solution. After eventual washes, dry out the bottom of the dishes, to promote a rapid attachment of the biopsies. 3. It is highly recommended to incubate the muscle fragments plated in 3.5 cm Petri dish in a sterile humid chamber. This can be generated by a 15 cm Petri dish containing uncovered 3.5 cm Petri dish filled with PBS. Given that the isolation may take up to 7 days, check the PBS level and eventually rinse it when necessary. 4. Low-oxygen (5%) and stable humidity/temperature conditions are critical steps during both isolation and expansion of MABs. 5. Floating fragments should be removed, because of possible bacterial contamination. 6. Usually, fibroblasts migrate first from the fragment and after 48–72 h, on top of them, MABs should start to spread out. MABs initially look round and small and then attach to the collagen as spindle-shaped cells. 7. Our current protocol is optimized on the antibodies listed on Subheading 2.2. In case of using different antibodies and isotypes, quantities must be scaled to the manufacturer’s protocol. 8. Depending on FACS-sorting sterility conditions in use, it may be necessary to supplement the collecting-DMEM-20 with 5% penicillin, 5% streptomycin, 0.5% gentamicin, and the subsequent plating-DMEM-20 with 0.1% gentamicin, to avoid bac-

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terial contamination. Remove the medium 24 h after plating, wash with PBS, and rinse with 2 mL of fresh DMEM-20. 9. MABs myogenicity should be evaluated as fusion index (percentage of myotube nuclei/total nuclei). Human MABs nuclei can be easily distinguished from C2C12 nuclei thanks to lamin A/C species specific antibody (Novocastra), which marks human nuclei and does not react with mouse cell lines. In case of murine MABs, we recommend to mark MABs by means of nuclear tracers, such as nuclear lacZ or GFP. 10. Only human MABs produce some myotubes in case of spontaneous differentiation, whereas murine MABs generally differentiate into smooth muscle cells. 11. We suggest Alizarin Red staining of calcium deposits, to test eventual osteogenic induction. Alkaline Phosphatase staining is not suitable, given that proliferating MABs are per se AP+. 12. We suggest Oil Red O staining of lipid-containing vacuoles. If the induction is not sufficient, repeat the AM/AMM alternant up to five times.

Acknowledgments This work was supported by: FWO-Odysseus Program n. G.0907.08; Research Council of the University of Leuven n. OT/09/053; the Nash Avery Stem Cell Research Wicka Fund, University of Minnesota; CARE-MI n. 242038 FP7-EC grant; the Italian Ministry of University and Scientific Research grant n. 2005067555_003, PRIN 2006–08, CARIPLO Foundation 2007.5639 and 2005–2008. References 1. Mauro A (1961) Satellite cell of skeletal muscle fibers. J Biophys Biochem Cytol 9: 493–495 2. Anversa P, Leri A, and Kajstura J (2006) Cardiac regeneration. J Am Coll Cardiol 47:1769–1776 3. Galvez BG, Covarello D, Tolorenzi R, Brunelli S, Dellavalle A, Crippa S, Mohammed SA, Scialla L, Cuccovillo I, Molla F, Staszewsky L, Maisano F, Sampaolesi M, Latini R, Cossu G (2009) Human cardiac mesoangioblasts isolated from hypertrophic cardiomyopathies are greatly reduced in proliferation and differentiation potency. Cardiovasc Res 83:707–716 4. Galvez BG, Sampaolesi M, Barbuti A, Crespi A, Covarello D, Brunelli S, Dellavalle A, Crippa S, Balconi G, Cuccovillo I, Molla F, Staszewsky L, Latini R, Difrancesco D, Cossu G (2008)

Cardiac mesoangioblasts are committed, selfrenewable progenitors, associated with small vessels of juvenile mouse ventricle. Cell Death Differ 15:1417–1428 5. Beltrami AP, Barlucchi L, Torella D, Baker M, Limana F, Chimenti S, Kasahara H, Rota M, Musso E, Urbanek K, Leri A, Kajstura J, NadalGinard B, Anversa P (2003) Adult cardiac stem cells are multipotent and support myocardial regeneration. Cell 114:763–776 6. Nadal-Ginard B, Anversa P, Kajstura J, Leri A (2005) Cardiac stem cells and myocardial regeneration. Novartis Found Symp 265: 142–154; discussion 155–147, 204–111 7. Quattrocelli M, Cassano M, Crippa S, Perini I, Sampaolesi, M (2010) Cell therapy strategies and improvements for muscular dystrophy. Cell Death Differ 17:1222–1229

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8. Sampaolesi M, Blot S, D’Antona G, Granger N, Tonlorenzi R, Innocenzi A, Mognol P, Thibaud JL, Galvez BG, Barthelemy I, Perani L, Mantero S, Guttinger M, Pansarasa O, Rinaldi C, Cusella De Angelis MG, Torrente Y, Bordignon C, Bottinelli R, and Cossu G (2006) Mesoangioblast stem cells ameliorate muscle function in dystrophic dogs. Nature 444:574–579 9. Sampaolesi M, Torrente Y, Innocenzi A, Tonlorenzi R, D’Antona G, Pellegrino MA, Barresi R, Bresolin N, De Angelis MG, Campbell KP, Bottinelli R, Cossu G (2003) Cell therapy of alpha-sarcoglycan null dystrophic mice through intra-arterial delivery of mesoangioblasts. Science 301:487–492 10. Cerletti M, Jurga S, Witczak CA, Hirshman MF, Shadrach JL, Goodyear LJ, Wagers AJ (2008) Highly efficient, functional engraftment of skeletal muscle stem cells in dystrophic muscles. Cell 134:37–47 11. Crisan M, Yap S, Casteilla L, Chen CW, Corselli M, Park TS, Andriolo G, Sun B, Zheng B, Zhang L, Norotte C, Teng PN, Traas J, Schugar R, Deasy BM, Badylak S, Buhring HJ, Giacobino JP, Lazzari L, Huard J, Péault B (2008) A perivascular origin for mesenchymal

stem cells in multiple human organs. Cell Stem Cell 3:301–313 12. Montarras D, Morgan J, Collins C, Relaix F, Zaffran S, Cumano A, Partridge T, Buckingham M (2005) Direct isolation of satellite cells for skeletal muscle regeneration. Science 309: 2064–2067 13. Zheng B, Cao B, Crisan M, Sun B, Li G, Logar A, Yap S, Pollett JB, Drowley L, Cassino T, Gharaibeh B, Deasy BM, Huard J, Péault B (2007) Prospective identification of myogenic endothelial cells in human skeletal muscle. Nat Biotechnol 25:1025–1034 14. Dellavalle A, Sampaolesi M, Tonlorenzi R, Tagliafico E, Sacchetti B, Perani L, Innocenzi A, Galvez BG, Messina G, Morosetti R, Li S, Belicchi M, Peretti G, Chamberlain JS, Wright WE, Torrent Y, Ferrari S, Bianco P, Cossu G (2007) Pericytes of human skeletal muscle are myogenic precursors distinct from satellite cells. Nat Cell Biol 9:255–267 15. Tonlorenzi R, Dellavalle A, Schnapp E, Cossu G, Sampaolesi M (2007) Isolation and characterization of mesoangioblasts from mouse, dog, and human tissues. Curr Protoc Stem Cell Biol Chapter 2:Unit 2B 1

Chapter 5 Direct Electrical Stimulation of Myogenic Cultures for Analysis of Muscle Fiber Type Control Eric J. Cavanaugh, Jennifer R. Crew, and Joseph X. DiMario Abstract Secondary skeletal muscle fiber phenotype is dependent upon depolarization from motor neuron innervation. To study the effects of depolarization on muscle fiber type development, several in vivo and in vitro model systems exist. We have developed a relatively simple-to-use in vitro model system in which differentiated muscle cells are directly electrically stimulated at precise frequencies. This allows for single cell analysis as well as biochemical and molecular analyses of the mechanisms that control skeletal muscle phenotype. Key words: Stimulation, Fiber type, Myosin heavy chain, Myotube, Cell culture

1. Introduction Skeletal muscle fibers can be broadly classified into two classes, “slow” twitch fibers and “fast” twitch fibers, which are based on that particular fiber’s metabolic and contractile properties (1). These two classifications can be subdivided further based upon the predominant isoform of myosin heavy chain (MyHC) present within the fiber. Mammalian muscles can consist of fast MyHCIIb, MyHCIId/x, MyHCIIa, and slow MyHCI (2). In contrast, avian muscles typically contain fast MyHC isoforms. It is the presence of slow MyHC isoforms that distinguishes fast and slow fiber types, providing a less complex repertoire of fiber types relative to mammalian species (3).

Joseph X. DiMario (ed.), Myogenesis: Methods and Protocols, Methods in Molecular Biology, vol. 798, DOI 10.1007/978-1-61779-343-1_5, © Springer Science+Business Media, LLC 2012

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The differentiation and maintenance of fiber type variation among secondary or fetal skeletal muscle fibers is influenced by innervation via complex signaling and molecular pathways (4–8). Physical denervation of the muscle via sciatic nerve ligation and functional denervation via administration of curare induce fiber type switching from a slow to fast fiber type (9, 10). Crossreinnervation of muscles, in which nerves that normally innervated fast muscle were transplanted to innervate slow muscle, indicated that innervation can alter muscle fiber type by regulating fast vs. slow MyHC gene expression. In addition, chronic stimulation in vivo using varying stimulation frequencies elicited fiber type changes based on the stimulation frequency (11). Interestingly, altered innervation was not typically sufficient to completely convert fiber type in these muscles indicating that intrinsic differences among muscle fiber types restricted adaptive ranges and patterns of gene expression in response to external stimuli (11–13). In addition to the in vivo experimental model systems above, a number of in vitro experimental model systems have been devised to investigate the mechanisms of fiber type regulation and corresponding patterns of contractile and metabolic gene expression. Coculture of spinal cord explants with myotubes is one method to study the effects of innervation in vitro (1, 9, 14). The explants contain functional motor neurons that innervate neighboring myotubes, providing a model system amenable to single muscle fiber analysis. However, there are some limitations to in vitro coculture systems. The numbers of innervated myotubes are restricted to regions near the cocultured explants. Therefore, the number of innervated myotubes is often limiting with respect to biochemical and molecular analyses. In addition, the possibility of a trophic effect(s) due to the presence of cocultured cells, rather than direct functional innervation, is a potentially complicating factor in the experimental design. Lastly, labor intensive, careful dissection of spinal cord tissue is required to avoid contamination of the cocultures with other cell types. Direct electrical stimulation of myotubes in vitro mimics innervation of muscle in vivo (15–18). Experimentally, direct stimulation circumvents some of the drawbacks of spinal cord explants. The procedure ensures that all of the myotubes within the culture are treated equally. More cells are available for biochemical and molecular analyses. There is no risk of confounding interpretation of experimental results due to the presence other cells types. Lastly, the direct electrical stimulation procedure is much less labor intensive. This makes using direct electrical stimulation ideal for the evaluation of mechanisms that regulate MyHC gene expression or other molecular mechanisms that are regulated by electrical activity. We have used direct electrical stimulation of avian muscle fibers to investigate both extrinsic and intrinsic control of MyHC

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gene expression and muscle fiber type identity. Direct in vitro stimulation of myotubes derived from fetal myoblasts of specific fast vs. fast/slow muscles yields fiber type differences that recapitulate development of muscle-specific, distinct fiber types in vivo (see Fig 1a) (15). The regulation of slow MyHC gene expression in stimulated myotubes is dependent on both the frequency of stimulation and the origin of the myoblasts that differentiate into the stimulated myotubes (see Fig 1b). In this protocol, we describe the isolation of myoblasts, direct electrical stimulation of differentiated cultures, and subsequent immunostaining of myotubes for fast and slow MyHC isoforms.

Fig. 1. Immunodetection of fast myosin heavy chain (MyHC) isoforms and slow MyHC2 in directly stimulated myotubes. (a) Embryonic day 13 chick pectoralis major (PM) and medial adductor (MA) muscles were isolated, and myoblasts from each were cultured until differentiated myotubes formed. Cultures were then stimulated (+Stim) with 50 V at 10 Hz for 7 days or remained unstimulated (−Stim). Myotubes were immunostained for fast MyHC and slow MyHC2 with F59 and S58 monoclonal antibodies, respectively. (b) Whole cell protein extracts from unstimulated and stimulated PM and MA myotube cultures were western blotted. Fast MyHC and slow MyHC2 were detected with F59 and S58 monoclonal antibodies, respectively. Cultures were stimulated for 7 days with the indicated stimulation parameters. Stimulation with 50 V at 10 Hz elicited the more slow MyHC2 gene expression (reprinted from Crew et al. (15) with permission from Elsevier).

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2. Materials 2.1. Myoblast Isolation and Myotube Stimulation

1. Collagen solution: Add 0.075 g Bovine Tendon Collagen (Worthington) to 100 mL ddH2O. Allow the collagen to reconstitute in the solution for 30 min with stirring. Autoclave the collagen solution. Filter the autoclaved collagen solution through Whatman #1 paper. Autoclave the collagen solution again. Store at 4°C. 2. Collagen coated slides: In a laminar flow hood, add 1.25 mL of collagen solution to the cell culture treated side of a coverslip (Nunc Thermanox 174942) (see Note 1). Allow the collagen solution to dry onto the coverslip overnight. 3. Chick Embryo Extract: Sterilize eggs with 70% ethanol. In a laminar flow hood, extract embryonic day 12–13 embryos from eggs. Remove the heads from the embryos. Wash the embryos twice with cold Hank’s Balanced Salt Solution (HBSS). Pass the embryos through a 60 cc syringe barrel and into a sterile graduated cylinder. Add an equal amount of HBSS to the embryos and stir at 4°C for 1 h. Centrifuge the extract at 12,000 × g for 10 min at 4°C. The extract can be stored at −20°C in aliquots. After thawing aliquots for use, centrifuge the extract at 1,000 × g for 5 min at room temperature. Collect the supernatant and filter through a 1.6 Pm and then a 0.8 Pm syringe filter. 4. Cell Culture Medium: In a graduated 100 mL cylinder combine 82 mL of F-10 base medium, 10 mL donor equine serum, 5 mL of filtered chicken embryo extract, 1 mL of 100× Penicillin/ Streptomycin/Fungizone (GIBCO 15240), 1 mL of 132 mM CaCl2, and 1 mL of 200 mM L-glutamine. Filter medium through a 0.22 Pm filter to sterilize. Store at 4°C. 5. 70% Ethanol. 6. Hank’s Balanced Salt Solution. 7. 35 mm Cell culture dish. 8. 10 cm Cell culture dish. 9. 0.125% Trypsin. 10. 15 mL Conical tubes. 11. Sterile microscissors. 12. 25 mm Stainless steel Swinney filter holder and filter (Fisher Scientific 30-025-00). 13. 13 mm Stainless steel Swinney filter holder and filter (Fisher Scientific 30-012-00). 14. 80 Pm Nitex filter (Sefar 03-80/37). 15. Sterile 10 mL syringes.

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16. Stimulation plates: Using the four well rectangular multidish with lid (Nunc 176597) drill a hole perpendicular to the wells near the edge of the dish penetrating each of the three walls dividing the wells (see Note 2), with a 1/16 in. drill bit. Repeat this procedure with the other side so that there are two holes on one side of the plate. Thread the platinum wires through the holes so there is a platinum wire on each side of the dish. Seal the remainder of the hole with Aquarium sealant to make a waterproof seal (see Note 3). Let the sealant dry overnight. 17. HSE-HA Stimulator CS Type 223 (Hugo Saks Electronik). 2.2. Immunocytochemistry

1. Phosphate buffered saline (PBS), 10×: Dissolve 87.7 g NaCl, 2.6 g Na2PO4·H2O, 22.5 g Na2HPO4·7H2O in 750 mL of ddH2O. Adjust volume to 1 L with ddH2O. 2. 100% Methanol. 3. Blocking Buffer: Add 5 mL of horse serum to 75 mL of 1× PBS. Add 2 g of bovine serum albumin. Adjust volume to 100 mL with PBS. 4. Primary antibodies: S58 is a monoclonal anti-slow MyHC2 IgA, and F59 is a monoclonal anti-fast MyHC IgG. The specificities of these antibodies have been previously described (19). Diluted monoclonal supernatants 1:10 in blocking buffer. 5. Secondary antibodies: Anti-mouse IgG TRITC and anti-mouse IgA FITC (Southern Biotech) diluted 1:100 in blocking buffer. 6. 1 mM DAPI (4c,6-diamidino-2-phenylindole): Weigh 3.5 g of DAPI and dissolve in 7.5 mL of PBS. Adjust volume to 10 mL with PBS. 7. 2.5% DABCO (1,4-diazabicyclo[2.2.2]octane): Weigh 0.25 g of DABCO and dissolve in 7.5 mL of a 90% glycerol, 10% PBS solution. Adjust volume to 10 mL with the glycerol/PBS solution.

3. Methods All steps are performed in a sterile laminar flow hood, and all components are sterile unless otherwise noted. 3.1. Myoblast Isolation

1. Place 500 PL HBSS into a 35 mm plate. 2. Sterilize eggs with 70% ethanol. Remove embryo from egg and decapitate the embryo. Place the embryo into a 10 cm plate and excise the tissue of interest. Place the tissue into the 35 mm plate with 500 PL of HBSS. 3. Mince the tissue by cutting it with the microscissors until only fine remnants are left (see Note 4). Place tissue into 0.125%

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trypsin. Incubate in a 37°C water bath for 40 min. Every 5 min vigorously shake the tissue suspension (see Note 5). 4. Add an equal volume of HBSS to the suspension to dilute the trypsin/cell suspension. 5. Triturate the suspension until undigested tissue is well dispersed. 6. Filter the suspension through a 25 mm Swinney filter to get rid of crude debris. Take filtered suspension and filter through a 13 mm Swinney filter with an 80 Pm Nitex filter insert. 7. Centrifuge the suspension at 1,000 × g for 5 min (see Note 6). 8. Remove the supernatant and resuspend the cell pellet in 10 mL of prewarmed cell culture medium. Place the suspension in a 10 cm cell culture dish and incubate in a 37°C incubator for 40 min. 9. Place the cell suspension (approximately 3 × 106 cells) onto collagen coated coverslips in a four well rectangular multidish (see Note 7). Incubate the cells at 37°C, replacing the medium every other day as needed. 3.2. Electrical Stimulation

1. Once myoblasts have begun to form myotubes (2–3 days), they are ready for stimulation. Remove the myotube covered coverslip and place coverslips into the stimulation dish. 2. Add 8 mL of prewarmed cell culture media to each well. 3. Place stimulation dish into a 37°C incubator. 4. Hook electrodes to the stimulation plate. 5. Stimulate cells with 50 V at a frequency of 10 Hz. The biphasic pulse width is 2 ms with a pulse gap of 100 Ps and a delay time of 2 ms. Carry out the stimulation regime for 30 min followed by a 15 min rest period for the duration of the experiment. 6. Change medium every day until myotubes are ready for future experiments. For results shown in Fig. 1, cultures were stimulated for 7 days.

3.3. Immunocytochemistry

1. Wash the myotubes with 8 mL of PBS 3 times for 3 min each. 2. Fix the myotubes with 5 mL of 100% methanol for 10 min. 3. Wash the myotubes with 8 mL of PBS 3 times for 3 min each. 4. Block the myotubes with 8 mL of blocking buffer for 1 h. 5. Add 4 mL of diluted primary antibodies to the myotubes for 1 h. 6. Wash the myotubes with 8 mL of PBS 3 times for 3 min each. 7. Add 4 mL of secondary antibodies to the myotubes for 1 h. 8. Add 4 mL of DAPI stain diluted 1:1,000 in PBS for 10 min. 9. Wash the myotubes with 8 mL of PBS 3 times for 3 min each.

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10. Add 1–2 drops of 2.5% DABCO to the myotubes. 11. Place a 24 × 50 mm cover glass on top of the myotubes.

4. Notes 1. To ensure complete coverage of the coverslip with the collagen solution, use pipette tip to push the collagen to the edges of the coverslip. If collagen solution goes beyond the edges of the coverslip, the coverslip will stick to the bottom of the plate. 2. If the hole is drilled too close to the edge of the multidish, it will crack the multidish. Also keep the hole near the bottom of the plate. The drill bit is generally not long enough to make all of the holes at once. For the last plate wall, drill the hole at the shallowest angle possible. 3. The platinum wire should be sticking out the hole so that an electrode can be attached to it. 4. When the pieces of the tissue are small enough to smoothly pass through a 1 mL pipette tip, then they have been minced enough. 5. As the incubation goes on the solution will become viscous, this is normal. 6. If there is no pellet, but a wispy web inside of the suspension, then add more HBSS to the suspension and centrifuge again. 7. Once the cells have been added to the coverslip, bubbles may form underneath the coverslip. To get rid of bubbles, tap the coverslip with a 1 mL pipette tip until the bubbles have been removed. If the bubbles are left there, then the cells may not adhere to the coverslip, because it is floating in the well instead of lying at the bottom of the well.

Acknowledgment This work was supported by NIH grant ARO45939. References 1. Zierath JR, and Hawley JA (2004) Skeletal muscle fiber type: Influence on contractile and metabolic properties. PLoS Biol. 2(10):e348 2. Pette D, and Staron RS (2001) Transitions of muscle fiber phenotypic profiles. Histochem Cell Biol. 115:359–372 3. Pette D, and Vrbova G (1999) What does chronic electrical stimulation teach us about muscle plasticity? Muscle Nerve 22:666–677

4. Condon K. Silbersein L, Blau HM, Thompson WJ (1990) Differentiation of fiber types in aneural musculature of the prenatal rat hindlimb. Dev. Biol. 138:275–295 5. Jordan T, Li J, Jiang H, and DiMario JX (2003) Repression of slow myosin heavy chain 2 gene expression in fast skeletal muscle fibers by muscarinic acetylcholine receptor and GDq signaling. J. Cell Biol. 162:843–850

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6. Jiang H, Jordan T, Li J, Li H and DiMario JX (2004) Innervation-dependent and fiber type specific transcriptional regulation of the slow myosin heavy chain 2 promoter in avian skeletal muscle fibers. Dev. Dyn. 231:292–302 7. Jordan T, Jiang H, Li H and DiMario JX (2005) Regulation of skeletal muscle fiber type and slow myosin heavy chain 2 gene expression by inositol trisphosphate receptor 1. J. Cell Sci. 118:2295–2302 8. Tothova T, Blaauw B, Pallafacchina G, Rüdiger R, Argentini C, Reggiani C, and Sciaffino S (2006) NFATc1 nucleocytoplasmic shuttling is controlled by nerve activity in skeletal muscle. J. Cell Sci. 119:1604–1611 9. DiMario JX, and Stockdale FE (1997) Both myoblast lineage and innervation determine fiber type and are required for expression of the slow myosin heavy chain 2 gene. Dev. Biol. 188:167–180 10. DiMario JX, and Funk PE (1999) Protein Kinase C activity regulates Slow Myosin Heavy Chain 2 gene expression in slow lineage skeletal muscle fibers. Dev. Dyn. 216:177–189 11. Ausoni S, Gorza L, Schiaffino S, Gundersen K, Lømo T (1990) Expression of myosin heavy chain isoforms in stimulated fast and slow rat muscles. J. Neurosci. 10:153–160 12. Gauthier GF, Burke RE, Lowey S, and Hobbs AW (1983) Myosin Isozymes in normal and cross-reinnervated cat skeletal muscle fibers. J. Cell Biol. 97:756–771

13. Hoh JFY and Hughes S (1991) Expression of superfast myosin in aneural regenerates of cat jaw muscle. Muscle Nerve 14:316–325 14. Wagner S, Dorchies OM, Stoeckel H, Warter JM, Poindron P, Takeda K. (2003) Functional maturation of nicotinic acetylcholine receptors as an indicator of murine muscular differentiation in a new nerve-muscle co-culture system. Pflugers Arch. 447:14–22 15. Crew JR, Falzari K, and DiMario JX (2010) Muscle fiber type specific induction of slow myosin heavy chain 2 gene expression by electrical stimulation Exp. Cell Res. 316:1039–1049 16. Thelen MH, Simonides WS, and van Hardeveld C (1997) Electrical stimulation of C2C12 myotubes induces contraction and represses thyroidhormone-dependent transcription of the fast-type sarcoplasmic-reticulum Ca2+-ATPase gene. Biochem. J. 321:845–848 17. Kubis HP, Scheibe RJ, Meissner JD, Hornung G, and Gros G (2002) Fast-to-slow transformation and nuclear import/export kinetics of the transcription factor NFATc1 during electrostimulation of rabbit muscle cells in culture. J. Physiol. 541:835–847 18. Elsner P, Grunner N, and Quistorff (2003) Effects of electrostimulation on glycogenolysis in cultured rat myotubes Pflugers Arch. 447:356–362 19. M.T. Crow, F.E. Stockdale (1986) Myosin expression and specialization among the earliest muscle fibers of the developing avian limb. Dev. Biol. 113:238–254

Chapter 6 Single Muscle-Fiber Isolation and Culture for Cellular, Molecular, Pharmacological, and Evolutionary Studies Judy E. Anderson, Ashley C. Wozniak, and Wataru Mizunoya Abstract The technique of single muscle-fiber cultures has already proven valuable in extending knowledge of myogenesis, stem cell heterogeneity, the stem cell niche in skeletal muscle, and satellite cell activation. This report reviews the background of the model and applications, and details the procedures of muscle dissection, fiber digestion and isolation, cleaning the fiber preparation, plating fibers, and extensions of the technique for studying activation from stable quiescence of satellite cells, mRNA expression by in situ hybridization and regulation of satellite cell activation in zebrafish muscle by nitric oxide, hepatocyte growth factor. Key words: FDB, Dissection, Collagenase, Single fibers, Satellite cells, Activation, Zebrafish, Electron microscopy, In situ hybridization, HGF, Nitric oxide, Drug screening

1. Introduction The study of satellite cell activation from quiescence, and more recently the study of myogenic signaling pathways and transcriptional profile, satellite-stem cell lineages and heterogeneity, asymmetric division, proliferation capacity and senescence, chemotaxis and migration capacity, the scope of differentiation and fiber type specificity, and the influence of cell-cycle, differentiation state, and components of the environmental niche (extracellular matrix, growth factors, etc.) have used many model systems. These include isolated satellite cell cultures (1–4), single muscle fibers (5–12) and in vivo studies in various experimental regeneration protocols (5–8). Each model affords the capability of studying satellite cells to a different degree of complexity. However, the maintenance of satellite cell quiescence during isolation procedures

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is essential to address the timing or nature of satellite cell withdrawal from G0 and entry to cycling during activation. This is accomplished in the cell culture model by isolating satellite cells from older animals, since those cells have a longer latent period of quiescence that is maintained after an activating stimulus, before they make DNA, compared with satellite cells from younger animals (1, 2). The use of single muscle fibers in culture is an alternative approach to study activation and subsequent events in early regeneration. The method allows for the isolation of single intact fibers with satellite cells in their characteristic position, still in a quiescent state beneath the external lamina, and not incorporating labeled nucleotides into new DNA under control conditions. Moreover, fibers in culture are recognized to be mature muscle in contrast to myotubes differentiated from cultures of dispersed muscle precursors in vitro (9). This method therefore usefully models many in vivo conditions of the satellite cell micro-environment while maintaining interactions between fibers and satellite cells (4, 10–13). The single fiber culture model involves isolating myofibers, usually from the flexor digitorum brevis (FDB) muscle of mice or rats (12, 14), although fibers can be prepared from many other muscles such as extensor digitorum longus and soleus (15, 16), tibialis anterior (17), and diaphragm (Anderson, unpublished). The process of isolating fibers should be able to maintain satellite cells in the normal, quiescent state, and in their characteristic position between the basal lamina and sarcolemma of skeletal muscle fibers. However, protocols for methods of isolation, digestion, plating, and culture vary among laboratories, and variations, particularly those from agitation during digestion, may cause inadvertent activation of satellite cells that may not be revealed by the markers of function, expression, or position that are selected for study. Other protocols, particularly use of sedimentation columns (see later) will tangle and damage fibers that are isolated from muscles with long fibers (17). The fiber-isolation method was pioneered by Bekoff and Betz (18) and later established by Bischoff (12). It allows for the isolation of intact, single, and entire muscle fibers with their satellite cells still attached or “resident” and in the quiescent state if prepared by careful dissection to minimize muscle trauma or stretching. Muscles are isolated and digested in collagenase-I, and then fibers are separated from one another using gentle trituration with a wide bore pipette. Fibers are separated from debris and dead fibers through gravity sedimentation. The clean fiber suspension is plated on dishes using a variety of adhesives (collagen, laminin, or a matrix containing growth factors, such as Matrigel). Over time, the basic method has changed and evolved in particular laboratories in relation to specific questions or hypotheses under investigation. One variation used extensively was established by Rosenblatt et al. (17). This isolation is not as concerned with keeping satellite cells quiescent

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as it employs collagenase digestion in a shaker. This shortens the time for digestion although the constant motion of the liquid activates satellite cells (19). In this method, gravity sedimentation is omitted and fibers are transferred individually to a fresh dish. This is required as the longer fibers (from EDL muscle, e.g.) tangle as they sediment in the columns, and this leads to fiber damage and death. Finally, in vivo studies can provide the most comprehensive picture of satellite cells during activation in situ in muscle (7, 20). However, the effects of treatments directed to satellite cells require careful interpretation, due to the complex contributions from non-muscle tissues, perfusion-dependent changes, constraints of tissue sampling, animal activity, and systemic physiology that each may introduce significant variability to indices under study. Using the single-fiber model, which maintains satellite cells in their normal in vivo (“satellite”) position on intact fibers, together with immunostaining for a wide variety of proteins such as muscle regulatory factors (e.g., myf5) in activated satellite cells, Pax7 or c-met receptor in both quiescent and activated satellite cells, or proliferation-marker proteins, demonstrates the range of applications for the technique (see Note 1). Fiber cultures can be used to screen unknown compounds for their ability to activate satellite cells, where activation is represented as the mean ± standard error (see Fig. 1a) or as a distribution of the proportion of fibers with different numbers of bromodeoxyuridine (BrdU)-labeled satellite cells per fiber (see Fig. 1b). Fiber cultures can also be used to characterize responses to one of the two-known activating molecules, hepatocyte growth factor (HGF, see Fig. 2). Very notably, satellite cells on dead fibers (such as induced

Fig. 1. (a) Counts of mean number of BrdU+ cells per fiber from an experiment on mouse flexor digitorum brevis (FDB) fibers, illustrating differential stimulation of satellite cell (SC) activation induced by two drugs, X and Y (**, p < 0.01, Student’s t-test). (b) A graph of the distribution of BrdU+ satellite cells per fiber (as a proportion of all fibers counted), illustrating information of the population of activation on fibers from the same dataset as in (a) (p < 0.01, Chi-square).

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Fig. 2. (a) Activation (p < 0.01, ANOVA) of satellite cells (SC) by increasing doses of hepatocyte growth factor (HGF) treatment in fiber cultures plated on a rigid substrate coated with collagen. (b) This dose–response curve of HGF-induced activation in stretched fiber cultures (incubated in a FlexCell system) is left-shifted (p < 0.01, Chi-square) toward smaller activating doses.

Fig. 3. Satellite cell activation (p < 0.01, ANOVA) by HGF (hepatocyte growth factor, 20 ng/mL) is prevented by pre-treating fiber cultures with the myotoxin, marcaine (0.025% in DMEM-HS for 5 min to study the role of an intact sarcolemma) (10, 23).

by treatment with the myotoxin, marcaine) will not respond the same way to HGF as those quiescent satellite cells resident on intact (live) fibers (see Fig. 3). One of the methods used to stimulate satellite cell activation in fiber cultures employs the FlexCell culture system. This system delivers a stimulus of cyclical stretching to cells in culture and was pioneered a decade prior to application with fibers (21, 22). FlexCell dishes have a flexible elastomer (silastic) substrate that enables experiments with mechanical-stretching stimuli that are programmable by intensity and frequency (23, 24) (see Note 2). Using this system, stretch was found to activate isolated quiescent satellite cells in culture within 2 h, as judged by significant increases in BrdU incorporation (2). Cells of other origins are also known to proliferate and enter tissue-specific processes as a result of stretching (22, 25).

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Although previous experiments without stretching report a low level of spontaneous activation of satellite cells (10), the frequency distribution of activation that would represent the population response by satellite cells was not reported. In our initial study of stretchactivation, we hypothesized that stretch would activate quiescent satellite cells on single fibers, after identifying appropriate conditions for maintaining the cultures. The in situ hybridization protocol can also be applied to single fiber cultures to enable observations of molecular events (e.g., transcript expression) at the level of individual satellite cells, as shown in studies of c-met mRNA expression and the distribution of quiescent satellite cells on single fibers (23, 26) after modifications to the procedure (see later). Application of the fiber-isolation technique is very effective in studies of satellite cell activation and quiescence as well as examinations of fiber morphology, mRNA and protein expression, and satellite-stem cell behavior in muscles of animals in other taxa in addition to mouse and rat. The approach supports investigations of the conservation of or disparity in the pathways related to activation and quiescence, myogenesis, and environmental niches that are driven by or drive evolution, since skeletal muscle function enables adaptive metabolism, behavior, and locomotion. For example, we perfected the single fiber technique for isolating zebrafish fibers with solutions based on medium specified for fish-cell culture. Figure 4 illustrates a dose–response of zebrafish-fiber satellite cells to isosorbide dinitrate (ISDN), a nitric oxide (NO)-donor compound, and studies comparing activation effects of treatment with ISDN, HGF, and HGF in the presence of neutralizing antibody to c-met.

2. Materials 2.1. Single-Fiber Isolation and Culture

1. Based on the experimental design, calculate the number of wells of fibers that are required to detect a difference in the particular parameter under study (see Note 3). 2. Basal growth medium: Dulbecco’s Minimum Essential Medium (DMEM) for mouse or L-15 for fish fibers containing 20% serum replacement (without growth factors, steroids, etc.), 1% fetal bovine serum, 1% antibiotic/antimycotic, and 0.1% gentamycin. Mix medium the day before and keep in the refrigerator overnight. 3. Proliferation medium (PM): DMEM containing 10% fetal bovine serum, 2% chick embryo extract, 1% antibiotic/antimycotic, and 0.1% gentamycin. At least 100 mL of PM is required to prepare fibers for culture. Mix medium the day before and keep in the refrigerator overnight. Warm the PM to 37°C just before use (see Note 4).

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Fig. 4. (a) Satellite cell activation on zebrafish fibers stimulated by nitric oxide (NO)-donor compound isosorbide dinitrate (ISDN; p < 0.02) and (b) HGF (p < 0.01) represented as mean (±SEM) of the number of BrdU+ satellite cells per fiber. (c) The distribution of activated satellite cells on zebrafish fibers treated in culture for 24 h with ISDN (0.5–2.5 mM) from the same dataset as in (a) (p < 0.01, Chi-square). (d) Activation of satellite cells (mean, SEM) in zebrafish fiber cultures indicates ISDN (2 mM) and HGF (5 ng/mL) stimulate activation above the control level, and in the presence of neutralizing antibody against the c-met receptor (1 Mg/mL anti-c-met or 2.5 Mg antibody/dish). Treatments with different letters (a, b) are significantly different (p < 0.01 ANOVA, p < 0.05 Tukey’s HSD method). (e) The distribution of activated satellite cells on zebrafish fibers treated in culture for 24 h with a ISDN, HGF and anti-c-met plus HGF, from the same dataset as (d) (p < 0.01, Chi-squared).

4. Mix 15 mL of 0.2% collagenase type 1 solution made in DMEM for mouse-fiber preparation or in L-15 for fish-fiber preparation (see Note 5). 5. Acid alcohol fixative: 90% absolute ethanol, 5% glacial acetic acid, 5% H2O.

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6. Tris-buffered saline (TBS): 2.425 g Tris base, 8.765 g NaCl with 1% horse serum (v/v) in 1 L, adjusted to pH 7.4. 7. Assemble and autoclave dissecting instruments in small foil packets, including: two pairs of fine scissors (0.5 cm blades), one pair of very fine spring scissors (8 cm, with 2–5 mm blades), two pairs of sharp forceps (0.3 × 0.25 mm tips), and two to four pairs of very fine (Dumont #5), straight forceps (0.1 × 0.06 mm tips). Check the tips of all forceps to ensure that they meet at the points and have no bends or nicks. 8. Assemble and autoclave glassware in individual foil packets, including four conical-bottom centrifuge tubes for use in gravity sedimentation (conical bottom, so fibers are visible when they settle), 60 mm Petri dishes (three to four, with lids), and small glass beakers (50 mL) for collecting waste solutions. 9. Prepare and autoclave in foil packets, glass spatulas for coating culture dishes with collagen or other substrate solution. Make these by heating the tip of a Pasteur pipette over a flame and bending the tip at 90° angle, approximately 15 mm from the end. Allow to harden for a few seconds and flame-polish the tip end to close it off. 10. Prepare and autoclave 10–15 wide-bore glass pipettes. To make these, use a diamond pencil to score the glass of a Pasteur pipette at an appropriate place along the tip and flame-polish the cut end. The opening should approximate twice the length of muscle fibers to be isolated. 11. Disposables: clean lint-free tissues, pipette tips, pipetters (to dispense aliquots of collagen and fibers), dental wax, and culture dishes (e.g., 35 mm plastic Petri dishes or FlexCell plates). 12. Prepare pipette tips to aliquot fibers. Tips to fit a mechanical pipettor that will dispense 50 Mm volumes, need to be cut so they are wide-bore. For FDB or zebrafish fibers, the tip opening should be approximately 2–2.5 mm in diameter. Cut the tip with a sterile razor blade and flame-heat it to smooth the cut edge (by slightly melting the plastic). 13. Locate a bench for dissection, preferably in a low-traffic area in the laboratory, and wipe down a dissecting microscope and the bench top with alcohol. 14. Collagen: an aseptic solution of type I collagen (2.9–3.5 mg/ mL) prepared from calf skin, bovine skin, rat tail, bovine tendon or comparable tissue, suitable for cell culture. 2.2. Electron Microscopy

1. 1% Osmium tetroxide (OsO4) in double-distilled water. 2. Sorenson’s phosphate buffer (40.5 mL of 0.2 M Na2HPO4·2H2O mixed with 9.5 mL of 0.2 M NaH2PO4·H2O, diluted to 100 mL with double-distilled water and brought to pH 7.4).

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3. Uranyl acetate (8% in water). 4. Graded series of ethanols (70, 95, 100%). 5. Methanol (anhydrous). 6. Propylene oxide. 7. Araldite or other embedding resin. 2.3. In Situ Hybridization

1. Diethylpyrocarbonate (DEPC)-treated water to inactivate RNAse: mix a stock (wear gloves, use a fume hood) by adding 25 mL of DEPC to 225 mL absolute ethanol. Add 10 mL of stock DEPC-treated water to 1 L double-distilled water (final DEPC concentration = 0.1%) in a glass bottle and swirl to mix. Let stand in fume hood overnight, then autoclave. Use fresh, in a fume hood. 2. Acid alcohol fixative (90% absolute ethanol, 5% glacial acetic acid, 5% H2O). 3. Phosphate-buffered saline (PBS): dissolve 8 g NaCl, 0.2 g KCl, 2.68 g Na2HPO4·7H2O, 0.24 g KH2PO4 in 800 mL of DEPCtreated water, pH to 7.4, and bring to a volume of 1 L; autoclave.

3. Methods 3.1. Mouse Muscle Dissection

1. Begin the dissection early in the day, as the procedures of dissection, cleaning, digestion, trituration, and plating can take many hours, depending on the number of muscles required to achieve sufficient fibers for the planned experiment. 2. Euthanize mice with CO2 or an anesthetic plus cervical dislocation. 3. Remove skin to minimize hair contaminating the preparation of fibers. Often skin will pull off the hind feet, leaving a “sock” of skin on each foot. Cut through the skin over the dorsum of the foot, and pull each side of the sock toward the sole of the foot. Separate skin from the underlying tissue for the full length of the plantar surface (bottom) of the foot. In particular, expose the first inter-phalangeal joints in the toes and the heel (see Note 6). 4. Stabilize the leg by pinning through the proximal muscles of the thigh or leg into a layer of dental wax, so that the plantar surface of the foot is facing straight upwards. 5. View the foot under a dissecting microscope and locate the calcaneous (heel bone). The yellow-white band running distally toward the toes is the proximal tendon of the FDB muscle. Grip it with a pair of fine forceps immediately distal to the heel (see Note 7).

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6. With a second set of fine forceps, begin to cut or gently tear the epimysial (outer fascial) covering along one side of the muscle (see Note 8). Once one border of the muscle is clean, clean the other side of the FDB, gently removing the fascia and separating the muscle from adjacent muscles on the medial and lateral sides. The muscle will become separable from the tissues that are deep to (underneath) the FDB. 7. Lift the proximal tendon of the FDB slightly from the muscle bed and put one blade of a second pair of forceps underneath the muscle. This will pierce through any remaining fascia. 8. Gently run this second pair of fine forceps once down the length of the FDB muscle from proximal to distal where the tendons enter the digits (see Note 9). 9. The distal end of the muscle has four small tendons, each running to a toe. Move the fine forceps along to the first interphalangeal joint on each toe (see Note 10). This process should separate the majority of fascia from the plantar surface of the muscle, as well as that on the undissected side. Use fine forceps (two pairs) or forceps and spring scissors to remove all visible fascia (see Note 11). 10. Gently separate the tendons from each other and the remaining fascia by running the forceps gently from distal to proximal, under one tendon at a time from its attachment point on the toe, until it joins the FDB muscle belly proper. After each tendon is isolated from surrounding tissue using this process, and the visible fascia removed, the FDB is ready to remove from the muscle bed (see Note 12). 11. Using dissecting scissors and viewing through a dissecting scope, hold the proximal tendon of FDB just distal to the calcaneous and cut it with spring scissors. Lift the muscle gently without pulling very hard or elongating the muscle, until the FDB is essentially vertical and you can see clearly the four tendons attaching to the bones in the digits. At this point, check to see if there are any large pieces of fascia remaining on the surface of the FDB. If so, remove them. Then cut all tendon attachments at the toe joints (without cutting through muscle fibers). 12. Remove the muscle and place in a 60 mm culture dish filled with proliferation medium (PM). 3.2. Muscle Cleaning

1. Take muscles one at a time to a clean 60 mm glass Petri dish containing approximately 5–6 mL of proliferation medium (PM). Use fine forceps to grip one tendon or a wide-bore pipette for a gentle transfer. 2. Using higher-power magnification on the dissecting microscope, inspect the muscle. Using very fine forceps (with no nicks or bends in the tips), remove any further connective tissue (including small nerve bundles or vessels) that is visible

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at high magnification. Clean between the distal muscle bellies of the FDB by gripping the end of one tendon with the very fine forceps, and then grip the other two tendons (or three if all four tendons were dissected with the muscle) together using a second pair of forceps. Gently begin to pull apart muscle bellies while you observe the muscle without completely separating them (see Note 13). 3.3. Collagenase Digestion

1. Place the cleaned muscle in a fresh 60 mm culture dish filled with DMEM with 0.2% collagenase and cover the dish (see Note 14). 2. Place the dish in a clean incubator (37°C, 5% CO2) for 2.5–3 h (see Note 15). 3. After approximately 2.5 h, remove the dish from the incubator. Remove all the muscles from the collagenase solution using a flamed wide-bore Pasteur pipette, and put them in fresh dishes of PM without collagenase.

3.4. Isolation of Fibers

1. Remove remaining connective tissue; this will start to separate the muscle bellies from each other (see Note 16). 2. Completely separate the muscle bellies from one another by gently pulling the distal tendons apart using very fine forceps. 3. Using a wide-bore pipette gently triturate each belly separately, without introducing bubbles into the medium (see Notes 17 and 18). 4. Triturate only until the muscle belly is nearly completely dissociated, or for 10 min to avoid prolonged agitation that will activate satellite cells. Use the very fine forceps (one in each hand) very carefully to separate bundles of fibers from the connective tissue near the tendons and neurovascular bundles that run among fibers. Repeat this process with all the muscle bellies (see Note 19). 5. Reduce the amount of medium in the dish and pool the fibers isolated from each muscle. Tilt the dish slightly and wait until the fibers sink to the bottom, then pipette medium off the top of the fluid level (see Note 20) to leave 3–4 mL over the fibers. 6. Inspect the preparation of fibers under the microscope. If they are alive, fibers will be translucent and have smooth membranes; if not, they will be cloudy in whole or part, frayed or hypercontracted (see Note 21).

3.5. Gravity Sedimentation

1. After removing damaged fibers and remnants of connective and other tissues, the preparation is ready for further cleaning by gravity sedimentation. Fill a 15 mL glass tube (“column”) with approximately 10 mL of fresh PM.

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2. Transfer the solution containing fibers into the tube using a wide bore pipette. Slowly pull up the entire volume of fluid in the dish and slowly pipette it onto the surface of the medium in the column (do not expel the fluid quickly). Allow the fibers to settle in the column for 20 min; broken fibers, single cells and small bits of connective tissue do not sink as quickly. 3. Once the fibers form a loose pellet at the bottom of the column, pull off and discard the medium in the column until about 2–3 mL remains over the fibers. Very gently, resuspend the pellet in the remaining medium and transfer the fibers to a second centrifuge tube filled with PM. Allow fibers to settle for only 5 min in this column to separate the intact fibers from those larger broken or damaged fibers and other debris (see Note 22). 4. Again, resuspend fibers in 2–3 mL of PM and using a widebore pipette, place them gently in a fresh 60 mm glass culture dish of PM. Look once again at the fibers under the microscope and remove any dead fibers, left over connective tissue and debris. Fibers are now ready to plate, in whichever way the experiment requires. 3.6. Plating Fibers

1. Prepare culture dishes for coating by placing them over ice for 5 min. Coat the dishes with collagen (to minimize stimulating satellite cells to activate) or other substrate. Pipette 80–120 ML of collagen solution into each 35 mm Petri dish or other culture plate and spread evenly across each plate using a glass spatula. 2. Pipette aliquots of the fiber preparation, gently and evenly dispersed in the PM, into each dish, to a maximum volume of 50 ML (see Note 23). 3. Place the dishes in an incubator (37°C, 5% CO2) for 20–25 min. 4. After the collagen has set or formed a gel that holds fibers (see Note 24), remove from the incubator and gently add medium and treatments to each dish before returning to the incubator. 5. After culturing for the appropriate time (e.g., approximately 24 h to detect stimulation of activation using BrdU incorporation into S phase), fiber cultures should be rinsed in PBS, fixed in acid alcohol, and air dried. Dishes can be wrapped or placed in an airtight container, and stored at 4°C for up to 2 weeks in TBS before immunostaining, as reported by numerous authors. Typically immunostaining of satellite cells on fibers requires application of a number of blocking strategies, such as those detailed in the IHCWorld protocol (27), to reduce background and increase detection of the epitope of interest.

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3.7. Electron Microscopy of Fibers

1. Fix fiber cultures for 20 min in acid alcohol, rinse in phosphate buffer (pH 7.4), post-fix in OsO4 and rinse in double-distilled water. 2. Stain with uranyl acetate (60 min), rinse in water, dehydrate in graded ethanols, rinse in anhydrous methanol (10 min) then propylene oxide (10 min), and infiltrate in resin (decreasing the proportion of propylene oxide in three to four steps of 10 min each). Fill the dish with 2 mL of resin and bake according to the resin instructions. 3. Separate resin-embedded fibers from the dish (see Note 25). 4. After separation from the dish, reembed the disc of resin containing fibers in methacrylate or other resin to protect the substrate layer of collagen or other coating and allow examination of the fiber membrane and extracellular matrix with the substratum. 5. Under a dissecting microscope, view the disc of hardened resin and identify fibers for further study; mark them for removal (cutting with a fine hacksaw blade) as small resin blocks for sectioning. 6. Cut blocks from the resin, trim and section for viewing by light microscopy (stain with toluidine blue (23)) or electron microscopy (stain with uranyl acetate and lead citrate or use immunogold labeling to study localization of an epitope of interest).

3.8. In Situ Hybridization of Fibers

1. Rinse fiber cultures with RNAse-free PBS and then fix in acidalcohol for 10 min (see Note 26). 2. Remove fixative and air dry dishes of fibers in a laminar-flow hood for 10–15 min before rinsing 3 times in RNAse-free PBS. Stored in the same solution at 4°C. 3. The remaining steps of the in situ protocol to detect transcripts in individual satellite cells on fibers (or around myonuclei) are carried out as reported, with attention to gentle handling to avoid loss of fibers from the collagen substrate on each dish (26, 28).

3.9. Isolating Fibers from Zebrafish

1. Anesthetize zebrafish (e.g., MS-222) and decapitate in PM prepared with fish medium. Remove internal organs, tail, fins, and skin (see Note 27). 2. Incubate cleaned fish in medium containing 0.2% collagenase at 37°C for 2 h. Incubate without shaking. 3. Pour off the collagenase and slide the fish into a clean 60 mm glass Petri dish of fresh PM. Using a very wide-bore pipette, triturate the whole fish to separate fibers from ribs. 4. Remove debris from PM (including bones, scales, fat droplets, etc.) and process through gravity sedimentation and plating with or without treatments.

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4. Notes 1. Experiments that require a control group of fibers in which satellite cells remain quiescent typically use a “basal growth” medium. Fibers in dishes are usually allowed to equilibrate overnight prior to addition of treatments or labeling compounds. Preparations that contain tissue debris or broken, hypercontracted fibers are discarded, since an intact sarcolemma is important for normal activation. In our hands, documenting that satellite cells are quiescent requires that dishes of fibers isolated and plated in the above fashion incorporate BrdU (0.002% w/v) into very few satellite cells and that the labeling (number of BrdU+ cells per fiber detected by immunostaining for BrdU, e.g.) does not change over time in culture for the control dishes. Dishes can be cultured for 5–8 days, or even longer, depending on the experiment. Staining by immune detection for proliferation markers (e.g., proliferating cell nuclear antigen, Ki67, phosphohistone H3 (H3P), minichromosome maintenance proteins (MCM), or geminin) are good alternatives to BrdU. Use of proliferation markers can confirm activation to the cell cycle in comparison with a positive and negative control, as the absence of these markers corroborates satellite cells are quiescent, which should be the case in untreated control dishes maintained over 3–5 days or longer (even up to 8–10 days). Cultures of control dishes in basal growth medium should prevent satellite cell migration off fibers; satellite cells will not proliferate as a clone on the bottom substrate of the dish if fibers were plated intact (without damage) to maintain quiescence and do not receive an activating treatment. 2. Label each dish on the bottom (typically a side edge) with a solvent-resistant permanent marker, and then spread the collagen around the entire surface of the dish; coat a number of dishes at the same time, and place them in a small plastic tray on top of crushed ice. 3. A typical experimental design, for example, with four treatment groups, will require 20 or 25 culture dishes (each dish is a population of 25–50 or more fibers). This provides sufficient information for a distribution study, with three to six replicates of each treatment. Experience with dissection and isolation will determine the number of muscles required to yield sufficient fibers. However, it is useful to have a “plan B” with fewer or alternative treatments, should the yield of fibers be lower than needed to plate the original number of dishes. Remember to plate a number of “test” culture dishes for working out any staining protocol, and for positive- and negative-staining controls.

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4. For basal growth medium, mix approximately 2 mL for each culture dish that will be plated. The volume depends on the experimental design and the yield of fibers from the dissection. For proliferation medium, mix approximately 100 mL for use in the isolation procedure. 5. We typically make aliquots and store in the freezer. 6. Try bending the toes to visualize the position of the joints between the metatarsals and the digits; remove the skin proximal to this joint. Pull the cut edge skin distally over the toes to expose the plantar surface of the foot. Move the mouse so the leg and foot are over a double or triple layer of wax. After all the skin is removed from the foot, pin the distal foot through its lateral or medial aspect and into the wax (using fine-gauge hypodermic needles or dissecting pins) without piercing the more centrally located muscles or tendons along the midline of the foot. 7. The next steps require fine motor control, typically with the dominant hand, so use the nondominant hand to hold this first pair of forceps. 8. It takes some practice to be able to visualize the anatomy of this muscle and locate its lateral and medial borders, so the first few attempts at dissection should be for orientation to the muscle architecture and practice cleaning. Consult an anatomy text for an overview of the other muscles in the hind limb. 9. Do not put upward pressure on the belly of the FDB with the forceps, repeatedly run the forceps under the muscle or stretch the muscle, as these maneuvers will activate the satellite cells. 10. The tiny tendon that runs to the smallest toe can be discarded as it has few fibers and is often damaged in dissection. 11. Through the microscope, the fascia looks thin and web-like as it separates from the muscle. 12. It is important to do this carefully because the thin tendons to the digits may tear and the attached muscle fibers will tear from the tendon or the muscle belly of the FDB. 13. Be careful not to pull too hard, or the muscle will tear. At this point, if the dissection in vivo was clean, there should be only a small amount of connective tissue. However, it is useful not to overdo the cleaning stage; do not try to remove absolutely everything now, as the collagenase will do a lot of that. The best approach is to clean the FDB as much as possible in vivo while the tendons are attached and the muscle is well anchored. Cleaning the muscle in a dish later is more difficult and it is useful to disturb the muscle as little as possible after removal from the animal. 14. Typically, this amount of collagenase will easily digest up to six or eight mouse muscles in a single small dish.

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15. The time required for effective digestion (one that will allow fibers to separate from tendons and one another) depends on a number of factors. The extent of the dissection (cleaning of the protective fascial coverings), the size (thickness) and number of muscles in the dish, and the muscle type, architecture, and connective tissue distribution (FDB vs. diaphragm; normal vs. more fragile fibers in dystrophic mdx muscle) all make a difference. It is also more difficult to isolate fibers from muscle of older mice. As well, the lot number of the collagenase, its starting temperature (should be approximately 37°C), and the temperature of the incubator affect the rate of digestion. If the experiment is designed to study the activation process itself, avoid shaking the dish and do not place muscles in a tube for incubation on a rolling platform during digestion; we demonstrated that agitation during digestion will activate the satellite cells compared with control cultures without agitation during isolation (19). However, if the aim is to study lineage or proliferation, or mechanisms of inducing quiescence, or to collect muscle satellite cells for dispersed cultures, then it may be advantageous to activate the satellite cells during isolation, and use other activated controls for comparison. 16. Fascia will become cloudy and sticky after digestion and be difficult to grip with forceps, and fat cells can resemble a soft cluster of little spheres. 17. Trituration pulls the muscle belly gently into the pipette and then expels it again into fluid in a repetitive fashion and serves to “shake” off the fibers loosened by digestion from the tendons and from each other. The bore of the pipette should be at least twice the length of the fibers; otherwise, trituration will damage the fibers very quickly. For muscles with fibers longer than FDB (e.g., extensor digitorum longus, tibialis anterior), it is useful to prepare Pasteur pipettes by removing the tapered end completely (flame polish this for safety), autoclave them, and attach the pipette rubber bulb to the cut end, to use the largest bore of the manufactured blunt end. It is also helpful to pull PM in and out of the pipette before using the pipette with the muscles or fibers to avoid fibers sticking to the glass. 18. Muscle fibers should begin to separate from the bellies after about five to ten in-out trituration cycles. If they do not, muscles can be returned to collagenase for 10–15 min for further digestion (in the incubator), and then removed into PM and triturated again. Depending on the collagenase activity of the lot in use, muscle (especially larger ones) may sometimes need a second period of digestion. 19. It is best to triturate one belly at a time, to avoid damaging fibers or stimulating satellite cells to activate by the shear forces generated in trituration. After 10 min, the majority of fibers

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that are going to come off a muscle belly will be isolated. Do not attempt to collect every fiber from each muscle belly; some are very resistant to separation. Since damage to the membrane of fibers may not be obvious until later in the isolation process or even after plating, as it can take some time for fibers to hypercontract (due to influx of calcium), be patient. Once fibers are hypercontracted, they will have released sarcoplasm containing cytokines and other soluble proteins that can damage adjacent fibers (as-yet intact) or stimulate the satellite cells resident on other fibers in the dish. Examine the very fine forceps under a dissecting microscope for bends or nicks that will damage fibers even with a very gently sweeping motion along a digested (jelly-like) tendon. Smooth the tips (the outside and inside surfaces) of forceps by “sanding” with black Emery paper, but after that process, they will lose their “very fine” status. The tips need to meet at a very tiny surface for a secure and finely controlled hold. 20. Pool the fibers from different dishes together, so the preparation uniformly treated between isolation until plating. At this stage, brief trituration (less than 5 min) with a wide-bore pipette is possible if there are still small bundles of fibers remaining in the dish. 21. Dead or dying fibers will begin to shorten or bend (hypercontract), become cloudy, and/or break into short chunky segments, mostly from damage during dissection and cleaning. At this stage, remove any tendons, muscle bundles or connective tissues from the dish using either very fine forceps or a flamepolished wide-bore pipette. 22. For thorough cleaning, repeat the gravity sedimentation process twice more, allowing the fibers to settle for 10 min each time (to minimize losing too many fibers since some get “lost” on the tube walls). Do not process long fibers isolated from muscles longer than FDB by sedimentation as fibers will tangle and damage each other. 23. A larger volume of medium will prevent the collagen from forming a gel that holds fibers on the dish. This step can use a mechanical pipettor and wide-bore disposable tips. 24. Tip a “test” dish sideways; if the fibers stay anchored to the center of the dish without running off into the “corner” at the edge of the dish, the collagen or other matrix has set. 25. If plated on a flexible substrate, the hardened resin disc separates easily from the substrate. If plated on a rigid dish, check the dishes in advance, for resistance to propylene oxide or alternatively embed fibers in aqueous resin (e.g., methacrylate). 26. Tissue sections are typically fixed for in situ hybridization using paraformaldehyde, which preserves RNA (28). However, since paraformaldehyde destroys the adhesion between collagen

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and the fibers (23), it is very important to avoid paraformaldehyde for fixation of fibers on a collagen substrate. Instead, use acid-alcohol fixation for this step of the in situ hybridization process. Modify the acid-alcohol fixation and rinses for in situ hybridization studies using DEPC water and handle with autoclaved instruments throughout, as described (10, 23, 29). 27. Be careful not to dislodge the tiny scales when removing skin as they can cut and damage muscle fibers during later trituration.

Acknowledgments The work of a dedicated research technician (Ritika Upadhaya), summer research students (Alyssa Janke, Stéphane Lenoski, Melody Ong, Jacqueline Richelle, and Colin Rumbolt) and a graduate student (Dr. Orest Pilipowicz, MSc, DMD) provided experiments illustrating effects of drug screening on mouse fibers (SL & CR, see Fig. 1), marcaine in mouse fibers (OP, see Fig. 3) and ISDN and HGF treatments in zebrafish fiber cultures (AJ, MO and JR, see Fig. 4). Funding for this work (to JEA) was from the Canadian Space Agency, the Manitoba Institute of Child Health (MICH), and the Natural Sciences and Engineering Research Council (NSERC). Student support was received from NSERC Undergraduate Summer Research Scholarships (AJ, MO, CR, JR), Faculty of Science University of Manitoba Undergraduate Scholarships (SL), a MICH Graduate Scholarship (OP), a Canada Graduate Scholarship (ACW) and a post-doctoral scholarship from the Canadian Bureau for International Education (WM). References 1. Tatsumi R, Hattori A, Ikeuchi Y, Anderson JE, Allen RE. (2002) Release of hepatocyte growth factor from mechanically stretched skeletal muscle satellite cells and role of pH and nitric oxide. Mol Biol Cell 13:2909–18. 2. Tatsumi R, Sheehan SM, Iwasaki H, Hattori A, Allen RE. (2001) Mechanical stretch induces activation of skeletal muscle satellite cells in vitro. Exp Cell Res 267:107–14. 3. Allen RE, Boxhorn LK. (1987) Inhibition of skeletal muscle satellite cell differentiation by transforming growth factor-beta. J Cell Physiol 133:567–72. 4. Allen RE, Temm-Grove CJ, Sheehan SM, Rice G. (1997) Skeletal muscle satellite cell cultures. Methods Cell Biol 52:155–76. 5. Anderson JE. (2000) A role for nitric oxide in muscle repair: nitric oxide-mediated activation of muscle satellite cells. Mol Biol Cell 11:1859–74.

6. Cooper RN, Tajbakhsh S, Mouly V, Cossu G, Buckingham M, Butler-Browne GS. (1999) In vivo satellite cell activation via Myf5 and MyoD in regenerating mouse skeletal muscle. J Cell Sci 112:2895–901. 7. Tatsumi R, Anderson JE, Nevoret CJ, Halevy O, Allen RE. (1998) HGF/SF is present in normal adult skeletal muscle and is capable of activating satellite cells. Dev Biol 194: 114–28. 8. Robertson TA, Grounds MD, Papadimitriou JM. (1992) Elucidation of aspects of murine skeletal muscle regeneration using local and whole body irradiation. J Anat 181:265–76. 9. Ravenscroft G, Nowak KJ, Jackaman C et al. (2007) Dissociated flexor digitorum brevis myofiber culture system – a more mature muscle culture system. Cell Motil Cytoskeleton 64: 727–38.

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10. Anderson J, Pilipowicz O. (2002) Activation of muscle satellite cells in single-fiber cultures. Nitric Oxide 7:36–41. 11. Bischoff R. (1986) A satellite cell mitogen from crushed adult muscle. Dev Biol 115:140–7. 12. Bischoff R. (1986) Proliferation of muscle satellite cells on intact myofibers in culture. Dev Biol 115:129–39. 13. Yablonka-Reuveni Z, Rivera AJ. (1997) Influence of PDGF-BB on proliferation and transition through the MyoD-myogenin-MEF2A expression program during myogenesis in mouse C2 myoblasts. Growth Factors 15:1–27. 14. Yablonka-Reuveni Z, Rudnicki MA, Rivera AJ, Primig M, Anderson JE, Natanson P. (1999) The transition from proliferation to differentiation is delayed in satellite cells from mice lacking MyoD. Dev Biol 210:440–55. 15. Shefer G, Van de Mark DP, Richardson JB, Yablonka-Reuveni Z. (2006) Satellite-cell pool size does matter: defining the myogenic potency of aging skeletal muscle. Dev Biol 294:50–66. 16. Day K, Shefer G, Richardson JB, Enikolopov G, Yablonka-Reuveni Z. (2007) Nestin-GFP reporter expression defines the quiescent state of skeletal muscle satellite cells. Dev Biol 304:246–59. 17. Rosenblatt JD, Lunt AI, Parry DJ, Partridge TA. (1995) Culturing satellite cells from living single muscle fiber explants. In Vitro Cell Dev Biol Anim 31:773–9. 18. Bekoff A, Betz W. (1977) Properties of isolated adult rat muscle fibres maintained in tissue culture. J Physiol 271:537–47. 19. Wozniak AC, Anderson JE. (2005) Single-fiber isolation and maintenance of satellite cell quiescence. Biochem Cell Biol 83:674–6. 20. Anderson JE. Murray L. (1998) Barr Award Lecture. Studies of the dynamics of skeletal muscle regeneration: the mouse came back! Biochem Cell Biol 76:13–26.

21. Anderson JE, Carvalho RS, Yen E, Scott JE. (1993) Measurement of strain in cultured bone and fetal muscle and lung cells. In Vitro Cell Dev Biol 29A:183–6. 22. Scott JE, Oulton MR, Anderson JE. (1994) Strain induces change in phospholipid and DNA synthesis, cAMP levels and cytoskeletal fibers in isolated fetal rabbit type II alveolar cells. Prog Respir Res 27:173–8. 23. Wozniak AC, Pilipowicz O, Yablonka-Reuveni Z et al. (2003) C-met expression and mechanical activation of satellite cells on cultured muscle fibers. J Histochem Cytochem 51: 1437–45. 24. Wozniak AC, Anderson JE. (2009) The dynamics of the nitric oxide release-transient from stretched muscle cells. Int J Biochem Cell Biol 41:625–31. 25. Li C, Xu Q. (2000) Mechanical stress-initiated signal transductions in vascular smooth muscle cells. Cell Signal 12:435–45. 26. Wozniak AC, Anderson JE. (2007) Nitric oxide-dependence of satellite stem cell activation and quiescence on normal skeletal muscle fibers. Dev Dyn 236:240–50. 27. IHCWorld Protocols. Immunohistochemistry Protocol for Mouse Antibody on Mouse Tissues. IHCWorld website 2008;Available at: URL: http://www.ihcworld.com/_protocols/general_ IHC/immuno_mom.htm. 28. Garrett KL, Anderson JE. (1995) Colocalization of bFGF and the myogenic regulatory gene myogenin in dystrophic mdx muscle precursors and young myotubes in vivo. Dev Biol 169: 596–608. 29. Yablonka-Reuveni Z, Seger R, Rivera AJ. (1999) Fibroblast growth factor promotes recruitment of skeletal muscle satellite cells in young and old rats. J Histochem Cytochem 47: 23–42.

Chapter 7 Somite Unit Chronometry to Analyze Teratogen Phase Specificity in the Paraxial Mesoderm Sara J. Venters and Charles P. Ordahl Abstract Phase specificity, the temporal and tissue restriction of teratogen-induced defects during embryonic development, is a poorly understood but common property of teratogens, an important source of human birth defects. Somite counting and somite units are novel chronometric tools used here to identify stages of paraxial mesoderm development that are sensitive to pulse-chase exposure (2 to >16 h) to 5-bromodeoxyuridine (BrdU). In all cases, it was the presomitic mesoderm (PSM) that was sensitive to BrdU induced segmentation anomalies. At high concentration (1.0 × 10−2 M BrdU), PSM presegment stages ss-IV and earlier were irreversibly inhibited from completing segmentation. At low concentration (2.6 × 10−6 M), BrdU induced periodic focal defects that predominantly trace back to PSM presegments between ss-V and ss-IX. Phase specificity is characteristic of both types of segmentation anomalies. Focal segmentation defects are phase-specific because they result from disruption of 2–3 presegments in the PSM while adjacent rostral and caudal presegments are (apparently) unaffected. Irreversible inhibition of segmentation is also phase-specific because only PSM presegments ss-IV or earlier were affected while older segments (ss-III to ss-I) were able to complete segmentation. The presegments predominantly affected have not yet passed the determination front, the point at which the segmentation clock establishes somite rostro-caudal polarity. Somite unit chronometry provides a means to identify specific PSM presegment stages that are susceptible to induced segmentation defects and the biological processes that underlie that vulnerability. Key words: Birth defect, 5-Bromodeoxyuridine, Cell cycle, Embryo, Intrinsic time, Paraxial mesoderm, Pre-somitic mesoderm, Segmentation defect, Segmentation clock, Somitogenesis

1. Introduction Teratogens account for approximately 10% of birth defects (see Note 1) by inducing developmental malformations in the human embryo during the first trimester of pregnancy (1). Teratogens include a broad spectrum of extrinsic influences that can disrupt embryonic development to produce birth defects (1, 2)

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(see Note 2). Teratogens are tissue-selective because they disrupt development of some embryonic tissues while leaving others apparently unaffected. Phase-specificity refers to the restricted window of teratogen sensitivity, which is a common feature of teratogeninduced defects. The underlying causes for phase-specificity are poorly understood (see Note 3), in part because embryonic development is gradual and most tissues develop through nonincremental phases. Paraxial mesoderm development, by contrast, involves the repetitive formation of somites. Somite formation, in turn, is controlled by the operation of a cell-intrinsic developmental clock. This unique intrinsic chronometry makes the paraxial mesoderm an ideal embryonic tissue in which to study the phase-specificity of teratogen action. In developing embryos, the bilateral strings of somites (see Fig. 1) represent the transient elements that are responsible for all aspects of body segmentation (3) including the formation of the segmented vertebrae and ribs (4, 5). Somites “bud” from the rostral tip of the PSM (see Note 4) as mesenchyme cells coordinately epithelialize into spherical somites. Somites form sequentially and with regular periodicity (see Note 5). After budding, each new somite reiterates a choreographed sequence of developmental events that are reflected in somite stages (ss) (6), which are transient values, designated by Roman numerals (see Fig. 1). The PSM consists of bilateral strips of unsegmented mesenchymal mesoderm immediately caudal to the epithelial somites (see Fig. 1). Mesoderm cells destined to comprise a paraxial mesoderm segment leave the primitive streak and node and enter the caudal PSM (7). As somites bud from the rostral tip of the PSM, the relative position of presegment cells in the caudal PSM regions is progressively advanced in the rostral direction. After completing the developmental stages represented by the entire caudal-to-rostral span of the PSM, each presegment ultimately buds from the rostral tip as a discrete epithelial sphere, a somite. In concert with each somite budding, every remaining presegment within the PSM sequentially advances one somite unit, both in intrinsic developmental time as well as caudalto-rostral position within the PSM, which are designated by negative Roman numerals (8). Thus, at any point in time, each somite or somite presegment of a particular embryo may be characterized by two different numerical values: 1. Its somite number (sn), which is a unique and fixed positive value (expressed as an Arabic numeral), reflecting its position along the axis of the body. Somite numbers increase sequentially in a rostral-to-caudal direction. 2. Its somite stage (ss), which is a transient value (expressed in Roman numerals). Somite stages reflect position relative to the budding somite, which is assigned ss0. For formed somites, ss values are positive and ascend in a caudal-to-rostral direction.

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Fig. 1. Somites and pre-somitic mesoderm (PSM) in a chick embryo. (a) Photograph in ovo. This photograph of a dorsal view of a chick embryo in ovo was taken after injection of contrast ink under the blastoderm. The orientation; horizontal, with its head (rostral) to the right; is used in all illustrations in this chapter. The somite and PSM regions of the paraxial mesoderm are indicated as well as the positions of embryonic primordia of the ear, eye, and neural tube. The bilateral somite strings are marked according to somite number (sn, on embryo’s left) or for somite stage (ss, on embryo’s right). Somite number is a fixed value reflecting position along the rostral-caudal axis. Somite stages are transient values reflecting developmental age (see (c)). (b) Chick embryo cartoon highlighting somites and PSM regions. Tracing of the embryo in A highlighting the location of the somites and PSM (dark gray ). New paraxial mesoderm cells continuously enter the caudal end of the PSM from the primitive node/streak (pn/s) (light gray at caudal end of blastoderm). The asterisk marks the position of the first two somites, which disappear rapidly after forming and are no longer discernable in embryos containing more than ten somites. (c) Somites as measures of intrinsic developmental time. This is a schematic snapshot of a single paraxial mesoderm strip from the embryo shown in (c). It is representative of the schematics used for illustrating experimental somite counting in this chapter. Somite number (sn) is a fixed Arabic numeral value representing the “address” of each new somite within the somite string. The sn of a given somite does not change. Somite number values ascend in a rostral-to-caudal sequence. Somite stages (ss), by contrast, are dynamic values that provide a means for comparing the developmental age of different somites. As each new somite buds, all of the older somites are displaced by one additional somite away from the rostral tip of the PSM. The relative developmental age of each somite can be quantified by the counting number of epithelial somites that lie between it and the rostral tip of the PSM. Somite stage (ss) values are expressed in Roman numerals that ascend from the caudal-most epithelial somite (assigned the value I ). In the PSM, by contrast, pre-somite stages are given negative Roman numeral values that decrease in a caudal direction from the point of somite formation. Somites in the process of budding are assigned the value 0. Somite units (su) are the measures of intrinsic time in the paraxial mesoderm. Each su spans the amount of time required for one somite to bud off of the tip of the PSM (~1.5 h for chick embryo). Somite units theoretically have no upper limit to their potential utility for chronometric analysis. In the experiments in this chapter, somite units are used to track development from PSM to late somite stages.

For presegments in the PSM, ss values are negative and descend in a rostral-to-caudal direction. The repetitive nature of the somite formation results in precise subdivision of the paraxial mesoderm in terms of both fixed body position (somite number) and transient intrinsic developmental staging (somite stage). The somite string, therefore, represents a

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finite chronometric record of paraxial mesoderm development because somite units represent intrinsic developmental time and are independent of “real” or sidereal time (see Note 5). The experimental protocols outlined below describe the “somite counting” methods necessary to track the timing of developmental defects. Somite counting entails meticulous observation and recording of the number of somite pairs present in all experimental and control embryos at multiple points during the experimental time course. Somite counting divides intrinsic developmental time into somite units (su) (see Fig. 1c). Details of somite counting methods have been reviewed (3, 6, 8, 9). Accurate somite counting and recording are the central tactics in the overall experimental strategy of establishing the place and time of teratogen sensitivity during paraxial mesoderm development. Paraxial mesoderm development is disrupted by many different teratogens including 5-bromodeoxyuriding (BrdU) (10). BrdU was recognized early as a teratogen (11), particularly in regard to dose- and time-dependent effects on paraxial mesoderm development (12, 13). The experimental protocols outlined below are designed to analyze the teratogenic effects of BrdU on paraxial mesoderm development to: 1. Identify the paraxial mesoderm development stages that abnormally segment after transient exposure to BrdU. 2. Compare the teratogenic effects of low and high concentrations of BrdU.

2. Materials Avian embryos, their acquisition, incubation in humidified 38°C incubators, using solutions and experimental manipulation as employed in the authors’ laboratories have been reviewed (14, 15), and many of the specific materials sources used in these studies were recently published (9). Briefly, they are given below: 1. Embryos: Obtain fertilized chicken eggs (Gallus gallus) from a local breeder and store in a chemical-free refrigerator between 10 and 12°C for up to 1 week. To obtain embryos at the target age, place eggs narrow end down, in a humid incubator at 38°C. The target age for the embryos in the experiments outlined here is approximately 2 days, or Hamburger Hamilton stages 10–12. The final determination of age is measured by somite counting (see Subheading 3.3). 2. General embryo manipulation: The following materials are used for embryo manipulations: curved scissors, iridectomy scissors, fine forceps, and a 10 mL syringe fitted with an

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18-gauge needle. Embryos are visualized in ovo using contrast medium (item 4, below) delivered with a hand-pulled glass capillary micropipette. Eggs are resealed with Parafilm. 3. Tyrodes solution: Prepare fresh from dehydrated Tyrode’s salts (Sigma) and distilled water according to the manufacturer’s instruction and the pH adjusted to pH 7.4. 4. Injectable contrast solution: Mix carbon-based ink 1:10 with Tyrodes solution and make up fresh for each experiment. 5. Explant cultures: Culture embryos ex-ovo using modified New Culture (16, 17). Punch clover-leaf holes into the center of No. 2 Filter Paper (Whatman) squares and then sterilize by autoclaving before being used to extract embryos from eggs. Prepare the culture media as described in (17) using 0.72 g Bacto-Agar heated in 120 mL of sterile simple saline (7.19 g NaCl in 1 L distilled water) until dissolved. Collect albumen aseptically into a 50 mL Falcon tubes. Equilibrate agar and albumen to 49°C. Mix the agar and saline solutions and add 5 U/mL penicillin/ streptomycin. Aseptically pipette the albumen/agar solution to half-fill 35-mm culture dishes and allow to set overnight at room temperature. Plates can be stored at 4°C. 6. Teratogen. Dissolve crystalline 5-bromodeoxyuridine (BrdU) (see Note 6) in Tyrode’s solution at a working concentration of (1.0 × 10−2 M/10 mM) and store frozen in aliquots. Employ the stock solution concentration (10 mM BrdU) without dilution for the “high” concentration BrdU experiments (see Subheading 3.2). “Low” concentration BrdU experiments (see Subheading 3.1) employed 2.6 × 10−6 M BrdU (2.6 PM) in Tyrode’s solution by dilution of the 10 mM BrdU stock solution.

3. Methods 3.1. Focal Segmentation Defects Caused by Transient Exposure to BrdU at Low Concentration

The degree of BrdU teratogenicity varies with concentration. At very low concentrations, BrdU induces focal defects within the right and/or left somite strings (9, 10). Because somites form as a continuous stream in the paraxial mesoderm, a focal defect is consistent with teratogen-sensitivity at a particular stage or phase of somitogenesis. The goal of the experiment outlined below is to identify the phases of paraxial mesoderm development that are sensitive to BrdU teratogenesis. The following is a step-by-step outline of a published experimental protocol (9) employed to analyze spacing between focal somite defects induced by transient exposure of chick embryos to BrdU (see Fig. 2a). Each step is applied to each experimental or control embryo.

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Fig. 2. Experiment #1: Backtracking focal segmentation defects. (a) Somite counting for a transient teratogen exposurechase experiment. Somite counts (i.e., determination of total number of somite pairs) are performed at the start the experiment (x ), after teratogen exposure (y ) and at the end of the experiment after the teratogen chase period (z ). (b) Somitogenesis and induction of focal segmentation defects. This illustrates the progress of one embryo from the beginning to end of the experiment and backtracking the location and developmental stage of presegments at the time of teratogen exposure as measured by somite units. (c) Backtracking defect exposure window. Example of data record for embryo illustrated in (b).

1. Record time and start somite number (see “x” in Fig. 2c). Remove egg from incubator and open egg shell to view blastoderm through stereomicroscope. Use somite counting to determine total number of somite pairs in embryo (see Note 7). 2. Micropipet 50 PL BrdU solution over the blastoderm of each experimental embryo (see Note 8). Micropipette same volume of Tyrode’s solution over blastoderm of each control embryo. Reseal egg shell and reincubate in ovo for 2 h after which BrdU is diluted by washout. 3. Excise embryo and transfer to New culture. Use iridectomy scissors to make a circular cut around the embryo (well outside of the blood ring) to excise blastoderm and transfer it to a holding dish with large excess of sterile Tyrode’s (see Note 9). 4. Transfer blastoderm to New culture (16, 17) and reincubate overnight in a humidified incubator (see Fig. 2a, step 4). This reincubation is the post-teratogen chase period, which allows

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development to continue in order to observe the location of downstream effects of transient teratogen exposure. 5. Using a stereomicroscope, carefully count and record total somite number for each experimental and control embryo (see z, Fig. 2c) and record sn values for all segments showing defects (see d, Fig. 2c; Note 10). Once somite counting is complete and recorded, the embryo may be harvested for histochemistry and further analysis, if required. 6. Although standard protocols call for recording sidereal times (h/min), in this experimental protocol, the passage of intrinsic developmental time for each embryo is measured in “somite units” (su, see Fig. 1b). The steps in calculating the teratogenic window are illustrated in Fig. 2b, which shows a single somite string of an embryo at the beginning and end of experiment outlined in Fig. 2a. (a) Measure elapsed time. In the example in Fig. 2b, the embryo has 14 somites (x = 14) at the beginning of the experiment. At the end, after being processed through steps 1–5 (see Fig. 2a), the embryo now has 24 somites (z = 24). Thus, the total elapsed time (z − x) is 10 su and is recorded as 'su = 10 (see Fig. 2c). (b) Identify defective segment(s). A defect appears as a fusion between sn 18 and sn 19 (see Fig. 2b). Those sn values are recorded under column “d” (see Fig. 2c). (c) Calculate relative segment age at start of exposure (d − x). Subtracting the number of somites in the embryo at the start of the experiment (x = 14) from the defect positions (d = 18, 19) is used to estimate the approximate age/position of those segments during the BrdU exposure period. This value (d − x, expressed in somite units) indicates that the affected segments were 4 and 5 somite stages younger than somite stage I (i.e., sn14) at the beginning of the BrdU exposure. (d) Exposure window calculation. The y − x value indicates the affected segments were 4–5 stages younger than somite stage I at the beginning of BrdU exposure. Because ss0 is a unit in somite counting, this means that, at the opening of the exposure window, the affected segments were approximately ss-IV to ss-V (after conversion to negative Roman numerals). Somitogenesis can be assumed to have advanced ~1+ su during the 2 h BrdU exposure (y value not determined in this experiment). PSM presegments younger than ss-V were unaffected (in this example) consistent with effective closure of the exposure window at the time BrdU was removed. The main conclusion is that the affected segments were transiently exposed to BrdU while they were in the PSM and that exposure resulted in defects that emerged

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during their transition from ss0 to ssI, the point at which a missegmentation defect would first be evident to an observer. 7. Example data of PSM and pre-PSM sites sensitive to BrdU teratogenic action: Experimental values for x, z, and d have been published for 11 embryos that were transiently exposed to BrdU using the protocol outlined above (9). Table 1 uses these values to estimate the time and place of teratogen exposure for those defects. Values for x, y, and z were calculated from published data (9). L and R refer to defects on the left and right sides, respectively. Defect origins shown in parentheses occurred at prePSM stages. 8. Conclusions from first experimental series: Focal defects in somite segmentation are observed after transient exposure of embryos to BrdU at low concentration (2.6 × 10−6 M BrdU). Focal defects in the somite string indicate that paraxial mesoderm cells pass through phases of elevated sensitivity to BrdU teratogenicity. Backtracking shows that defects observed in somite segmentation result from exposure to BrdU at the PSM or earlier stage of development. The chick PSM contains approximately 12 presegments ((18), and refs therein). Therefore, defect origin values (dL − x and dR − x)

Table 1 Localizing BrdU-sensitive zones in paraxial mesoderm Emb

Somite counts

#

x

Defect locations

Defect origins

Teratogenic window

z

'su dL

dR

dL − x

dR − x

PSM stages

Pre-PSM

1

8

26

16

13–16, 21

–

5–8 (13)

–

-IV to -VII

Yes

2

10

28

18

25

26–28

(15)

(16–18)

−

Yes

3

10

31

21

24

–

(14)

–

−

Yes

4

10

31

21

21, 27

25–26

11 (17)

(15–16)

-X

Yes

5

11

30

19

17–18

–

6–7

–

-V, -VI

–

6

12

28

16

–

17, 22–23

–

5, 10–13

-IV

Yes

7

12

30

18

19, 26–27

26–27

7 (14–15)

(14, 15)

-VI

Yes

8

17

33

16

28

17, 28

11

0, 11

0, -X

–

9

18

35

17

–

19–21

1–3

1–3

0 to -II

–

10

19

33

34

25–28

25–28

6–9

6–9

V–VIII

–

11

21

38

17

–

–

–

–

–

–

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greater than 12 (i.e., lower than -XII) are attributable to BrdUsensitivity in the pre-PSM (i.e., prior to entering the caudal PSM). Defect origin values <12 signify PSM presegment regions that are sensitive to BrdU exposure (see Fig. 4b). The results in Table 1 show that 10 of 11 embryos transiently exposed to 2.6 × 10−6 M BrdU show single or multiple defects in PSM or pre-PSM stages. Six embryos (#1–4, 6, 7) had focal defects that resulted from BrdU-exposure prior to entry into the caudal PSM (Table 1, pre-PSM stages). All regions of the PSM gave rise to segmentation defects in response to transient exposure to BrdU. In those embryos with PSM defect origins, 7 out of 8 showed defects that originated in “young” presegments that had not yet passed the determination front (ss-IV/ss-III boundary) (18, 19). 3.2. Inhibition of New Somite Formation Caused by Transient Exposure to BrdU at High Concentration

This experiment differs from the previous one chiefly in that the embryos are transiently exposed to 10 mM BrdU, a concentration approximately 400 times higher than the concentration used in Subheading 3.1. That is also the generally recommended concentration for BrdU DNA labeling in embryos for antibody detection (20), and which continues in widespread use (9, 21). Preliminary experiments (not shown) showed that embryos exposed continuously to 10 mM BrdU typically formed fewer than five somites overnight while controls formed more than 15 somite pairs during the same period. Transient exposure to high concentration BrdU combined with somite counting shows that the inhibition of new somite formation is irreversible and is essentially stopped during the chase period after BrdU is removed. Experimental steps for control and experimental embryos (see Fig. 3a, b): This is a repeat of steps 1–5 from Subheading 3.1 (see Fig. 2). Each step is applied to each experimental or control embryo. 1. Record time and start somite number (see “x” in Fig. 2c). Remove egg from incubator and open egg shell to view blastoderm through stereomicroscope. Use somite counting to determine total number of somite pairs in embryo (see Note 7). 2. Micropipet 50 PL BrdU solution over the blastoderm of each experimental embryo (see Note 8). Micropipette same volume of Tyrode’s solution over blastoderm of each control embryo. Reseal egg shell and reincubate in ovo for 2 h after which BrdU is diluted by washout. 3. Excise embryo and transfer to New culture. Use iridectomy scissors to make a circular cut around the embryo (well outside of the blood ring) to excise blastoderm and transfer it to a holding dish with large excess of sterile Tyrode’s. Perform intermediate somite count (“y”) (see Note 9). 4. Transfer blastoderm to New culture (16, 17) and reincubate overnight in a humidified incubator (see Fig. 2a, step 4).

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Fig. 3. Experiment #2: Inhibition of segmentation. (a) High concentration BrdU-transient exposure experiment. Shown are illustrations of processing control embryo (upper cartoons) through steps 1–5 as detailed in Subheading 3.1. Note that the control embryo formed 17 new somites by the end of the experiment. Shown are illustrations of processing BrdU-treated embryo (lower cartoons) through steps 1–5 as detailed in Subheading 3.1. Note that the BrdU-treated embryo formed only three new somites by the end of the experiment while the PSM has become elongated and thin. (b) Photograph of control embryo with 33 somite pairs at experiment end. The last somite at experiment start (x ) and end (z ) are indicated. The bracket marks the 17 somites formed during the course of the experiment. (c) Photograph of embryo transiently exposed to 10 mM BrdU after overnight incubation in the absence of BrdU. The last somite at experiment start (x ) and end (z ) are indicated. The short bracket marks the three new somites formed during the course of the experiment. The remaining PSM stopped segmenting and has become elongated and thin.

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Fig. 4. Summary of results of segmentation teratogenesis by transient BrdU exposure. (a) The two experiments outlined in this chapter employed transient (2 h) exposure to BrdU at two concentrations to induce segmentation defects in paraxial mesoderm. Somite unit counting was used to identify PSM stages that are susceptible to segmentation defects induced by this teratogen. Exposure to BrdU at high concentration (1.0 × 10−2M, exp #2) irreversibly inhibits segmentation at PSM stages ss-IV and earlier, while more rostral PSM segments are able to complete somite budding. Exposure to BrdU at low concentration (2.6 × 10−6M, exp #1) induces focal defects in a few PSM presegments but not others. The incidence of focal defects traced to the PSM are enumerated by the plus signs above the somite stages estimated by somite counting to have been affected during the exposure period (see Table 1). (b) Somite stage diagram (see Fig. 1). (c) The solid vertical arrow shows the estimated time of transcriptional activation of the titin gene (70) the first differentiated cell-type (myotome myocyte) specific gene product to appear during paraxial mesoderm development.

Somite formation during the post-teratogen chase period represents presegments that may have escaped the effects of transient teratogen exposure. 5. Using a stereomicroscope, carefully count and record total somite number for each experimental and control embryo (z). Observe and record sn values for last somite formed (see Note 10). Once somite counting is complete and recorded, the embryo may be harvested for histochemistry and further analysis, if required. 6. Use somite count values and the following formulae to calculate the number of somites formed during each experimental interval for each embryo. y − x = number of somites formed during teratogen exposure z − y = number of somites formed during chase period z − x = number of somites formed overall during experiment 7. Use somite count values and the following formulae to calculate the rate of somite formation during each experimental interval for each embryo. t/ yx

rate during teratogen exposure period

t/ yx

rate during chase period

t/ yx

rate overall during course of experiment

8. The averaged values for 3 control and 10 BrdU-exposed embryo is tabulated in Table 2.

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Table 2 Averaged data for experiment #2 During experiment

During chase

End of experiment

#

Rate

#

Rate

#

Rate

Embryo

N

y−x

t/y − x

z−y

t/z − y

z−x

t/z − x

Control

3

2.6

1.2

14.7

1.5

17

1.4

10

1.6

1.7

3.2

7.3

5

5.3

BrdU

9. Conclusions from second experimental series: Pre-somite segments in the rostral PSM (0, -I, -II and sometimes -III) are able to complete segmentation in the presence of 10 mM BrdU. Presegments at ss-IV or earlier, by contrast, are unable to complete somite formation after transient exposure to 10 mM BrdU. The average rate of somite formation during BrdU exposure was slightly slower than the control rate during this period. Somite formation during the chase period, however, was much slower in BrdU-treated embryos (7.3 h/somite) vs. the normal rate of 1.5 h/ somite for controls. Therefore, there are two major conclusions from the second experiment: First, the greatest slowing in somitogenesis rate occurs after transient exposure to 10 mM BrdU, not during exposure; and second, rostral PSM presegments are able to complete segmentation by forming discrete somites, while PSM presegments ss-IV and earlier do not complete somite formation. 3.3. General Conclusions and Discussion

1. The first major conclusion is that somite presegments within the PSM are highly sensitive to the teratogenic action of BrdU. This proved to be true when BrdU segementation teratogenesis was tested at two concentrations, micromolar and millimolar. At both concentration extremes, BrdU affected the subsequent segmentation of presegments in the central PSM at the time of exposure. Rostral PSM segments (ss-III to 0), by contrast, were able to complete segmentation after exposure to BrdU at high concentration. (Note: exposed rostral PSM segments may be affected in other ways not scored in these experiments, such as terminal cell differentiation; data not shown). The potential/possible effects of BrdU on cell structure/ function remain undefined at this time. BrdU is incorporated into replicating DNA, where it efficiently replaces thymidine (reviewed in (22)). BrdU-substituted DNA binds regulatory proteins with higher affinity than does ordinary DNA (23–25). BrdU is readily available intracellularly for DNA synthesis because it is actively and passively transported into cells via the

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same mechanisms as thymidine (26). Intracellular BrdU floods intracellular thymidine metabolic pathways (27, 28) and forms intracellular pools (29). In other words, all cells are permeable to BrdU, some cells actively transport BrdU and S-phase cells incorporate it into newly-replicated DNA. BrdU replacement of thymidine residues in DNA was proposed early as the underlying cause of the teratogenic action of BrdU (30) but a mechanistic cause–effect relationship has yet to be demonstrated. Although BrdU incorporates into DNA only in S-phase cells, soluble BrdU is likely to be present in all cells until it is washed out for the chase period. Therefore, consideration of potential teratogenic targets of BrdU-exposure should include cellular structures and/or systems in addition to DNA. For example, the chromatin-associated enzyme poly(ADP) ribose polymerase (PARP1) is strongly inhibited by BrdU (31–35). In any event, it is likely that all paraxial mesoderm cells were exposed to intracellular BrdU during the 2 h BrdU pulse, and washout removed soluble BrdU from these cells with similar efficacy. A subset of these cells retained BrdU residues in DNA until the end of the experiment. These differences in intracellular location and persistence must be taken into account in distinguishing among potential hypothetical mechanisms that might emerge from analysis of developmental correlates such as those outlined below and illustrated in Fig. 4c. 2. The segmentation clock and underlying developmental maturation of PSM segments may be vulnerable to teratogen action. Developmental progression of presegments in the PSM is dependent upon their continuous participation in rhythmic activity of a segmentation clock (see (36, 37) and references therein), which in turn is dependent upon FGF signaling among PSM presegment cells (18, 38). FGF-dependence ceases, and presegment rostro-caudal polarity becomes fixed, as presegments mature from ss-IV to ss-III, a transition named the “determination front” (19). Presegments also undergo a coherent transcriptome shift at this point (39). Specific human birth defect syndromes affecting vertebral and rib development have now been linked to genetic defects that disrupt segmentation clock function (40, 41) suggesting that these mutation act during the PSM stages of development. 3. Teratogens may disrupt cell cycle synchronization during development of PSM presegments. A surge in mitoses at ss 0 indicates that cell-cycle synchrony becomes established among a large cohort of presegment cells as they pass the G1-to-S phase transition coincident with the determination front at the ss-IV/ss-III boundary (9) (see Fig. 4c, M0 cohort). Cell cycle

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inhibition blocks segmentation in zebrafish without affecting the segmentation clock (42). Mitotic rate has recently been shown to increase dramatically as presegments reach ss 0 (9) indicating that a large cohort of presegment cells enter S phase at roughly the same time they cross the determination front (M0 cohort, see Fig. 4c). Early workers suggested that BrdU inhibits a special type of cell division, named a “quantal” cell cycle (43) and BrdU treatment reduces the number of mitoses in somites (44). More recently, extended exposure to 10 mM BrdU was shown to slow cell cycle rate in the PSM ((21) supplementary data).

4. Notes 1. Birth defects, irrespective of cause, are a significant cause of infant mortality, and infants that survive birth defects require disproportionate pediatric medical care and are at ongoing risk for disability and continuing medical care into adulthood (45). The prevention, or mitigation, of the causes and consequences of birth defects are important goals from a public health perspective and, more importantly, as a means of alleviating human suffering. Unfortunately, the causes of birth defects remain obscure even after systematic epidemiologic studies spanning decades (45). As many as 20–25% of human birth defects have been attributed to genetic/chromosomal causes (2). 2. Embryos exposed to ethanol (46) and tobacco (47) are known to be at increased risk for birth defect. Exposure to industrial toxins is not necessarily teratogenic at low doses (48–50). However, teratogenic chemicals are present in common household products such as phthalates in plasticwares (51, 52) and triethanolamine in cosmetics (53). Many prescription drugs also pose significant teratogenic risk of human birth defects (reviewed in (1)). 3. Phase-specificity is typically defined in terms of teratogen sensitivity during specific gestational days, usually for outcomes assessed at birth (see e.g., (54, 55)). Thalidomide, for example, causes shortened limbs (phocomelia) through the selective disruption of proximal limb bone development (e.g., humerus/ femur, which forms early in limb development), while laterforming bones (carpals/tarsals) in the same limb may be almost normal. Phase-specificity is a tissue-intrinsic property because thalidomide-exposure early will selectively affect the upper but not lower limb and vice versa for later exposure. Mechanistic explanations for thalidomide’s phase-specificity on forelimb development are a subject of scientific speculation (56). Recently, a hypothesis-tested model indicates that

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endothelial progenitor cells in the early limb mesenchyme have high-thalidomide-sensitivity whereas later endothelial precursor populations are less sensitive (57). Thus, the impaired development of the early blood vessels can selectively lead to nondevelopment of proximal bones, which are the earliest to form, while leaving distal bones relatively unaffected because their formation is dependent upon later blood vessels. Interestingly, early limb endothelial precursor cells migrate from the somites into the somatic mesoderm well before the onset of limb outgrowth (58). 4. Nonstandard (or specialized) terms and abbreviations used in the paper. BrdU = 5-bromodeoxyuridine; PSM = presomitic mesoderm Somite budding-epithelialization of somites at the tip of the PSM. 5. The average time interval for somite budding in chicken embryos is approximately 1.5 h/somite at the standard incubation temperature (38°C). That rate drops to ~3 h/somite in embryos incubated at 28°C (59), indicating that the rate of somite budding is ultimately governed by cell metabolic rate. Somite number also provides a good general metric to compare the age of different embryos. Thus, two chicken embryos with the same number of somite pairs are within about 1.5 h of each other in intrinsic developmental age of the paraxial mesoderm 6. Halogenated pyrimidines were early recognized as teratogenic (60) and have been employed as experimental teratogens in a wide spectrum of avian and mammalian embryo systems (11, 61–66). BrdU has excellent attributes as an experimental teratology drug. First, as shown here, it is teratogenic over a wide concentration range; from micromolar to millimolar. Second, because BrdU is water-soluble, it can easily be delivered at a wide range of concentrations and, at the end of exposure periods, readily diluted and removed. Third, cells actively take up BrdU (26) ensuring rapid onset of exposure. Finally, BrdU is incorporated into DNA in place of thymidine (67) allowing immunodetection of those cells exposed during S phase of the cell cycle. At extremely low concentrations (i.e., 2.6 × 10−6 M BrdU as employed in experiment #1 here, see Subheading 3.1) BrdU causes focal defects (9, 10, 68) indicating that it affects some paraxial mesoderm segments and not others, the property of phase-specificity under study here. Finally, focal defects induced by BrdU resemble those produced by other teratogenic agents (10, 68) consistent with the possibility of a common set of cellular/molecular mechanism(s) sensitive to teratogenic disruption. In addition, the material properties of BrdU also contribute to its utility as an experimental teratology drug for paraxial mesoderm. BrdU is water-soluble and easily dissolved at a wide range of concentrations and, to end the exposure period,

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readily diluted and removed. Importantly, cells actively take up BrdU (26) ensuring rapid onset of intracellular exposure. Finally, BrdU is incorporated into DNA in place of thymidine (67) allowing immunodetection of those cells exposed during S phase of the cell cycle. BrdU is also toxic and mutagenic (67, 69). Material safety data sheets for BrdU are available at many internet sites. 7. The values for somite counts are: x = somite count at start of experiment (immediately before administering teratogen) y = intermediate somite count (immediately after removing teratogen) z = final somite count (after chase period) In practice, the following tips may be valuable aids to accurate somite counting. (a) The vitelline membrane must be removed from the 2-day embryo for clear stereomicroscopic view of somites in ovo. Injection of contrast media under the blastoderm is essential for accurate somite counts. Counts in vitro are also conducted using a contrast medium. (b) Somites #1 and #2 disappear quickly after forming (see asterisk in Fig. 1b) and are greatly diminished in embryos with more than 10 somites. Notice that somite #3 is the rostral-most somite with three well-defined borders: caudal, medial, and rostral. (c) The embryo twists and turns as it acquires more somites so only one side may be visible for counting in some regions of the embryo. It is sometimes necessary to switch back and forth between the left and right somite strings to get an accurate count of total somite number. (d) In older embryos (>25 somites), embryo turning and the growing amnion together obscure rostral somites, which interferes with somite counting. In such cases, the right omphalomesenteric/vitelline vein can be used as an approximate marker of somite ~#21 because this vein enters the embryo at that level. This can be helpful in counting somites in older embryos. The appearance of hind limbs can also be a guide. They are adjacent to somite numbers 31–34. (e) In general, it is helpful to double count somites, ideally by two different operators or by counting on two different occasions by the same operator (e.g., before and after excising blastoderm, see later). This discipline enhances accuracy of counts. 8. Teratogen exposure periods are best controlled by good delivery and washout methods. Our preferred method of delivery is to “puddle” the aliquot (~50 PL) of teratogen solution directly

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onto the upper surface of the embryo under the torn/removed vitelline membrane using a micropipette. The egg is then resealed with Parafilm, tape, or other sealant and returned to the incubator for the specified teratogen-exposure period. Termination of exposure is accomplished by removing and unsealing the egg, and flushing the surface of its blastoderm with 2–5 mL of fresh Tyrode’s solution. This rapidly dilutes the BrdU and thoroughly wets the embryo blastoderm in preparation for excision and transfer into New culture for the chase. That method is easily employed in early embryos (i.e., <20 somite pairs) because there is no amnion to interfere. Injections under the surface of the blastoderm also cause similar defects. No systematic comparison of these two routes of delivery has been conducted but both yield equivalent defect patterns. The amnion is a potential barrier to this route in older embryos. 9. To excise the embryo, the whole egg may be removed from the shell into a dish (taking care to keep the yolk intact) or excision can be performed while the embryo is in the egg shell, this is optional and depends on the egg. The cut-out paper ring is placed over the blastoderm with the embryo positioned in the center of the cutout. The periphery of the blastoderm adheres to the paper as it is lifted up and together they are transferred to a holding dish containing excess (2–5 mL) Tyrode’s solution. The transfer of the embryo blastoderm to a holding dish with fresh Tyrode’s further dilutes extracellular BrdU concentrations and, over time, will decrease intracellular BrdU levels (except for BrdU incorporated into DNA). Excised embryo blastoderms must be gently transferred by spoon or wide mouth pipette to maintain integrity of fragile paraxial mesoderm. At this point, it is convenient to perform an “intermediate somite count” (value “y,” see Note 7) because it is easy to view the embryo while in the holding dish and accurately count somites. The y value is used to provide a record of the number of new somite pairs formed during the period of exposure to teratogen. This value (y) is not recorded in experiment #1 but is an important value for the second experimental series below (see Subheading 3.2). 10. The total number of somite pairs recorded at the end of the experiment allows calculation of both the rate of formation and total number of new somite pairs formed. Included in this final count is observation and recording of defects. The axial position (sn) of defects (d) are recorded and converted to somite stage (ss) as necessary using formulas outlined in the main text. Focal defects, such as those observed with low concentration BrdU (Subheading 3.1), affect/comprise one, two, or three adjacent somites, which might have fused borders or are of abnormal size. Other somites appear normal (i.e., somites located rostral and caudal to the affected somites).

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Continuous inhibition of somitogenesis, such as that observed with high concentration BrdU (see Subheading 3.2), results in PSM elongation (apparent) without somite formation. In such embryos, the last somite formed (z) is the only quantitative measure of inhibition. By comparing the value y − x and z − x in controls vs. BrdU-exposed embryos, it is possible to discern whether inhibition of somite formation occurred concomitant with or after exposure.

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Colarusso, T., Siffel, C., Gambrell, D., Thompson, D., Atkinson, M., and Chitra, J. (2007) Reporting birth defects surveillance data 1968–2003, Birth Defects Res A Clin Mol Teratol 79, 65–186. 46. Armant, D. R., and Saunders, D. E. (1996) Exposure of embryonic cells to alcohol: contrasting effects during preimplantation and postimplantation development, Semin Perinatol 20, 127–139. 47. Chiriboga, C. A. (2003) Fetal alcohol and drug effects, Neurologist 9, 267–279. 48. Frakes, R. A. (1988) Drinking water guideline for ethylene thiourea, a metabolite of ethylene bisdithiocarbamate fungicides, Regul Toxicol Pharmacol 8, 207–218. 49. Hardin, B. D., Kelman, B. J., and Brent, R. L. (2005) Trichloroethylene and dichloroethylene: a critical review of teratogenicity, Birth Defects Res A Clin Mol Teratol 73, 931–955. 50. Keller, J. M., Zelditch, M. L., Huet, Y. M., and Leamy, L. J. (2008) Genetic differences in sensitivity to alterations of mandible structure caused by the teratogen 2,3,7,8-tetrachlorodibenzo-pdioxin, Toxicol Pathol 36, 1006–1013. 51. Ritter, E. J., Scott, W. J., Jr., Randall, J. L., and Ritter, J. M. (1985) Teratogenicity of dimethoxyethyl phthalate and its metabolites methoxyethanol and methoxyacetic acid in the rat, Teratology 32, 25–31. 52. Ritter, E. J., Scott, W. J., Jr., Randall, J. L., and Ritter, J. M. (1987) Teratogenicity of di(2-ethylhexyl) phthalate, 2-ethylhexanol, 2-ethylhexanoic acid, and valproic acid, and potentiation by caffeine, Teratology 35, 41–46. 53. Jurand, A. (1958) Action of triethanomelamine (TEM) on early stages of chick embryos, J Embryol Exp Morphol 6, 357–362. 54. Bolon, B., Dorman, D. C., Janszen, D., Morgan, K. T., and Welsch, F. (1993) Phasespecific developmental toxicity in mice following maternal methanol inhalation, Fundam Appl Toxicol 21, 508–516. 55. Horton, V. L., Sleet, R. B., John-Greene, J. A., and Welsch, F. (1985) Developmental phasespecific and dose-related teratogenic effects of ethylene glycol monomethyl ether in CD-1 mice, Toxicol Appl Pharmacol 80, 108–118. 56. Stephens, T. D., Bunde, C. J., and Fillmore, B. J. (2000) Mechanism of action in thalidomide teratogenesis, Biochem Pharmacol 59, 1489–1499. 57. Therapontos, C., Erskine, L., Gardner, E. R., Figg, W. D., and Vargesson, N. (2009)

Thalidomide induces limb defects by preventing angiogenic outgrowth during early limb formation, Proc Natl Acad Sci USA. 58. Eichmann, A., Marcelle, C., Breant, C., and Le Douarin, N. M. (1993) Two molecules related to the VEGF receptor are expressed in early endothelial cells during avian embryonic development, Mech Dev 42, 33–48. 59. Bucciante, L. (1935) Ulteriori richerche sulla velocita della mitosi nelle cellule cotivato ‘in vitro’ in funzione della temparatura [Further research on the velocity of mitosis as a function of temperature in cells cultured in vitro]. Arch Entw-Mech Bd 115 S, 396. 60. Karnofsky, D. A., and Basch, R. S. (1960) Effects of 5-fluorodeoxyuridine and related halogenated pyrimidines on the sand-dollar embryo, J Biophys Biochem Cytol 7, 61–71. 61. Nakamura, N., Fujioka, M., and Mori, C. (2000) Alteration of programmed cell death and gene expression by 5-bromodeoxyuridine during limb development in mice, Toxicol Appl Pharmacol 167, 100–106. 62. Ogawa, T., Kuwagata, M., Muneoka, K. T., and Shioda, S. (2005) Neuropathological examination of fetal rat brain in the 5-bromo-2cdeoxyuridine-induced neurodevelopmental disorder model, Congenit Anom (Kyoto) 45, 14–20. 63. Rizki, T. M., Rizki, R. M., and Douthit, H. A. (1972) Morphogenic effects of halogenated thymidine analogues on Drosophila. I. Quantitative analysis of lesions induced by 5-bromodeoxyuridine and 5-fluorouracil, Biochem Genet 6, 83–97. 64. Sahambi, S. K., and Hales, B. F. (2006) Exposure to 5-bromo-2c-deoxyuridine induces oxidative stress and activator protein-1 DNA binding activity in the embryo, Birth Defects Res A Clin Mol Teratol 76, 580–591. 65. Skalko, R. G., Packard, D. S., Jr., Schwendimann, R. N., and Raggio, J. F. (1971) The teratogenic response of mouse embryos to 5-bromodeoxyuridine, Teratology 4, 87–93. 66. Zamenhof, S., Grauel, L., and Van Marthens, E. (1971) The effect of thymidine and 5-bromodeoxyuridine on developing chick embryo brain, Res Commun Chem Pathol Pharmacol 2, 261–270. 67. Goz, B. (1977) The effects of incorporation of 5-halogenated deoxyuridines into the DNA of eukaryotic cells, Pharmacol Rev 29, 249–272. 68. Primmett, D. R., Stern, C. D., and Keynes, R. J. (1988) Heat shock causes repeated segmental

7 anomalies in the chick embryo, Development 104, 331–339. 69. Ashman, C. R., Kaufman, E. R., and Davidson, R. L. (1985) Bromodeoxyuridine mutagenesis and deoxyribonucleotide pool imbalance in mammalian cells, Basic Life Sci 31, 391–408.

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Part II Non-Mammalian Models of Myogenesis

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Chapter 8 Analysis of Skeletal Muscle Development in Drosophila Ginny R. Morriss, Anton L. Bryantsev, Maria Chechenova, Elisa M. LaBeau, TyAnna L. Lovato, Kathryn M. Ryan, and Richard M. Cripps Abstract The Drosophila system has been invaluable in providing important insights into mesoderm specification, muscle specification, myoblast fusion, muscle differentiation, and myofibril assembly. Here, we present a series of Drosophila protocols that enable the researcher to visualize muscle precursors and differentiated muscles, at all stages of development. In doing so, we also highlight the variety of techniques that are used to create these findings. These protocols are directly used for the Drosophila system, and are provided with explanatory detail to enable the researcher to apply them to other systems. Key words: Drosophila, Muscle, Embryo, Larva, Pupa, Adult, Founder cell, Myoblast, Development, Myogenesis, Method, Protocol, Staining, Hybridization

1. Introduction The mechanisms involved in invertebrate muscle development are similar to those for vertebrate muscle formation, making the animal model Drosophila melanogaster well suited for studies of myogenesis (1). In flies, the basic patterning and specification of the somatic, or skeletal, musculature is similar for all muscles. Moreover, early embryonic myogenesis in Drosophila is completed within a few hours (2). These facts, alongside the utility of Drosophila to apply a genetical approach to the study of biological processes, make this system highly amenable to uncovering basic and broadly relevant aspects of muscle development. The mechanisms of Drosophila muscle development can be considered as a series of consecutive events, and the outcome of

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these steps are usually the measures through which the investigator assesses the extent of myogenesis in wild-type and mutant combinations. Embryonic skeletal muscle development initiates upon specification of a unique founder cell (FC) for each skeletal muscle fiber, and upon the genetical segregation of that cell from unspecified fusion-competent myoblasts (FCMs) within the mesoderm (2). FCs are specified by the Ras signaling pathway (3, 4) and are differentiated from FCMs by expression of specific genetic markers, such as Kin of Irre/Dumbfounded, Roughest/Irregular Chiasm, and Rolling Pebbles/Antisocial. The FCs then attract the unspecified FCMs to the site of muscle formation, for fusion to generate precursors of the individual muscles of the somatic musculature (reviewed in ref. (5)). The FCs are critical determinants of muscle fate, since it is widely thought that the FC is responsible for conferring upon the resulting muscle many of the characteristics unique to that muscle: sites of muscle attachment to the cuticle, orientation of the muscle in the embryo, and muscle size (6). Importantly for the investigator, the specification of all FCs can be inferred by using transgenic lines, such as rP298-lacZ, a lacZ enhancer trap of the dumbfounded gene described by (7). The transgenic line rP298-lacZ is expressed in all skeletal muscle FCs in the embryo (see Fig. 1a). Specific subsets of FCs can alternatively be visualized based upon the accumulation of a number of different markers, many of them are transcription factors that are thought to participate in the specification of individual muscle fiber characteristics (8). Following specification of FCs and FCMs, myoblast fusion is mediated by genetic factors governing orientation, adhesion, and eventually fusion of the cells to form myotubes (reviewed in ref. (9)). The fusion process includes the previously-mentioned FC-specific markers, as well as FCM-specific markers, such as Sticks and Stones, Hibris, and Lame Duck (3). Other markers expressed during myoblast fusion are molecules of the Rac GTPase signaling pathway and the Ras activator myoblast city (10), Loner, kette (11), and blown fuse (12). Multiple rounds of fusion between FCs and FCMs are required for growth of muscles in the embryo (2, 11). The progression of fusion is analyzed by similar methods as founder specification, using genetic markers expressed during this process. Ectodermal cues are also necessary for specification of cell type, myoblast fusion, and differentiation (2, 6, 13, 14). The success of the fusion process can be assessed by the investigator, by analyzing the expression of muscle terminal differentiation markers. Specific muscle gene sets are selectively activated in the individual myoblasts and myotubes by myogenic regulatory proteins such as MEF2 (15), including the contractile proteins myosin heavy chain, Troponins I, T, and C, and muscle-specific actins (16–18). Accumulation of these contractile proteins, or their mRNA transcripts, is indicative of terminal muscle differentiation (see Fig. 1b).

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Fig. 1. Examples of how muscle development can be visualized during the Drosophila life cycle. (a) At stage 12, skeletal muscle founder cells expressing rP298-lacZ (red ) are distinguished amongst all other myoblasts expressing MEF2 (green), using fluorescently labeled antibodies and confocal microscopy of whole-mount embryos. (b) The mature skeletal muscle pattern of a stage 16 embryo, visualized using immunohistochemistry with an anti-Myosin heavy chain antibody in wholemount embryos. (c) Ultrastructure of a single muscle fiber in the late larval stage, observed using double-label immunofluorescence of whole animal fillets and confocal microscopy. Red: F-actin visualized using a fluorescent conjugate of phalloidin; Green: Tropomyosin visualized by a monoclonal antibody (AbCam Inc.) and fluorescently-labeled secondary antibody (Molecular Probes, Inc.). (d) The adult muscle precursor myoblasts visualized in whole-mount preparations of dissected imaginal discs, detected using in situ hybridization to a myoblast-specific gene. (e) The adult muscle pattern observed in transverse paraffin sections, stained with Hemotoxylin and Eosin. Note the six pairs of dorsal longitudinal muscle fibers in the medial region of the thorax. (f) Muscle diversity within a horizontal section of the adult thorax, visualized by activity of a tubular muscle-specific enhancer controlling lacZ expression, and stained with X-gal. The tubular jump muscle is visualized here.

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During the larval stage, muscles specified in the embryo undergo a profound degree of hypertrophy, without overt addition of new myoblasts, or of nuclear division within the muscle syncitium (19). Myofibrillar organization, muscle growth, and muscle degeneration can be readily assessed during the larval stage in whole-mount preparations where the animal is filleted and immobilized for fixation and antibody staining (see Fig. 1c). By contrast, the pattern of adult muscles bears little resemblance to the pattern of larval muscles. Do adult muscles form by restructuring of larval muscles, or are adult muscles formed de novo during metamorphosis? Studies have shown that precursors for adult muscles are specified in the embryonic mesoderm, and can be detected in whole-mount embryos at stage 13 or later by persistent expression of the mesodermal determinant twist (20). These precursors for adult muscles postpone differentiation, and for those muscles that are formed in the adult head and thorax, the myoblasts are stored in the imaginal discs until the muscles are formed during pupal development (20, 21). During the larval stage, the myoblasts proliferate extensively, and can be readily observed in whole-mount preparations of larval imaginal discs (see Fig. 1d). To form the adult muscles, at metamorphosis most larval muscles histolyze and adult muscles are formed de novo by migration and fusion of adult muscle precursor cells (13, 22). Within these migrating populations, the new adult muscles develop in much the same manner as is observed for embryonic/larval muscles: founder cells are specified early during the pupal stage, and myoblast fusion occurs, presumably through a mechanism similar to that defined for the embryo (21, 23, 24). The final pattern of adult muscles is far more complex than that of the embryo (reviewed in ref. (25)). Being significantly larger, the adult is not amenable to whole-mount preparations, thus much of adult muscle development and patterning is assessed through analysis of sections. These sections might be generated from paraffin embedded samples (see, e.g. (26); Fig. 1e). The adult is also characterized by a vast diversity in adult muscle types, which differ from each other ultrastructurally, physiologically, and at the level of gene expression. Patterns of gene expression in the adult muscles are most easily visualized on sections from frozen animals (see Fig. 1f). On the basis of the findings described earlier, we present protocols that will enable the researcher to assess each of the stages of muscle development in the Drosophila system: analysis of muscle founder specification and of muscle differentiation in the embryo; analysis of muscle structure and patterning in the larva; analysis of adult muscle precursors, and of developing adult muscles, in the larval and pupal stages, respectively; and analysis of the pattern and differentiation of adult muscles in mature pupae or adults. The protocols that we describe include antibody staining

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with fluorescent or colorimetric detection, detection of transcripts by in situ hybridization of a labeled riboprobe, and histochemical methods for evaluation of sectioned material.

2. Materials 1. Grape juice agar plates: Combine in a 1-L beaker 9 g Bacto Agar, 150 mL of red grape juice, and 450 mL of dH2O. Bring to a boil in the microwave, stirring regularly. Allow to cool until hand-hot, then pour into Petri dishes. 2. NaCl/Triton: 0.7% NaCl, 0.02% Triton ×-100. 3. 50% bleach. 4. 4% formaldehyde/PBS solution: 3.960 mL of dH2O, 540 PL of 37% formaldehyde, 500 PL of 10× PBS. 5. Paraformaldehyde solution: To prepare 10% stock, dissolve 25 g paraformaldehyde in 200 mL of dH2O, add 15 PL of 10 N NaOH, add 25 mL of 10× PBS, and heat to 65°C with stirring, until dissolved (solution may remain partly cloudy); do not overheat; work under a fume hood; bring volume to 250 mL with dH2O; store at −20°C. To make working 4% solution, take 4 mL of 10% paraformaldehyde, add 1 mL of 10× PBS and 5 mL of dH2O (see Note 1). 6. PEM: 100 mM Pipes pH 6.95, 2 mM EGTA, 1 mM MgSO4. 7. Heptane. 8. Methanol. 9. PBTx: 1× PBS, 0.2% Triton ×-100, 0.2% w/v Blocking Reagent (Roche11096176001). Autoclave, mix thoroughly while hot, and allow to cool. The solution will remain slightly cloudy. 10. PBTxN: 1 mL of PBTx, 50 PL of normal goat serum. 11. PBTw: 1× PBS containing 0.1% Tween-20. 12. Sylgard plates: Sylgard is a polymer available from Dow Corning Inc., which is prepared as a liquid, poured into 50 mm Petri dishes, and allowed to cure overnight at 37°C into a rubberlike substrate. 13. X-gal staining solution (prepared fresh immediately use before from stock solutions): 1× PBS, 100 mM K4[Fe(CN)6], 100 mM K3[Fe(CN6)], 5 M NaCl, 1 M MgCl2, 0.2% X-Gal (see Note 2). 14. Hybridization solution for in situ hybridization: 50% formamide, 5× SSC, 0.1% Tween-20, 0.1 mg/mL sonicated salmon sperm DNA, 0.1 mg/mL heparin. Filter sterilize through a 0.2 Pm filter, and store at −20°C.

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15. Alkaline phosphatase buffer: 100 mM NaCl, 50 mM MgCl2, 100 mM Tris–HCl pH 9.5, 0.1% Tween. 16. Mowiol mounting medium: Add 4.8 g Mowiol 4–88 (Sigma) and 12 g glycerol to a 50-mL Falcon tube. Mix well using a glass rod. Add 12 mL of ddH2O and continue stirring occasionally for several hours at room temperature. Add 24 mL of 0.2 M Tris–HCl (pH 8.5). Continue stirring. Heat by microwaving or using a waterbath to approximately 50°C; continue stirring until most of the Mowiol is dissolved. Once dissolved, add DABCO (1,4-diazabicyclooctane) to the final concentration of 100 mg/mL, and centrifuge the solution at 500 × g for 15 min to clarify from persistently undissolved particles. Carefully remove the supernatant; store aliquots at −20°C. 17. QIAquick PCR Purification Kit (Qiagen). 18. DIG RNA Labeling Kit (SP6/T7) (Roche). 19. Vectastain Elite ABC kit (Vector Laboratories). 20. Alkaline Phosphatase Substrate Kit IV BCIP/NBT (Vector Laboratories). 21. DAB substrate kit (Vector Laboratories). 22. Glycerol solution: 80% (v/v) glycerol in 1× PBS. 23. DEPC ddH2O. 24. 4 M LiCl. 25. 20 mg/mL tRNA. 26. 70 and 100% ethanol. 27. 5 Pg/mL and 12.5 Pg/mL proteinase K in PBTw. 28. 2 mg/mL glycine in PBTw. 29. 50% ethanol/50% xylenes. 30. OCT freezing medium. 31. 1% BSA in PBTx.

3. Methods In this chapter, we provide techniques that enable the researcher to visualize all major aspects of muscle development in Drosophila, throughout its life cycle. These aspects of development are specification of embryonic muscles, including visualizing founder cells; patterning and differentiation of muscles in the embryo; analysis of muscle integrity in the larva; identification of adult muscle precursors in larval imaginal discs; development of adult muscles during the pupal stage; and patterning of imaginal muscles as visualized in the adult.

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Many of the techniques that we present can be applied to different stages of the life cycle, with relatively minor modifications. Therefore, as we describe methods to visualize and analyze muscles and myoblasts at different stages of development, we do so through the depiction of different methods that can be applied. We also provide alternate protocols that enable the researcher to adapt a method description from one stage of the life cycle to other applications or stages of development. Representative images from some of the procedures described are presented in Fig. 1. 3.1. Immunohistochemical Staining of Drosophila Embryos

3.1.1. Collection and Fixation of Embryos

This is the standard protocol, adapted from (27), that is used to visualize the distribution of specific epitopes during embryonic development. Detection of the signal is via enzyme-linked colorimetric detection. A representative image is shown in Fig. 1. Fluorescent secondary antibodies can also be used for detection, for which a representative image is shown in Fig. 1. 1. Set up fly cages for egg collection. Cages are plastic beakers, over the opening of which is attached a grape-juice agar plate. The plate is either 50 or 90 mm diameter, depending upon the beaker used. Flies added to the cage lay eggs next to a smear of yeast paste that is placed on the plate. Ventilation holes are drilled into the sides of the beaker. 2. Collect embryos. Allow flies to lay eggs on the plate for the desired period of time (typically an overnight collection of 18 h at 25°C), then replace the plate with a fresh one. The removed plate will contain eggs that are 0–18 h old if the collection period was 18 h. 3. Remove yeast and dead flies from the plate. Scrape the debris from the plate using the blunt end of a paintbrush, being careful not to gouge the agar. 4. Gently remove embryos from grape juice agar plate. Add ~1–3 mL of NaCl/Triton to plate and gently loosen embryos into solution by rubbing the surface of the plate with finger, without breaking or gouging the agar. Pour solution and embryos through a microfilter sieve, and set the filter on clean paper towel to drain. Microfilters can be made using a cut-off 50 or 15 mL Falcon tube and mesh sieve. 5. Dechorionate with 50% bleach for 3 min. To remove the chorion (eggshell), fill a Petri dish with a 50% bleach solution. Place the microfilter(s) in the solution and mix to disperse the embryos. Wait for 3 min, and then rinse with dH2O. 6. Rinse well. Remove the microfilter(s) from the bleach solution and gently rinse with dH2O for at least 1 min. Be sure to rinse all embryos onto the sieve. At this point, the embryos will have

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lost their egg shells, but will still be surrounded by the hydrophobic vitelline membrane. 7. Fix embryos. Using a paintbrush, transfer embryos into glass scintillation vial containing 4.5 mL of PEM, 0.5 mL of 37% formaldehyde, and 5 mL of heptane. Close cap tightly. 8. Shake embryos for 20–30 min on rotator. Vigorous shaking is required (500 rpm for 20 min, see Note 3). 9. Remove the aqueous layer containing PEM and formaldehyde (bottom) with a pipette. Remove as much fixative as you can, as well as embryos that have fallen into the aqueous phase. These embryos either did not get fully dechorionated, or had already hatched from the egg. In either case, they are not suitable for subsequent steps and should be excluded. Most of the bottom layer can be removed without losing any embryos from the interface (see Note 4). 10. Add 5 mL of methanol (MeOH) and vigorously shake for 1 min. This step will fracture the vitelline membranes, so embryos should pop out and fall to the bottom of the MeOH. Perform this step in the fume hood. Remove the top (heptane) layer and much of the bottom layer without disturbing the fallen embryos. Allow a tiny amount of solution to remain, to protect the embryos from drying out (see Note 5). 11. Wash embryos three times with MeOH to remove residual heptane. Remove as much of the MeOH as possible, leaving a tiny amount of solution to protect the embryos from drying out (see Note 6). 3.1.2. Treatment of Embryos with Antibodies, and Detection

1. Transfer embryos to 1.5 mL microcentrifuge tube and remove MeOH from embryos. Ideally, there will be 50–100 PL of settled embryos for each staining reaction (see Note 7). 2. Wash quickly with 800 PL of PBTx. 3. Wash 5 min with 800 Pl of PBTx. 4. Wash 30 min in 800 PL of PBTx. 5. Incubate 30 min in 100 PL of PBTxN at room temperature on rotator. 6. Remove PBTxN and add 200 PL of primary antibody diluted in PBTxN. Incubate on the rotator 2 h to overnight. If there are not a lot of embryos, make 100 PL of a primary antibody dilution at 2× the desired final concentration, and add this straight to the embryos already immersed in 100 PL of PBTxN. This will dilute the antibody to the desired final concentration (see Note 8). 7. Remove primary antibody and wash quickly 5 min with 800 PL of PBTx.

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8. Wash four times at 30 min each in 800 PL of PBTx on the rotator. 9. Incubate 30 min in 100 PL of PBTxN on the rotator. Make sure there is enough PBTxN for embryos to move around. 10. Remove PBTxN and add 200 PL of secondary antibody diluted with PBTxN. Incubate for 2 h to overnight. If there are not a lot of embryos, make secondary antibody dilution 2× the desired final concentration and add this straight to the embryos already immersed in 100 PL of PBTxN reagent of the same volume. This will dilute the antibody to the final desired concentration (see Note 9). 11. Remove and discard secondary antibody, and wash quickly 5 min with 800 PL of PBTx. 12. Wash four times at 30 min each in 800 PL of PBTx on the rotator. 13. During the last wash, mix up a batch of Vectastain ABC reagent (0.5 mL PBTx, 10 PL reagent A, 10 PL reagent B). Mix (but do not vortex) after addition of each component, and let stand 30 min at room temperature. 14. Remove all PBTx and add 500 PL ABC reagent to each tube of embryos. Rotate 15 min. 15. Remove ABC reagent, and wash five times over 1 h with PBTx. Remove as much PBTx as possible after last wash. 16. Wash 2× 5 min with 1× PBS to remove traces of PBTx. 17. Mix DAB staining solution from DAB staining kit: combine 2.5 mL dH2O, 2 drops DAB (supplied), 1 drop buffer (supplied), and 1 drop H2O2 (supplied). Mix well after addition of each reagent (see Note 10). 18. Remove 1× PBS from the embryos and add 500 PL of Vectastain DAB staining solution to embryos. Allow the reaction to occur in the dark (usually 2–3 min) at room temperature. Do not overstain. The investigator can closely monitor the development of the reaction under a microscope using a small representative sample of embryos to determine when the reaction should be stopped. 19. Stop reaction by removing stain and washing with PBTx (see Note 11). 20. If double-DAB staining, wash embryos 30 min in PBTx to remove any residual stain, and repeat process starting at step 6 for second stain. 21. Store embryos in PBTx at 4°C. In preparation for mounting, remove PBTx, and add 800 PL of glycerol solution (80% (v/v) glycerol, 1× PBS). Allow glycerol to infiltrate embryos for 2 h at room temperature, or overnight at 4°C.

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3.1.3. Mounting Embryos for Photomicroscopy

1. Select a single representative stained sample and put on a microscope slide. 2. Add 2 drops of 80% glycerol/1× PBS to slide. Be careful with amount of glycerol, as having too much will allow the sample to roll, and too little will cause the coverslip to crush the sample. 3. Gently put a coverslip over the sample and use fingers to gently push the coverslip around, maneuvering the tissue into the appropriate orientation. The sample is now ready for photodocumentation using DIC optics.

3.2. Immunofluorescent Analysis of Larval and Pupal Whole-Mount Samples

This protocol is used for dissected preparations of larvae and pupae, and can be adapted for adults. Given the need to dissect the samples to expose the developing muscle tissue, and then keep that area exposed during subsequent staining steps, we have developed a procedure where animals are dissected in a Sylgard-coated dish, and all incubations are carried out in that same dish (28). At the end of the procedure, the samples are removed from the dish and mounted for photodocumentation. A representative image is shown in Fig. 1c.

3.2.1. Preparation and Fixation of Larval and Pupal Samples

Some photographs of specific dissection steps are included here to guide the researcher in this process (see Fig. 2). 1. Select desired larvae/pupae at correct stage of development. Larvae are generally dissected at the third instar stage. Pupae are generally selected at the onset of pupariation (0 h; white prepupal stage) and dissected at the appropriate stage after puparium formation (APF). 2. For larvae only: Heat kill. Place larva on a small cheese cloth or microfilter, and dip into 60°C water bath for 20–30 s. Immediately cool by placing filter directly on ice for 2–3 min (see Note 12). 3. Place larva/pupa on dissecting dish (Petri dish with Sylgard on bottom) containing 1× PBS. Using dissecting pins, pin sample at anterior end. Pin dorsal side up for pupae, dorsal or ventral side up for larvae, depending on what structures will be viewed. 4. Cut off posterior end of animal with dissecting scissors (see Note 13). 5. Make straight cut from posterior to anterior end of larva/pupa using short snips and never removing scissors, stopping short of the anterior pin. If working with pupa, make sure scissors are cutting into the underlying animal cuticle, and not just the pupal case. Be careful not to cut too deep into the sample.

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Fig. 2. Steps for fillet preparation of larval or pupal samples. A pupal dissection is depicted in this image, but similar steps are used for larval dissections. (a) Immobilize the larva at the anterior end with a dissecting pin, facing ventral side down. (b) With a pair of fine scissors, puncture the posterior tip in order to relieve hemolymph pressure. (c) Remove the posterior tip of the abdomen, including cuticle and tissue of the animal. Then, insert the lower prong of the scissors into the orifice produced, and cut anteriorly along the dorsal side of the animal. (d) Once you reach the head, turn the scissors and cut at an angle to either side of the pin (this step aids in opening up the fillet). Not shown: once the cuts have been completed, place a pin through the posterior and ventral side of the animal, to allow it to be immobilized at both ends. (e) Using pins and forceps, carefully open up the sides of the cuticle and pin laterally to generate a flat fillet. Not shown: usually, the posterior pin is removed once this is complete. Internal organs and tissues can be carefully removed to enable the researcher to visualize the tissue of interest.

6. From the anterior end of the long cut, make an incision at a 45° angle, toward each side of the sample, angling anteriorly and laterally. 7. Using forceps to hold the dissecting pins, pin the body (and pupal case) open at all four corners, exposing regions to be stained. 8. Gently pull out fat body (milky white in appearance) from the larva, leaving muscles exposed. If working with pupa, use a Pasteur pipette to clear away any debris from around the sample. 9. Remove 1× PBS and fix tissues in 4% formaldehyde/1× PBS solution on ice for 30 min.

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3.2.2. Treatment of Larval and Pupal Samples with Primary Antibodies and Fluorescently-Linked Secondary Antibodies

The steps for primary and secondary antibodies are essentially similar to those for immunohistochemical staining of embryos. We modify the protocol here to provide details of how to do it in the dishes, and to illustrate the use of fluorescent secondary antibodies. 1. Remove fixative to appropriate waste container. 2. Wash quickly with 5 mL of PBTx. 3. Wash 5 min with 5 mL of PBTx. This and all subsequent steps should be carried out on a flat-bed shaker, at a low setting (~30 rpm). Overnight steps should be carried out on a shaker at 4°C; shorter steps (up to 2 h) can be at room temperature on the shaker. 4. Wash 30 min in 5 mL of PBTx. 5. Incubate 30 min in 5 mL of PBTxN. 6. Remove PBTxN and add 5 mL of primary antibody diluted in PBTxN. Incubate 2 h to overnight (see Note 14). 7. Remove primary antibody and wash 5 min with 5 mL of PBTx. 8. Wash four times at 30 min each in 5 mL of PBTx. 9. Incubate for 30 min in 5 mL of PBTxN. 10. Remove PBTxN and add 5 mL of secondary antibody diluted with PBTxN. Incubate for 2 h to overnight in a darkened environment (usually we place the lid of a cardboard freezer box over the dish, see Note 15). 11. Remove and discard secondary antibody and wash 5 min with 5 mL of PBTx. 12. Wash four times at 30 min each in 5 mL of PBTx in a darkened environment. Your samples are now ready for viewing.

3.2.3. Mounting Larval/ Pupal Samples for Photomicroscopy

1. Put a small amount of 80% glycerol on a slide so the fillet can be sufficiently covered. 2. Remove pins from the larval/pupal fillet. For pupae, reinsert pins so that pupal case is still pinned down but carcass can be moved. Under the dissecting fluorescent microscope, additional dissection can be carried out to fully remove unnecessary tissue. 3. Gently move fillet to microscope slide and orient. 4. Place one coverslip on each side of fillet, creating a platform. (This can be done prior to adding glycerol). 5. Cover, but do not smash, the fillet by setting a third coverslip on top of platform created with other coverslips. The sample can now be viewed and documented using a fluorescence or confocal microscope.

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3.3. In Situ Hybridization Analysis of Imaginal Discs

Imaginal myoblasts, those giving rise to the adult skeletal muscles, are segregated in the larva. Those in the thorax are associated with the imaginal discs, and the most prominent example of this is the myoblasts associated with the wing imaginal disc (20). These cells give rise to the indirect flight muscles, as well as to some direct flight muscles associated with the base of the wing. The wing discs are located in the anterior portion of the larvae. They are usually attached to the trachea, unless dislodged during the initial opening of the larvae. One wing disc lies on either side of the brain. A series of images during larval dissection is provided for guidance (see Fig. 3). Here, we present a procedure to visualize gene expression in the wing imaginal discs using in situ hybridization of a labeled riboprobe to mRNA targets. The protocol follows that of (29) for embryos, as modified by (30) for imaginal discs. Gene expression is visualized in whole-mount discs that have been dissected from the larva. A representative image is shown in Fig. 1d.

3.3.1. Dissection of Discs from the Larva

1. Prepare 1 mL of fixative solution (3.7% formaldehyde, in 1× PBS) in a 1.5 mL microcentrifuge tube, and place on ice. 2. Fill Sylgard-coated dissecting dish with 1× PBS. Place larvae into PBS.

Fig. 3. Steps for isolation of imaginal discs from larvae. (a) Immobilize the larva at the posterior end using a pair of forceps. (b) Once immobilized, grasp cuticle at the posterior end with a second pair of forceps, and peel in an anterior direction. (c) Locate finger-like projections on tracheal ends (arrowhead ) near the head, in order to roughly locate the imaginal discs. (d) Clear fat and gut debris away from carcass; discs should now be visible (arrow indicates wing disc). (e) Follow trachea (arrowhead ) to locate disc of interest (arrow ); they will be posterior to the tracheal finger projections, and located close to the brain. (f) Clean away remaining debris from disc. The disc can now be gently pipetted into a tube containing fixative.

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3. Under the dissecting microscope, immobilize the larva at its posterior end with forceps. With the other set of forceps, take hold of the cuticle adjacent to the immobilizing forceps, and peel the tissue anteriorly in a single motion. Remove fat body and debris, utilizing trachea and brain as markers for the discs’ approximate location. Separate the discs from surrounding tissue. The wing disc frequently is associated with the haltere and T3 leg discs. At this stage, we do not seek to separate them from this association, as the larger complex of discs is more easily visible and easier to handle during the subsequent steps. 4. Carefully pipette cleaned discs into the formaldehyde solution. Once you have dissected the last pair of discs, allow fixation for an additional 30 min. It is wise to obtain at least ten discs for antibody stains and at least 20 for in situ hybridization, as many are lost during the subsequent steps (see Note 16). 3.3.2. RNA Probe Synthesis

Labeled RNA probes are synthesized in vitro by transcription of a plasmid containing a cDNA of the gene of interest. The plasmid should contain promoter regions for RNA-polymerases (T7, T3, or SP6) on either side of the Multiple Cloning Site, to permit RNA synthesis from both DNA strains. 1. Linearize 10 Pg of plasmid DNA with an appropriate enzyme. Two parallel reactions should be set up. The first reaction should cut the plasmid containing an insert at the 5c end of the insert, for generation of the antisense probe. The second reactions should cut the plasmid on the 3c end of the insert for generation of the control sense probe. 2. Analyze 1 PL of digests on a 0.8% agarose gel to ensure linearization. 3. Clean-up the reactions by processing them through the QIAquick PCR Purification Kit (Qiagen). Alternatively, the digests can be phenol/chloroform extracted and then precipitated. 4. Use 0.5–1 Pg of linearized template DNA for the DIG RNA labeling reaction (per the Roche DIG RNA labeling kit). Incubate at 37°C for 2 h. 5. Precipitate probe by adding the following per 10 PL synthesis reaction: 15 PL of DEPC ddH2O, 4 PL of 4 M LiCl, 5 PL of 20 mg/mL tRNA, 100 PL of 100% ethanol. Mix well after addition of each reagent. 6. Incubate at −80°C for 20 min. 7. Spin at 4°C for 15 min at maximum speed in a benchtop centrifuge. 8. Remove supernatant, and wash pellet with 100 mL of 70% ethanol.

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9. Spin at 4°C for 5 min at maximum speed. 10. Resuspend pellet in 75 PL of hybridization solution. 3.3.3. Imaginal Disc In Situ Hybridization

1. Remove fixative from dissected imaginal discs, and wash with PBTw. 2. Add 500 PL of 12.5 Pg/mL proteinase K in PBTw, and incubate on rotator for 2 min (see Note 17). 3. Remove proteinase K solution and quench in 1 mL of 2 mg/ mL glycine in PBTw by mixing on the rotator for 2 min. 4. Fix again with 800 PL of 4% formaldehyde in PBTw on rotator for 20 min. 5. Remove fixative and wash several times with 800 PL of PBTw. 6. Wash with 800 PL of 50% PBTw/50% hybridization solution for 10 min at room temperature on the rotator. 7. Wash 3× with 800 PL of hybridization solution. 8. Add 800 PL of hybridization solution, and prehybridize at 55°C for 30–60 min. Samples do not need to be on the rotator for steps at 55°C, and these steps are carried out in heating blocks. 9. Dilute probe to approximately 50 ng per 100 PL of hybridization solution. This concentration may have to be adjusted, depending on the abundance of the transcripts being investigated. 10. Remove prehybridization solution from embryos, and add 100 PL of diluted probe. Incubate overnight at 55°C. 11. Remove probe/hybridization solution and wash with hybridization solution 3× over the course of an hour at 55°C. 12. Wash with 50% hybridization solution and 50% PBTw for 10 min on room temperature rotator (the remaining steps are all carried out at room temperature). 13. Wash 4× with PBTw over the course of an hour. 14. Remove last wash and add 200 PL of PBTwN (1:20 Normal Goat Serum: PBTw). Incubate on rotator for 30 min. 15. Add 200 PL of Anti-Digoxigenin-AP (Roche) diluted to a final concentration of 1:1,000 in PBTwN, and incubate on rotator for 2 h. Alternatively, this incubation can be carried out overnight on a rotator at 4°C. 16. Remove antibody and wash four times with PBTw over the course of an hour on rotator. 17. Wash three times in Alkaline Phosphatase Buffer for 15 min. 18. Remove last APB wash and add BCIP stain. 19. Monitor reaction (staining may come up as soon as 10 min or take as long as overnight).

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20. Stop reaction by washing samples with 100% ethanol, and then PBTw. Store in PBTw at 4°C. 21. For viewing, stained discs should be placed on a slide, add two drops of 80% glycerol/1× PBS, and mount under a coverslip. We do not recommend clearing discs in glycerol prior to mounting (as is performed for stained embryos), since the discs become highly transparent and difficult to orient. 3.3.4. Alternate Protocol: In Situ Hybridization to Drosophila Embryos

This is also a common procedure, whose main steps correspond to those described already for in situ hybridization to imaginal discs. Here, we describe the early steps of the embryo procedure, to the point where the embryo and imaginal disc protocols converge. 1. Place approximately 75 PL of fixed embryos Subheading 3.1.1) into a microcentrifuge tube.

(see

2. Remove methanol and wash with 1 mL of 100% ethanol. 3. Add 1 mL of 50% ethanol/50% xylenes. Incubate on rotator for 30 min. 4. Wash 5 times with 1 mL of 100% ethanol. 5. Wash 5 times with 1 mL of PBTw. 6. Add 5 Pg/mL Proteinase K in PBTw, incubate on rotator for 8 min. 7. Wash with 1 mL of 2 mg/mL glycine in PBTw. 8. Wash four times with 1 mL of PBTw. 9. Fix in 1 mL of 4% formaldehyde in PBTw on rotator or 25 min. 10. Wash 5× with 1 mL of PBTw. 11. Next, follow the same steps as for in situ hybridization to imaginal discs, starting at Subheading 3.3.3, step 6. Stained embryos should be cleared in glycerol prior to photodocumentation, as described for immunohistochemistry of embryos. 3.4. Analysis of Reporter Gene Expression in Adult Cryosections

This protocol describes how cryosections of pupae or adults can be generated, and then analyzed for expression of a lacZ reporter gene using X-gal staining. A representative image is shown in Fig. 1f. Simple modifications, allowing for cryosections to be subjected to immunofluorescent staining or in situ hybridization, are described in Subheading 3.4.3.

3.4.1. Mounting and Sectioning Samples

1. Anesthetize adult flies with CO2, submerge into a large drop of OCT freezing medium, and gently toss about in the media to remove air pockets and bubbles. 2. Put a piece of double-sided sticky tape (Scotch) onto the flat tongue of a metal spatula; attach a paper strip with sample’s description to one side of the tape.

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3. After removing excess freezing medium, spread a fly on its dorsal side on the sticky tape piece, spread wings and legs along both sides to acquire perfect central positioning and prevent rolling over; allow sample to dry slightly (see Note 18). 4. Cover the fly with fresh layer of freezing media, and submerge into liquid nitrogen. Once the sample is fully frozen (about 20 s), detach the frozen block with the sticky tape from the spatula (see Note 19). 5. Mount the frozen block containing the specimen onto sample holder of a cryotome, so the sticky tape is facing up; peel the sticky tape off and trim the block into a truncated pyramid shape using a razor blade (see Note 20). 6. Align the cryotome blade with the plane of the block’s top surface. Begin sectioning 10–12 Pm thick sections. 7. Collect sections individually onto a slide, and air-dry for at least 15 min. Sections can be stored in a dry box at 4°C for a few days before processing (see Note 21). 3.4.2. Histochemical Staining of Cryosections for b-Galactosidase Activity

1. Wash sections by immersing the slide in 1× PBS solution for 10 s (see Note 22). 2. Fixation: You can either immerse slide into 4% formaldehyde solution, or add 200 PL of fixative per slide and cover with 18 × 60 mm coverslip. We use both methods with no visible difference. Incubate for 5 min at room temperature (see Note 23). 3. Wash fixed sections three times, 5 min each, by immersing the slide into 1× PBS solution. 4. Add 200 PL of X-gal staining solution to the slide and carefully cover with coverslip. Incubate at 37°C in a humid box until a blue stain is developed (see Notes 24 and 25). 5. Wash the slide three times by 5 min with 1× PBS; rinse with PBTx. 6. Add 100 PL of Mowiol mounting medium to the slide and carefully cover with coverslip, avoiding bubbles. Observe and document using a compound microscope.

3.4.3. Alternate Protocol: Immunofluorescent Staining of Cryosections

Since diffusion of reagents into cryosections occurs rapidly, antibody staining of these sections can be readily achieved in a single day. 1. Wash dried sections by immersing slide into 1× PBS solution for 10 s (see Note 26). 2. Fixation: immerse microscope slide with sections into 4% formaldehyde solution in 1× PBS for 10 min (see Note 27). 3. Wash sections three times, 5 min each, by immersing the slide into PBTx solution.

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4. Add 200 PL of blocking solution (1% BSA in PBTx) to each slide, and cover with a cover slip. Incubate in a humid box for 30 min. 5. Dilute primary antibody in 200 PL PBTx containing 1% BSA. BSA blocks antibody binding to nonspecific epitopes (see Note 28). 6. Add 200 PL of antibody solution to the slide and gently cover with a coverslip. Incubate for 1 h at 37°C or overnight at 4°C (see Note 29). 7. Wash sections three times, 5 min each, by immersing slide into PBTx solution. 8. Dilute secondary antibody in PBTx containing 1% BSA; include fluorochrome-labeled phalloidin (1:500) and DAPI (1:1,000) to counter-stain muscles and nuclei, respectively (see Note 30). 9. Add 200 PL of secondary antibody solution to a slide and carefully cover with a coverslip. Incubate for 1–2 h at room temperature, protect from light. 10. Remove coverslip, and wash sections three times, 5 min each, by immersing slide into PBTx solution. 11. Add 100 PL of Mowiol mounting medium to slide, and carefully add a coverslip avoiding bubbles. Dry at room temperature for 1–2 h, and visualize stain via fluorescence or confocal microscopy. 3.5. Analysis of Adult Muscle Patterning via Hematoxylin and Eosin-Stained Paraffin Sections

3.5.1. Fixation and Embedding of Samples in Paraffin

This procedure is from (30), as modified from (31). This allows for the generation of sections with particularly good preservation of structures, and is used to assess the overall arrangement and patterning of adult muscles. A representative image is shown in Fig. 1e. Given the extensive fixation and embedding steps that are required, paraffin sections are not ideal for antibody staining, although a method to achieve immunostaining of paraffin sections is included in the next section. 1. Anesthetize adult flies with CO2 on a sleeping pad; prepare thorax specimens by chopping off heads, abdomens, legs, and wings with fine scissors or razor blade. 2. Fix specimens with 4% paraformaldehyde in 1× PBS at 4°C overnight in a scintillation vial (see Note 31). 3. Remove fixative and dispose using appropriate means. Wash embryos with 1× PBS at 4°C for at least 30 min. 4. Embed specimens in agarose: pour warm (but not hot) 1% agarose prepared in 1× PBS into a Petri dish. Add the thoraces, and quickly orient with forceps. Allow the agarose to solidify. Next, cut agarose blocks containing thoraces in the shape of isosceles triangles making the base of the triangle align with the dorsal side of thoraces (see Note 32).

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5. In a scintillation vial, dehydrate agarose-embedded specimens through the following sequential washes: 0.85% saline, 25 min; saline/ethanol (1:1), 25 min; 70% ethanol, 25 min, twice; 85% ethanol, 25 min; 95% ethanol, 25 min; absolute (100%) ethanol, 30 min, twice (see Note 33). 6. Displace ethanol with xylene and, subsequently, with paraffin, as follows: Xylene, 30 min, twice; xylene/paraffin (1:1) at 63°C, 45 min; paraffin at 63°C, 30 min, twice (see Note 34). 7. After the second paraffin incubation, place specimens into embedding base moulds, orient the base of the agarose triangle against the side of the mould, and let solidify overnight at 4°C (see Note 35). 8. Remove solidified, specimen-containing paraffin block from the mould. Trim the paraffin into the shape of a pyramid and mount on the microtome holder, so that the base of the agarose triangle is now on top and facing the microtome blade. 9. Cut sections from the block at 8–10 Pm thickness. If the block has been trimmed so that the top and bottom sides of the pyramid are parallel to each other, straight ribbons of sections should be generated. 10. Transfer the ribbons to a prewarmed water bath set at 40°C, to remove folds and jams, put onto a slide, and let dry overnight on warming plate. Dried sections can be stored indefinitely. 11. For hematoxylin and eosin staining, sections need to be deparaffinized and rehydrated through the following series of solvents (2 min each step): Xylene, twice; absolute ethanol, twice; 90% ethanol; 80% ethanol; 70% ethanol; 50% ethanol; 30% ethanol; ddH2O. 3.5.2. Staining Paraffin Sections with Hematoxylin and Eosin

1. Once sections have been rehydrated to ddH2O, stain sections by immersion in hematoxylin stain; time of staining to be determined empirically. Stop staining by washing several times in ddH2O. 2. Dehydrate sections with 30, 50, and 70% ethanol, incubating for 2 min at each step. 3. Stain sections with eosin, times to be determined empirically. Wash away excess eosin using 70% ethanol. 4. Finish dehydration with 2-min incubations in 70, 80, 90%, and absolute ethanol (twice). 5. Rinse slides with sections in two changes of xylene, and mount a coverslip over sections using Cytoseal-XYL (VWR). Gently push the coverslip down over the samples, and allow mountant to dry overnight in the fume hood.

3.5.3. Antibody Staining of Paraffin Sections

Although this procedure is not always successful (depending upon the activity of the antibody being used), it can often be necessary to obtain immunohistochemical data from paraffin sections.

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The basic protocol is to deparaffinize the sections, then to reverse the fixation crosslinks through a procedure known as “antigen retrieval,” and then to follow the antibody staining procedure described above for cryosections, but adapted to achieve detection using the ABC reagent and immunohistochemistry. Stained samples are then usually partially dehydrated, counterstained with eosin, and permanently mounted in Cytoseal-XYL. 1. Paraffin sections, which have been dried overnight, are deparaffinized as described for hematoxylin and eosin staining of sections (Subheading 3.5.1, step 10). Once the sections are in diH2O, the researcher should immediately proceed to the steps described below. 2. Place slides in a glass slide bath, containing 250 mL of working strength Antigen Retrieval Citra (Biogenex Inc.). 3. Fill the base of a microwaveable pressure cooker with 600 mL of dH2O. Place the slide bath in the water and ensure that the water does not over-flow into the slide bath. Apply the lid of the pressure cooker and tighten. Some models also have a pressure regulator weight, which should also be applied. 4. Microwave at 800–850 W for 15 min. During this time, the pressure indicator button on the oven should rise. 5. Reduce the power level to 300–350 W, and heat for a further 15 min. 6. After this time, remove the pressure cooker from the microwave, and allow to cool until the pressure indicator drops. If there is a pressure regulator weight, remove that and determine whether any steam is still escaping through that aperture. If there is, then there is still pressure inside the cooker, and it should be allowed to cool for longer. If no steam escapes, then you can proceed with the next step (see Note 36). 7. Once the pressure has reduced, use oven mitts to carefully open the cooker (and avoid burns from escaping steam). Carefully lift the slide bath from the cooker (recall that the slide bath will be sitting in 600 mL of water), and set on the bench to dry. 8. Allow the immersed slides to cool to approximately room temperature, and then proceed with the antibody staining as described in the next section. 9. Once the samples have cooled, the slides can be washed in 1× PBS. Antibody stains can proceed, through a combination of the cryosectioning protocol, starting at Subheading 3.4.2, step 3 (to include incubations with primary and secondary antibodies, and the associated washes); and the embryo immunohistochemistry protocol, starting at 3.1.2, step 13 (to include the ABC reagent steps) and detection using DAB. For the latter

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protocol, adapt the volumes from those indicated for embryos to those used for slide staining. 10. DAB-stained sections should be washed with PBTx, 1× PBS, 0.85% saline (2 min each), before immersion in dH2O. At this point, samples can be partially dehydrated to 70% ethanol, stained with eosin, and then fully dehydrated for mounting in Cytoseal-XYL.

4. Notes 1. (Para)formaldehyde is a known irritant and carcinogenic compound; use caution working with its powder and solution. Use gloves and fume hood for protection. Collect and utilize formaldehyde waste in accordance with institute’s policy. 2. 10% (w/v) X-gal stock solution is made in formamide and stored liquid at −20°C, protected from light; ferro- and ferricyanides are stored at 4°C as 500 mM aqueous stocks. 3. The PEM/Formaldehyde/Heptane solution will separate into an aqueous and an organic phase. The embryos will collect at the interface and remain suspended. The formaldehyde will act as a fixative on the cellular structures in the embryo, and the heptane facilitates entry of the fixative into the embryo through the vitelline membrane. 4. Discard PEM/formaldehyde waste according to the appropriate guidelines. 5. A significant number of embryos do not pop out of their vitelline membranes, and therefore do not fall into the methanol. These cannot be retrieved and should be discarded. Methanol and heptane waste can be discarded together but must be separated from PEM/formaldehyde waste. 6. After removal of heptane, embryos can be stored in MeOH in a vial at −20°C indefinitely. 7. All washes and incubations should be carried out using a rotator to keep the embryos suspended in the solutions. Steps are carried out at room temperature, unless they are overnight steps, which are carried out at 4°C on the rotator. For each change of solution, simply remove the tube of embryos from the rotator, allow them to settle for about 1 min, then remove as much of the solution as possible using a glass Pasteur pipette. During all of these steps, it is important that the samples are not allowed to dry out. 8. For low affinity antibodies, use final dilutions in the range between 1:5 and 1:200. For high affinity antibodies, use final

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dilutions in the range between 1:1,000 and 1:5,000. If doublefluorescent antibody staining, both primary antibodies can be used at the same time. If double-staining with DAB colorimetric detection, use only one primary antibody at a time. Primary antibodies raised in closely related animals may cross-react when adding secondary antibodies (step 11). Used primary antibody can be saved after incubation, and stored at 4°C with 2 PL of 3% sodium azide solution. 9. Secondary antibodies are typically manufactured and can be used in dilution ranges of 1:1,000–1:5,000. Use secondary antibodies appropriate for the animal in which primary antibodies were raised. 10. If double-DAB staining, use also 1 drop of NiCl2 with the first primary antibody, do not use it with the second primary antibody. Wear gloves when handling DAB, which is highly toxic. 11. DAB is toxic, discard in an appropriate waste container. 12. Heat killing of larvae is recommended since it prevents the larva from squirming during the dissection process. However, heat-killing is not compatible with staining for endogenous (or introduced) enzyme activity, such as X-gal staining to assess transgenic expression of a lacZ reporter. In the latter instance, larvae can be temporarily immobilized by placing on ice. 13. For pupae, it is important to first make a small snip at the posterior end of animal prior to cutting off the entire end. This will allow some hemolymph to leak out, thus relieving pressure from the pupal case. After pressure is released, end may be cut off. 14. When double-fluorescent antibody staining, both primary antibodies can be used at the same time. Primary antibodies are raised in a specific host animal, if staining multiple targets, make sure antibodies raised in different hosts are used. Closely related host species used for primary antibody generation may cross react when adding secondary antibodies (step 10). If primary antibody is to be saved after incubation, remove from embryos/larvae/pupae and store at 4°C with 35 PL of 3% sodium azide solution. Caution: sodium azide is highly toxic. 15. Secondary antibodies are typically purchased from commercial sources and can be used in ranges of 1:1,000–1:5,000. Use secondary antibodies appropriate for the animal in which primary antibodies were raised. 16. This same dissection and fixation procedure can be used for immunostaining of imaginal discs. After fixation, immunostaining will commence at the first PBTx wash step and continue as described for embryos. 17. The proteinase K solution, if too concentrated, can disrupt the samples. Some titration of different batches of proteinase K may be necessary.

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18. Drying the freezing media around the fly will ensure better adhesion to the sticky tape and prevent displacement of the fly during freezing. 19. Samples can be indefinitely stored at −80°C in a plastic zip bag or tightly closed container. Before sectioning, the samples should be removed from the −80°C freezer, and equilibrated to the temperature of the cryomicrotome. Typically, we set this to −18°C. 20. Before starting to section, transfer the information from sample label to the slide that is used for section collection. 21. Avoid storing sections that need to be stained with X-gal, as this may adversely affect beta-galactosidase activity and ruin reproducibility. Moisture negatively affects the quality of the section; to avoid condensation on slides, prewarm the storage box to ambient temperature before opening. 22. Once you have started the staining procedure, do not allow sections to dry out. Always keep the slide in solution until you are ready to move it to the next solution. 23. Longer fixation times can interfere with beta-galactosidase enzymatic activity, so avoid keeping your slide in formaldehyde solution for more than 10 min. 24. For high levels of ß-galactosidase expression, we observe staining in 5 min; for low expression levels, it can take 1 h or more. If blue stain is not visible after 1 h of incubation, the reaction can be left at room temperature overnight. Overnight incubation may lead to significant nonspecific staining and thereby should be interpreted with caution. 25. To create a humid box, lay filter paper at the bottom of a 150-mm Petri dish, soak with dH2O, and place into the dish two halves of a 2-mL disposable serological pipette. The pipette halves serve as rails on which slides sit in the chamber up above the filter paper. 26. Once you immerse sections in solution, do not let them dry out. Always keep your slide in solution until you are ready to move to a fresh solution. 27. For some epitopes, these parameters should be shortened to 2% formaldelyde for 5 min. 28. The final dilution of antibody depends on its affinity. If you are using antibody for the first time, it is better to titer it in the range between 1:100 and 1:1,000 and in the following experiments use the dilution that came out the best (i.e., strongest staining signal to lowest background signal). The researcher can stain different targets with different antibodies at the same time. In this case, make sure that primary antibodies were raised in different animal hosts, and use proper secondary antibodies with different fluorochromes in step 8.

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29. The cover slip from the blocking step (step 4) can be removed by immersing the slide in a beaker containing PBTx, at which point the coverslip should simply slide off the sections. 30. Secondary antibodies are sold by different manufacturers. Follow the manufacturer’s suggestions for working dilution, or titer the antibody as described earlier. Phalloidin binds to and highlights polymerized actin, serving as a general muscle marker; DAPI is useful as a counter-stain for nuclear antigens. 31. Samples can be placed on a rotator or any other agitating device to keep a constant fluid movement around specimens. Thoraces generally do not sink in the fixative, but this does not hamper fixation. 32. Although this step is optional, agarose embedding will substantially help to properly orient samples when sectioning. The base of the agarose triangle may be aligned to any part of the thorax, depending on what sectioning plane the investigator would like to achieve. 33. During the dehydration steps, specimens can be stored for several days in 70% ethanol. For complete dehydration, the quality of absolute ethanol is important: traces of water will cause white precipitation during paraffin embedding and artifacts on sections (e.g., specimen crumbling). To prevent absolute ethanol from absorbing water from the air, make sure all lids on ethanol containers are always tightly closed. Before the absolute ethanol wash, change the cap on the scintillation vial for a new one. 34. Specimens first float near the surface of the xylene, but then sink down as the xylene penetrates. Paraffin can be kept liquid in an incubator set to 63°C. Watch for white precipitation in xylene washes. If they appear, repeat the last dehydration steps using fresh absolute ethanol. Paraffin can be obtained from a number of sources, e.g., Paraplast Plus (Sigma). 35. To avoid bubbles, orient specimens after 20 min, keeping the mould on warming plate. 36. Pressure cookers generate high temperatures, steam, and high pressures. Use due caution when handling, and observe all safety guidelines included with the cooker.

Acknowledgments Research in the Cripps laboratory is funded by grants from the National Institutes of Health, the American Heart Association, the March of Dimes Birth Defects Foundation, and the Muscular Dystrophy Association. GRM and EML were supported by the NIGMS/IMSD award GM060201. EML is supported by a predoctoral fellowship from the AHA.

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References 1. Baylies MK and Michelson AM (2001) Invertebrate myogenesis: looking back to the future of muscle development. Curr Opin Gen Dev 11:431–439 2. Bate M. (1990) The embryonic development of larval muscles in Drosophila. Development 110:791–804 3. Artero R, Furlong EE, Beckett K, Scott MP, and Baylies, M. (2004) Notch and Ras signaling pathway effector genes expressed in fusion competent and founder cells during Drosophila myogenesis. Development 130:6257–6272 4. Stute C, Schimmelpfeng K, Renkawitz-Pohl R, Palmer RH, and Holz A (2004) Myoblast determination in the somatic and visceral mesoderm depends on Notch signaling as well as on milliways (milliAlk) as receptor for Jeb signaling. Development 131:743–754 5. Chen EH and Olson EN (2004) Towards a molecular pathway for myoblast fusion in Drosophila. Trends Cell Biol 14:452–460 6. Rushton E, Drysdale R, Abmayr SM, Michelson AM, and Bate M (1995) Mutations in a novel gene, myoblast city, provide evidence in support of the founder cell hypothesis for Drosophila muscle development. Development 121:1979–1988 7. Ruiz-Gomez M, Coutts N, Price A, Taylor MV, and Bate M (2000) Drosophila dumbfounded: a myoblast attractant essential for fusion. Cell 102:189–198 8. Tixier V, Bataillé L, and Jagla K (2010) Diversification of muscle types: recent insights from Drosophila. Exp Cell Res 316:3019–3027 9. Haralalka S, and Abmayr SM (2010) Myoblast fusion in Drosophila. Exp Cell Res 316:3007–3013 10. Laurin M, Fradet N, Blangy A, Hall A, Vuori K, and Cote J (2008) The atypical Rac activator Dock180 (Dock1) regulates myoblast fusion in vivo. Proc Natl Acad Sci 105:15446–15451 11. Menon SD, Osman Z, Chenchill K, and Chia W (2005) A positive feedback loop between Dumbfounded and Rolling pebbles leads to myotube enlargement in Drosophila. J Cell Biol 169:909–920 12. Schroter RH, Lier S, Holz A, Bogden S, Klambt C, Beck L, and Renkawitz-Pohl R (2004) kette and blown fuse interact genetically during the second fusion step of myogenesis in Drosophila. Development 131:4501–4509 13. Currie DA and Bate M (1991) The development of adult abdominal muscles in Drosophila: myoblasts express twist and are associated with nerves. Development 113:91–102

14. Baylies MK, Martinez Arias A, and Bate M (1995) wingless is required for the formation of a subset of muscle founder cells during Drosophila embryogenesis. Development 121:3829–3837 15. Sandmann T, Jensen LJ, Jakobsen JS, Karzynski MM, Eichenlaub MP, Bork P, and Furlong EE (2006) A temporal map of transcription factor activity: mef2 directly regulates target genes at all stages of muscle development. Dev Cell 10:797–807 16. Lin M-H, Nguyen HT, Dybala D, Stroti RV (1996) Myocyte-specific enhancer factor 2 acts co-operatively with a muscle activator region to regulate Drosophila tropomyosin gene muscle expression. Proc Natl Acad Sci USA 93:4623–4628 17. Kelly KK, Meadows SM and Cripps RM (2002) Drosophila MEF2 is a direct regulator of Actin57B transcription in cardiac, skeletal and visceral muscle lineages. Mech Dev 110: 39–50 18. Kelly Tanaka KK, Bryantsev AL and Cripps RM (2008) Myocyte enhancer factor-2 and Chorion factor-2 collaborate in activation of the myogenic program in Drosophila. Mol Cell Biol 28:1616–1629 19. Demontis F and Perrimon N (2009) Integration of Insulin receptor/Foxo signaling and dMyc activity during muscle growth regulates body size in Drosophila. Development 136:983–993 20. Bate ME, Rushton E and Currie DA (1991) Cells with persistent twist expression are the embryonic precursors of adult muscles in Drosophila. Development 113:79–89 21. Rivlin PK, Schneiderman AM, and Booker R (2000) Imaginal pioneers prefigure the formation of the adult thoracic muscles in Drosophila melanogaster. Dev Biol 222:450–459 22. Fernandes J, Bate M, and VijayRaghavan K (1991) Development of the indirect flight muscles of Drosophila. Development 113: 67–77 23. Dutta D, Anant S, Ruiz-Gomez M, Bate M, and VijayRaghavan K (2004) Founder myoblasts and fibre number during adult myogenesis in Drosophila. Development 131:3761–3772 24. Atreya KB and Fernandes JJ (2008) Founder cells regulate founder number but not fiber formation during adult myogenesis in Drosophila. Dev Biol 321:123–140 25. Bernstein SI, O’Donnell PT, and Cripps RM (1993) Molecular genetic analysis of muscle development, structure, and function in

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Drosophila., pp. 63–152 in International Review of Cytology. Academic Press, Inc. 26. Cripps R.M. and Olson EN (1998) Twist is required for muscle template splitting during adult Drosophila myogenesis. Dev Biol 203:105–116 27. Patel NH (1994) Imaging neuronal subsets and other cell types in wholemount Drosophila embryos and larvae using antibody probes. Methods Cell Biol. 44:445–487 28. Molina MR and Cripps RM (2001) Ostia, the inflow tracts of the Drosophila heart, develop from a genetically distinct subset of cardial cells. Mech Dev 109:51–59

29. O’Neill JV and Bier E (1994) Double-label in situ hybridization using biotin and digoxigenintagged RNA probes. BioTechniques 17: 874–875 30. Cripps RM, Black BL, Zhao B, Lien C-L, Schulz RA, and Olson EN (1998) The myogenic regulatory gene Mef2 is a direct target for transcriptional activation by Twist during Drosophila myogenesis. Genes Dev 12: 422–434 31. Lyons GE, Schiaffino S, Barton P, Sassoon D, and Buckingham M. (1990) Developmental regulation of myosin gene expression in mouse cardiac muscle. J Cell Biol 111:2427–2436

Chapter 9 Immunocytochemistry to Study Myogenesis in Zebrafish Nathan C. Bird, Stefanie E. Windner, and Stephen H. Devoto Abstract During myogenesis, cells gradually transition from mesodermal precursors to myoblasts, myocytes, and then to muscle fibers. The molecular characterization of this process requires the ability to identify each of these cell types and the factors that regulate the transitions between them. The most versatile technique for assaying cell identities in situ is immunocytochemistry, because multiple independent molecular markers of differentiation can be assayed simultaneously. The zebrafish has developed into a popular model for the study of myogenesis, and immunocytochemical techniques have been critical. We have adapted existing protocols to optimize immunocytochemistry in zebrafish, and have tested many antibodies developed against mouse, chick, and frog muscle antigens for their cross-reactivity in zebrafish. Here, we present protocols for whole mount immunocytochemistry on both formaldehyde and Carnoy’s fixed embryos as well as on sectioned zebrafish tissue. We include a table of antibodies useful for experiments on the molecular biology of myogenesis in zebrafish. Key words: Muscle, Development, Myotome, Dermomyotome, Differentiation, Slow fiber, Fast fiber

1. Introduction Proteins are the principal players that carry out cellular function, and thus the spatial and temporal distribution of proteins specific to differentiation and physiological functions shows the patterns of myogenesis and the distribution of different types of muscle fibers. The characterization of protein distribution in the animal is best done by the technique of immunocytochemistry. This is especially true in small model organisms such as zebrafish. The zebrafish has emerged as a powerful model for studying myogenesis. It is easy to observe embryonic myogenesis in a live, developing embryo, it is possible to do large-scale genetic screens for mutations affecting muscle development, and it is

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easy to do embryological manipulations such as lineage tracing and transplantations. Immunocytochemistry in zebrafish has both advantages and challenges, when compared with that done in other species. The small size and optical transparency make it optimal for microscopy, and the relative simplicity of tissue organization simplifies cell type identification. On the contrary, the small size poses challenges for making sections for immunolabeling, and few publicly available antibodies have been made against zebrafish myogenic proteins. In addition, antibody penetration into the embryos and larvae is more challenging, probably because zebrafish develops functional muscle very early and because they need to shield themselves from the external environment in which they develop. Immunocytochemical techniques have been developed for work both in whole mount and on sections for many different species used to study myogenesis, including mouse (1), chick (2), frog (3), and Drosophila (4). Below, we present an immunocytochemistry protocol for labeling multiple components of muscle development in whole mount and sectioned zebrafish embryos. These protocols have been adapted from these and other earlier immunohistochemistry protocols (5–7). The two most commonly used fixatives for zebrafish embryos are Carnoy’s fixative and 4% paraformaldehyde. Carnoy’s fixative preserves cellular architecture by denaturing proteins via dehydration and acidification, causing them to become insoluble. Paraformaldehyde, in contrast, covalently modifies amine residues on proteins, causing them to become insoluble and in many cases crosslinked to other amine-containing compounds. The preservation of cellular structure varies between fixatives, but both preserve the essential cytoarchitecture of muscle and muscle precursors sufficiently for light microscopic observations. However, the two fixatives sometimes have different effects on the tertiary structure of individual proteins, and the epitope recognized by a specific antibody may be destroyed by one fixative but not by another (e.g., the fast muscle marker zm4 works with Carnoy’s, but not with paraformaldehyde fixation). Thus, identification of appropriate fixatives is important, and influences further aspects of the protocol. Another key issue for whole mount immunocytochemistry is permeabilization. Live, the external vitelline layer (EVL) forms a protective barrier around the embryo. After fixation, the EVL as well as the dense cytoskeletal and extracellular matrices, especially prevalent in differentiated muscle, block large molecules such as immunoglobulins from entering the embryo and from diffusing within it. Thus, successful whole mount immunocytochemistry in zebrafish requires methods to enhance penetration. We use methanol to dissolve lipids away, it may also loosen aggregations between proteins, making bigger holes in the embryo and the intra and extra-cellular protein meshworks. Detergents (Tween-20, Triton-X)

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have a similar effect, as well as reducing nonspecific binding of antibodies to the embryo. Proteases digest proteins, loosening up the dense protein meshworks. We find that methanol followed by Tween-20 treatment is sufficient to permeabilize embryos, until some time between 36 and 60 h, after which we use partial Proteinase K digestion. In addition to the protocols outlined here, we include a table of antibodies that label muscle tissue in zebrafish, and the conditions under which each gives clear and repeatable results. We and others post information on antibodies, including those that did not work in our lab, in an antibody Wiki (available as part of the Zebrafish Model Organism Database, http://zfin.org).

2. Materials 2.1. Fixatives

1. 4% Paraformaldehyde: 1 vial 16% formaldehyde (10 mL), 10 mL 0.2M Phosphate Buffer (see below), 20 mL ddH20 in a 50-mL conical tube (see Note 1). Store at 4ºC. Good for up to 1 month. Do not autoclave. 2. Carnoy’s Fixative: 60% ethanol, 30% chloroform, 10% glacial acetic acid. Do not autoclave.

2.2. Buffers and Reagents

1. Methanol (100 and 50%): dilute with ddH2O. 2. Ethanol series (95, 75, 50, 25%): dilute with ddH2O. 3. 10× Phosphate buffered saline (PBS): for 1 L, add 80 g NaCl, 2.0 g KCl, 11.4 g Na2HPO4, 2.6 g KH2PO4 to ddH2O. pH to 6.8 with either 5M HCl or 5M NaOH. Autoclave. Store at room temperature (RT). Good for several months. 4. 1× PBS: Dilute 10× PBS (above) tenfold in ddH20. Store at RT. Good for several months. 5. PBS-Tween-20 (PBS-Tw): 1× PBS + 0.1% Tween-20. Store at RT. Good for several months. 6. PBS-Tw-Bovine serum albumin (PBS-Tw-B): PBS-Tw + 2% bovine serum albumin (BSA). Fill 50 mL conical tube to 50 mL mark with PBS-Tw. Add 1 g BSA. Store at 4ºC. Good for up to 2 months. 7. PBT-B-Normal goat serum (PBS-Tw-B-N): PBS-Tw-B + 5% normal goat serum (NGS). Make up 2–10 mL of PBS-TwB-N (e.g., 3 mL PBS-Tw-B + 150 ML NGS). Store at 4ºC. Should be used within 2 weeks. 8. Normal goat serum: Sigma G-9023 (see Note 2). 9. Proteinase K (1,000× stock, 10 mg/mL): Add 100 mg of Proteinase K to 10 mL ddH2O. Store at −20ºC in 250 ML aliquots.

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10. Proteinase K (working, ProtK, 10 Mg/mL): Dilute stock ProtK 1:1,000 in PBS-Tw (10 ML stock added to 10 mL PBS-Tw). 11. 0.2M phosphate buffer: Combine 80 mL of 0.2M Na2HPO4 with 20 mL NaH2PO4. Adjust the pH to between 7.0 and 7.3, autoclave, and store at RT; good for several months. 12. Stock Tricaine (MS-222; 3-aminobenzoid acid ethyl ester): Add 2.1 mL of 1M Tris buffer (pH 9.5, below) to a 100 mL graduated cylinder. Fill to 100 mL with ddH2O. Add solution to a bottle, then add 0.4 g Tricaine. Mix well, and check pH, should be around 7. Do not autoclave. Store at 4ºC. Good for 2–4 weeks. 13. 50% Glycerol: Add molecular grade glycerol to PBS-Tw. Rock gently to combine. Lasts for several months. 14. Hoechst solution: Used to fluorescently label nuclei. Dilute stock Hoechst 1:10,000 in PBS-Tw. Store at 4ºC, protected from light. 15. Embedding agar for cryostat sectioning: Weigh 1.5 g agarose and 5.0 g sucrose and add to an Erlenmeyer flask. In a graduated cylinder, measure 100 mL of 0.1M phosphate buffer. Add to flask and swirl gently to mix. Microwave for 1 min on high setting, then swirl to mix (contents will be hot, so wear heat resistant gloves at this point). Microwave an additional 50 s. If solids are not dissolved, microwave in additional 10 s increments until dissolved. Distribute 4 mL aliquots into small glass vials, label, and store at 4ºC. Good for several months. 16. 30% Filtered sucrose: Add 150 g sucrose to 300 mL of 1× PBS in an Erlenmeyer flask. Stir on moderate speed and low temperature until the sucrose has completely dissolved. Pour solution into graduated cylinder, and add 1× PBS to a final volume of 500 mL. Filter and store at 4ºC. Good for up to a month. 17. 1M Tris buffer: Add 121.14 g Tris base to 800 mL ddH2O, and stir until dissolved. Adjust the pH to desired level with HCl, then add ddH2O to final volume of 1 L, autoclave; Good for several months. 2.3. Antibodies

1. Primary antibodies: Stock primary antibodies should be diluted in PBS-Tw-B-N to 5 Mg/mL concentration for use (see Note 3). Primary antibodies are stable at 4ºC for at least several months if the antibody contains sodium azide. See Table 1 for a list of myogenesis-related primary antibodies that work in zebrafish. 2. Secondary antibodies: These secondary antibodies are available from several sources, depending on the desired detection method (see Note 4). Secondary antibodies should be diluted in PBS-Tw-B-N immediately prior to use (see Note 4).

MyHC (bovine)

BA-D5 (19)

Myotome borders (23)

Myotome borders (29) Fast myofibrils (16)

A-Dystroglycan (human)

A-Sarcoglycan (human)

Cardiac actin (synthetic)

Desmin (chicken)

Dystrophin (human)

MyHC (chicken)

MyHC (chicken)

Anti-A-dystroglycan (22)

Anti-A-sarcoglycanb

Anti-cardiac actin (25)

Anti-desminb

Dystrophin (28)

EB165 (30)

F59 (31)

Slow myofibrils early, fast myofibrils late (14)

Muscle cytoskeleton (18, 27)

Myofibrils (26)

Myotome borders (24)

Cell membranesa (21)

B-Catenin (chicken)

Cell membranes (20)

a

Anti-B-catenina

B-Catenin (human/mouse)

Myofibrils (18)

Sarcomeric A-actinin

Anti-A-actinin (17)

Anti-B-catenin

Myofibrils (16)

MyHC (human)

A4.1025 (15)

Slow myofibrils (16)

Fast muscle (12, 14)

Ca2+ channel (Newt)

12/101 (13)

a

Slow and fast muscle membranes (12)

Titin (bovine)

9 D10 (11)

Mouse IgG1

Mouse IgG1

Mouse IgG1

Rabbit Ig

Mouse IgG1

Mouse IgG1

Mouse IgG1

Mouse IgG1

Rabbit Ig

Mouse IgG2b

Mouse IgG1

Mouse IgG2a

Mouse IgG1

Mouse IgM

Mouse IgG1

Muscle pioneer nucleia (9, 10)

Engrailed (Drosophila)

4D9 (8)

Ab isotype

Labeling in Zebrafish muscle

Antigen

Primary antibody

Table 1 Listing of available antibodies used in the study of zebrafish myogenesis

DSHB (F59)

DSHB (EB165)

Sigma (D8043)

Sigma (D8281)

Progen Biotechnik (65175)

Novocastra (NCL-L-b-SARC)

DSHB (MANDAG2 clone 7D11)

Sigma (C7207)

Sigma (C2206)

DSHB (BA-D5)

Sigma (A7811)

DSHB (A4.1025)

DSHB (12/101)

DSHB (9 D10)

DSHB (4D9)

Company (Cat #)

(continued)

PFA

PFA

PFA

PFA

PFA

PFA

PFA

PFA

PFA

PFA

PFA

PFA

PFA

Carnoys

PFA

Fixative

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Mouse IgM Mouse IgG1

Mouse IgA Mouse IgG1

Differentiated muscle nucleia (26) Myofibrils (12)

Differentiated muscle nuclei (37) Preferentially slow myofibrils (12) Dermomytomea (37) Slow muscle nuclei (41) Slow myofibrils (14) Synaptic vesiclesa (43, Devoto Lab Unpublished)

Mef-2 (human)

MyHC (chicken)

Myf-5 (human); MyoD (zebrafish)

Myogenin (rat)

MyHC (human)

Pax-7 (chicken)

Prox 1 (mouse)

Slow MyHC (chicken)

Keratan sulfate proteoglycan (Ray)

Tropomyosin (chicken)

MF 20 (35)

Myf-5

Myogenin

N1.551 (15)

Pax-7 (38, 39)

Prox 1 (40)

S58 (31)

SV2 (42)

Anti-tropomyosinb

Myofibrils (26)

a

Differentiated muscle nuclei (36)

Mouse IgG1

Rabbit Ig

Rabbit Ig

Rabbit Ig

Mouse IgG2b

Rabbit Ig

Rabbit Ig

Mef-2 (34)

Myotome bordersa (21)

Laminin (rat)

Rabbit Ig

Rabbit Ig

Mouse IgG1

Anti-lamininb

Myotome bordersa (23)

Myotome borders (21)

a

Ab isotype

Laminin (mouse)

Fibronectin (human/mouse)

Myofibrils (33)

Labeling in Zebrafish muscle

Anti-lamininb

Anti-fibronectin

MyHC (chicken)

F310 (32)

c

Antigen

Primary antibody

Table 1 (continued)

Sigma (T9283)

DSHB (SV2)

DSHB (S58)

Millipore (AB5475)

DSHB (Pax7)

DSHB (N1.551)

Santa Cruz (sc-576)

Santa Cruz (sc-302)

DSHB (MF 20)

Santa Cruz (sc-313)

Lab Vision (RB-082-A0)

Sigma (L9393)

Lab Vision (RB-077-A0)

DSHB (F310)

Company (Cat #)

PFA

Any Fixative

Carnoys or PFA

PFA

PFA

Carnoys

PFA

PFA

Any Fixative

PFA

PFA

PFA

PFA

PFA

Fixative

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Utrophin (human)

Utrophin (human)

Unknown

Activated leukocyte cell adhesion molecule A

Anti-utrophin (44)

Anti-utrophin (45)

Zm4d

Zn5 (48)

Mouse IgG1 Mouse IgG1

Fast muscle fibers (47) Slow muscle membranesa (14)

Mouse IgG1

Goat Ig

Myotome bordersa (24) Myotome borders (46)

Ab isotype

Labeling in Zebrafish muscle

ZIRC (zn-5)

ZIRC (zm-4)

Novocastra (NCLDRP2)

Santa Cruz (sc-7460)

Company (Cat #)

PFA

Carnoys

PFA

PFA

Fixative

Included are the antibody names (with reference to its developer or first known use), antigen, brief description of zebrafish muscle labeling pattern (with first use in zebrafish), antibody isotype, source and product number, and recommended fixative. Many of the included antibodies have been used extensively in zebrafish (e.g., F59, MF20, S58) and due to space limitations, it was not possible to include all available references. More complete reference lists, along with lists of other antibodies used in zebrafish, can be found at zfin.org PFA paraformaldehyde a This antibody also labels non-myogenic cells b First antibody usage not readily available from manufacturer c Anti-fibronectin (A-10) no longer appears in the catalog of the referenced supplier (Lab Vision) d Developed by Monte Westerfield at the University of Oregon

Antigen

Primary antibody

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2.4. Equipment (Nonstandard)

1. Single plane rotator, capable of slow rotation (single rotation every 10–15 s, see Note 5). 2. Vacuum apparatus (vacuum port, tubing, collection flask). 3. PAP pen. 4. Humidified chamber (for sectioned immunocytochemistry). Construct a humidified chamber by taping trimmed wooden applicator sticks to the bottom of a 150-mm petri dish, and add moistened Kim-wipes to the edges opposite the slides. Wrap the top and bottom of the dish in foil. 5. Plastic molds (for agar embedding and cryostat sectioning). 6. OCT compound (Tissue Tek).

3. Methods Embryos should be dechorionated first if they are older than 14 h, as fixation after this point in the chorion will lock the tails into a curled position (see Note 6). Embryos older than 24 h should be anesthetized prior to fixation using Tricaine (7). Volumes are 1 mL unless stated otherwise below. 3.1. Fixation and Storage

1. Move embryos from petri dish into an appropriately labeled 1.5-mL Eppendorf tube. 2. Remove embryo medium, and replace with desired fixative (4% paraformaldehyde, Carnoy’s, etc.). 3. Fix at RT for 2 h, rotating (see Note 7). 4. If Carnoy’s was used, embryos can be stored at −20°C indefinitely. If formalin-based fixative is used, continue to step 5. 5. Remove paraformaldehyde from all tubes and rinse with PBS (see Notes 8 and 9). 6. Remove PBS and replace with fresh PBS. Incubate at RT for 5–15 min, rotating. 7. Repeat step 6. 8. Remove PBS, and replace with 100% methanol. 9. Store embryos at −20°C for at least 5 min. Embryos can be stored indefinitely at −20°C at this point.

3.2. Whole Mount Immunocytochemistry: Paraformaldehyde Fixed (Fig. 1)

1. Remove tubes from −20°C storage. If all the embryos will not be used for one labeling, transfer to separate tube(s) for labeling. 2. Remove 100% methanol, replace with 50% methanol. Incubate at RT for 1–5 min, rotating.

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Fig. 1. Example of whole-mount immunocytochemistry. Twenty-four hour embryo showing trunk somite. Dermomyotome nuclei are green, labeled with anti-Pax7 (Alexa 488), all myofibrils are red, labeled with MF20 (Alexa-546), and differentiated muscle nuclei are teal, labeled with anti-MEF2 (Alexa 647). Scale bar = 100 Mm.

3. Remove 50% methanol, and rinse with PBS-Tw. 4. Remove PBS-Tw, replace with PBS-Tw-B. Incubate at RT for 5 min, rotating. 5. Repeat step 4. 6. Remove the last PBS-Tw-B wash and replace with 200 ML of PBS-Tw-B-N. Incubate at RT for 10 min, rotating. 7. Remove PBS-Tw-B-N, replace with 200 ML of primary antibody diluted in PBS-Tw-B-N. Incubate for 2 h at RT, rotating (see Note 7). 8. Remove primary antibody and SAVE (see Note 3). Rinse specimens 3× in PBS-Tw. 9. Remove last PBS-Tw rinse, replace with fresh PBS-Tw. Incubate at RT for 10 min, rotating (see Note 10). 10. Repeat step 9 twice. 11. Remove the last PBS-Tw wash and add 200 ML of PBS-TwB-N. Incubate at RT for 10 min, rotating. 12. Remove PBS-Tw-B-N, replace with 200 ML of secondary antibody diluted in PBS-Tw-B-N (see Note 4). Incubate at RT for 2 h, covered in foil (in all subsequent steps), rotating. 13. Remove secondary antibody and discard. Rinse specimens 3× in PBS-Tw. 14. Remove last PBS-Tw rinse and replace with fresh PBS-Tw. Incubate at RT for 10 min, rotating. 15. Repeat step 14 twice (see Note 11). 16. Remove PBS-Tw, replace with 50% glycerol in PBS-Tw. Incubate at RT for 10 min, rotating.

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17. Store at 4°C in foil until imaging. 18. Embryos can be mounted between double-bridged cover slips, as described in Westerfield (7). 3.3. Whole Mount Immunocytochemistry: Carnoy’s Fixed

1. Remove embryos from −20°C. If all the embryos will not be used for one labeling, transfer to separate tube(s) for labeling. 2. Remove Carnoy’s (see Note 8), and replace with 95% ethanol (see Note 12). Incubate at RT for 1–5 min, rotating (see Note 10). 3. Remove 95% ethanol, and replace with 75% ethanol. Incubate at RT for 1–5 min, rotating. 4. Remove 75% ethanol, and replace with 50% ethanol. Incubate at RT for 1–5 min, rotating. 5. Remove 50% ethanol, and replace with 25% ethanol. Incubate at RT for 1–5 min, rotating. 6. Remove 25% ethanol and replace with PBS-Tw. Incubate at RT for 1–5 min, rotating. 7. If embryos are older than 24 h, partial proteinase digestion can help antibody penetration (see Note 13). 8. Remove PBS-Tw-B and replace with fresh PBS-Tw-B. Incubate at RT for 5 min, rotating. 9. Repeat step 10 twice. 10. Remove PBS-Tw-B and replace with 200 ML of PBS-Tw-B-N. Incubate at RT for 15 min, rotating. 11. Remove PBS-Tw-B and replace with 200 ML of primary antibody diluted in PBS-Tw-B-N. Incubate at RT for 2 h, rotating (see Note 7). 12. Remove primary antibody and SAVE (see Note 3). Rinse specimens 3× in PBS-Tw. 13. Remove last PBS-Tw rinse, replace with PBS-Tw. Incubate at RT for 10 min, rotating (see Note 10). 14. Repeat step 14 twice. 15. Remove the last wash and replace with 200 ML of PBS-TwB-N. Incubate at RT for 10 min, rotating. 16. Remove PBS-Tw-B-N and replace with 200 ML of secondary antibody diluted in PBS-Tw-B-N. Incubate at RT for 2 h, in foil (in all subsequent steps), rotating. 17. Remove secondary antibody (discard). Rinse 3× in PBS-Tw. 18. Remove last PBS-Tw rinse, replace with PBS-Tw. Incubate at RT for 10 min, rotating (see Note 11). 19. Repeat step 19 twice. 20. Remove PBS-Tw, replace with 50% glycerol in PBS-Tw. Incubate at RT for 10 min, rotating. 21. Store at 4°C in foil until imaging.

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1. To make sections, modified from ref. (7). (a) Remove embryos from fixative. If in Carnoy’s, rehydrate through ethanol series. (b) Wash 2–3 times in 1× PBS, 5 min each. (c) Melt embedding agarose aliquot by placing in boiling water bath, then place in 50ºC heating block or water bath to maintain temperature. (d) Transfer specimen(s) to a flexible plastic well (we use trimmed paraffin molds). (e) Remove buffer and add agar to cover. (f) Position the specimen(s) using wire tool or other fine instrument. (g) Allow block to solidify (approximately 2 min). (h) Pop the block out of the mold and trim using razor blade leaving about 2 mm around the specimen. (i) Place block in a vial filled with 30% filtered sucrose (in PBS), and store at 4°C until the block is fully infiltrated and sinks. (j) Mount the block on a layer of OCT on an object holder resting in crushed dry ice for rapid freezing. When block is frozen, place in cryostat, and allow to equilibrate to cryostat temperature for several minutes.

Fig. 2. Example of sectioned immunocytochemistry. Twenty-four hour embryo, sectioned through an anterior somite. Fast myofibrils are green, labeled with F59 (Alexa 488), all myofibrils are red, labeled with MF20 (Alexa-546), differentiated muscle nuclei are teal, labeled with anti-MEF2 (Alexa 647), and all nuclei are blue, labeled with Hoechst counterstain. Scale bar = 50 Mm.

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(k) Section on cryostat to desired thickness (typically between 12 and 20 Mm) and collect sections on coated slides. (l) Dry slides on hot plate (26°C). (m) Store slides at −20°C. 2. Remove slide(s) from −20°C storage. 3. Once dry, draw a hydrophobic barrier to separate individual sections on the slide, using PAP pen (see Note 14). 4. Place slide in humidified chamber. 5. Rehydrate section with PBS-Tw (see Note 15). 6. Remove PBS-Tw with vacuum apparatus, apply PBS-Tw-B-N blocking solution (do not let dry). 7. Remove PBS-Tw-B-N, apply primary antibody diluted in PBSTw-B-N (same concentration as in whole mount). Incubate at RT, covered, for 45 min. 8. Remove antibody, and rinse 3× with PBS-Tw. 9. Wash in PBS-Tw 2× for 3 min each. 10. Remove PBS-Tw, replace with PBS-Tw-B-N for 2 min. 11. Remove PBS-Tw-B-N, add secondary antibody diluted in PBS-Tw-B-N. Incubate at RT, covered, for 30 min (see Note 16). 12. Remove secondary antibody, rinse 3× with PBS-Tw. 13. Remove PBS-Tw, and replace with Hoechst Solution. Incubate at RT, covered, for 5 min. 14. Remove Hoechst Solution, and rinse 3× with PBS-Tw. 15. Wash in PBS-Tw 2× for 3 min each (see Note 11). 16. Remove PBS-Tw, and replace with 50% glycerol in PBS-Tw. 17. Coverslip (see Note 17). 18. Store in humidified slide box at 4°C (see Note 18). The labeling is stable for months, but the quality of the sections decays in time.

4. Notes 1. Formaldehyde varies in quality and ease of use. Premade 4% paraformaldehyde is convenient, but it often contains impurities, which may increase background labeling. Electron microscopy grade paraformaldehyde powder is ultra pure, but requires extra care to prepare a 4% solution. 2. Upon arrival, we distribute normal goat serum into 1 mL aliquots and store at −20ºC to minimize freeze–thaw cycles.

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Upon need, we thaw a single aliquot, store at 4ºC, and use for a few months. 3. For most primary antibodies, a concentration of 5 Mg/mL is sufficient to give strong labeling with minimal background. However, when using an untested antibody, we test a series of concentrations ranging from 0.5 to 50 Mg/mL to determine the concentration which gives the highest ratio of specific to nonspecific labeling. Used, diluted primaries give less background than unused primaries. 4. Upon arrival from the company, we distribute secondary antibodies into 25–50 ML aliquots, and store at −20ºC. Individual aliquots are thawed when needed, stored at 4ºC, and used for about a month. Immediately prior to every use, the aliquot should be micro-centrifuged at top speed for 5 s before dilution to pellet any aggregates, which might decorate section with flecks. Secondaries can also be filtered. Before using a new secondary antibody, we test a series of dilutions ranging from 1:50 to 1:5,000 to determine the dilution which gives the highest ratio of specific to nonspecific labeling. We most commonly use Alexa conjugated secondary antibodies (Invitrogen, Molecular Probes) at a dilution of 1:800 for both whole mount and sectioned immunocytochemistry with fluorescence. A standard combination in our research is a triple label with Alexa-488 (green), Alexa-546 (red), and Alexa-647 (far red). 5. Rotation helps the washing and incubation processes by continually turning over specimens to expose all surfaces, if it is slow and gentle there is little damage to the delicate embryos. 6. We prefer dechorionation by hand using forceps, but dechorionation can also be accomplished using a protease (7). 7. This step can be done at 4ºC overnight for convenience. 8. Paraformaldehyde is toxic, and should be disposed of in a proper waste receptacle, along with at least the first rinse post fixation. The chloroform in Carnoy’s fixative also must be properly disposed. 9. Embryos should have sunk to the bottom of the tube, but if any embryos remain on the top due to surface tension, take care to avoid losing embryos at wash steps. 10. The times of most washes in whole mount immunocytochemistry can be easily tripled without affecting the experiment if timing is an issue. We see no benefit from longer blockings or washings, however, neither does it hurt; labeling is completely unaffected. The only risk is that with very extensive incubations (e.g., if it takes 4 days from rehydration to the final wash), fragile embryos will begin to look bedraggled. If washes or incubations go for more than triple the times indicated, do them at 4ºC.

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11. At this point, if doing a color (diaminobenzidine, DAB) reaction instead of fluorescence: (i) After last PBS-Tw wash, apply DAB solution without H2O2 for 5 min. (ii) Remove DAB and apply DAB solution with H2O2. (iii) Monitor reaction under a dissecting scope. (iv) Stop reaction by washing with PBS-Tw. L

DAB stock: 4% DAB (40 mg/mL, or 1 g/25 mL; filtered and kept at −20ºC).

L

DAB solution without H2O2: 0.05% DAB in PBS-Tw (13 ML DAB stock in 1 mL of PBS-Tw).

L

DAB solution with H2O2: 0.05% DAB in PBS-Tw (13 ML DAB stock + 13 ML 0.3% H2O2 in 1 mL of PBS-Tw).

12. The concentrations of the ethanol series during the rehydration of Carnoy’s fixed embryos are not critical. For example, 70 or 80% ethanol can be substituted for 75%. 13. For embryos older than about 24 h, antibody penetration into the central part of the myotome can be incomplete. To enhance permeability, we use partial digestion with proteinase K. After rehydrating embryos, we incubate in working ProtK solution for a length of time that depends both on the age and the batch of Proteinase K. These serve as rough guidelines: 30 h, 5 min; 48 h, 40 min; 72 h, 75 min; 96 h, 90 min, 120 h, 105 min. Some antibodies no longer recognize their epitope if ProtK is used. 14. PAP pens are easy to use, reliable, and quick drying. We have also used nail polish or rubber cement (applied with a syringe) to isolate sections on a slide. 15. Most sections require between 30 and 50 ML of solutions (rinses and washes); however, for reagents in limited supply, 20 ML is usually more than necessary to cover a section. 16. Incubation in secondary antibody for longer than 1 h will increase background. 17. Place the slide onto a paper towel and tip slide up to wick out extra glycerol. Wipe excess glycerol from the slide with a wet paper towel. Excess glycerol both hinders the cover slip from making a good seal to the slide, and risks creating a mess at the microscope. 18. We use small black slide boxes with a moistened Kimwipe placed in the bottom to maintain humidity and prevent photobleaching.

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Acknowledgments This work was supported by an NIH grant to SHD (R01 HD044929). We thank other labs for sharing protocols, and all members of the Devoto Lab, past and present, who helped to test and refine this protocol. References 1. Joyner, A., and Wall, N. (2008) Immunohistochemistry of Whole-Mount Mouse Embryos, Cold Spring Harb. Protoc. doi:10.1101/pdb.prot4820. 2. Psychoyos, D., and Finnell, R. (2009) Method for whole mount antibody staining in chick, JoVE. 24. http://www.jove.com/index/details. stp?id=956, doi: 10.3791/956. 3. Lee, C., Kieserman, E., Gray, R. S., Park, T. J., and Wallingford, J. (2008) Whole-mount fluorescence immunocytochemistry on Xenopus embryos, Cold Spring Harb. Protoc. doi:10.1101/pdb.prot4957. 4. Ramachandran, P., and Budnik, V. (2010) Immunocytochemical staining of Drosophila larval body-wall muscles, Cold Spring Harb. Protoc. doi:10.1101/pdb.prot5470. 5. Harlow, E., and Lane, D. (1999) Using Antibodies: A Laboratory Manual, Cold Spring Harbor Laboratory Press, Cold Spring Harbor, NY. 6. van Raamsdonk, W., Tekronnie, G., Pool, C. W., and van de Laarse, W. (1980) An immune histochemical and enzymic characterization of the muscle fibres in myotomal muscle of the teleost Brachydanio rerio, Hamilton-Buchanan, Acta Histochem 67, 200–216. 7. Westerfield, M. (2000) The Zebrafish Book. A guide for the laboratory use of zebrafish (Danio rerio), 4th ed., University of Oregon Press, Eugene. 8. Patel, N. H., Martin-Blanco, E., Coleman, K. G., Poole, S. J., Ellis, M. C., Kornberg, T. B., and Goodman, C. S. (1989) Expression of engrailed proteins in arthropods, annelids, and chordates, Cell 58, 955–968. 9. Hatta, K., Schilling, T. F., BreMiller, R. A., and Kimmel, C. B. (1990) Specification of jaw muscle identity in zebrafish: correlation with engrailed-homeoprotein expression, Science 250, 802–805. 10. Hatta, K., Bremiller, R., Westerfield, M., and Kimmel, C. B. (1991) Diversity of expression of engrailed-like antigens in zebrafish, Development 112, 821–832. 11. Wang, S. M., and Greaser, M. L. (1985) Immunocytochemical studies using a monoclo-

nal antibody to bovine cardiac titin on intact and extracted myofibrils, J Muscle Res Cell Motil 6, 293–312. 12. Barresi, M. J., D’Angelo, J. A., Hernandez, L. P., and Devoto, S. H. (2001) Distinct mechanisms regulate slow-muscle development, Curr Biol 11, 1432–1438. 13. Kintner, C. R., and Brockes, J. P. (1984) Monoclonal antibodies identify blastemal cells derived from dedifferentiating limb regeneration, Nature 308, 67–69. 14. Devoto, S. H., Melancon, E., Eisen, J. S., and Westerfield, M. (1996) Identification of separate slow and fast muscle precursor cells in vivo, prior to somite formation, Development 122, 3371–3380. 15. Webster, C., Silberstein, L., Hays, A. P., and Blau, H. M. (1988) Fast muscle fibers are preferentially affected in Duchenne muscular dystrophy, Cell 52, 503–513. 16. Blagden, C. S., Currie, P. D., Ingham, P. W., and Hughes, S. M. (1997) Notochord induction of zebrafish slow muscle mediated by Sonic hedgehog, Genes Dev 11, 2163–2175. 17. Lazarides, E., and Burridge, U. (1975) A-Actinin: immunofluorescent localization of a muscle structural protein in non-muscle cells, Cell 6, 289–298. 18. Costa, M. L., Escaleira, R. C., Jazenko, F., and Mermelstein, C. S. (2008) Cell adhesion in zebrafish myogenesis: distribution of intermediate filaments, microfilaments, intracellular adhesion structures and extracellular matrix, Cell Motil Cytoskeleton 65, 801–815. 19. Schiaffino, S., Gorza, L., Sartore, S., Saggin, L., Ausoni, S., Vianello, M., Gundersen, K., and Lomo, T. (1989) Three myosin heavy chain isoforms in type 2 skeletal muscle fibres, J Muscle Res Cell Motil 10, 197–205. 20. Moore, C. A., Parkin, C. A., Bidet, Y., and Ingham, P. W. (2007) A role for the Myoblast city homologues Dock1 and Dock5 and the adaptor proteins Crk and Crk-like in zebrafish myoblast fusion, Development 134, 3145–3153.

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21. Henry, C. A., McNulty, I. M., Durst, W. A., Munchel, S. E., and Amacher, S. L. (2005) Interactions between muscle fibers and segment boundaries in zebrafish, Dev Biol 287, 346–360. 22. Pereboev, A. V., Ahmed, N., thi Man, N., and Morris, G. E. (2001) Epitopes in the interacting regions of B-dystroglycan (PPxY motif) and dystrophin (WW domain), Biochim Biophys Acta 1527, 54–60. 23. Hall, T. E., Bryson-Richardson, R. J., Berger, S., Jacoby, A. S., Cole, N. J., Hollway, G. E., Berger, J., and Currie, P. D. (2007) The zebrafish candyfloss mutant implicates extracellular matrix adhesion failure in laminin A2-deficient congenital muscular dystrophy, Proc Natl Acad Sci USA 104, 7092–7097. 24. Bassett, D. I., Bryson-Richardson, R. J., Daggett, D. F., Gautier, P., Keenan, D. G., and Currie, P. D. (2003) Dystrophin is required for the formation of stable muscle attachments in the zebrafish embryo, Development 130, 5851–5860. 25. Franke, W. W., Stehr, S., Stumpp, S., Kuhn, C., Heid, H., Rackwitz, H. R., Schnölzer, M., Baumann, R., Holzhausen, H. J., and Moll, R. (1996) Specific immunohistochemical detection of cardiac/fetal A-actin in human cardiomyocytes and regenerating skeletal muscle cells, Differentiation 60, 245–250. 26. Hinits, Y., and Hughes, S. M. (2007) Mef2s are required for thick filament formation in nascent muscle fibres, Development 134, 2511–2519. 27. Rowlerson, A., Radaelli, G., Mascarello, F., and Veggetti, A. (1997) Regeneration of skeletal muscle in two teleost fish: Sparus aurata and Brachydanio rerio, Cell Tissue Res 289, 311–322. 28. Ellis, J. M., Man, N. T., Morris, G. E., Ginjaar, I. B., Moorman, A. F., and van Ommen, G. J. (1990) Specificity of dystrophin analysis improved with monoclonal antibodies, Lancet 336, 881–882. 29. Parsons, M. J., Campos, I., Hirst, E. M., and Stemple, D. L. (2002) Removal of dystroglycan causes severe muscular dystrophy in zebrafish embryos, Development 129, 3505–3512. 30. Cerny, L. C., and Bandman, E. (1987) Expression of myosin heavy chain isoforms in regenerating myotubes of innervated and denervated chicken pectoral muscle, Dev Biol 119, 350–362. 31. Miller, J. B., Crow, M. T., and Stockdale, F. E. (1985) Slow and fast myosin heavy chain content defines three types of myotubes in early muscle cell cultures, J Cell Biol 101, 1643–1650.

32. Crow, M. T., and Stockdale, F. E. (1986) Myosin expression and specialization among the earliest muscle fibers of the developing avian limb, Dev Biol 113, 238–254. 33. Elworthy, S., Hargrave, M., Knight, R., Mebus, K., and Ingham, P. W. (2008) Expression of multiple slow myosin heavy chain genes reveals a diversity of zebrafish slow twitch muscle fibres with differing requirements for Hedgehog and Prdm1 activity, Development 135, 2115–2126. 34. De Angelis, L., Borghi, S., Melchionna, R., Berghella, L., Baccarani-Contri, M., Parise, F., Ferrari, S., and Cossu, G. (1998) Inhibition of myogenesis by transforming growth factor B is density-dependent and related to the translocation of transcription factor MEF2 to the cytoplasm, Proc Natl Acad Sci USA 95, 12358–12363. 35. Bader, D., Masaki, T., and Fischman, D. A. (1982) Immunochemical analysis of myosin heavy chain during avian myogenesis in vivo and in vitro, J Cell Biol 95, 763–770. 36. Hammond, C. L., Hinits, Y., Osborn, D. P., Minchin, J. E., Tettamanti, G., and Hughes, S. M. (2007) Signals and myogenic regulatory factors restrict pax3 and pax7 expression to dermomyotome-like tissue in zebrafish, Dev Biol 302, 504–521. 37. Devoto, S. H., Stoiber, W., Hammond, C. L., Steinbacher, P., Haslett, J. R., Barresi, M. J., Patterson, S. E., Adiarte, E. G., and Hughes, S. M. (2006) Generality of vertebrate developmental patterns: evidence for a dermomyotome in fish, Evol Dev 8, 101–110. 38. Ericson, J., Morton, S., Kawakami, A., Roelink, H., and Jessell, T. M. (1996) Two critical periods of Sonic Hedgehog signaling required for the specification of motor neuron identity, Cell 87, 661–673. 39. Kawakami, A., Kimura-Kawakami, M., Nomura, T., and Fujisawa, H. (1997) Distributions of PAX6 and PAX7 proteins suggest their involvement in both early and late phases of chick brain development, Mech Dev 66, 119–130. 40. Bagri, A., Gurney, T., He, X., Zou, Y. R., Littman, D. R., Tessier-Lavigne, M., and Pleasure, S. J. (2002) The chemokine SDF1 regulates migration of dentate granule cells, Development 129, 4249–4260. 41. Liew, H. P., Choksi, S. P., Wong, K. N., and Roy, S. (2008) Specification of vertebrate slowtwitch muscle fiber fate by the transcriptional regulator Blimp1, Dev Biol 324, 226–235. 42. Buckley, K., and Kelly, R. B. (1985) Identification of a transmembrane glycoprotein specific for secretory vesicles of neural and endocrine cells, J Cell Biol 100, 1284–1294.

9 43. Müller, J. S., Jepson, C. D., Laval, S. H., Bushby, K., Straub, V., and Lochmüller, H. (2010) Dok-7 promotes slow muscle integrity as well as neuromuscular junction formation in a zebrafish model of congenital myasthenic syndromes, Hum Mol Genet 19, 1726–1740. 44. Monaco, A. P. (1989) Dystrophin, the protein product of the Duchenne/Becker muscular dystrophy gene, Trends Biochem Sci 14, 412–415. 45. Bewick, G. S., Nicholson, L. V., Young, C., O’Donnell, E., and Slater, C. R. (1992) Different distributions of dystrophin and related proteins at nerve-muscle junctions, Neuroreport 3, 857–860.

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46. Böhm, S., Jin, H., Hughes, S. M., Roberts, R. G., and Hinits, Y. (2008) Dystrobrevin and dystrophin family gene expression in zebrafish, Gene Expr Patterns 8, 71–78. 47. Barresi, M. J., Stickney, H. L., and Devoto, S. H. (2000) The zebrafish slow-muscle-omitted gene product is required for Hedgehog signal transduction and the development of slow muscle identity, Development 127, 2189–2199. 48. Trevarrow, B., Marks, D. L., and Kimmel, C. B. (1990) Organization of hindbrain segments in the zebrafish embryo, Neuron 4, 669–679.

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Chapter 10 Immunofluorescent Localization of Proteins in Caenorhabditis elegans Muscle Kristy J. Wilson, Hiroshi Qadota, and Guy M. Benian Abstract Caenorhabditis elegans is a premier model genetic system for discovering new information about the assembly and maintenance of striated muscle. The localization of a protein within a nematode muscle cell can reveal important clues to its function. In C. elegans, proteins can be localized by two different methods at the light microscopy level: GFP tagged proteins and indirect immunofluorescence. Although there are advantages and disadvantages of each method, antibodies can be used to localize proteins expressed at endogenous levels and without tags that might interfere with function. Immunolocalization requires efficient and effective methods of fixation. Here, we describe in detail two different methods for fixation of adult worms, the Nonet method and the Constant Spring method. We also discuss the advantages and the disadvantages of each, and how to choose between them. These methods are also useful for localizing proteins expressed in other cell types. Key words: C. elegans, Muscle, Fixation, Immunostaining, Methods

1. Introduction Sarcomeres, highly ordered assemblages of several hundred proteins, perform the work of muscle contraction. Despite ever increasing knowledge of the components and functions of sarcomeric proteins, we do not have a clear picture about how sarcomeres are assembled, and maintained in the face of muscle contraction. Caenorhabditis elegans is an excellent model genetic system in which to investigate these questions. With this system, most often, a muscle component is identified by a mutation and the eventual molecular identification of the encoded protein, or by RNAi screens of known genes. The localization of a protein within C. elegans adult body wall muscle is an important component of this analysis (1–9). This is true for both determining the localization of a new component of the sarcomere, as well as characterizing the phenotype of a mutant Joseph X. DiMario (ed.), Myogenesis: Methods and Protocols, Methods in Molecular Biology, vol. 798, DOI 10.1007/978-1-61779-343-1_10, © Springer Science+Business Media, LLC 2012

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by localizing already known sarcomeric components. Localization can also validate protein–protein interactions that are determined by other methods. Proteins may be localized by two different methods at the light microscopy level: GFP tagged proteins and indirect immunofluorescence. Generation and localization of a GFP tagged protein can usually be completed sooner than generating and localizing an antibody, and GFP tagged proteins can be visualized in live animals. A disadvantage of GFP tagging is that the usual way the required transgenic worms are created leads to overexpression of the GFP fusion protein, and consequently the danger that the protein may localize to places other than its endogenous location. In addition, overexpression may sometimes affect function. Antibodies can be used to localize native proteins expressed at normal levels. Many monoclonal and polyclonal antibodies are available to known C. elegans sarcomeric proteins (10), and as new antibodies are discovered they are detailed on http://www.wormbase.org/ with the information found for each individual gene or protein. Immunostaining requires efficient and effective methods of fixation. Here, we describe in detail two different methods for C. elegans fixation, the Nonet method (11) and the Constant Spring method (12, 13). Although both of these fixation methods were originally developed for neuronal staining, they are also effective for the fixation of body wall muscle and many other tissues and cell types. The Nonet method of fixation has many advantages including ease of use and results in usually sharper sacromeric striations upon immunostaining, possibly due to faster penetration and fixation. One disadvantage of the Nonet method, however, is that it destroys the fluorescence signal from GFP. In contrast, the Constant Spring method preserves the GFP signal, but often results in less sharp localization by immunostaining (see Fig. 1) (4). Table 1 compares and contrasts the Nonet and Constant Spring methods. Thus, whenever possible we use Nonet fixation but we have found that this method does not always work with monoclonal antibodies. For example, when staining with six different monoclonal antibodies (KT3, KT6, KT9, KT10, KT11, and KT12) all worked with the Nonet method except for KT11 (14). In these situations, we perform Constant Spring Fixation. Thus, both Nonet method and Constant Spring fixation methods are important tools for evaluating C. elegans muscle and for understanding protein function.

2. Materials Prepare all solutions using deionized water and analytical grade reagents. Prepare and store all reagents at 4°C (unless otherwise indicated). Follow all local waste disposal regulations when disposing of waste materials.

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Fig. 1. Comparison of immunolocalization after Nonet or Constant Spring fixation. (a) Nonet method fixation of wild type worms with anti-UNC-94 (TMD-1) and either anti-UNC-89 (obscurin) or anti-A-actinin. The Nonet method shows clearly that UNC-94 (TMD-1) localizes to two closely spaced parallel lines flanking the M-line. (b) Constant-spring method fixation of wild type worms with anti-UNC-94 (TMD-1) shows broad I-band localization with no hint of the lines revealed by the Nonet method. The localization obtained with the Nonet method is consistent with likely localization to the pointed ends of the thin filaments and the known biochemical activity of other tropomodulins. Figure modified from (4).

Table 1 A comparison of two methods for Caenorhabditis elegans fixation Nonet method

Constant Spring method

Freezing worms during fixation

Yes

Yes

Time for experienced user

~5 h

~7 h

Preserves GFP signal

No

Yes

Sharpest striations

Yes

No

Store worms for at least a month after fixation

Yes

Yes

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2.1. Solutions

1. 1M MgSO4: Weigh out 246.48 g MgSO4 .7H2O and dissolve in water, bringing the final volume to 1 L. Sterile filter and store at room temperature. 2. M9: Weigh out 6 g Na2PO4, 3 g KH2PO4, and 5 g NaCl, and dissolve in water to a final volume of 1 L. Autoclave, let solution cool, and then add 1 mL of sterile 1 M MgSO4, and store at room temperature. 3. Bouin’s fixative: Mix together 75 mL saturated picric acid (see Note 1), 25 mL formalin, and 5 mL glacial acetic acid. 4. 20% Triton X-100: Weigh out 50 g Triton X-100 liquid into a 250 mL bottle (see Note 2). Using a graduated cylinder add 200 mL of water. Using a stir rod allow Triton X-100 to dissolve. This may take as long as overnight, so if that is the case tighten the lid to prevent evaporation. 5. 50× Borate buffer (BO3): 1M H3BO3 and 0.5M NaOH with the pH being not less than 9.5 (see Note 3). Add approximately 70 mL of water to a small plastic beaker. Weigh out 3.1 g boric acid (H3BO3) and add to the beaker, using a stir bar to dissolve. Add 5 mL of 5M NaOH and then using a plastic 100 mL cylinder bring solution to 100 mL with water. 6. BT: 1× borate buffer and 0.5% Triton X-100. Add 10 mL of 50× borate buffer and 12.5 mL of 20% Triton X-100 and using a graduate cylinder bring volume to 500 mL with water. 7. BTB: 1× BT + 2% B-mercaptoethanol. 8. Phosphate buffered saline (PBS): For 1 L of 10× PBS add 2 g KCl, 2 g KH2PO4, 11.5 g Na2HPO4·7H2O, and 80 g NaCl. Check that the pH is 7.2. Sterile filter. Do not autoclave. 9. Antibody Buffer A (AbA): 1× PBS, 1% bovine serum albumin (BSA), 0.5% Triton X-100, 1 mM NaN3, and 1 mM EDTA (see Note 4). 10. 4× MRWB: 320 mM KCl, 80 mM NaCl, 40 mM EGTA pH 7.4, 20 mM spermidine, 60 mM PIPES pH 7.4. Add 5.3 mL of 3M KCl, 800 ML of 5M NaCl, 20 mL of 0.1M EGTA, 1 mL of 1M spermidine, and 6 mL of 0.5M PIPES pH 7.4. Bring to 50 mL with water. 11. TTB (Tris-Triton buffer): 100 mM Tris–HCl pH 7.4, 1% Triton X-100, 1 mM EDTA. Into a graduated cylinder add 50 mL of 1M Tris pH 7.4, 25 mL of 20% Triton X-100, and 1 mL of 0.5M EDTA. Bring solution to 500 mL, filter sterilize, and store at room temperature. 12. Antibody Buffer B (AbB): 1× PBS, 0.1% BSA, 0.5% Triton X-100, 1 mM NaN3, and 1 mM EDTA (see Note 4). Add approximately 150 mL of water to a beaker. Add 25 mL of 10× PBS, 6.25 mL of 20% Triton X-100, 1.25 mL of 200 mM NaN3, 500 ML of 0.5M EDTA, and 0.25 g BSA. Add BSA last

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by sprinkling BSA powder on top of the liquid and allow to dissolve slowly before stirring (otherwise lumps will form). Using a graduate cylinder, bring volume to 250 mL. 13. 0.1M Dithiothreitol (DTT): Shortly before it is needed, measure 15.4 mg DTT and dissolve in 1 mL of water. 14. DABCO: 20 mM Tris–HCl pH 8.0, 0.2M 1,4-diazabicyclo2,2,2-octane (DABCO), and 90% glycerol. 2.2. Additional Materials

1. Methanol 2. Liquid nitrogen 3. Glass centrifuge tube with screw cap closure 4. 15 mL polypropylene conical tube 5. Saturated picric acid 6. Formalin (formaldehyde solution, 36.5%) 7. Paraformaldehyde, 16% solution (10 mL ampoules). We aliquot ~300 ML into 1.5 mL Eppendorf tubes and store at −20°C. For each fixation, thaw 1 aliquot. If crystals appear, simply heat at 60°C for several minutes and vortex until dissolved. 8. 30% H2O2 solution 9. BSA 10. Secondary antibodies: Donkey anti-Rabbit Alexa 488 and Goat anti-Mouse Alexa 594. 11. Microscopy slides (3p × 1p × 1 mm) and coverslips (22 × 22 mm).

3. Methods Carry out all procedures at room temperature unless otherwise indicated. 3.1. Collect Worms

1. Collect unstarved worms from four 100 mm plates using M9 buffer (see Note 5). Use 3–5 mL of M9 per plate and tilt back and forth to resuspend the worms. Transfer them to a glass tube using a glass pipette (see Note 6). 2. Spin down worms for 1 min at 282 × g in a table top centrifuge. 3. Wash two more times by adding 10–14 mL of M9 buffer, wait for 15 min for worms to settle by gravity (see Note 7), and then aspirate M9 buffer off, until 50 ML remains. 4. Proceed to either Nonet method fixation or Constant Spring fixation (see Subheading 1 about which method to chose).

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3.2. Nonet Fixation Method

1. Put worms on ice for 2 min (see Note 8). During this time prepare fixative solution (10 ML of B-mercaptoethanol, 400 ML of Bouin’s fixative, and 400 ML of methanol) in a separate tube. 2. Remove excess M9 buffer and add fixative to worms. Transfer worms from glass tube into a 1.5 mL Eppendorf-type tube, using a glass Pasteur pipette (see Note 9). 3. Mix by rotation on nutator for 30 min. Obtain liquid nitrogen during this time. 4. Freeze worms quickly by dipping tube into liquid nitrogen (see Note 10). Let it stay frozen for at least 5 min. This step can be used as a stopping point by placing the tube in a −80°C freezer overnight or longer (up to 3 months). 5. Thaw tube in warm tap water until fully thawed (see Note 11). 6. Mix by rotation on nutator for 30 min. 7. Centrifuge in microfuge for ~12 s (enough time for centrifuge to reach full speed) and pipette supernatant into waste container. 8. Wash at least three times by adding 1.4 mL of BTB solution, centrifuging for ~12 s, and pipetting supernatant into waste container (see Note 12). The BTB solution needs to be made fresh every time. 9. After washing, resuspend worms in 1 mL of BTB, and mix by rotation on nutator for 1 h (see Note 13). 10. Centrifuge for ~12 s and pipette supernatant into waste container. Add 1 mL of BTB and rotate on nutator for 2–3 h. 11. Centrifuge for ~12 s and pipette supernatant into waste container. Wash by adding 1 mL of BT (not BTB), inverting 3–4 times, centrifuging, and pipetting supernatant into waste container. 12. Wash twice by adding 1 mL of AbA buffer (without EDTA), inverting 3–4 times, centrifuging for ~12 s, and removing supernatant. Be very careful when aspirating off supernatant since the small worm pellet may become loose after washing with the PBS-based AbA. 13. After washing, resuspend worms in 1 mL of AbA buffer (without EDTA) and rotate on nutator for at least 30 min. 14. Centrifuge for ~12 s, and pipette off supernatant. Leave worms in ~100 ML (at least a 1:1 ratio) of AbA buffer (without EDTA). Fixed worms can be kept for up to a month at 4°C.

3.3. Constant Spring Fixation

1. Put worms on ice for 2 min (see Note 8). During this time prepare fixative solution (900 ML of 4× MRWB buffer, 2,850 ML of methanol, and 250 ML of 16% paraformaldehyde) in a separate tube.

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2. Remove excess M9 buffer and add fixative to worms. Transfer worms from glass tube into a 15 mL polypropylene conical tube (see Note 14). Mix by inversion. 3. Freeze worms in liquid nitrogen and keep frozen for at least 3 min (see Note 10). 4. Thaw worms by hand (or in tap water) until reaching glacial stage instead of completely liquid stage (see Note 15). 5. Repeat the freeze/thaw cycle 4 more times (only need to freeze until frozen; no need to wait 3 min in between). 6. After the final thaw, incubate worms on ice for 1 h, inverting 3–4 times approximately every 10 min. 7. Spin at 282 × g for 1 min and aspirate off supernatant. 8. Wash twice by resuspending in 5 mL of 1× TTB, inverting three times, spinning at 282 × g for 1 min, and aspirating off supernatant. 9. Resuspend in 5 mL of 1× TTB with 1% (50 ML) B-mercaptoethanol. Incubate at 37°C, rotating on nutator for 2 h (see Note 16). 10. Spin at 282 × g for 1 min and aspirate off supernatant. Resuspend in 5 mL of 1× BO3 buffer (see Note 17). Respin and remove supernatant. 11. Resuspend in 6.3 mL of 1× BO3 and 700 ML of 0.1M DTT (this DTT solution needs to be made fresh). Rotate on nutator for 15 min. Spin at 282 × g for 1 min and aspirate off supernatant. 12. Resuspend in 7 mL of 1× BO3, invert three times, resuspend, and aspirate off supernatant. 13. Resuspend in 7 mL of 1× BO3 and 70 ML of 30% H2O2 (see Note 18). Incubate for 15 min while inverting every 5 min (see Note 19). 14. Spin at 282 × g for 1 min and aspirate off supernatant. Resuspend in 7 mL of 1× BO3. 15. Spin at 282 × g for 1 min and aspirate off supernatant. Resuspend in 5 mL of AbB and rotate on nutator for 20 min. 16. Spin at 282 × g for 1 min and aspirate off supernatant. Resuspend in 1 mL of AbA and transfer to an Eppendorf 1.5 mL tube. 17. Centrifuge in microfuge for ~12 s and remove all but 100 ML of buffer. Fixed worms can be kept for up to a month at 4°C. 3.4. Brief Description of Immunostaining Using Either Nonet or Constant Spring Fixed Worms

1. Using a cut down p200 tip with a p20 pipetteman, dispense 5 ML of packed fixed worms into a 1.5 mL Eppendorf tube (see Note 20). 2. Add at least 20 ML of primary antibody in AbA to the worms and incubate overnight at room temperature. Either rocking

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horizontally on a flat bed orbital shaker, or rocking in a tube rack sitting on the top of a moving nutator is effective. 3. Wash four times by adding 1 mL of 1× PBS, 0.5% Triton X-100, rocking on nutator for 15 min, centrifuging in microfuge for ~12 s, and pipetting off supernatant (see Note 21). 4. After the last wash, remove as much buffer as possible. Add at least 20 ML of secondary antibody in AbA for 2 h without azide at room temperature in the dark (see Note 22). 5. Repeat washing step detailed in step 3. 6. After the last wash leave some buffer with the worms and with a cut down p200 tip using a p20 pipetteman, dispense 5 ML slightly suspended (by repipetting) (see Note 23) stained worms onto a glass slide. Apply on top of a 5 ML drop of DABCO on a glass coverslip then gently place coverslip onto slide (such that the DABCO and worms touch). Then seal with nail polish. Alternatively, apply 5 ML of worms directly into a 50 ML drop of Prolong (Invitrogen), obviating the need for nail polish. 7. Evaluate body wall muscle using confocal microscopy (see Note 24).

4. Notes 1. Saturated picric acid solution can be made from the solid but we do not recommend this as the powder can be explosive. Also wear gloves anytime handling solutions containing picric acid. 2. Preparing this solution as 20% weight per volume is critical because the high viscosity of Triton X-100 makes it extremely difficult to be accurate in measuring out a volume. 3. The pH of the Borate Buffer is critical for both fixation methods but especially the Constant Spring method. 4. Sodium azide (NaN3) is a poison, so wear gloves. 5. Most important factor in good fixation (and thus good immunostaining) is that of the worm stage when the worms are fixed. The worms ideally should be young adults. Worm growth and stage is highly variable upon strain, temperature, etc. 6. Worms stick to plastic tips and tubes, so it is best to use glass whenever possible. 7. Letting the worms settle by gravity is meant to remove bacteria from the solution and from inside the worm to reduce background when staining. Thus, it is important for this to be done at room temperature so the worms can swim around. Some of

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the worms may not settle during the 15 min. These worms could be immature/larval worms that do not stain well anyway, so aspirate them off. 8. Ice makes the worms stop moving allowing for a better packing of the worms. 9. Once the worms are in fixative, they will no longer significantly stick to plastic. 10. The worms can settle quickly, so mix the worms before dipping, then every couple of seconds take out and invert a couple of times until frozen. If the worms settle, the fixation will not be as good. 11. Be careful to wear gloves because tubes tend to leak at this point. After thawed, open lid, and wipe off to prevent leaking during next step. 12. The goal is to have the solution be relatively clear of yellow (picric acid) and the worms should be bright yellow. Bright yellow worms indicate that the fixation went well. 13. The purpose of this step and subsequent washing steps is to remove picric acid from the worms. If the solution is still yellow, it is necessary to do additional washes with BTB solution. 14. It is very important to use polypropylene not polystyrene, because during freezing a polystyrene tube will crack and break. 15. Thawing worms only until the glacial stage reduces worm fragmentation during fixation that can be very significant if thawing all the way to the liquid stage. 16. This step is critical for the permeabilization of the cuticle (by breaking down disulfide bonds of the cysteines in the cuticular collagens), and the elevated temperature aids the kinetics. 17. During these washes without detergent it is normal for many of the worms to become attached to the plastic conical tube. Upon the addition of detergent in later steps some will detach. 18. The H2O2 solution needs to be fresh (less than 1 year since opening). The H2O2 solution can burn badly, so do not touch without gloves. 19. Oxidation by H2O2 prevents reformation of disulfide bonds among cysteines, and consequent collagen cross-linking, thus maintaining the “holes” in the cuticle. During the 15 min incubation, one should observe small bubbles, so it is important to keep the cap ajar between mixing, otherwise the tube might explode. 20. The cut-down tip prevents worm breakage.

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21. Pipetting off the wash prevents the loss of worms that could occur if using aspiration. If worms are taken up into the tip they can be expelled and centrifuged again. 22. We have found that Alexa 488 conjugated secondary antibodies produce more stable signals than FITC conjugated secondary antibodies. Also, nonspecific signals from secondary antibodies can be eliminated using preabsorption with wild type worm acetone powder overnight at 4°C. For specific details see ref. (10). 23. The worms need to be slightly suspended to allow worms to be spread out on slide. 24. We have found that confocal microscopy tends to provide better images than deconvolution microscopy for adult body wall muscle.

Acknowledgments We thank the NIH for research support (grant AR052133), and for a Fellowship in Research and Science Teaching (FIRST) postdoctoral fellowship (K12GM000680), also from the NIH, for supporting K.J.W. References 1. Xiong G, Qadota H, Mercer KB, McGaha LA, Oberhauser AF, Benian GM (2009) A LIM-9 (FHL)/SCPL-1 (SCP) complex interacts with the C-terminal protein kinase regions of UNC-89 (obscurin) in Caenorhabditis elegans muscle. J Mol Biol 386:976–988 2. Miller RK, Qadota H, Stark TJ, Mercer KB, Wortham TS, Anyanful A, Benian, GM (2009) CSN-5, a component of the COP9 signalosome complex, regulates the levels of UNC-96 and UNC-98, two components of M-lines in Caenorhabditis elegans muscle. Mol Biol Cell 20:3608–3616 3. Qadota H, McGaha LA, Mercer KB, Stark TJ, Ferrara TM, Benian, GM (2008) A novel protein phosphatase is a binding partner for the protein kinase domains of UNC-89 (Obscurin) in Caenorhabditis elegans. Mol Biol Cell 19:2424–2432 4. Stevenson TO, Mercer KB, Cox EA, Szewczyk NJ, Conley CA, Hardin JD, Benian, GM (2007) unc-94 encodes a tropomodulin in Caenorhabditis elegans. J Mol Biol 374: 936–950

5. Qadota H, Mercer KB, Miller RK, Kaibuchi K, Benian GM (2007) Two LIM domain proteins and UNC-96 link UNC-97/pinch to myosin thick filaments in Caenorhabditis elegans muscle. Mol Biol Cell 18:4317–4326 6. Miller RK, Qadota H, Landsverk ML, Mercer KB, Epstein HF, Benian, GM (2006) UNC-98 links an integrin-associated complex to thick filaments in Caenorhabditis elegans muscle. J Cell Biol 175:853–859 7. Mercer KB, Miller RK, Tinley TL, Sheth S, Qadota H, Benian, GM (2006) Caenorhabditis elegans UNC-96 is a new component of M-lines that interacts with UNC-98 and paramyosin and is required in adult muscle for assembly and/or maintenance of thick filaments. Mol Biol Cell 17:3832–3847 8. Small TM, Gernert KM, Flaherty DB, Mercer KB, Borodovsky M, Benian GM (2004) Three new isoforms of Caenorhabditis elegans UNC89 containing MLCK-like protein kinase domains. J Mol Biol 342:91–108 9. Mercer KB, Flaherty DB, Miller RK, Qadota H, Tinley TL, Moerman DG, Benian, GM (2003)

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Caenorhabditis elegans UNC-98, a C2H2 Zn finger protein, is a novel partner of UNC-97/ PINCH in muscle adhesion complexes. Mol Biol Cell 14:2492–2507 10. Miller DM, Shakes DC (1995) Immunofluorescence microscopy. Methods Cell Biol 48:365–394 11. Nonet ML, Grundahl K, Meyer BJ, Rand, JB (1993) Synaptic function is impaired but not eliminated in C. elegans mutants lacking synaptotagmin. Cell 73:1291–1305 12. Benian GM, Tinley TL, Tang X, Borodovsky M (1996) The Caenorhabditis elegans gene

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unc-89, required fpr muscle M-line assembly, encodes a giant modular protein composed of Ig and signal transduction domains. J Cell Biol 132:835–848 13. Finney M, Ruvkun G (1990) The unc-86 gene product couples cell lineage and cell identity in C. elegans. Cell 63:895–905 14. Takeda K, Watanabe C, Qadota H, Hanazawa M, Sugimoto A (2008) Efficient production of monoclonal antibodies recognizing specific structures in Caenorhabditis elegans embryos using an antigen subtraction method. Genes Cells 13:653–665

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Part III Experimental Models and Analysis of Skeletal Muscle Exercise and Disuse

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Chapter 11 Resistance Loading and Signaling Assays for Oxidative Stress in Rodent Skeletal Muscle Stephen E. Alway and Robert G. Cutlip Abstract Resistance loading provides an important tool for understanding skeletal muscle responses and adaptations to various perturbations. A model using anesthetized rodents provides the means to control the input parameters carefully, and to measure the output parameters of each muscle contraction. Unilateral models of anesthetized loading also provide the advantage of comparing an unloaded and loaded muscle from the same animal. Voluntary models for resistance loading arguably provide a more “physiological response” but it also introduces more variability in the input parameters, which can be affected by the stimulus used to motivate the animal to exercise. After either acute or chronic periods of muscle loading, the loaded muscles can be removed and various signaling proteins can be determined by sodium dodecyl sulfate polyacrylamide gel electrophoresis (SDS-PAGE) or enzyme assays. Several assays are described, which provide an indication of downstream markers for oxidative stress. Key words: Exercise, Evoked contractions, Isometric contractions, Stretch-shortening contractions, Electrophoresis, Oxidative stress, Skeletal muscle

1. Introduction Resistance loading has been shown to be an effective means of increasing muscular mass and force output in rodents, although the extent of the increase is attenuated with aging (1). Loading also increases oxidative stress in skeletal muscle in humans (2, 3) and rodents (1, 4, 5). Although this is not a problem in young rodents (1), loading elevates the oxidative stress levels in muscles of aged rodents (1, 4) beyond the heightened levels that occur as a part of aging per se (6). Oxidative stress increases in skeletal muscle after acute exercise (2, 5); however, chronic exercise enhances the endogenous

Joseph X. DiMario (ed.), Myogenesis: Methods and Protocols, Methods in Molecular Biology, vol. 798, DOI 10.1007/978-1-61779-343-1_11, © Springer Science+Business Media, LLC 2012

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antioxidant defenses and decreases production of prooxidants resulting in lower indices of oxidative stress. The reduced buffering capacity may compromise the muscles’ abilities to hypertrophy and/or to improve muscle function in aged animals. In addition, increased oxidative stress may contribute to muscle injury or maladaptation in response to resistance loading (7). However, given that oxidative stress may also be an important stimulus for skeletal muscle adaptation (8) in the absence of myofiber degeneration, modulating the level of loading inputs may be important to maximize the training response in muscles of old and young animals. Thus, models of resistance loading provide the means to accurately select and control the input parameters and quantify the resultant muscle responses. This is a necessary schema for identifying potential mechanisms, which regulate muscle remodeling/adaptation, or maladaptation and injury (4, 9, 10). Appropriately designed in vivo loading models have the latitude to vary the input parameters for systematic study of the effects of different biomechanical loading signatures on the target muscle’s response in real-time and temporally after the exposure or exposures (11). Varying the mechanical inputs has been shown to affect the magnitude of skeletal muscle injury and/or adaptation following repetitive loading contractions (reviewed in ref. (12)). These include muscle length (and whether it changes) during a contraction (13, 14), the velocity of the contraction (15), number of repetitions, (16) and the rest intervals between contractions (17). For example, isometric exercise increases acute levels of oxidative stress (5), without invoking muscle injury (13, 18). Muscle lengthening via eccentric loading results in large degrees of muscle damage (19–21). Stretch-shortening contractions (SSCs), which include both eccentric and concentric movements, can induce injury or result in adaptation without injury, depending on the velocity of the contraction and/or the rest intervals between contractions (1, 17). There is some utility in studying each type of contraction, but the case could be made that investigating the response to multiple contractions with different parameters are necessary under controlled loading conditions to systematically study muscle injury and/or adaptation/maladaptation. It is during activities that are similar to those observed in athletic and occupational settings that skeletal muscle encounters periods of alternating, repetitive lengthening and shortening contractions along with periods of isometric contractions that appear to magnify the extent of muscle injury (12). Therefore, it is necessary to develop physiologically relevant models to best study skeletal muscle mechanics. Furthermore, there are some intrinsic benefits to investigating stretch–shortening exercise in the context of muscle damage that cannot be addressed by eccentric-only or concentric-only protocols. First, real-time changes in peak eccentric forces, isometric pretest

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forces, and force enhancement during injurious stretch–shortening cycles can be quantified during a muscle contraction. These changes in “real-time” parameters allow for the detection of muscle damage as it occurs, and muscle force can be tracked noninvasively to identify repair/recovery strategies. Second, synergistic effects of combined concentric and eccentric muscle actions can be studied in the context of muscle damage and real-time changes in mechanical response. Third, SSCs increase the versatility of sequential eccentric repetitions, because the muscle is not neutrally deactivated during the concentric phase preceding each stretch. Thus, the effects of cycle frequency on real-time force response and muscle damage can be studied. Fourth, real-time changes in elastic and viscoelastic properties of muscle and the frequency dependence of these properties can be examined during cyclic loading. Finally, stretchshortening allows for the investigator to study work loops, power absorption, and generation in real-time, and changes in those parameters during an injury-producing protocol. The work loop approach allows for the distinction between damage assessed as a decrease in positive work and power from damage assessed as a decrease in negative work and power (11). Oxidative stress is elevated with aging in most tissues, including skeletal muscle (1, 22, 23). Increased reactive oxygen species (ROS) production may contribute to aging-induced skeletal muscle wasting (i.e., sarcopenia) (1, 24). We have used rodent models of stretch-shortening and isometric exercise to show that while acute levels of oxidative stress are elevated with loading, long-term adaptations are often possible, albeit more attenuated in aged rodents. Although exercise is a useful approach to counter aging-induced sarcopenia, it also increases oxidative stress levels within exercising muscles (23–25). The additive effects of an increase in oxidant production and an attenuated antioxidant buffering capacity potentially leave aged skeletal muscles vulnerable to oxidative damage. Buffering oxidative stress may improve muscle responses to repetitive loading, at least in muscles from old rodents (1, 4, 5). Oxidative stress in skeletal muscles can arise from several sources (26, 27). NAD(P)H oxidase is one potentially important contributor to oxidative stress in skeletal muscles because NAD(P) H oxidase activity increases in skeletal muscle with aging animals and long-duration exercise (27). The xanthine oxidase system is another important contributor to oxidative stress. Xanthine oxidase has been shown to be an important source of oxidant production in the vascular endothelium (28) and also a contributing factor to oxidative stress during strenuous exercise (29–31). A high demand on anaerobic metabolism, coupled with intermittent localized obstruction of blood flow and subsequent reperfusion within contracting muscles raises the potential for xanthine oxidase to be an important source of oxidant production during intense resistance

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exercises. Previous studies have observed that after resistance training, type II fibers are preferentially hypertrophied in both young and aged muscle (32, 33). Additionally, type II fibers tend to be more susceptible to oxidative damage than type I fibers (34). Therefore, the increased oxidant production associated with aging could preferentially limit the ability of type II fibers to adapt to exercise training.

2. Materials Resistance loading in rodents can be accomplished using similar types of materials as described below. We have constructed a custombuilt rodent dynamometer (35) with improved software and data acquisition system to assess muscle function in rats (1, 10, 15, 16, 35–37). However, we have also found that commercially available hardware and software works very well for this purpose in our mouse systems (e.g., Aurora Scientific) (5). 2.1. Rodent Anesthesia Station

Whether working with rats or mice, it is necessary to have an anesthesia station, so that the animals can be properly positioned, the electrodes can be placed appropriately, and the muscle parameters can be controlled in a precise fashion. 1. Small animal anesthesia machine (Smith Medical). (a) Isoflurane vaporizer (b) Tabletop research anesthesia machine (c) Flowmeter (isoflurane and oxygen) (d) Gas tubes and rodent mask diaphragms for anesthesia system 2. Lab evacuation system (Smith Medical) to collect isoflurane and filter through a charcoal to prevent gas buildup in the lab. 3. Oxygen cylinder, pressure valve with gauge. 4. Plexiglass “induction” box large enough to fit the size of rat that you work with. Add ports to the chamber and attach a hose to the isoflurane unit.

2.2. Custom Rat Dynamometer

The apparatus used to conduct loading in rat dorsiflexor or plantar flexor muscles is shown in Fig. 1 and it is also described below. This custom made system is described in greater detail by Cutlip et al. (35). Vertical forces applied to an aluminum sleeve fitted over the dorsum of the foot will be translated to a load cell transducer in the load cell fixture. The force produced by the dorsiflexor or plantar flexor muscles (depending on your experiment) will be measured at the interface of the aluminum sleeve and the dorsum of the foot.

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Fig. 1. Anesthetized rat model for resistance loading one limb of a rat as described by Cutlip et al. (35). This unit can be used for studies in dorsiflexor or plantar flexor muscles. The positioning of the electrodes is for dorsiflexion. An aged Fisher Brown Norway rat (32 months) is shown.

It is also possible to obtain other suitable hardware selections from other commercial sources (see Note 1). 1. Labview-based virtual instrument developed to interface with a data acquisition board (PCI-MIO-16XE-10, National Instruments). 2. Motion controller (Unidex 100, Aerotech Inc.) for precise control of an Aerotech servomotor. 3. A brushless DC servomotor (1410DC, Aerotech). This DC servomotor and gearbox produce a peak torque ~100-fold greater than the maximum torque that a rat can produce. 4. A data acquisition board (PCI-MIO-16XE-10, National Instruments). 5. Develop or purchase software for data storage of experimental position, force, and velocity data (see Note 2). 6. Develop or purchase software for data analyses (see Note 3). 7. Obtain X–Y positioning table (preferably heated to control the animal’s body temperature; an alternative approach is to use a heat lamp over the animal). 2.3. Custom Dynamometer for the Mouse

The apparatus used to conduct loading in mouse dorsiflexor or plantar flexor muscles is shown in Fig. 2a, b. The apparatus is similar in design to the rat dynamometer system, but it is scaled down to

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Fig. 2. (a) Anesthetized mouse model for resistance loading one limb of a rat as described by Ryan et al. (5). This unit can be used for studies in dorsiflexor or plantar flexor muscles. The positioning of the electrodes is for plantar flexion. An aged C57BL/6 mouse (30 months) is shown. (b) The leg brace is shown in the anesthetized mouse.

the appropriate size of the much smaller mouse (see Note 4). This system is also modified from an in vitro model that has been described by Warren et al. (38–40). 1. Servo motor, 300C (Aurora Scientific). This unit has a capacity of 1N of peak force. An aluminum footplate must be custom made to fit the mouse and attach to the lever arm of the servomotor. 2. Custom or commercial temperature-controlled plate, with capacity to clamp the mouse limb and hold the servomotor. Obtain a metal rod that will be positioned on the lateral side of the knee. 3. Data acquisition Instruments).

board

(PCI-MIO-16XE-10,

National

4. Cables and computer system. 5. Software for data storage of experimental position, force, and velocity data. 6. Data analysis software. 2.4. Electrical Stimulation

1. Platinum stimulating electrodes (Grass Medical Instruments) (see Note 5). 2. Model SD9, Grass Medical Instruments (see Note 6). 3. Computer with appropriate board slots, and capabilities for data acquisition and analysis (see Notes 2 and 3). 4. BNC cables for interfacing computer to stimulator.

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2.5. Glutathione (GSH) to Oxidized Glutathione (GSSG): GSH/GSSG

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1. 5% Metaphosphoric acid (MPA) in deionized water (dH2O). Make fresh and keep on ice. 2. 3.3 mM 1-methyl-2-vinylpyridinium trifluoromethanesulfonate (M2VP) (a GSH scavenger) in hydrochloric acid (HCl). 3. Assay buffer: 143 mM sodium phosphate (Na·PO4) and 6.3 mM sodium-EDTA (pH 7.5). 4. 50 U/mL GSH reductase (e.g., type HI, from Saccharomyces cerevisae, Sigma) in assay buffer. 5. 0.3 mM E-Nicotinamide adenine dinucleotide phosphate (NADPH) in assay buffer. Make fresh and use the same day. 6. 6 mM Chromogen 5,5c-dithiobis-(2-nitrobenzoic acid) (DTNB) in assay buffer. 7. Six standards of GSH in assay buffer. Each GSSG molecule is equivalent to two GSH molecules; therefore, the values are expressed as PM GSH:

2.6. Hydrogen Peroxide (H2O2 ) Levels

GSSG (mM)

GSH (mM)

1

0.000

0.00

2

0.050

0.10

3

0.125

0.25

4

0.250

0.50

5

0.750

1.50

6

1.500

3.00

1. 300 PM N-acetyl-3,7-dihydroxyphenoxazine (Amplex Red; Molecular Probes/Invitrogen) in dimethyl sulfoxide (DMSO). This can be stored at −20°C for up to 6 months. 2. Black 96-well plates. 3. 30% hydrogen peroxide (H2O2). 4. Horseradish peroxidase (HRP). 5. 50 mM Tris, pH 7.4. 6. 20 mM phosphate-buffered saline, pH 7.4 (1× PBS).

2.7. 8-Hydroxy-2 cDeoxyguanosine (8-OHdG)

1. 96-Well protein binding plate. 2. Pure 8-hydroxy-2c-deoxyguanosine (8-OHdG) and 8-hydroyguanosine (8-OHG). 3. Bovine serum albumin (BSA). 4. 20 mM phosphate-buffered saline, pH 7.4 (1× PBS). 5. 100 mM phosphate-buffered saline, pH 7.4 (5× PBS). 6. Blocking buffer: PBS containing 1% BSA, 0.05% casein, 0.05% Tween 20, 2% sucrose, and 0.5 nM thimerosal.

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7. 1M phosphoric acid. 8. 3,3c,5,5c-Tetramethylbenzidine (Sigma). 9. HRP secondary antibody, 1 Pg/mL. 10. 8-OHG antibody – 1 Pg/mL mouse monoclonal anti-8OHdG antibody (clone 1F7; Trevigen). 2.8. Malondialdehyde (MDA)

1. 0.5M butylated hydroxytoluene in acetonitrile. 2. 1,1,3,3-Tetramethoxypropane (TMP) as a source of malondialdehyde (MDA) in 1% sulfuric acid. 3. Concentrated (37%) HCl. 4. 1-methyl-2-phenylindole (MP) in a mixture of acetonitrile/ methanol (3:1) made to a final concentration of 10 mM. 5. Homogenization buffer: ice-cold 1× PBS, pH 7.4.

2.9. Catalase Activity

1. 1× Assay buffer: 100 mM potassium phosphate, pH 7.0. When stored at 4°C this will be stable for 2 months. 2. 1× Sample buffer: 25 mM potassium phosphate, pH 7.5, containing 1 mM EDTA and 0.1% BSA. 3. 4.25M formaldehyde. 4. 8.82M H2O2. 5. Bovine liver Catalase (Sigma). Standards equal to 0, 2, 4, 6, 8, 10 mU activity per mg protein should be adequate for most muscle experiments. The diluted enzyme is stable for 30 min. The reconstituted catalase is stable for 1 month at −20°C. 6. 10M potassium hydroxide (KOH). The KOH solution is stable for at least 3 months if stored at 4°C. 7. 100% methanol. 8. 35.28 mM H2O2 diluted in HPLC-grade water. 9. 5% 4-amino-3-hydrazino-5-mercapto-1,2,4-triazole (Purpald) in 0.5M HCl. 10. 0.5M KOH. 11. 96 Well microplate and plate sealer. 12. 100 mM potassium periodate (KIO4). 13. Homogenization buffer: ice-cold 50 mM potassium phosphate, containing 1 mM EDTA, pH 7.0.

2.10. MnSOD and CuZnSOD

1. Phosphate buffered saline (PBS). 2. Bovine CuZnSOD. 3. 0.05M potassium phosphate buffer, pH 7.8. 4. Homogenization buffer: 20 mM HEPES buffer, pH 7.2, containing 1 mM EGTA, 210 mM mannitol, and 70 mM sucrose.

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5. Assay buffer: 0.05M potassium phosphate, 0.1 mM diethylenetriaminepentaacetic acid (DTPA), 0.13 mg/mL BSA, 1.0 U catalase, 5.6 × 10 PM nitroblue tetrazolium (NBT), 0.1 mM hypoxanthine, 50 PM bathocuproinedisulfonic acid (BCS) disodium salt. Adjust to pH 7.8. 6. 12 mM potassium cyanide (KCN). 7. 1.18 mM Xanthine oxidase in 0.05M potassium phosphate buffer, pH 7.8. Keep on ice.

3. Methods Choose the appropriate rat species, age, and sex for your experiments. Obviously, the noise levels, temperature, and light–dark cycle should be held constant for all animals over the course of the experiment. 3.1. Positioning the Rat for Dorsiflexion or Plantar Flexion

1. Place a rat in an “induction” tank filled with 4% isoflurane gas diluted with 100% oxygen. Normally, the rat will be anesthetized within 3–4 min at this concentration of isoflurane. 2. Place the rat supine on the heated X–Y positioning table of the rodent dynamometer (35). 3. Place an anesthetic mask over the nose and mouth of the rat. 4. Adjust the anesthesia to 2% isoflurane to maintain anesthesia, but closely monitor the animal. You may need to adjust the isoflurane to maintain anesthesia during the loading session. This is somewhat dependent on the size of the rodents. 5. Secure the rat’s knee with a knee holder. 6. Secure the foot (we use the left foot but either foot is fine) onto the load cell fixture with the ankle axis (assumed to be between the medial and lateral malleoli) aligned with the axis of rotation of the load cell fixture. The joint position of the ankle should be defined by the angle between the tibia and the plantar surface of the foot. 7. Carefully monitor each animal during the protocol to ensure proper anesthetic depth and body temperature. Reduce the concentration of isoflurane if the depth of breathing becomes shallow and frequent.

3.2. Stimulation Parameters for Obtaining Maximal Contractions

1. For obtaining dorsiflexion, position the two platinum stimulating (needle) electrodes near the head of the fibula, to span the peroneal nerve. For plantar flexor experiments, place the platinum stimulating electrodes subcutaneously to span the tibial nerve in the popliteal fossa.

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2. Set the stimulator frequency to 120 Hz to generate maximal dorsiflexion or plantar flexion force. Set the stimulator to deliver a square wave pulse duration of 200 Ps, and set the stimulus to 4 V. The stimulus duration should be from 300 ms to 1 s, depending on your experimental outcome and frequency of stimulation. You should check the stimulus parameters on your preparation to maximize dorsiflexor or plantar flexor contractile performance. If force output is not maximal at the frequencies suggested, you should increase the stimulus voltage and/or the frequency of stimulation until a maximum force is obtained at a given stimulus frequency. It is also possible that you may need to adjust the position of the stimulating electrodes. You should not increase the pulse duration. 3. First, obtain isometric contractions on the dorsiflexor or plantar flexor muscle group to determine maximal force and to ensure that your setup is appropriate. 4. It may be necessary to reposition the stimulating electrodes, if inappropriate muscles are contracting (e.g., based on palpating the hind limb muscles). Examine the hind limb carefully during the contraction. If there is excessive eversion or inversion of the foot during the stimulus, the needle electrodes must be repositioned. 5. Set the ankle angle to 90° for isometric contractions. Stimulate the appropriate nerve 1 min before and after stretch-shortening exposures (1, 4, 10, 17, 36, 41). 6. If desired, you can obtain isometric contractions at 10° ankle increments throughout the range of motion of the dorsiflexors (13) or plantar flexors. 3.3. Preparing and Positioning the Mouse in the Dynamometer

The principals for both the apparatus and the animal set up and experiments in the mouse dynamometer (5) are similar to that in the rat (11, 13, 17, 41). 1. Place a mouse in an “induction” tank filled with 4% isoflurane gas and oxygen. If mice are not anesthetized in 3–4 min, increase the isoflurane to a maximum of 5%. 2. Place the mouse on its right side on the heated table. 3. Place an anesthetic mask of the appropriate size over the mouse’s nose and mouth (see Fig. 2a). 4. Reset the anesthesia station to deliver 2% isoflurane, but adjust it to 3% as needed to maintain anesthesia during the loading session. 5. Secure the mouse’s leg with a brace (see Fig. 2b). 6. Secure the left foot on the custom made footplate. You can use a small paper clip or a piece of nonstretch tape to secure the foot.

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7. Carefully monitor each animal during the protocol to ensure proper anesthetic depth and body temperature. Adjust the level of isoflurane to ensure that the depth of breathing is appropriate. 3.4. Repetitive Loading Parameters in Rat Muscles

1. Program your software so that the stimulator and dynamometer will conduct the desired number of sets and cycles, velocity of contraction, etc. to accomplish the goals of your experiment. Typically, you should begin with an isometric contraction, and follow this with the type of training/repetitive loading contractions to suit your study (e.g., isometric, stretch shortening to induce injury, noninjurious contractions). 2. You should first obtain an isometric pretest contraction. Fix the ankle angle at 90°. Activate the dorsiflexors maximally using a 300 ms stimulation duration in a similar fashion as Davis et al. (42) and Willems and Stauber (43). 3. Obtain a single SSC from the dorsiflexor or plantar flexor muscle group 2 min preceding and following training with any isometric or SSC protocol that will be used as a training or injuring stimulus (13). Conduct the single SSC by activating the dorsiflexor muscles for 300 ms then moving the load cell fixture from 70 to 140° at an angular velocity of 500°/s. The load cell fixture should then be immediately returned to 70°, at 500°/s. Continue activation of the muscle (via the nerve) for 300 ms after cessation of the movement. 4. Repetitive SSC should be used to induce muscle injury or to induce oxidative stress in the rodent skeletal muscles. In addition, SSC should be used to evaluate the muscle’s ability to generate dynamic forces and to perform positive and negative work in vivo during dynamic SSCs of the target muscle. Follow the methods, and illustration of force and position outputs for stretch-shortening cycles for resistance loading as described in detail in Geronilla et al. (11). These methods are outlined below: (a) One minute after the single SSC contraction, activate the dorsiflexor or plantar flexor muscles for 100 ms and then move the load cell fixture from 70 to 120° angular position at a velocity of 500°/s, in a reciprocal fashion, for ten oscillations and a duration of 2.4 s (allowing for motor ramp up and ramp down time). The load cell fixture should then be immediately returned to an ankle angle of 70°. (b) Continue the muscle activation via the nerve for 300 ms after cessation of the movement. (c) After a 2 min rest period, repeat the exposure a total of 10 times (i.e., in sets of ten repetitions). Continue with the contractions until you have obtained a total of 30 repetitions to 300 repetitions (depending on your experimental goals) (16).

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(d) After a 2 min rest period after the last SSC, conduct a post-test single SSC in the same fashion as the pretest single SSC. (e) After a 2-min rest period, conduct a post-test isometric contraction in the same fashion as the pretest isometric. 3.5. Loading Parameters in the Mouse

1. Adjust the stimulator settings to maximize dorsiflexor or plantar flexor contractile performance. Increase the stimulus until a maximum force is obtained at a given stimulus frequency (see Notes 7 and 8). 2. For obtaining dorsiflexion in the mouse, the platinum stimulating electrodes should be placed subcutaneously near the head of the fibula, to span the peroneal nerve. For plantar flexor experiments, the platinum stimulating electrodes should be placed subcutaneously to span the tibial nerve in the popliteal fossa (see Notes 9 and 10). 3. Isometric contractions should be performed on the dorsiflexor or plantar flexor muscle group to determine maximal force and ensure that your setup is appropriate. An example of the force record for one plantar flexion contraction is shown in Fig. 3. Isometric contractions should be conducted at an ankle angle of 90°, 1 min before and after any stretch-shortening or

Fig. 3. Force tracing of a single maximal isometric plantar flexion contraction from the hindlimb of an aged (30 month) old C57BL/6 mouse. The frequency of stimulation was 120 Hz. The duration of the contraction was for 4.0 s. This contraction shows significant “sag” in the force record. Sag would be minimal if a shorter contraction duration (<2.0 s) had been selected.

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repetitive loading exposures. Conduct the desired number of sets and cycles, velocity of contraction, etc. to accomplish the goals of your experiment. The parameters below are given for dorsiflexors, but identical parameters can be used to stimulate and study the plantar flexor muscle group. Rest 2–3 min between each isometric contraction to prevent fatigue. 4. Conduct continuous stretch shortening cycles (SSC) to study repetitive loading, injury, or oxidative stress in skeletal muscles of the mice (5). Always begin with an isometric contraction to determine the maximal force (Fig. 3). Conduct the single SSC by activating the dorsiflexor muscles for 300 ms then moving the load cell fixture from 70 to 140° at an angular velocity of 500°/s. The load cell fixture should then be immediately returned to 70°, at 500°/s. Continue activation of the muscle (via the nerve) for 300 ms after cessation of the movement. (a) For repetitive loading studies to induce muscle injury, activate the dorsiflexor or plantar flexor muscles for 0.1 s, 1 min after the single SSC contraction, and then move the load cell fixture from 70 to 120° angular position at a velocity of 500°/s, in a reciprocal fashion, for ten oscillations and a duration of 2.4 s (allowing for motor ramp up and ramp down time). The load cell fixture is immediately returned to 70°. (b) Continue the muscle activation via the nerve for 300 ms after cessation of the movement. Repeat this activation sequence for 10 times for one set of activation. (c) After a 2 min rest period, repeat the exposure a total of 10 times (i.e., in sets of ten repetitions) to induce muscle injury. Continue with the contractions until you have obtained a total of 30 repetitions to 300 repetitions (depending on your experimental goals) (16). If your research goals are not to induce muscle injury, you should conduct repetitive isometric loading (5), or use a lower velocity of contractions as described in Subheading 3.7. (d) After a 2 min rest period after the last SSC, conduct a post-test single SSC in the same fashion as the pretest single SSC. (e) After a 2-min rest period, conduct a posttest isometric contraction in the same fashion as the pretest isometric. 3.6. Using Eccentric Contractions in Rat and Mouse Muscles

1. To measure eccentric muscle force/torque or repetitively load the plantar flexors or dorsiflexors with eccentric training, set the foot plate to move at a velocity of 40 mm/s during full tetanic stimulation of the plantar flexors or dorsiflexors. 2. The frequency of stimulation should be sufficient to induce full tetanic fusion (e.g., 120 Hz should be adequate for these contractions).

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3. Increase the muscle length for 200 ms after initiation of the stimulation train. This will begin the eccentric contraction during the isometric plateau of the tetanus. 4. Set the foot angle to 30°. This is well inside the physiological limits of movement for the mouse or rat foot. Nevertheless, you may wish to choose longer excursions. 5. To limit the degree of force sag, the contractions should be reduced to 600 ms for either plantar flexors or dorsiflexors. 6. You should be able to obtain a ~20–30% decline in isometric force (when compared with maximal isometric force that is measured before initiation of the stretch) if you set the parameters to three sets of 20 repetitions with 60 s rests between each set of repetitive activity. 3.7. Changing Velocity Parameters for Grading Injury or Training Without Injury in Stretch-Shortening Contractions in Rat and Mouse Muscles

1. The velocity of movement during SSCs has a significant impact on the muscle response temporally after the exposure. If the goal of your experimental outcome is to induce muscle injury, you should conduct the single or acute exposures at 500°/s using continuous muscle activation as described earlier. You should modulate the extent of desired muscle injury by varying the exposure from seven sets of continuous SSCs (70 repetitions) at 500°/s (less severe injury) to 15 sets (150 repetitions), which will induce much greater injury severity (11, 16, 37). 2. If you wish to induce overload (or exercise) of the hindlimb muscles without inducing injury, you should slow the velocity of the SSCs to 60°/s, and intermittently activate the dorsiflexor or plantar flexor muscles (2 s rest between each repetition), with an exposure up to eight sets (80 repetitions) (9). These “training” exposures should be conducted for at least 80 intermittent SSC/exposure for 14 sequential exposures over a 4.5 week duration. This training protocol will increase muscle strength and mass without histological damage or inflammation, especially in young adult rats (10). These same parameters can be applied to senescent rats without obtaining histological muscle damage (10); however, the intensity and duration may be too high in old animals to see large improvements in muscle mass or strength. Therefore, you may have to reduce the stimulus duration and intensity if it is your goal to obtain improvements in muscle function from old rodents (44).

3.8. Duty Cycle Parameters for SSC Loading

1. If you wish to regulate oxidative stress, injury, or training outcomes to a muscle, you should also adjust the duty cycle of the repetitive loading parameters of your study. 2. To maximize the metabolic load to the mitochondria, and increase oxidative stress and thereby increase the rate of muscle fatigue, you should shorten the rest between the duty cycles

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(e.g., 15–30 s). This will induce rapid muscle fatigue with sequential sets, reduce muscle output force, and thereby reduce biomechanical stresses in the subsequent sets of activation. 3. If instead, you wish to maximize the biomechanical load, so that muscle fatigue is minimized and the target muscle can generate higher forces in subsequent sets, you should increase the duration of rest between the duty cycles (e.g., to 5 min) and/or reduce the duration of each duty cycle (100–200 ms). You will be able to achieve the greatest isometric force deficit over an exercise session with exposures that contain either short rest intervals, or with long duty cycles and long durations between each duty cycle (17). 3.9. Muscle Preparation for Measuring Oxidative Stress in Loaded Skeletal Muscle

1. At the end of the acute exercise (5) or chronic exercise (1, 4), the appropriate muscles should be removed and the animals euthanized. For dorsiflexor studies in rats, remove the tibialis anterior, and or the extensor digitorum longus muscles. Both heads of the gastrocnemius should be removed as the major contributions to the plantar flexor function. If you are also interested in biochemical (e.g., fiber type differences), the gastrocnemius muscles can be separated into red (more oxidative) and white (more glycolytic) parts. In addition, you can obtain the soleus as a muscle with primarily a type I myosin contribution in rats, or a mixed muscle in mice. 2. Remove the muscles (keeping them moist with 0.9% saline or phosphate buffered saline during dissection), and clean them of excess connective tissue. 3. It is important to weigh the muscles for any chronic loading experiments. Acute studies of one session or less than 2 weeks of repetitive loading will likely not result in much of a change in muscle wet weight, so this measurement is not needed for very acute studies. 4. For measures of oxidative stress, the muscles should be minced and washed in phosphate buffered saline (PBS) to remove blood in the muscle. 5. Obtain an appropriate (e.g., 100 mg) section of the muscle and prepare fresh tissue for assays to determine muscle reduced glutathione (GSH) to oxidized glutathione (GSSG). You must combine several muscles to obtain enough tissue when working with mice. 6. Quickly freeze the remainder of the muscle in liquid nitrogen and keep it in a −80°C freezer to preserve enzyme activity. 7. Determine the best way to prepare tissue for your assays, because some assays require fresh muscle to be homogenized, and in other assays, the homogenates can be frozen.

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8. In addition to the assays described below, most of the proteins in the oxidative stress pathway can also be assessed by routine sodium dodecyl sulfate polyacrylamide gel electrophoresis (SDS-PAGE) analyses (1, 5, 41). 3.10. Determining the GSH/GSSG Ratio

The assay for quantitative determination of amounts of total glutathione (GSH + GSSG) was modified from that originally described by Tietze (45). The method employs Ellman’s reagent (5,5c-dithiobis-2-nitrobenzoic acid or DTNB), which reacts with GSH to form a spectrophotometrically detectable product at 412 nm. GSSG can be determined by the reduction of GSSG to GSH, which is then determined by the reaction with Ellman’s reagent. The change in color development is monitored during the reaction at 412 nm, and the reaction rate is proportional to the GSH and GSSG concentrations. For additional information see Afzal et al. (46) and Note 11. 1. For GSH, homogenize 40 mg of fresh (not frozen) muscle tissue immediately after dissection in 530 PL of cold 5% MPA. 2. For GSSG, homogenize 40 mg of fresh (not frozen) muscle tissue immediately after dissection in 500 PL of cold 5% MPA and 30 PL of M2VP to scavenge GSH. 3. Freeze GSH and GSSG homogenates in liquid nitrogen and store them at −80°C until analyzed. 4. Thaw the samples on wet ice on the day you will run the assay. 5. Add 290 PL cold 5% MPA to 500 PL of the GSSG sample. 6. Add 350 PL cold 5% MPA to the 500 PL sample for GSH. 7. Vortex homogenates and then centrifuge them at 1,000 × g for 10 min. 8. For the GSSG sample, add 25 PL of MPA extract and 350 PL of GSSG to each tube, then place them on ice until use. 9. Add the MPA extract (10 PL) and assay buffer (600 PL) to the GSH sample, then place them on ice. 10. Mix 50 PL of GSH or GSSG homogenate sample and 50 PL of DNTB (chromagen) in a cuvette. 11. Incubate the cuvette and contents for 5 min at room temperature (20°C). 12. Add 50 PL of NADPH to each cuvette. 13. Read the absorbance of each sample every 60 s at 412 nm for 3 min. 14. Determine the protein concentration for each sample via a DC Protein Assay (Bio-Rad). Signals from each sample should then be normalized to the corresponding protein content of that sample.

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15. Calculations (a) The calculation of the GSH and GSSG concentrations and the GSH/GSSG ratio requires (1) Determination of the reaction rate, (2) construction of calibration curves, (3) calculation of the concentrations of GSH and GSSG, and (4) calculation of the GSH/GSSG ratio. (b) The change in absorbance at 412 nm is a linear function of the GSH concentration in the reaction mixture, is described by: A412 = slope × minutes + intercept. The slope of the regression equation is equal to the rate. The intercepts for these rate curves are ignored because they are dependent on the DTNB background and the time interval between the addition of the NADPH (reaction start) and the first recorded A412 measurement. (c) The general form of the regression equation describing the calibration curve is: Net rate

slope u GSH  intercept.

To calculate the sample concentration from the GSH calibration curve: GSH

Net rate  intercept u dilution factor . Slope

(d) The GSH/GSSH Ratio is calculated by dividing the difference between the GSH + GSSG concentrations (reduced GSH) by the concentration of GSSG. GSSG ratio 3.11. Hydrogen Peroxide (H2O2) Levels

GSH  2GSSG . GSSG

Superoxide is converted to H2O2 either spontaneously or by superoxide dismutase. H2O2 is very stable and it is also membrane permeable. Although it is not an initiator of oxidative stress, it is frequently chosen as a marker for the general level of oxidants in resistance-loaded muscle cells (1, 5, 41). N-acetyl-3, 7-dihydroxyphenoxazine (Amplex Red) is a highly selective compound that is used to measure H2O2 in muscle samples. H2O2 oxidizes the detection reagent in a 1:1 stoichiometric ratio to produce a fluorescent product resorufin (see Note 12). This assay is based on the method of Zhou et al. (47) (see also Note 13). 1. Prior to use, dilute Amplex red stock to 10 PM with 50 mM Tris, pH 7.4. 2. Prepare a standard curve of HRP from 0 to 20 mU/mL (100 PL volumes of each concentration) in 50 mM Tris HCl.

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3. Homogenize 50–75 mg of muscle in 1× PBS. 4. Mix 10 PL H2O2 and 1 U/mL HRP in 50 mM Tris HCl, pH 7.4. Incubate at room temperature for 5 min. Dilute this by threefold and this will become the assay reagent. 5. Mix 50 PL of controls, samples, or H2O2 dilutions with 50 PL of the assay reagent in each well to initiate the reaction. 6. Incubate the plate in the dark for 10 min at 20°C. 7. Detect the fluorescence with an excitation at 563 nm and emission at 587 nm. 8. Perform all analyses in duplicate or triplicate. 9. Obtain both positive and negative (no HRP) controls if a standard curve is not used. 10. Calculate the average absorbance of each standard and sample. This is a linear relationship from the standards. 11. Correct for background (from the blank). 12. Obtain the protein concentration of the sample via a DC Protein Assay (Bio-Rad). Normalize the data from the muscle homogenates to muscle protein concentration in each sample. 3.12. 8-Hydroxy-2 cDeoxyguanosine (8-OHdG)

Although multiple oxidative lesions are possible in DNA, 8-hydroxylation of the guanine base is one of the most abundant. Measurement of 8-OHdG in loaded muscle tissue is obtained by a competitive enzyme-linked immunosorbent (ELISA). This is based on the method of Yin et al. (48) and Chiou et al. (49). Although it is possible to design your own ELISA plate as described below, we have found that commercially available ELISA for this purpose provides a sensitive measure of 8-OHdG in muscle samples (see Note 13). 1. Conjugate 8-OHG with BSA according to the method described by Senapathy et al. (50). 2. Add 100 PL of the 1 Pg/mL 8-OHG-BSA (15 ng BSA/mL) conjugate to each well and incubate overnight at 4°C. 3. Remove the 8-OHG coating solution and wash once with distilled H2O. 4. Block the wells with 350 PL of blocking buffer for 6 h at room temperature. 5. Wash the microplate with PBS immediately before conducting the assay. 6. Make standards from pure 8-OHdG. Dilute the standards in PBS at concentrations between 0.5 and 100 ng/mL (see Note 14). 7. Extract DNA from the muscle tissue by using a DNeasy Tissue Kit (Qiagen).

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8. Measure DNA purity via a plate reader. Use the DNA for the assay only if it has a minimum 260:280 ratio of 1.8. 9. Add 50 PL of purified DNA (or standard) to each well of the ELISA plate. 10. Add 50 PL of primary 8-OHdG antibody (diluted 1:10,000 in PBS containing 1% BSA) to each plate. Do not add the antibody to the blank well. 11. Incubate the samples at 37°C for 1 h. 12. Wash the wells with 5× PBS. 13. Add 100 PL of HRP secondary antibody in 1× PBS, at 37°C for 1 h. 14. Add 100 PL of chromagen (3,3c,5,5c-tetramethylbenzidine) to each well. 15. Cover the plate with a plate seal. Shake the plate and incubate it at room temperature in the dark for 15 min. 16. Stop the reaction with 100 PL 1M phosphoric acid per well. 17. Let the plate sit for 3 min at room temperature, then read the plate at an absorbance of 450 nm. 18. Normalize the absorbance of each well to the DNA concentration of the sample in the well. 19. Do analyses in triplicate. If a standard curve is not generated, the data can be expressed as relative absorbance units normalized to DNA. 3.13. Lipid Peroxidation

MDA and 4-hydroxyalkenals (HAE) are abundant components of reactive carbonyl compounds. Therefore, measurement of MDA is widely used as an indicator of lipid peroxidation (51). The MDA assay (52) is based on the reaction of a chromogenic reagent, N-methyl-2-phenylindole (R1, NMPI), with MDA (see Note 15). One molecule of MDA reacts with two molecules of NMPI to yield a stable carbocyanine dye. 1. Homogenize 75 mg of muscle in 500 PL of homogenization buffer and 5 PL of 0.5M butylated hydroxytoluene in acetonitrile. 2. Centrifuge the homogenate at 3,000 × g for 5 min at 4°C for 10 min. 3. Collect the supernatant for the subsequent assay. 4. To a clean tube, add 200 PL of the sample (or standard) to 650 PL of MP. 5. Prepare a sample blank in which 200 PL of water is added to 650 PL of MP. 6. Add 150 PL of 37% HCl to initiate the reaction. Cap tube, vortex briefly.

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7. Incubate at 45°C for 60 min. 8. Centrifuge at 12,000 × g for 10 min. Collect the supernatant. 9. Measure the 586 nm absorbance (A) of the supernatant (see Note 16). 10. Normalize the samples for differences for protein in each sample as determined by a DC Protein Assay (Bio-Rad). 11. Conduct the calculations for the assay. To do this, subtract the average A586 value for the zero concentration standard from the average A586 values of the other standards and the average sample A586 value to give corrected absorbance. Use the following equation for calculating (MDA): [MDA]

(Sample A586 corrected for blank)  b u dilution factor, a

(MDA) = concentration of MDA in the sample A586 = absorbance at 586 nm of sample a = regression coefficient (slope) b = intercept 3.14. Catalase Activity

Catalase is an antioxidant enzyme, which catalyzes the ROS product of H2O into water. Catalytic activity

catalase 2H 2O2   o O2  2H 2O

The catalase enzyme activity (EC 1.11.1.6; 2H2O2 oxidoreductase) assay is based on the reaction of the enzyme with methanol in the presence of an optimal concentration of H2O2 (53). The formaldehyde produced is measured spectrophotometrically with 4-amino-3-hydrazino-5-mercapto-1,2,4-triazole (Purpald) as the chromogen (54). Purpald specifically forms a bicyclic heterocycle with aldehydes, which upon oxidation changes from colorless to a purple color in the reaction: H 2O2  CH3OH o CH 2O  2H 2O2 (see Note 17). 1. Mince 50–75 mg of muscle in PBS to remove blood. 2. Homogenize tissue on ice in cold phosphate buffer (e.g., 50 mM potassium phosphate, ph 7.0 containing 1 mM EDTA). 3. Centrifuge at 10,000 × g for 10 min. 4. Collect the supernatant. Proceed to step 5, or freeze the supernatant at −80°C until analysis. 5. Equilibrate all reagents (but not the samples) to room temperature before beginning the assay. 6. Prepare the formaldehyde standards by diluting 10 PL of 4.25M formaldehyde with 9.99 mL of 1× sample buffer to obtain a 4.25 mM formaldehyde standard stock solution.

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7. Make formaldehyde standard wells by adding 100 PL of 1× assay buffer, 30 PL of methanol, and 20 PL of formaldehyde standards in designated wells on the plate. 8. Dilute 40 PL of 8.82M H2O2 in 9.96 mL of HPLC grade H2O. This dilute solution is stable for 2 h. Keep on ice. 9. Add 100 PL of 1× assay buffer, 30 PL of methanol, and 20 PL of diluted bovine catalase to two designated positive control wells. 10. In duplicate, add 100 PL of 1× Assay Buffer, 30 PL of methanol, and 20 PL of muscle homogenate to the sample wells. To obtain reproducible results, the amount of catalase added to the well should result in an activity between 0.25 and 4 nmol/ min/mL. When necessary, samples should be diluted with 1× sample buffer, or if it is below this level, it should be concentrated (e.g., with an Amicon centrifuge concentrator with a molecular weight cut-off of 100,000 to bring the enzymatic activity to this level). 11. Add 20 PL of H2O2 to each well to initiate the reaction. Make sure to note the precise time the reaction is initiated and add the H2O2 as quickly as possible with a multipipettor. 12. Cover the plate with the plate sealer and incubate on a shaker for 20 min at room temperature. 13. Add 30 PL of KOH to each well to terminate the reaction. 14. Add 30 PL of Purpald (Chromogen) to each well. 15. Cover the plate with the plate sealer and incubate for 10 min at room temperature on a shaker. 16. Add 10 PL of KIO4 to each well. Cover with the plate sealer and incubate for 5 min at room temperature on a shaker. The final volume of the assay is 240 PL in all the wells. 17. Read the absorbance at 550 nm using a plate reader. 18. Complete the calculations for the assay. (a) Purpald specifically forms a bicyclic heterocycle with aldehydes, which upon oxidation changes from colorless to a purple color. One unit of catalase activity is defined as the amount of enzyme that will cause the formation of 1 Pmol formaldehyde per min at 25°C (Pmol/min/mL). (b) Average the OD at 550 nm of replicate wells of each catalase standard, sample, and blank. Subtract the average 550 nm OD value of the blank from the average 550 nm OD values obtained with all other samples. (c) Make a standard curve by plotting 550 nm OD as a function of catalase concentration. Determine the equation and R2 value of the trend line.

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(d) Use the equation of the trend line of the standard curve: y = Ax + B, to calculate the catalase concentration of samples as follows: [Catalase]

(OD550 nm  B) / A.

(e) Normalize the samples to the muscle protein in each sample via a DC Protein Assay (Bio-Rad). 3.15. MnSOD and CuZnSOD

Both total SOD activity and MnSOD activity are obtained using modifications of the methods of Spitz and Oberley (55) and Beauchamp and Fridovich (56). CuZnSOD is determined by subtracting MnSOD activity from total SOD activity (see Note 18). All analyses are measured in duplicate and the samples are normalized to Pg of protein per PL of muscle homogenate. The reaction mixture of the assay (200 PL) will contain 0.1 mM xanthine, 0.025 mM NBT, 0.1 mM EDTA, 0.06M sodium carbonate buffer (pH 10.2), and xanthine oxidase. 1. Prepare blanks and standards. Standards will be 5 Pg/mL bovine CuZnSOD, with dilutions in 0.05M potassium phosphate buffer, pH 7.8. 2. Mince 75 mg of muscle tissue with scissors. Rinse in PBS. Centrifuge at 200 × g and remove supernatant. 3. Homogenize pellet in homogenization buffer. 4. Centrifuge the homogenate at 1,000 × g for 10 min. 5. Collect the supernatant in a fresh tube. 6. Add 10 PL of homogenate to each well of a 96-well plate. Add samples in quadruplicate; 2 for MnSOD and 2 for CuZnSOD. 7. Add 180 PL of assay buffer to each well. 8. Add 10 PL of 12 mM KCN to inhibit CuZnSOD and extracellular SOD activity to two wells for each sample. 9. Add 10 PL of phosphate buffer to the two wells containing the samples to be assayed for MnSOD. 10. Add 20 PL of xanthine oxidase. Use multipipettor so all of the wells are started at the same time. 11. Protect the reagents and samples from white light and incubate them at 26°C for 20 min with periodic shaking. 12. Read the absorbance at 450 nm using a 96-well plate reader. 13. Calculate the average absorbance of each standard and sample. This is a linear relationship from the standards. Normalize to the protein content of each sample. One unit of SOD is defined as the amount of enzyme that inhibits 50% of the rate of NBT reduction at room temperature. Calculate MnSOD from the absorbance from wells containing KCN, because KCN inhibits CuZnSOD activity (57).

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4. Notes 1. Aurora Scientific sells an in situ rat system (1305A) that should serve the same general purposes for most experiments. 2. We use custom Labview software to control the stimulator, the servomotor, and the stimulation parameters. However, commercial software (DMA, Aurora Scientific) could also be used for this purpose. 3. We use the commercially available software for off-line analysis of the force data (DMC, Aurora Scientific). 4. An in situ system is commercially available by Aurora Scientific (1300A, Aurora). The footplate will have to be manufactured to fit on the servomotor (see Fig. 2b). 5. We have found that longer stimulating electrodes are needed for some experiments, especially in old or obese rats. 6. It is convenient to develop or purchase software to control the electrical stimulator. We use DMC software (Aurora Scientific) and it works well for this purpose. You will need an A/D board for this to interface with the stimulator and computer. 7. Twitches consist of one single electrical pulse, so the frequency is irrelevant. 8. Submaximal contractions could be used for training purposes. For example, determine what the maximal force output is, and then reduce the electrical stimulation parameters (voltage and/ or frequency of stimulation) to obtained the desired effort for training efforts (e.g., at 60% of maximal force output). 9. The correct positioning of the electrode is an art. It may be necessary to reposition the stimulating electrodes, if inappropriate muscles are contracting (e.g., based on palpating the hindlimb muscles). Also, look to avoid excessive eversion or inversion of the foot during the stimulus. 10. Long-term exercise studies can be conducted with internal dwelling wire electrodes placed around the appropriate nerve (e.g., tibial nerve or common peroneal nerve). This method has been described by Warren et al. (58). In brief, a shielded electrode wire (e.g., platinum) is threaded from the dorsal scapular region, subcutaneously along the spine, down the hip to the area of the knee. The externalized wire and plug (on the back) is connected to the stimulator during the loading sessions. Although it works well, there is also a ~30% failure rate with this approach. Nevertheless, when it works, it does make stimulus location constant and reduces the variability associated with electrode placement.

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11. A BIOXYTECH GSH/GSSG-412 assay system (Percipio Biosciences, Inc.) can be purchased to determine the GSH/ GSSG ratio. 12. High levels of HRP (100 mU/mL, final concentration) will produce lower fluorescence than 1 mU/mL, because the excess HRP oxidizes the fluorescent reaction product, resorufin to nonfluorescent resazurin. 13. H2O2 can be determined by a commercial fluorescent H2O2 detection kit (Fluoro H2O2, Cell Technology). 14. 8-OHdG can be measured and quantified by a commercial competitive ELISA kit (Percipio Biosciences, Inc.). Otherwise, the data can be expressed as relative absorbance units normalized to DNA. 15. MDA can be measured using reagents from Percipio Biosciences, Inc. 16. In the MDA assay, the color from the MDA assay is stable for at least 2 h at room temperature (52). 17. Catalase can be assayed by a commercially available kit (EMD Biosciences). 18. A commercially available SOD Assay Kit II (EMD/Calbiochem) can be used to measure total and MnSOD activity. References 1. Ryan, M. J., Dudash, H. J., Docherty, M., Geronilla, K. B., Baker, B. A., Haff, G. G., Cutlip, R. G., and Alway, S. E. (2008) AgingDependent Regulation of Antioxidant Enzymes and Redox Status in Chronically Loaded Rat Dorsiflexor Muscles, J Gerontol A Biol Sci Med Sci 63, 1015–1026. 2. McBride, J. M., Kraemer, W. J., TriplettMcBride, T., and Sebastianelli, W. (1998) Effect of resistance exercise on free radical production, Med. Sci. Sports Exerc. 30, 67–72. 3. Gianni, P., Jan, K. J., Douglas, M. J., Stuart, P. M., and Tarnopolsky, M. A. (2004) Oxidative stress and the mitochondrial theory of aging in human skeletal muscle, Exp. Gerontol. 39, 1391–1400. 4. Baker, B. A., Hollander, M. S., Kashon, M. L., and Cutlip, R. G. (2010) Effects of glutathione depletion and age on skeletal muscle performance and morphology following chronic stretch-shortening contraction exposure, Eur. J. Appl. Physiol. 108, 619–630. 5. Ryan, M. J., Jackson, J. R., and Alway, S. E. (2010) Suppression of oxidative stress by resveratrol after isometric contractions in gas-

trocnemius muscles of aged mice, J Gerontol A Biol Sci Med Sci 65, 815–831. 6. Harman D. (1956) Aging: a theory based on free radical and radiation chemistry, J Gerontol. 11, 298–300. 7. Uchiyama, S., Tsukamoto, H., Yoshimura, S., and Tamaki, T. (2006) Relationship between oxidative stress in muscle tissue and weightlifting-induced muscle damage, Pflugers ArchivEuropean Journal of Physiology 452, 109–116. 8. Urso, M. L. and Clarkson, P. M. (2003) Oxidative stress, exercise, and antioxidant supplementation, Toxicology 189, 41–54. 9. Baker, B. A., Hollander, M. S., Mercer, R. R., Kashon, M. L., and Cutlip, R. G. (2008) Adaptive stretch-shortening contractions: diminished regenerative capacity with aging, Appl Physiol Nutr Metab 33, 1181–1191. 10. Cutlip, R. G., Baker, B. A., Geronilla, K. B., Mercer, R. R., Kashon, M. L., Miller, G. R., Murlasits, Z., and Alway, S. E. (2006) Chronic exposure to stretch-shortening contractions results in skeletal muscle adaptation in young rats and maladaptation in old rats, Appl Physiol Nutr Metab 31, 573–587.

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11. Geronilla, K. B., Miller, G. R., Mowrey, K. F., Wu, J. Z., Kashon, M. L., Brumbaugh, K., Reynolds, J., Hubbs, A., and Cutlip, R. G. (2003) Dynamic force responses of skeletal muscle during stretch-shortening cycles, Eur J Appl Physiol 90, 144–153. 12. Cutlip, R. G., Baker, B. A., Hollander, M., and Ensey, J. (2009) Injury and adaptive mechanisms in skeletal muscle, J Electromyogr Kinesiol 19, 358–372. 13. Cutlip, R. G., Geronilla, K. B., Baker, B. A., Kashon, M. L., Miller, G. R., and Schopper, A. W. (2004) Impact of muscle length during stretch-shortening contractions on real-time and temporal muscle performance measures in rats in vivo, J Appl Physiol 96, 507–516. 14. Gosselin, L. E. and Burton, H. (2002) Impact of initial muscle length on force deficit following lengthening contractions in mammalian skeletal muscle, Muscle Nerve 25, 822–827. 15. Cutlip, R. G., Baker, B. A., Geronilla, K. B., Kashon, M. L., and Wu, J. Z. (2007) The influence of velocity of stretch-shortening contractions on muscle performance during chronic exposure: age effects, Appl Physiol Nutr Metab 32, 443–453. 16. Baker, B. A., Mercer, R. R., Geronilla, K. B., Kashon, M. L., Miller, G. R., and Cutlip, R. G. (2007) Impact of repetition number on muscle performance and histological response, Med Sci Sports Exerc 39, 1275–1281. 17. Cutlip, R. G., Geronilla, K. B., Baker, B. A., Chetlin, R. D., Hover, I., Kashon, M. L., and Wu, J. Z. (2005) Impact of stretch-shortening cycle rest interval on in vivo muscle performance, Med Sci Sports Exerc 37, 1345–1355. 18. McCully, K. K. (1986) Exercise-induced injury to skeletal muscle, Fed. Proc 45, 2933–2936. 19. Lieber, R. L., Woodburn, T. M., and Friden, J. (1991) Muscle damage induced by eccentric contractions of 25% strain, J Appl Physiol 70, 2498–2507. 20. Brooks, S. V. and Faulkner, J. A. (1990) Contraction-induced injury: recovery of skeletal muscles in young and old mice, Am J Physiol 258, C436–C442. 21. Koh, T. J. and Brooks, S. V. (2001) Lengthening contractions are not required to induce protection from contraction-induced muscle injury, Am J Physiol Regul Integr Comp Physiol 281, R155–R161. 22. Figueiredo, P. A., Powers, S. K., Ferreira, R. M., Appell, H. J., and Duarte, J. A. (2009) Aging impairs skeletal muscle mitochondrial bioenergetic function, J Gerontol A Biol Sci Med Sci 64, 21–33.

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Chapter 12 Analysis of Skeletal Muscle Hypertrophy in Models of Increased Loading Sue C. Bodine and Keith Baar Abstract Skeletal muscle is a highly adaptive tissue that modifies its size in response to a variety of external stimuli. In adult mammals, skeletal muscle hypertrophy occurs primarily as a response to increases in external loading. Here, we describe the methods that should be used for a comprehensive assessment of muscle hypertrophy in animal models. The methods include the measurement of muscle mass, fiber cross-sectional area, contractile function, and protein concentration. Key words: Resistance exercise, Muscle hypertrophy, Physiological cross-sectional area, Specific tension, Isometric maximal force

1. Introduction The ability to quantify skeletal muscle hypertrophy and detect enhanced or defective growth in adult mammals is a valuable tool for assessing genetically altered mice, therapeutic interventions for the treatment of disease, and the effects of aging. Skeletal muscle hypertrophy in adult mammals occurs in response to an increase in mechanical loading (1). In humans, resistance exercise training consisting of intermittent bouts (2–3 times per week) of low frequency repetitions (3–10 sets of 6–8 repetitions/set) with high loads (65–85% of maximum voluntary contraction) is used to illicit increases in muscle size and strength. The rapid increase in strength that occurs in humans is primarily the result of neural adaptation, while muscle hypertrophy occurs more slowly with a time course of weeks to months (2).

Joseph X. DiMario (ed.), Myogenesis: Methods and Protocols, Methods in Molecular Biology, vol. 798, DOI 10.1007/978-1-61779-343-1_12, © Springer Science+Business Media, LLC 2012

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A number of animal models have been developed over the years in an attempt to understand the mechanisms responsible for muscle growth (see refs. (2–4) for review). These models mimic human resistance training in that they increase the load on the skeletal muscle. However, the increase in load is often more severe than occurs in humans, and thus muscle growth (especially in rodents) occurs more rapidly and to a greater extent than in humans. That said, the advantage of animal models over human models is that: (1) nutritional and environmental factors can be tightly regulated, (2) pharmacologic agents and tracers can be used, (3) functional properties of single muscles can be directly determined either in situ or in vitro, (4) whole muscles can be removed for morphological, histochemical, biochemical, and molecular analysis; and finally (5) genetic manipulation can be performed to assess the contribution of specific genes to the regulation of muscle size. Two commonly used animal models of overload are compensatory hypertrophy (CH) (also known as functional overload and synergist ablation) and electrical stimulation (ES). The CH model was developed in the 1970s (5, 6) as a modification of the tenotomy model (7), and involves the complete removal of one or more synergists in a group of muscles that perform the same physiological function at a single joint. The plantaris muscle, an ankle extensor, is most commonly overloaded through the removal of the gastrocnemius (both medial and lateral heads) and the soleus (8, 9). Following surgical overload, the plantaris experiences an increase in both neural activity and loading during normal cage activity resulting in a very strong growth stimulus. Subsequently, significant hypertrophy of the plantaris occurs within 7 days (~30%) and reaches a plateau at approximately twice its original size following 8 weeks in rodents (see Fig. 1 and ref. (10)). Another muscle that can be studied is the extensor digitorum longus (EDL) with the removal of the primary dorsiflexor of the ankle, the tibialis anterior (TA) muscle. Since the EDL and TA experience less force than the plantar flexor muscle during the step cycle, this model results in a slower rate of hypertrophy (30% in 4 weeks) (11). ES of the hindlimb muscles while the animal is under anesthesia is another model that has been utilized to induce skeletal muscle hypertrophy. The ES model, first developed by Wong and Booth (12), has gone through several modifications. The method most commonly used today is one in which the sciatic nerve is electrically stimulated resulting in maximal activation of all of the muscles of the lower limb. Upon stimulation, the ankle joint extends (plantar flexion) due to the greater muscle mass in the posterior vs. the anterior compartment of the lower leg. Since all the muscles below the knee are maximally activated, the posterior (extensor) muscles undergo shortening (concentric) contractions, while the anterior (flexor) muscles undergo lengthening (eccentric) contractions.

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Fig. 1. (a) The mass of the rat plantaris muscle was measured following no treatment (control) or 1, 3, 7 14, and 21 days following functional overload and is presented as both absolute wet weight, in milligrams, and (b) percent change from control. The increase in mass at day 1 post overload is primarily due to edema. (c) The mass of the rat tibialis anterior (TA) and medial gastrocnemius (MG) was measured in a cohort of control rats at the start of the experiment and at the completion of the 28-day experiment. The hind limbs of young adult female rats (8 weeks old) were unloaded for 14-days, followed by 0, 1, 3, 7, or 14 days of reloading. Data are means ± SD, n = 8–10 per time point.

A typical resistance exercise training protocol consists of 6–10 sets of six repetitions delivered twice per week for 6 weeks. This protocol produces approximately 14% hypertrophy in the flexor muscles (TA and EDL) at the end of 6 weeks (13). Additional models of overload have been utilized and are described in detail elsewhere (2). The models described above are generally used to induce muscle growth in otherwise “normal” resting muscle, although they also can be performed in muscle subjected to atrophy inducing treatments such as disuse or glucocorticoids. Muscle growth or “regrowth” following a period of atrophy induced by unloading is another effective model of hypertrophy. Here, an animal is simply returned to normal locomotor loading patterns, i.e., reloaded, following a period of unloading. Reloading or recovery of muscle mass following unloading has been studied following cast immobilization (14, 15) and hindlimb suspension (16, 17) (see Note 1).

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The final model of muscle hypertrophy that should be mentioned is the genetic induction model. Spontaneous mutations followed by breeding of animals with increased muscle mass in commercial farming have resulted in animals with extreme increases in muscle size (e.g., Belgian Blue Cattle (18) and Callipyge lambs (19)). Identification of the genes underlying these mutations and more has resulted in the development of mice with genetic alterations (both deletions and over-expression) resulting in animals with larger muscles (18, 20). As our understanding of the genetic basis for hypertrophy improves, genetic models of increased muscle mass are becoming more common. Since in many cases the genetically altered gene is expressed not only in adult muscle, but also in developing muscle cells, the basis for the increase in muscle size may be very different than what is seen in adult human muscle undergoing resistance exercise. Therefore, it is important to use the proper tools to assess fiber size, muscle mass and function in an attempt to determine the mechanism(s) underlying the increase in muscle size. Regardless of the model used, comprehensive assessment of hypertrophy should include the following: (1) measurement of muscle wet weight, (2) histological measurement of fiber cross-sectional area and number, (3) measurement of contractile function, and (4) determination of protein concentration and content. Additionally, biochemical markers of hypertrophy can be assessed to identify potential defects or enhancements in the growth response.

2. Materials 2.1. Mass

1. Surgical tools for dissection of the muscles. 2. Dissecting scope or surgical loops. 3. Physiological saline (0.9% NaCl). 4. Balance capable of weighing masses of 5–200 mg in mice or 50–2,000 mg in rat.

2.2. Fiber CrossSectional Area

1. Cork, minutin pins (10 mm), parafilm. 2. Tissue Freezing Medium (example, OCT [optimal cutting temperature]). 3. Isopentane (2-methyl-butane). 4. Thermos or dewer, metal crucible for isopentane. 5. Liquid nitrogen. 6. Cryostat with rotary microtome. 7. Light microscope equipped with digital camera and image acquisition/measurement software.

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1. Clamps and frame for securing limb. 2. Nerve Stimulator, stimulating electrode. 3. Force transducer, silk thread for tying tendon. 4. Thermister to monitor body and muscle temperature. 5. Computer with analog-to-digital converter and software for data acquisition and analysis.

2.4. Protein Concentration/Content and Biochemistry

1. Motor and pestle. 2. Homogenizer. 3. Cold room shaker. 4. Micropipettes. 5. Bradford reagent. 6. Spectrophotometer.

3. Methods 3.1. Muscle Mass

1. Determine muscle mass by weighing whole muscles that have been carefully dissected free from their proximal and distal attachment sites in the limb. 2. Clean the dissected muscle belly of fat and connective tissue. Cut long tendons as close as possible to the end of the fibers. 3. Wash the dissected muscle in saline to remove fur and blood, and then blot dry prior to weighing. 4. Wet weight is most often expressed in absolute units of mass (e.g., milligrams or grams), but can also be expressed relative to body mass as a ratio: mg MW/g BW (see Fig. 1a; Note 2). 5. Changes in muscle mass, as the result of overload, can be assessed by converting absolute mass to a percent change relative to control ((overload-control)/control × 100] (see Fig. 1b; Note 3).

3.2. Fiber CrossSectional Area

Fiber cross-sectional area is measured from serial cross-sections taken from frozen muscle, and provides information regarding changes in the size of individual muscle fibers of different myosin (type I, IIa, IIx, or IIb) (21) or metabolic (slow oxidative [SO], fast oxidative glycolytic [FOG], or fast glycolytic [FG]) (22) phenotypes. Cross-sectional area measurements can also be correlated to maximum force measurements and used to calculate specific tension (force per cross-sectional area, see later). 1. Rapidly freeze the whole muscle, or a portion taken from the belly of the muscle, in isopentane chilled with liquid nitrogen

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Fig. 2. (a) The mouse gastrocnemius muscle (MG and lateral gastrocnemius [LG]) is shown pinned at a fixed length using minutin pins onto cork. To prevent the muscle from sticking to the cork, a piece of parafilm is placed between the muscle and the cork. (b) Isopentane is place in a metal crucible, which is lowered into a dewer with liquid nitrogen. When a frozen layer of isopentane forms on the bottom of the crucible, the isopentane is ready to be used for freezing of the muscle as shown in (c). At a later time, a “block” of the frozen muscle can be taken from the mid belly, as shown in (d) for frozen sectioning.

(see Fig. 2). Freeze muscle by one of two methods: (1) the whole muscle is pinned at a fixed length, approximately the in vivo resting length, onto cork followed by rapid freezing (see Note 4); or (2) a portion of the muscle is cut from the mid-belly of the muscle (making sure that the cut is perpendicular to the length of the fibers), positioned with the distal end down on a piece of cork, submerged in OCT (or tissue freezing medium), and rapidly frozen (see Note 5). 2. Cut serial cross-sections (6–10 Mm thick) from the frozen tissue block using a rotary microtome in a cryostat at −20°C and placed on glass slides. 3. Stain tissue sections with hemotoxylin and eosin to assess general morphology. Make fiber cross-sectional area measurements from sections stained for H&E or from sections stained with anti-laminin antibodies that detect the basal lamina surrounding the individual fibers (see Note 6). 4. Measure fiber cross-sectional area using a light microscope equipped with a digital camera and software for image acquisition and analysis. To obtain an accurate measure of mean fiber cross-sectional area, and detect changes resulting from the hypertrophic stimulus, a representative sampling of the fibers in the muscle must be made. Since many muscles have regional

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Fig. 3. (a) Serial cross-section of the mouse triceps surae complex (contains MG, LG, soleus [Sol], and plantaris [PL] muscles) stained with an anti-myosin heavy chain fast (MHC-f) antibody. For the measurement of fiber cross-sectional area of the gastrocnemius muscle, multiple regions, as delineated by the rectangles, should be analyzed. Analysis of the soleus and plantaris muscles can also be made from this cross-section. (b) Muscle cross-section stained with anti-laminin antibody for identification of the basal lamina of individual muscle fibers. (c) Serial cross-section of the mouse triceps surae complex histochemically stained for succinate dehydrogenase (SDH). Note the regional differences in staining density that indicates differences in oxidative capacity. The inset illustrates the typical fiber size difference between oxidative and nonoxidative fibers in a muscle of mixed fiber type.

variations in fiber types (fast vs. slow and oxidative vs. nonoxidative), it is recommended that 100–200 fibers be sampled from multiple regions throughout the muscle cross-section to obtain an unbiased estimate of mean fiber cross-sectional area (see Fig. 3a, Note 7). 5. To assess differences in the response of slow and fast fibers to overload, use commercially available primary antibodies against slow (type I) and fast (all type II) myosin heavy chains (MHC) to broadly classify fibers as type I (slow) or type II (fast) fibers. Additional primary antibodies are available to detect the different type II (IIa, IIx, and IIb) isoforms for more specific classification of the fast fibers (21) (see Note 8). 6. To identify differences in the metabolic characteristics of individual fibers, use histochemical techniques to classify fibers as SO, FOG, and FG. For metabolic classification, stain serial

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Fig. 4. (a) Histogram of mean fiber cross-sectional area (in Mm2) in the rat plantaris muscle following no treatment (CON) and 14-days of compensatory hypertrophy (CH). Muscle fibers were classified as slow or fast based on staining with an anti-MHC-f antibody. Data are means ± SD, n = 10 per group. (b) Histogram showing the distribution of fiber cross-sectional areas (in Mm2) in the TA muscle following injection of a DNA plasmid expressing either EGFP (CON, white bars) or activated AKT-EGFP (PLASMID, black bars) for 7 days.

sections for the following contractile and metabolic enzymes: myosin adenosine triphosphatase (myofibrillar ATPase with alkaline or acid preincubation), alpha-glycerophosphate dehydrogenase, and succinate dehydrogenase (23). It is important to note that metachromatic staining is only a qualitative measure of a fiber’s phenotype. 7. Calculate mean fiber cross-sectional area for all fibers and separately for different fiber types in response to overload. In addition, plot the distribution of cross-sectional area as a histogram to identify subtle or significant shifts in fiber size and variability within the population (see Fig. 4). 8. The cross-sectional area of a muscle can change due to “hyperplasia” due to either the splitting of existing fibers or the addition of new fibers (24, 25). Hyperplasia is not thought to be a major mechanism for increasing muscle mass in response to increased loading in mammalian muscles. However, increases in the number of fibers per muscle have been documented in genetic models of increased muscle mass (18). Estimate muscle fiber number from a single cross-section taken from the mid belly of the muscle. The accuracy of this measurement depends on the architecture of the muscle (26) (see Note 9). 3.3. In Situ Contraction Testing

Although muscle cross-sectional area is generally related to the ability to generate force, pathological forms of hypertrophy can result in a greater increase in muscle mass than in force. Therefore, it is important to determine the extent to which increases in muscle

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mass and fiber cross-sectional area reflect improvements in tension or strength. To determine the contractile properties of individual muscles, isometric contractions are elicited at various frequencies of stimulation either in situ (in the animal via indirect nerve stimulation) or in vitro (in an oxygenated bath via direct muscle stimulation). The decision to use in situ vs. in vitro techniques to determine contractile function is dependent on (1) the muscles to be tested, and (2) the set up available to the investigator. In vitro testing of skeletal muscle is restricted to small, relatively thin muscles, such as the soleus and EDL, which can be adequately perfused and oxygenated in a bath. The thickness of the muscle can also be a factor in achieving maximal activation of the muscle fibers via direct stimulation (see Note 10). 1. For in situ testing of the lower limb muscles, mobilize the sciatic nerve from the sciatic notch to the popliteal fossa, isolating the tibial, peroneal, and sural branches of the sciatic nerve. 2. Isolate, cut, and attach in series the distal tendon of the muscle to be studied to a force transducer with silk suture. 3. Securely clamp the tibia and femur to a solid frame. 4. Maintain the nerve and muscle at 35 ± 2°C with mineral oil and heating lamps. 5. Place a bipolar electrode on the sciatic nerve to activate the muscle. Crush the nerve with forceps proximal to the stimulating electrode to prevent antidromic stimulation of the muscle. Cut the unwanted branches of the nerve distal to the crush injury so as not to stimulate the remaining muscles and introduce mechanical vibrations. 6. Amplify and record isometric tension outputs using an analogto-digital acquisition system (e.g., PowerLab, AD Instruments, Inc or LabVIEW, National Instruments). 7. At the start of the experiment, stimulate the nerve at increasing voltage until there is no further increase in force with an increase in voltage. This value, the minimum voltage amplitude necessary to induce maximal contraction, is termed the rheobase. For testing purposes, set the voltage at twice the rheobase to assure maximal activation of the muscle. 8. Next set the muscle to a length that produces maximum twitch tension (Lo). Measure twitch time to peak tension (TPT) and half relaxation time (HRT) from twitches produced at Lo and 35°C. 9. Obtain the force-frequency profile of a muscle by stimulating the muscle at frequencies between 10 and 150 Hz with contractions of 500 ms in duration (see Notes 11 and 12). 10. Record maximal isometric tension (Po) as the maximum tension achieved during the force-frequency protocol.

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11. Upon completion of the contractile testing, measure the length of the muscle (excluding tendons) with calipers. 12. Specific tension is a measure of the force produced per unit area of active contractile tissue, and usually has units of Newtons per centimeter squared (N/cm2) (see Note 13). Calculate specific tension by dividing maximum isometric force by physiological cross-sectional area (PCSA). 13. PCSA depends on muscle volume, fiber length, and angle of pennation. Calculate PCSA as the product of wet weight (g) and pennation angle (cos F) divided by the product of fiber length (cm) and muscle density. Assume the density of the muscle to be 1.056 gm/cm3, while the effects of the angle of pennation are minimal given the angles measured for mammalian muscle (26). Therefore, PCSA is primarily dependent on fiber length and muscle mass (see Notes 14 and 15). 3.4. Protein Concentration/Content

One reason that specific tension of a muscle can decrease is an increase in fluid within the fibers (edema) as a result of load-induced muscle injury. A simple and highly accurate method for assessing whether an increase in mass is due to hypertrophy or swelling is to determine protein concentration/content. If the increase in mass is due to edema, protein concentration will decrease and content will not change. If the increase in mass is due to hypertrophy, the protein concentration will remain constant and the content will increase. 1. To accurately determine protein concentration, powder the muscle of interest with a mortar and pestle on dry ice/liquid nitrogen. Remove large pieces of sinew and create a homogenous mixture of muscle to prevent local differences in fiber type, oxidative capacity, etc. from biasing the results. 2. Next place a scoop of powder into a preweighed tube and reweigh the tube to determine the exact amount of muscle used for the assay. Since the calculation of protein content is dependent on the mass of muscle used, and one scoop of muscle provides 20–40 mg, a highly sensitive scale is required for this measure. 3. Homogenize the powder using either a dounce or polytron homogenizer in 20 volumes of a high salt buffer (see Note 16). A high salt buffer is used to weaken the association of actin and myosin allowing the muscle to dissolve more completely. Shake the samples for 30 min at 4°C to further enhance the dissolution, and permit the complete extraction of the contractile proteins. 4. Following extraction, mix the samples well, dilute, and load in triplicate onto a 96-well plate in parallel with an ovalbumin standard curve (0–1 mg/mL). 5. Then, add Bradford reagent to the samples and read the plate at 595 nm on a spectrophotometer. Calculate the protein concentration of the homogenate from the standard curve and

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use this number to calculate the protein concentration (mg/mL in homogenate divided by the mg of the muscle/ml of buffer added) and content (concentration multiplied by the mass of the muscle). Muscle should normally have a protein concentration of ~20% and this fact can be used as an internal control to confirm the validity of the assay. 3.5. Experimental Design

The experimental design is critical for obtaining an accurate assessment of muscle growth in models of altered loading. 1. First, there must be the appropriate control groups and adequate sample size per group (see Note 17). For most experimental models, the ideal is to have an age-matched untreated group as the control for the experimental group (see Note 18). In some cases, it may be appropriate to treat one limb and use the contralateral limb in the same animal as the control. This should be done with caution, and intra-animal controls should be used only if the investigator has determined that the experimental procedure has no effect on the untreated limb. 2. If comparing the effect of loading on wild type and genetically altered mice, it is critical that both groups have the same genetic background. The use of wild type and genetically altered litter mates is the optimal experimental design. 3. The number of time points to be analyzed is dependent on the hypothesis being tested. In many cases, the collection of muscle samples at multiple time points is required to determine the mechanisms underlying load induced growth or alterations in the growth response to loading (e.g., the effect of the deletion of a specific gene) (see Note 19). 4. Measurement of mass, fiber cross-sectional area, and protein content can be obtained from a single muscle sample. The ability to make additional biochemical measurements depends on the size of the muscle and often requires a second set of muscles. In situ measurement of contractile function leads to fluid uptake into the muscle; consequently, the muscle wet weight measured after contractile testing is not an accurate reflection of muscle mass.

4. Notes 1. A potential advantage of the hindlimb suspension model over the immobilization model is that periods of limb immobilization can result in the development of joint stiffness and loss of joint mobility (27), whereas in the tail suspension model the joints remain mobile and do not develop contractures (unpublished observations). Consequently, following casting,

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movement at the knee and ankle joints may be restricted, especially in old rats that may be more susceptible to developing joint contractures. A loss of joint mobility will alter overall locomotor activity and the loads placed on the muscle resulting in a reduction in growth signals. 2. Caution should be used in the interpretation of changes in muscle mass if significant alterations in body weight have occurred during the course of an experiment or if the experimental groups vary significantly in body weight. 3. This measure is useful for making comparisons across experiments, and adjusts for large differences in the initial size of the animals that may occur due to age or genetic manipulations. 4. When the whole muscle is frozen at length, a portion of the frozen muscle (~5 mm block) is later cut from the mid-belly using a sharp blade, and mounted distal end down on cork in OCT prior to sectioning (see Fig. 2). Care must be taken during the blocking of the tissue to ensure that the muscle remains frozen. Thawing and refreezing of the tissue can cause ice crystal formation and the formation of large holes in the muscle fibers. 5. Both methods are appropriate for the determination of fiber cross-sectional area, but can yield different absolute values (28). Consequently, a consistent procedure should be used for an entire experiment. If using the fiber CSA measurements to calculate specific tension, freezing the muscle at a fixed length that approximates the optimal physiological length should be used. 6. If using laminin, visualization of the antibody–antigen complex with a diaminobenzidine peroxidase reaction results in a dark outline surrounding the individual fibers and assists with the measurement of cross-sectional area when using computer software that allows for threshold detection (see Fig. 3b). 7. For example, in many rodent muscles, the deep region of the muscle tends to have a high percentage of type I and IIa fiber types while the superficial region has a high percentage of type IIx and IIb fibers. Since type IIb and IIx fibers tend to be larger than type IIa and I fibers, a biased mean cross-sectional area measurement will be recorded if only one region is sampled. In a large muscle, such as the gastrocnemius muscle, sampling from four to five regions and measuring 100–200 fibers per region is suggested for an unbiased estimate of mean fiber cross-sectional area (see Fig. 3). 8. Serial sections stained with MHC antibodies that recognize the different isoforms will also identify fibers that express multiple MHC proteins, i.e., “hybrid” fibers. 9. Estimation of fiber number from a single cross-section is best done in a unipennate muscle. For muscles with short fibers and high angles of pennation, multiple cross-sections throughout

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the length of the muscle will be required to get an accurate count of fiber number. 10. The isometric contractile properties of the mouse soleus and EDL are often tested using in vitro methods, while larger muscles such as the gastrocnemius and TA must be tested in situ. In most instances, rat hindlimb muscles are tested using in situ methods; however, some labs have been successful in getting accurate tension measurements from the rat soleus and EDL using in vitro methods (29, 30). 11. Muscle force measurements should be presented as Newtons. The formula to convert tension measurements obtained from the transducer in grams to Newtons is: Force (N) equals mass (kg) × acceleration (9.81 m/s2). 12. The force-frequency profile can be presented as absolute force (Newtons) or relative force (percent of maximum tension) (see Fig. 5). The absolute force-frequency profile can reveal differences in absolute force capabilities between the control and overloaded muscles, while the relative force-frequency profile can reveal changes in the kinetics of the contractions and potential fiber type changes. A leftward shift in the relative force-frequency curve suggests a slowing of the muscle and an increase in the percentage of muscle fibers expressing slow contractile proteins (e.g., myosin ATPase and sarcoplasmic reticulum ATPase). Following resistance training in humans, skeletal muscle does not become slower (2). CH is a unique model in that the muscle experiences increases in both loading and neural activity (recruitment). In rats, there is a significant slowing of the contractile properties and increases in slow MHC in the plantaris muscle with CH (8, 9). In contrast, in mice there is no significant slowing of the plantaris in mature muscles in response to CH (10). 13. In normal mammalian muscle, the specific force of slow and fast muscle fibers is similar and ranges between 20 and 25 N/ cm2 (31). An accurate calculation of specific tension depends on the accuracy of the maximum isometric tension and PCSA measurements. Much of the variability in the literature for specific tension is related to inaccuracies in the measurement of PCSA. Calculation of PCSA is easier for unipennate muscles, such as the rodent soleus and EDL, than for bipennate or multipennate muscles. Consequently, many investigators have chosen the soleus or EDL for measurement of specific tension. 14. Following an increase in loading, if muscle strength, as measured by the maximum isometric tension, increases in proportion to muscle mass then specific tension should not change from control values. The specific tension of the rat plantaris muscle has been reported to decrease in response to 8–12 weeks of

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Fig. 5. Plot of frequency–tension relationship in the rat plantaris in response to no-treatment (normal) or 12–14 weeks of CH (overloaded). Force is expressed as absolute (N) (top), per cross-sectional area (N/cm2) (middle), and as percent maximum isometric force (bottom). Following overload, the frequency–tension profile of the plantaris shifted toward that of a normal soleus (bottom graph ). Data are means ± SE (figure is taken from Roy et al. (8)).

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CH (8, 32). In contrast, in mice and cats, there is no change in specific tension following extended periods of CH (10). 15. In most mammalian muscles, fiber length is not equal to muscle length (26, 33, 34). An accurate determination of fiber length is done through the dissection of individual fibers from muscles that have been formalin fixed at optimal length (33). Measurement of the optimal muscle length (Lo) upon completion of the contractile testing and knowledge of the ratio of fiber to muscle length will allow for a more accurate calculation of PCSA. 16. An example of a high salt buffer is: 500 mM KCl, 2 mM MgCl2, 2 mM EGTA, 2 mM Na4P2O7, and 100 mM Na-phosphate buffer, pH 6.8. 17. Sample size should be calculated using a power analysis. In general, a sample size of five or more is desirable to identify significant muscle hypertrophy. 18. In this experiment, two control groups were used to control for the growth of the young, juvenile (8 weeks old) rats that occurred over the 28-day experimental protocol. If using growing or aging animals to examine muscle hypertrophy, one must take into consideration the growth or atrophy of the animal during the course of the experiment. This experiment also shows how different muscles can respond differently to reloading, and illustrates how collecting data at multiple time points can provide valuable information (note initial decrease, followed by increase in mass in TA upon reloading). 19. The immediate time points (days 1–3) post loading often involve some period of edema, and can provide information on the inflammatory response, injury, and early transcriptional and signaling events. The intermediate time points (days 7–14 post loading) usually measure the period of rapid growth, and can provide information on the activation of signal transduction pathways and changes in muscle mass and morphology. Analysis of growth at 30 days relative to 7 or 14 days provides a stable growth phase and can provide information on adaptive changes that may be occurring in the muscle. References 1. McDonagh, M. J., and Davies, C. T. (1984) Adaptive response of mammalian skeletal muscle to exercise with high loads, Eur J Appl Physiol Occup Physiol 52, 139–155. 2. Booth, F. W., and Thomason, D. B. (1991) Molecular and cellular adaptation of muscle in response to exercise: perspectives of various models, Physiol Rev 71, 541–585.

3. Timson, B. F. (1990) Evaluation of animal models for the study of exercise-induced muscle enlargement, J Appl Physiol 69, 1935–1945. 4. Lowe, D. A., and Alway, S. E. (2002) Animal models for inducing muscle hypertrophy: are they relevant for clinical applications in humans?, J Orthop Sports Phys Ther 32, 36–43.

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5. Armstrong, R. B., Marum, P., Tullson, P., and Saubert, C. W. T. (1979) Acute hypertrophic response of skeletal muscle to removal of synergists, J Appl Physiol 46, 835–842. 6. Baldwin, K. M., Cheadle, W. G., Martinez, O. M., and Cooke, D. A. (1977) Effect of functional overload on enzyme levels in different types of skeletal muscle, J Appl Physiol 42, 312–317. 7. Goldberg, A. L. (1972) Mechanisms of growth and atrophy of skeletal muscle, Muscle Biol 1, 89–118. 8. Roy, R. R., Meadows, I. D., Baldwin, K. M., and Edgerton, V. R. (1982) Functional significance of compensatory overloaded rat fast muscle, J Appl Physiol 52, 473–478. 9. Roy, R. R., Baldwin, K. M., Martin, T. P., Chimarusti, S. P., and Edgerton, V. R. (1985) Biochemical and physiological changes in overloaded rat fast- and slow-twitch ankle extensors, J Appl Physiol 59, 639–646. 10. Roy, R. R., and Edgerton, V. R. (1995) Response of mouse plantaris muscle to functional overload: comparison with rat and cat, Comp Biochem Physiol A Physiol 111, 569–575. 11. Hamilton, D. L., Philp, A., Mackenzie, M. G., and Baar, K. (2010) A Limited Role for PI(3,4,5)P(3) Regulation in Controlling Skeletal Muscle Mass in Response to Resistance Exercise, PLoS One 5, e11624. 12. Wong, T. S., and Booth, F. W. (1990) Protein metabolism in rat tibialis anterior muscle after stimulated chronic eccentric exercise, J Appl Physiol 69, 1718–1724. 13. Baar, K., and Esser, K. (1999) Phosphorylation of p70(S6k) correlates with increased skeletal muscle mass following resistance exercise, Am J Physiol 276, C120–127. 14. Booth, F. W. (1978) Regrowth of atrophied skeletal muscle in adult rats after ending immobilization, J Appl Physiol 44, 225–230. 15. Childs, T. E., Spangenburg, E. E., Vyas, D. R., and Booth, F. W. (2003) Temporal alterations in protein signaling cascades during recovery from muscle atrophy, Am J Physiol Cell Physiol 285, C391–398. 16. Bodine, S. C., Stitt, T. N., Gonzalez, M., Kline, W. O., Stover, G. L., Bauerlein, R., Zlotchenko, E., Scrimgeour, A., Lawrence, J. C., Glass, D. J., and Yancopoulos, G. D. (2001) Akt/mTOR pathway is a crucial regulator of skeletal muscle hypertrophy and can prevent muscle atrophy in vivo, Nat Cell Biol 3, 1014–1019. 17. Hwee, D. T., and Bodine, S. C. (2009) Agerelated deficit in load-induced skeletal muscle growth, J Gerontol A Biol Sci Med Sci 64, 618–628.

18. McPherron, A. C., Lawler, A. M., and Lee, S. J. (1997) Regulation of skeletal muscle mass in mice by a new TGF-beta superfamily member, Nature 387, 83–90. 19. Lewis, A., and Redrup, L. (2005) Genetic imprinting: conflict at the Callipyge locus, Curr Biol 15, R291–294. 20. Nakatani, M., Takehara, Y., Sugino, H., Matsumoto, M., Hashimoto, O., Hasegawa, Y., Murakami, T., Uezumi, A., Takeda, S., Noji, S., Sunada, Y., and Tsuchida, K. (2008) Transgenic expression of a myostatin inhibitor derived from follistatin increases skeletal muscle mass and ameliorates dystrophic pathology in mdx mice, FASEB J 22, 477–487. 21. Schiaffino, S., and Reggiani, C. (1994) Myosin isoforms in mammalian skeletal muscle, J Appl Physiol 77, 493–501. 22. Peter, J. B., Barnard, R. J., Edgerton, V. R., Gillespie, C. A., and Stempel, K. E. (1972) Metabolic profiles of three fiber types of skeletal muscle in guinea pigs and rabbits, Biochemistry 11, 2627–2633. 23. Barnard, R. J., Edgerton, V. R., Furukawa, T., and Peter, J. B. (1971) Histochemical, biochemical, and contractile properties of red, white, and intermediate fibers, Am J Physiol 220, 410–414. 24. Antonio, J., and Gonyea, W. J. (1993) Skeletal muscle fiber hyperplasia, Med Sci Sports Exerc 25, 1333–1345. 25. Gonyea, W., Ericson, G. C., and BondePetersen, F. (1977) Skeletal muscle fiber splitting induced by weight-lifting exercise in cats, Acta Physiol Scand 99, 105–109. 26. Sacks, R. D., and Roy, R. R. (1982) Architecture of the hind limb muscles of cats: functional significance, J Morphol 173, 185–195. 27. Okita, M., Yoshimura, T., Nakano, J., Motomura, M., and Eguchi, K. (2004) Effects of reduced joint mobility on sarcomere length, collagen fibril arrangement in the endomysium, and hyaluronan in rat soleus muscle, J Muscle Res Cell Motil 25, 159–166. 28. Roy, R. R., Pierotti, D. J., and Edgerton, V. R. (1996) Skeletal muscle fiber cross-sectional area: effects of freezing procedures, Acta Anat (Basel) 155, 131–135. 29. Brooks, S. V., and Faulkner, J. A. (1988) Contractile properties of skeletal muscles from young, adult and aged mice, J Physiol 404, 71–82. 30. Ryall, J. G., Plant, D. R., Gregorevic, P., Sillence, M. N., and Lynch, G. S. (2004) Beta 2-agonist administration reverses muscle wasting and improves muscle function in aged rats, J Physiol 555, 175–188.

12 31. Fitts, R. H., and Widrick, J. J. (1996) Muscle mechanics: adaptations with exercise-training, Exerc Sport Sci Rev 24, 427–473. 32. Kandarian, S. C., and White, T. P. (1990) Mechanical deficit persists during long-term muscle hypertrophy, J Appl Physiol 69, 861–867. 33. Powell, P. L., Roy, R. R., Kanim, P., Bello, M. A., and Edgerton, V. R. (1984) Predictability

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of skeletal muscle tension from architectural determinations in guinea pig hindlimbs, J Appl Physiol 57, 1715–1721. 34. Burkholder, T. J., Fingado, B., Baron, S., and Lieber, R. L. (1994) Relationship between muscle fiber types and sizes and muscle architectural properties in the mouse hindlimb, J Morphol 221, 177–190.

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Chapter 13 Protein Overexpression in Skeletal Muscle Using Plasmid-Based Gene Transfer to Elucidate Mechanisms Controlling Fiber Size Chia-Ling Wu and Susan C. Kandarian Abstract Plasmid DNA electrotransfer is a direct method of gene delivery to skeletal muscle commonly used to identify endogenous signaling pathways that mediate muscle remodeling or pathological states in adult rodents. When plasmids encoding a protein to be overexpressed are fused to a fluorescent protein or an epitope-tag, plasmid electrotransfer permits visualization of the expressed protein in muscle fibers. Here, we demonstrate the use of electrotransfer of plasmids encoding mutant or wild type proteins to identify the role of the endogenous protein in regulating muscle fiber atrophy. The plasmids used encode a dominant negative form of the inhibitor of kappaB kinase beta (IKKB) fused to green fluorescent protein (GFP), a constitutively active form of IKKA fused to GFP, and a wild type IKKB fused to an HA tag. We show the effects of overexpression of these proteins on rat or mouse fiber size either with disuse atrophy or in normal weight bearing muscle. The effects of overexpressed proteins on myofiber size are assessed by comparing cross-sectional area of the transfected, fluorescent myofibers to the nontransfected, nonfluorescent myofibers. Using optimized intramuscular plasmid DNA injection and electroporation, we illustrate high transfection efficiency with no overt muscle damage using medium sized fusion proteins (105 kDa). Key words: Plasmid DNA, Electrotransfer, In vivo gene transfer, Electroporation, Transfection efficiency, Fiber cross-sectional area

1. Introduction Since the discovery that excitable cells such as striated muscle fibers can be transiently transfected by direct injection of supercoiled plasmid DNA (1), the use of plasmid-based gene transfer in muscle has been widespread (e.g., refs. (2–6)). Direct muscle gene transfer has been used to deliver therapeutic genes, most often for

Joseph X. DiMario (ed.), Myogenesis: Methods and Protocols, Methods in Molecular Biology, vol. 798, DOI 10.1007/978-1-61779-343-1_13, © Springer Science+Business Media, LLC 2012

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those encoding secretory proteins (7–10). It has also been used as a facile method of in vivo promoter analysis (11–13) or to assess in vivo transcriptional activity using transcription factor specific reporter plasmids (4, 5, 14, 15). The plasmid-based gene transfer method described here is optimized for the study of muscle fiber size regulation in health or disease, or for protein localization studies. The overexpression of wild type, dominant negative (d.n.), or constitutively active (c.a.) proteins is a widely used method to understand the mechanisms of biological processes in a specific cell or tissue type. This is often done by the construction of transgenic mice. However, the use of plasmid-based gene transfer for protein overexpression in muscle fibers can circumvent the time consuming process where a gene of interest is randomly inserted in the genome to produce transgenic mice (16, 17). Plasmid DNA transfected into adult muscle also has the advantage that it is ectopic, it is muscle fiber specific, the amount of protein overexpressed can be controlled, the protein expression in post-mitotic muscle fibers is stable for at least several weeks, and it minimizes the potential confounding effects of developmental or whole body physiological compensation seen in germ line genetic manipulation techniques (1, 13, 18, 19). Plasmid gene transfer also avoids safety concerns associated with viral transduction such as immune responses or recombination events (19, 20). Among the different gene delivery methods, intramuscular injection of endotoxin free plasmid DNA is one of the simplest genetic tools to study muscle cell biology. The drawback of direct plasmid DNA injection is that the transfection efficiency is sometimes low. However, three groups of investigators showed that combining plasmid injection with the local application of electrical pulses, referred as electrotransfer, increased transfection efficiency by at least 25-fold and reduced inter-muscle transfection variability (21–23). With optimal stimulation parameters, high levels of transgene expression can be detected in muscle while keeping muscle damage to a minimum (17, 24, 25). The in vivo plasmid DNA electrotransfer protocol described here is a powerful tool to study mechanisms of physiological/pathological processes in adult skeletal muscle by overexpressing mutant proteins that activate or inhibit signaling pathways or transcription factors (e.g., refs. (2–6)). We describe several examples of plasmid DNA electrotransfer in rat and mouse skeletal muscle to study the regulation of muscle fiber size by overexpression of enhanced green fluorescent protein (EGFP) fusion or epitope tagged proteins, which are subsequently visualized histologically. First, we show injection of a plasmid encoding a d.n. form of the inhibitor of kappaB kinase beta fused to enhanced green fluorescent protein (d.n. IKKB-EGFP) into rat soleus muscle to determine whether endogenous IKKB is required for unloading atrophy by measuring fluorescent vs. non-fluorescent muscle fiber cross-sectional area, 8 days after plasmid injection. We also show a histological section of mouse tibialis anterior (TA)

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muscle expressing c.a. IKKA-EGFP, which elicits atrophy in normal weight bearing muscle. Lastly, we show a section from a rat soleus muscle expressing wild type IKKB fused to an HA tag (w.t. IKKB-(A). Fibers expressing w.t. IKKB-(A are identified by an HA antibody. Overexpression of this wild type protein does not affect fiber size (5).

2. Materials 2.1. Plasmid DNA Preparation

1. Plasmids: d.n. IKKB-EGFP, c.a. IKKA-EGFP, w.t. IKKB-(A (see Notes 1 and 2). 2. Maximum efficiency DH5A-T1R competent cells (Invitrogen). 3. Qiagen EndoFree Plasmid Mega kit. 4. 4 M NaCl. 5. 100% EtOH. 6. Sterile diluted saline (0.45% NaCl) or sterile saline (0.9% NaCl). 7. Disposable 28-gauge 3/10 cc insulin needles. 8. UV spectrophotometer.

2.2. Surgery, Plasmid Injection, and Electroporation

1. 8-Week old female Wistar rats weighing ~150 g (Charles River). 2. 100 mg/mL Ketamine and 13 mg/mL xylazine hydrochloride cocktail. 3. 7.5% Povidone-iodine scrub solution. 4. 70% Isopropyl alcohol. 5. Dermalon 5.0 suture. 6. Ethilon 699G 4.0 suture. 7. Sterile saline (0.9% NaCl). 8. BTX Electro Square Porator ECM 830 (Harvard Apparatus). 9. BTX Tweezertrodes (stainless steel electrodes, 10 mm diameter, Model 522) or spatula-like electrodes (0.5 cm wide, 2 cm long) (see Fig. 1 in ref. (24)). 10. Surgical instruments (small scissors, scalpel, delicate tissue forceps). 11. Surgical tape to secure foot during surgery. 12. Glass petri dish (or equivalent) for positioning foot and lower leg during surgery. 13. Fur shaver.

2.3. Muscle Harvesting

1. Ketamine-xylazine cocktail (as above). 2. Surgical instruments (as above). 3. Tissue-Tek OCT compound.

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4. Wooden tongue depressors. 5. Isopentane. 6. Liquid nitrogen. 2.4. Immunohistochemistry

1. Cryostat. 2. Glass microscope slides. 3. 1 and 10% Bovine serum albumin (BSA). 4. 4% Paraformaldehyde. 5. Phosphate buffered saline (PBS). 6. Rabbit anti-laminin antibody. 7. Texas Red®-X goat anti-rabbit IgG. 8. Fluorescent microscope equipped with filters for detecting Texas Red and EGFP. 9. SPOT RT camera and the SPOT Advanced software. 10. MetaMorph Imaging software (Molecular Devices).

3. Methods 3.1. Plasmid DNA Preparation

1. Transform plasmids into DH5A competent cells (or appropriate bacterial strain) for plasmid DNA replication in a large scale culture using standard procedures ((26) or see chapter by Senf and Judge in this issue. 2. Isolate plasmid DNA from bacteria using a Qiagen EndoFree Mega kit, suspend in TE, and store at −20°C until use. 3. Check the integrity of the plasmid DNA (i.e., supercoiled vs. open-circular) by agarose gel electrophoresis (see Note 3). 4. Measure the plasmid concentration by UV spectrophotometry at 260 nm (see Note 4). 5. Calculate the amount of plasmid DNA required for plasmid injection. Using the measured concentration, determine the volume needed. Pipet this volume of plasmid into a sterile microcentrifuge tube (see Note 5). 6. For TE to sterile saline buffer exchange, add 4 M NaCl to the plasmid-TE mixture to a final concentration of 0.2 M (see Note 6). 7. Gently invert the mixture 4–6 times. 8. Precipitate the plasmid DNA by adding 2.5 volumes of ice-cold 100% EtOH. Invert to mix. 9. Precipitate the plasmid DNA on ice for 10 min. 10. Pellet the precipitated plasmid DNA by centrifugation in a microfuge at maximum speed for 10 min at 4°C.

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11. Decant the supernatant without disturbing the pellet. 12. Air dry the pellet for 10 min and resuspend the plasmid DNA in the desired volume of sterile saline (or half sterile saline) by gently inverting the tube. The volume of sterile saline depends on the amount of plasmid needed (the number of muscles and the microgram per muscle to be injected). For injection of ten rat soleus muscles (~100 mg each), inject 50 Mg plasmid/muscle. For 1 Mg/1 ML plasmid concentration, redissolve 500 Mg of the plasmid in 500 ML sterile saline. In practice, prepare 10% more plasmid to account for dead space in needles and to measure plasmid integrity using agarose gel electrophoresis. 13. Quantify the concentration of plasmid DNA by UV spectrophotometry. Reserve 300 ng of plasmid DNA aliquot on ice and check the plasmid integrity by agarose gel electrophoresis. For optimal uptake most of the plasmid preparation (>80%) should be in the supercoiled conformation. 14. Fill a sterile disposable 28-gauge 3/10 cc insulin syringe with 100 Mg of IKK plasmid (e.g., d.n. IKKB-EGFP) in 100 ML sterile saline solution. 3.2. Surgery, Plasmid Injection, and Electroporation

1. Anesthetize the rat with an intraperitoneal injection of 0.1 mL/200 g body weight of ketamine-xylazine cocktail. 2. Shave the hindlimb and scrub the shaved skin with povidoneiodine then remove excess with 70% isopropyl alcohol. 3. Use a scalpel to make a small incision through the skin along the lateral side of the rat lower limb. 4. With the rat lying on its side, tape the foot in a lateral position to a small stage to elevate the foot and to provide a clear dorsallateral view of lower leg (we use a small glass petri dish). 5. In the incision just made, cut through the fascia along the posterior-lateral edge of gastrocnemius muscle. Carefully isolate the soleus muscle by blunt dissection, taking care to avoid cutting blood vessels, nerves, or muscle. 6. Insert the needle of a 28-gauge insulin syringe containing the expression plasmid near the distal myotendinous junction of the soleus, and slowly push it ~1 cm rostrally to the proximal end of the soleus. 7. Inject 50 ML of the plasmid DNA evenly along the longitudinal axis of soleus while slowly withdrawing the syringe (see Note 7). The remaining 50 ML of plasmid is for the contralateral muscle. For the optimized mouse DNA injection protocol, see Note 8. 8. Wait 1 min before electroporation. 9. Moisten the surface of the soleus muscle using a sterile salinesoaked cotton applicator.

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10. Place the electrodes on either side of the muscle. For rat soleus, the electrodes are generally placed 4 mm apart to avoid pinching the tissue (see Note 9). 11. Apply 5 square-wave electrical pulses at 75–100 V/cm, 20 ms, 1 Hz, with 200 ms interpulse interval, using BTX Electro Square Porator (see Note 10). 12. Close the incision through the subcutaneous fascia layer using 5.0 Dermalon sutures using a running stitch (this step is not necessary in mice). Close the dermal layer using 4.0 Ethilon suture to make discontinuous square knots, distal to proximal. 13. If necessary, electrotransfer 33 Mg of pEGFP-N1 into the contralateral leg (see Note 11). 14. Before recovering from anesthesia, the rat is tail-casted as previously described (27), although the experimental condition to be used is project specific. 15. Subject rat or mouse to control or experimental condition. In our illustrated case with the rat, 24 h after plasmid injection the rat is suspended by a tail cast so that the hind limbs, when fully extended, are 1 mm off the ground (see Note 12). 3.3. Muscle Harvesting

1. Following the period of physiological/pathological intervention, here we used 7 days of hind limb unloading, anesthetize the rat with 0.2 mL/200 g body weight of ketamine-xylazine cocktail by intraperitoneal injection. 2. Excise soleus muscles from both legs of the rat. 3. Quickly weigh harvested muscle and pin it horizontally (at approximately optimum length) on a wooden tongue depressor and embed in OCT compound so that a thin layer covers the muscle. 4. Immediately snap freeze muscle in isopentane prechilled by liquid nitrogen. Cover with foil and store at −80°C until sectioning.

3.4. Immunohistochemistry

1. Serially section the frozen muscles in 10 Mm sections starting from the muscle mid-belly. 2. Mount sections on microscope slides and fix the sections in 4% paraformaldehyde solution for 20 min at room temperature (see Note 13). 3. Wash muscle sections for 10 min by immersion in three changes of PBS. 4. Block the sections in 10% BSA in PBS for 45 min. 5. Incubate the sections in a 1:200 dilution of anti-laminin antibody in 1% BSA for 1 h at room temperature (see Note 14). 6. Wash the sections in PBS for 10 min.

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7. Incubate sections with a 1:200 dilution of Texas Red goat anti-rabbit secondary antibody in 1% BSA for 40 min in a dark chamber. 8. After washing in PBS for 10 min, visualize muscle sections through a fluorescent microscope using Texas Red and FITC filters. Acquire images with a SPOT RT camera and merge the images of fluorescent signals for Texas Red and EGFP using SPOT Advanced software. 9. Measure fiber cross-sectional area using MetaMorph (or equivalent) Imaging software where the fibers taking up the expression plasmid (i.e., fluorescent) are compared with those in the same field that did not take up the plasmid (i.e., nonfluorescent). Typical results of fluorescent fibers from muscles overexpressing either d.n. IKKB-EGFP (Fig. 1) or c.a. IKKA-EGFP (Fig. 2) show high transfection efficiency. An example of immunohistochemistry to show muscle fibers overexpressing wild type IKKB-HA is visualized by colorimetric (HRP) staining (Fig. 3). 10. To confirm overexpression of full-length EGFP fusion protein or HA-tagged protein, we recommend performing an immunoblot from a homogenate of a separate plasmid injected muscle as previously demonstrated (4, 5).

Fig. 1. Fluorescent image of rat soleus muscle cross section expressing dominant negative (d.n.) inhibitor of kappaB kinase beta fused to enhanced green fluorescent protein (IKKBEGFP) and counterstained with anti-laminin. Fifty microgram of d.n. IKKB-EGFP plasmid was electroporated into rat soleus muscle using BTX Tweezertrodes for 5 square-wave pulses, 20 ms each, at 100 V/cm. Muscle was removed 8 days after plasmid electrotransfer; 1 day of weight bearing plus 7 days of hind limb unloading. Mean cross-sectional area of fibers expressing d.n. IKKB-EGFP inhibited unloading induced atrophy by 50% (5).

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Fig. 2. Fluorescent image of mouse TA muscle cross-section expressing constitutively active (c.a.) IKKA-EGFP and counterstained with anti-laminin. Forty micrograms of c.a. IKKA-EGFP was electroporated into mouse TA muscle by spatula electrodes at 75 V/cm, 5 square-wave pulses, each with a 20 ms duration. Muscle from a normal weight bearing mouse was removed 10 days after plasmid electrotransfer. Mean cross-sectional area of fibers expressing c.a. IKKA-EGFP are 50% smaller than nonfluorescent black fibers. Similar data were shown for weight bearing rat soleus muscle overexpressing c.a. IKKA-EGFP for 8 days (5).

Fig. 3. Immunohistochemistry staining of rat soleus muscle cross section. Rat soleus muscle was electrotransfered with 50 Mg w.t. IKKB-HA. After 1 day of weight bearing followed by 7 days of hind limb unloading, muscle was harvested, sectioned, and incubated with biotinylated anti-HA antibody (Vector Labs). Biotinylated w.t. IKKB-HA complexes were then visualized by avidin-biotin peroxidase labeling (ABC reagent, Vector Labs) with diaminobenzidine (DAB, Vector Labs) colorization under bright field microscopy. This serves as an example of expression and visualization of a HA-tagged plasmid in muscle fibers when the use of fluorescent fusion protein is not feasible or desirable. Mean fiber cross-sectional area of w.t. IKKB-HA stained fibers was not different from nonstained fibers (5).

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4. Notes 1. The d.n. IKKB-EGFP was constructed by inserting a d.n. IKKB fragment, subcloned from the d.n. IKKB plasmid provided by M. Karin (28), into a CMV-driven EGFP-N1 plasmid (4.7 kb, Clontech) (5). The resulting 7.0 kb d.n. IKKB-EGFP plasmid contains a mutation at K44M in the IKKB sequence, and is therefore catalytically inactive. To construct c.a. IKKA-EGFP, mutations in w.t. IKKA were made at Ser176 and Ser180 to produce glutamic acid; this construct was cloned in-frame into pEGFP-N1 (5). The w.t. IKKB-(A plasmid was constructed by inserting the wild type IKKB sequence into a CMV-driven HA-tagged vector as previously described (5). All plasmid inserts were sequenced. We prefer the EGFP on the C-terminus of the protein because it is a visual check that a full-length protein was synthesized. 2. To verify that the function of d.n. IKKB or c.a. IKKA is not altered by fusion to green fluorescent protein (GFP), d.n. IKKB-EGFP and c.a. IKKA-EGFP were separately transfected with an NF-KB-luciferase reporter plasmid in C2C12 cells to confirm that d.n. IKKB-EGFP abolished TNFA-induced NF-KB activity and that c.a. IKKA-EGFP activated NF-KB activity in the absence of treatment. 3. Injections of supercoiled DNA result in ~10–20 times higher ectopic gene expressions compared with injections of linear DNA (18). It is essential to check for maintenance of supercoiled DNA after the plasmid is suspended in sterile saline before injection. 4. We find that plasmid concentrations between 0.8 and 3 Mg/ML are easiest to work with and are less susceptible to nicking or degradation. 5. The amount of the plasmid DNA for injection depends largely on the size of the muscle to be injected and the size of the encoded protein. For example, we inject between 40 and 100 Mg of expression plasmid/rat soleus muscle (~100 mg). This range depends on the size of the expressed protein and the size of the plasmid. Fifty microgram of EGFP-N1 plasmid (4.7 kb) contains more copies of the encoded protein (25 kDa) than 50 Mg of d.n. IKKB-EGFP plasmid (7 kb), which encodes a 105 kDa protein. We have observed that plasmids encoding larger EGFP fusion proteins are more difficult to detect than plasmids encoding smaller EGFP fusion proteins (compare refs. (4, 5)). When we coinjected d.n. Foxo3-DsRed (4.8kb, encodes a 41 kDa protein) and d.n. IKKB-EGFP into the same soleus muscle, we used 40 Mg of the former and 60 Mg of the latter

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plasmid in an attempt to express similar amounts of the protein encoded in each plasmid (29). A linear increase in numbers of transfected fibers in rat TA muscle was observed when the amount of electroporated plasmid DNA increased from 10 to 100 Mg (30). Other studies have found that plasmid DNA expression reached a plateau if an excess amount of plasmid DNA was injected (24, 31). Caution should be taken to avoid injecting too much plasmid, especially using viral promoters where depletion of basal transcription factors could be problematic (32). 6. We resuspend the plasmid into sterile saline solution or sterile PBS on the day of plasmid DNA electrotransfer. Injection of plasmid in TE causes muscle damage (33). Several investigators have shown that the use of half saline as an injection vehicle improved transfection efficiency when compared with normal saline (17, 34). 7. Although the increased hydrostatic pressure caused by injecting large volume of plasmid DNA has been postulated as one of the mechanisms by which plasmid DNA enters cell membranes in vivo, injecting an excess volume of plasmid may cause tissue swelling and muscle damage. Therefore, it is advisable to use a smaller injection volume relative to the size of the muscle; for example, plasmid DNA is generally suspended in a 50 ML sterile saline for injections of soleus or extensor digitorum longus in a 150 g rat. However, if too little volume is injected, transfection efficiency is low and variability is high (35). 8. The procedure for mouse plasmid DNA injection is similar to that of the rat protocol, although the mouse TA muscle is more easily accessible than the soleus. For the mouse TA shown here, we injected 40 Mg plasmid (c.a. IKKA-EGFP) in 35 ML sterile half saline (0.45% NaCl). 9. We use either stainless steel BTX Tweezertrodes or spatula plate electrodes (24) for plasmid DNA electrotransfer. Both plate electrodes have the advantage of providing a more homogeneous electrical field between the electrodes (36), whereas needle electrodes have been reported to apply higher field intensities at the tips of the needles. We achieve at least 50% transfection efficiency with either spatula or Tweezertrodes (see Fig. 3 in ref. (37) where BTX Tweezertrodes are applied to the rat TA for electroporation). 10. The efficiency of intramuscular gene electrotransfer generally increases as a function of voltage intensity and the total pulse duration (pulse number × pulse duration) (21–23). However, care must be taken to avoid high electrical stimulation parameters

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so that one is studying the underlying muscle physiology and not muscle damage. For example, Schertzer et al. (17) found significant damage by applying eight 20 ms pulses at 150– 200 V/cm to the mouse TA (using spatula electrodes), and therefore, the use of a lower voltage (75–100 V/cm) for electroporation was suggested. Dona et al. (24) delivered 5 squarewave pulses of 50 V/cm, 20 ms duration, and 200 ms interpulse interval, with spatula electrodes and reported that 80% of mouse TA or rat soleus fibers were transfected. It is noted that an EGFP plasmid was used that alone encodes a small and thus easily synthesized fluorescent protein. Our lab has established an optimal protocol of electroporating 5 square-wave pulses at 75–100 V/cm, 20 ms, 1 Hz, with 200 ms interpulse interval, directly on the rat soleus or mouse TA muscle using BTX Tweezertrodes or spatula electrodes, both of which show high transfection efficiency and cause minimal muscle damage. 11. The parental vector of the expression plasmid is sometimes injected into the contralateral leg of the animal to control for any effect that the vector alone has on fiber size (e.g., EGFP expressing vector). We have demonstrated that EGFP does not have an effect on fiber size in weight bearing or unloaded rat soleus muscle (4), so this control plasmid injection does not always need to be repeated. If this were done for the IKK-EGFP expression plasmids, in order to achieve similar copy numbers of different sized plasmids with different sized encoded protein, 33 Mg (in 50 ML) of EGFP-N1 and 50 Mg (in 50 ML) of d.n. IKKB-EGFP would be injected into the right and left muscle of the same animal, respectively. 12. The reason for waiting 24 h before hind limb suspending the rats is that we found lower transfection efficiency if rats are not allowed to bear weight after plasmid injection (32). 13. To avoid quenching GFP signals encoded by EGFP-N1 expression plasmids, keep slides in a dark humid chamber. 14. We have also had good success with staining the muscle fiber periphery using wheat germ agglutinin (WGA) conjugated to Texas Red-X; sections are first incubated with a 1:50 dilution of WGA in PBS for 2 h, washed, and are visualized by fluorescent microscopy.

Acknowledgment This work was supported by NIH grant AR41705.

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References 1. Wolff JA, Malone RW, Williams P et al (1990) Direct gene transfer into mouse muscle in vivo. Science 247:1465–1468 2. Bodine SC, Stitt TN, Gonzalez M et al (2001) Akt/mTOR pathway is a crucial regulator of skeletal muscle hypertrophy and can prevent muscle atrophy in vivo. Nat Cell Biol 3:1014–1019 3. Sartori R, Milan G, Patron M et al (2009) Smad2 and 3 transcription factors control muscle mass in adulthood. Am J Physiol Cell Physiol 296:C1248–1257 4. Judge AR, Koncarevic A, Hunter RB et al (2007) Role for I{kappa}B{alpha}, but not c-Rel, in skeletal muscle atrophy. Am J Physiol Cell Physiol 292:C372–382 5. Van Gammeren D, Damrauer JS, Jackman RW et al (2009) The IkappaB kinases IKKalpha and IKKbeta are necessary and sufficient for skeletal muscle atrophy. Faseb J 23:362–370 6. Senf SM, Dodd SL, McClung JM et al (2008) Hsp70 overexpression inhibits NF-kappaB and Foxo3a transcriptional activities and prevents skeletal muscle atrophy. Faseb J 22:3836–3845 7. Kessler PD, Podsakoff GM, Chen X et al (1996) Gene delivery to skeletal muscle results in sustained expression and systemic delivery of a therapeutic protein. Proc Natl Acad Sci USA 93:14082–14087 8. Raz E, Watanabe A, Baird SM et al (1993) Systemic immunological effects of cytokine genes injected into skeletal muscle. Proc Natl Acad Sci USA 90:4523–4527 9. Rizzuto G, Cappelletti M, Maione D et al (1999) Efficient and regulated erythropoietin production by naked DNA injection and muscle electroporation. Proc Natl Acad Sci USA 96:6417–6422 10. Yin D, Tang JG (2001) Gene therapy for streptozotocin-induced diabetic mice by electroporational transfer of naked human insulin precursor DNA into skeletal muscle in vivo. FEBS Lett 495:16–20 11. Mitchell-Felton H, Hunter RB, Stevenson EJ et al (2000) Identification of weight-bearingresponsive elements in the skeletal muscle sarco(endo)plasmic reticulum Ca2+ ATPase (SERCA1) gene. J Biol Chem 275:23005–23011 12. Esser K, Nelson T, Lupa-Kimball V et al (1999) The CACC box and myocyte enhancer factor-2 sites within the myosin light chain 2 slow promoter cooperate in regulating nervespecific transcription in skeletal muscle. J Biol Chem 274:12095–12102

13. Kitsis RN, Leinwand LA (1992) Discordance between gene regulation in vitro and in vivo. GeneExpr 2:313–318 14. Hunter RB, Stevenson E, Koncarevic A et al (2002) Activation of an alternative NF-kappaB pathway in skeletal muscle during disuse atrophy. Faseb J 16:529–538 15. Hunter RB, Kandarian SC (2004) Disruption of either the Nfkb1 or the Bcl3 gene inhibits skeletal muscle atrophy. J Clin Invest 114:1504–1511 16. Bartlett RJ, Secore SL, Singer JT et al (1996) Long-term expression of a fluorescent reporter gene via direct injection of plasmid vector in to mouse skeletal muscle: comparison of human creatine kinase and CMV promoter expression levels in vivo. Cell Transplantation 5:411–419 17. Schertzer JD, Plant DR, Lynch GS (2006) Optimizing plasmid-based gene transfer for investigating skeletal muscle structure and function. Mol Ther 13:795–803 18. Wolff JA, Ludtke JJ, Acsadi G et al (1992) Long-term persistence of plasmid DNA and foreign gene expression in mouse muscle. HumMolGenet 1:363–369 19. Dickson G (1996) Gene transfer to muscle. BiochemSocTrans 24:514–519 20. Davis HL, Demeneix BA, Quantin B et al (1993) Plasmid DNA is superior to viral vectors for direct gene transfer into adult mouse skeletal muscle. HumGenTher 4:733–740 21. Mir LM, Bureau MF, Gehl J et al (1999) High-efficiency gene transfer into skeletal muscle mediated by electric pulses. Proc Natl Acad Sci USA 96:4262–4267 22. Mathiesen I (1999) Electropermeabilization of skeletal muscle enhances gene transfer in vivo. Gene Ther 6:508–514 23. Aihara H, Miyazaki J (1998) Gene transfer into muscle by electroporation in vivo. Nat Biotechnol 16:867–870 24. Dona M, Sandri M, Rossini K et al (2003) Functional in vivo gene transfer into the myofibers of adult skeletal muscle. Biochem Biophys Res Commun 312:1132–1138 25. Taylor J, Babbs CF, Alzghoul MB et al (2004) Optimization of ectopic gene expression in skeletal muscle through DNA transfer by electroporation. BMC Biotechnol 4:11 26. Sambrook J, Fritsch EF, Maniatis T (1989) Molecular cloning: A laboratory manual. Cold Spring Harbor Laboratory, Cold Spring Harbor, NY 27. Peters DG, Mitchell-Felton H, Kandarian SC (1999) Unloading induces transcriptional

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Southern blot: application to SERCA1 promoter analysis. Am J Physiol 277:C1269–C1276 Wolff JA, Williams P, Acsadi G et al (1991) Conditions affecting direct gene transfer into rodent muscle in vivo. BioTechniques 11: 474–485 Lee MJ, Cho SS, Jang HS et al (2002) Optimal salt concentration of vehicle for plasmid DNA enhances gene transfer mediated by electroporation. Exp Mol Med 34:265–272 Davis HL, Whalen RG, Demeneix BA (1993) Direct gene transfer into skeletal muscle in vivo: factors affecting efficiency of transfer and stability of expression. Hum Gene Ther 4:151–159 Gehl J, Sorensen TH, Nielsen K et al (1999) In vivo electroporation of skeletal muscle: threshold, efficacy and relation to electric field distribution. Biochim Biophys Acta 1428: 233–240 Komamura K, Miyazaki J, Imai E et al (2008) Hepatocyte growth factor gene therapy for hypertension. In: Li S (ed) Methods in Molecular Biology, 1st edn. Human Press, New Jersey

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Chapter 14 In Vivo Measurement of Muscle Protein Synthesis Rate Using the Flooding Dose Technique Marta L. Fiorotto, Horacio A. Sosa Jr., and Teresa A. Davis Abstract Skeletal muscle mass is determined by the balance between rates of protein synthesis and degradation. Protein synthesis rates can be measured in vivo by administering an amino acid as a tracer that is labeled with an isotope (radioactive or stable) of C, H, or N. The rate at which the labeled amino acid is incorporated into muscle protein, as a function of the amount of labeled amino acid in the precursor pool at the site of translation, reflects the rate of protein synthesis. There are a number of approaches for performing this measurement depending on the question being addressed and the experimental system being studied. In this chapter, we describe the “flooding dose” approach using L-[3H]-phenylalanine as the tracer and that is suitable for determining the rate of skeletal muscle protein synthesis (total and myofibrillar proteins) over an acute period (ideally less than 30 min) in any size animal; details for working with mice are presented. The method describes how to administer the tracer without anesthesia, the tissue collection, and the preparation of muscle and blood samples for analysis of the tracer and tracee amino acids in the precursor pool and in muscle proteins. Key words: Protein synthesis, Translation, Protein degradation, Skeletal muscle, Amino acid tracer, Phenylalanine

1. Introduction Protein mass is determined by the balance between protein synthesis and degradation (1–3). Over a relatively chronic time-frame (days), the two processes are in balance in the skeletal muscles of the full grown adult at steady state and muscle mass remains constant. However, the balance in vivo shifts from positive to negative during a day due to the responsiveness of protein synthesis and breakdown to changes in nutrients, activity, hormones, and growth factors. Net gain of muscle protein occurs when the rate of protein

Joseph X. DiMario (ed.), Myogenesis: Methods and Protocols, Methods in Molecular Biology, vol. 798, DOI 10.1007/978-1-61779-343-1_14, © Springer Science+Business Media, LLC 2012

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synthesis is greater than the rate of protein breakdown, whereas muscle mass is lost when the rate of protein degradation exceeds synthesis rates. Therefore, the quantification of these processes in vivo, and how the balance changes over time, is critical for understanding how muscle protein mass is regulated by biological variables. There are a number of approaches for quantifying the rates of skeletal muscle protein synthesis and degradation in vivo, and the one that is most suitable depends on the question being addressed and the experimental system being studied. The differences among these approaches and their advantages and disadvantages have been addressed in a number of reviews (4–8). As is evident from these reviews, it is not possible to cover the technical details for all of these in a single chapter as they can entail quite different in vivo protocols and subsequent analytical techniques. The majority of the techniques involve the administration of an amino acid as a tracer that is labeled with an isotope (radioactive or stable) of C, H, or N. The rate at which the labeled amino acid is incorporated into muscle protein, as a function of the amount of labeled amino acid in the precursor pool at the site of translation, reflects the rate of protein synthesis. The approaches for measuring skeletal muscle protein degradation in vivo (5, 9–12) are more limited, technically difficult, and not practical to perform in small animals such as rats or mice. They also require the use of amino acid tracers either to estimate arteriovenous differences across a muscle bed, or to label muscle proteins, and then measure the rate at which the label is lost from the muscle due to degradation. An alternative approach that can be used in experimental animals and measures protein degradation rate averaged over a period of days will be presented in this chapter (13, 14). Additionally, it should be remembered that at steady state, the rates of synthesis and degradation are equal. In this chapter, the protein synthesis technique known as the “flooding” or “large” dose technique developed originally by Garlick et al. (15) is presented. It is suitable for measuring protein synthesis in any size animal, and to determine the synthesis rate of both constitutive and secreted proteins in any tissue (4). The basic procedure involves administering relatively rapidly (<30 s), as a single bolus, a large dose of the tracee amino acid along with the tracer amino acid. The total amount of the labeling amino acid administered should be 5 to 10 times the body’s total free pool of the essential amino acid chosen as tracee, and ideally, the amino acid should not be metabolized by the tissue of primary interest to minimize the introduction of error from the recycling and dilution of the label; the use of amino acids that are posttranslationally modified in skeletal muscle (e.g., histidine, lysine) also could introduce errors and are best avoided. The amount of tracer administered (i.e., the specific radioactivity or enrichment) depends on the synthesis rate of the protein (higher amounts for slower rates), and

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the sensitivity of the detection system. The objective of using a large dose is to “flood” the precursor amino acid pool with the tracer amino acid so that the specific radioactivity or enrichment (in the case of stable isotope-labeled tracers) of the extracellular, intracellular, and acylated tRNA pools come into equilibrium instantaneously (ideally), and remain in equilibrium throughout the labeling period (16). To ensure that this condition is satisfied, the labeling period is restricted to less than 30 min, ideally, during which time the organism should be in a relatively physiological “steady state.” The proteins to be assessed are then isolated and the amount of tracer incorporated into the protein is determined. The data are expressed as fractional rates of protein synthesis (FSR or Ks), which represent the fraction of the total protein (TP) pool synthesized per unit time. This provides a measure of the rate at which translation is occurring in the cells, and is independent of the TP mass. Absolute synthesis rate (ASR) is the product of FSR and the total mass of the protein; because TP mass reflects the product of the long-term protein balance of the tissue, ASR does not reflect the activity of the protein synthetic process in the tissue at that moment in time. When all proteins in a muscle are analyzed, the synthesis rate of mixed muscle proteins is determined; this includes the synthesis rate of all cell and protein species in the tissue including both extracellular and intracellular proteins. Alternatively, individual proteins, or groups of proteins (e.g., the myofibrillar, sarcoplasmic, and stromal fractions in muscle), or organelles (e.g., mitochondria) can be isolated and the FSR of their component proteins determined (14, 17–20). Over the years, the “flooding dose” technique has received criticism for a number of reasons (reviewed in ref. (21)). However, as with all indirect approaches for measuring a process, it is difficult to be certain of the approach that gives the “correct” value. Nonetheless, there is general agreement that if the technique is used adhering strictly to conditions that ensure that the underlying assumptions are not violated, the results obtained are valid. Thus, the measurement described provides a brief snapshot of the in vivo FSR. In this chapter, the described protocol uses L-[43H]-phenylalanine as the tracer and it is geared toward measurements of skeletal muscle proteins (total and myofibrillar) in rats or mice.

2. Materials The procedure is divided into two parts: the in vivo animal labeling component, and the analytical component. The latter describes the preparation of the samples and it is suitable for the measurement of both specific radioactivity and enrichment (if the amino acid tracer is labeled with a stable-isotope) of the

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amino acid tracer incorporated into muscle proteins. The actual determination of specific radioactivity or enrichment is not described here as there are a number of possible options depending on the technology available to the investigator. Care should be taken to ensure that all solutions, glassware, and disposables that come into contact with the proteins to be analyzed are protein and amino acid-free. Thus, high purity water (18 M7 cm at room temperature) should be used for everything; all glassware for making up solutions should be soaked in 5% HNO3 for at least 6 h and rinsed thoroughly before using. Glassware that comes into contact with samples from the animals should be soaked overnight in Chromerge® solution and then rinsed thoroughly. Extreme precaution should be taken when handling this solution that contains concentrated H2SO4; i.e., use only in a chemical fume hood; wear a rubber apron over lab coat, heavy duty acid-resistant rubber gloves over regular latex gloves, eye protection, and preferably handle glassware with tongs. The solution should be stored in an acid-proof cabinet and can be reused until it turns green at which point it should be disposed as biohazard waste. When handling radioactive materials, take all standard precautions and ensure that you have biosafety approval for using the radioactive materials in the lab and in the animals. 2.1. In Vivo Protein Synthesis Protocol

1. Isotope injection solution: Each animal is administered 1.5 mmol phenylalanine/kg body weight. The amount of radioactive tracer added depends on the FSR of the proteins of interest, the duration of the labeling period, and the sensitivity of the detection system, and requires a rough idea a priori of what these values are (see Notes 1 and 2). For an experiment with 16 adult mice (approximately 30 g body weight each) and a labeling time of 30 min (note that this is just an example and is unlikely to be appropriate for all applications or the same application in different labs), a total volume of 5.3 mL will be needed ([30 × 16]/100 = 4.8 mL + 10% for losses), containing 600 MCi L-[43H]-phenylalanine/mL in 0.9% NaCl. The final concentration is 150 mM with a specific radioactivity of 4 mCi/mM; each mouse receives approximately 200 MCi. In a 10 mL flat-bottomed clean, screw-cap, glass vial weigh 47.7 mg NaCl (analytical grade); and 131.175 mg L-phenylalanine (TraceCERT®, see Note 3). Pipette in 2.12 mL water and 3.18 mL L-[4-3H] phenylalanine (1 mCi/mL; see Note 4). Add a magnetic stir bar, cap, and Parafilm® the vial, and place on a stir plate to mix until the phenylalanine is completely dissolved (see Note 5). Unless you are going to use it immediately, store the solution at 4°C. Before using it, bring it to room temperature and ensure that any phenylalanine that has precipitated out is redissolved. For long-term storage (>4 weeks), the injection solution should be frozen at −20°C.

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2. Injection line: 1 mL syringe (with 0.01 mL calibration markings); 5” piece of PE 50 tubing; 23G Luer-stub adaptors; 26G × 5/8p hypodermic needles with hub removed (see Note 6). Attach the Luer-stub adapter to one end of the tubing, and the white end of the hypodermic needle to the other. At both ends, the tubing should be pushed until it abuts the white flare. This will be attached to the syringe and used for tail injection. 3. For injection, large pieces of cheese cloth (triple layer, approximately 8 × 8p); water in a beaker heated to 47–49°C; lamp; timer. 4. Large scissors or a guillotine with very sharp blades (see Note 7); various sized sharp scissors and fine tipped forceps for dissection. 5. Polystyrene and aluminum weigh dishes (small); 3p × 3p aluminum foil pouches (see Note 8); 1.5 mL microfuge tubes. 6. 0.2M chilled perchloric acid (PCA, 2.2 mL 60% PCA and 97.8 mL mQ water; see Note 9). 7. 95 × 12 mm Pyrex Petri dish bottoms filled with black dissecting wax; pins. 8. Liquid nitrogen, styrofoam containers. 9. Analytical and pan balance. 10. Screw capped tissue storage tubes suitable for liquid nitrogen. 11. One data recording worksheet per mouse. 12. Metric Vernier calipers. 2.2. Preparation of Tissue and Blood for Measurement of L-[4 3H]Phenylalanine Specific Radioactivity

1. 4M KOH; use chilled. 2. 1M acetic acid: 57 mL of glacial acetic acid (HPLC grade)/L (with mQ water). 3. 3M NH4OH: 48 mL of 14.8M NH4OH/200 mL (with mQ water). 4. 1M HCl: 16.6 mL of 6M HCl (ULTREX II Ultrapure)/100 mL (with mQ water). 5. Dowex®-50 resin (AG-50W-X8; 100–200 mesh, H+ form) cleaned and stored in 1M NaOH (see Note 10). 6. X2 low salt/sucrose homogenizing buffer: 100 mM K2HPO4 (17.42 g/L); 100 mM KH2PO4 (13.61 g/L); 0.5M sucrose; Triton X-100. Mix 61.5 mL of K2HPO4 with 38.5 mL of KH2PO4 and pH to 7.0 with KOH or phosphoric acid. To a 100 mL volumetric flask add 68.4 g sucrose, add approximately 90 mL of phosphate buffer and dissolve sucrose. Add 2 mL of Triton X-100 and bring to volume with phosphate buffer; good for <2 weeks; use chilled.

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7. X1 low salt/sucrose homogenizing buffer, pH 7: dilute X2 buffer 1:1 with mQ water; store at 4°C; use chilled. 8. 1.2M PCA: 13 mL of 60% PCA/100 mL (with mQ water); use chilled. 9. 0.2M PCA; use chilled. 10. 0.1M and 0.3M NaOH. 11. 6M HCl: concentrated HCl (ULTREX II Ultrapure) diluted 1:1 with mQ water (see Note 11); use fresh. 12. BCA protein assay reagents (22). 13. pH paper tape. 14. Centrifugal concentrator (e.g., Savant Speedvac) with rotors that will accept all tube sizes to be used in the analyses (see Note 12). 15. Syringe filters (0.22 or 0.45 Mm). 16. Disposable filter columns for ion exchange resin (individually fitted with funnel) and racks. 17. Glass vials (17 × 60 mm, flat-bottomed) with screw caps. 18. Steel mortar and pestle for powdering frozen muscles. 19. Homogenizer (e.g., an Ultra-Turrax®T-25) with stainless steel dispersion probes (S25N-8G) chilled on ice. 20. Glass rods (4 × 125 mm). 21. Polypropylene tubes (12 × 75 mm). 22. Tube shaker (e.g., Tomy shaker). 23. Kimble KIMAX® borosilicate glass (13 × 100 mm) with Teflon®-lined caps.

screw

top

tubes

24. Evaporator with heating blocks that accommodate glass tubes and with a gassing manifold equipped with removable stainless steel needles. 25. Oven set at 110°C. 26. N2 gas.

3. Methods 3.1. In Vivo Protein Synthesis Measurement

1. Values for skeletal muscle protein FSR are influenced by whether the animal is in the postprandial or fasted state (23). Because different information is conveyed by the FSR values in these two conditions, careful thought should be given to determining which is most appropriate, and the protocol should then be standardized to ensure that all mice are in the same condition. Access to food should be denied for 4–8 h for FSR

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measurements performed in the fasted state, whereas measurements should be made within 3 h of the start of the light cycle to ensure that FSR values reflect the “fed” or postprandial condition. Alternatively, animals can be fasted for 6–8 h during the dark cycle, and then given access to food for 90–120 min preceding the FSR measurement (see Note 13). 2. Before beginning the experiment, the lab should be set up with four work stations: (a) an injection station: this should have a pan balance for weighing the mouse, a lamp, the isotope injection solution, the injection line, and the supply of hub-less needles; (b) a euthanasia station: this should have an ice bucket into which three labeled blood collection tubes, one aluminum foil pouch, and a beaker with chilled ice water are placed before each mouse is injected; a large polystyrene weigh boat, large and medium scissors, and forceps; (c) a dissection station: with a shallow rectangular filled ice tray containing the dissection dishes (keep surface in ice until ready to use); dissection instruments and pins; a styrofoam bucket containing liquid nitrogen on which one labeled aluminum weigh boat per type of muscle is floated; (d) a weighing and storage station: with an analytical balance, prelabeled blood, and tissue storage tubes; liquid nitrogen in a styrofoam box for temporary sample storage; a microfuge (ideally refrigerated) for processing blood samples. 3. At least 2 or, ideally, 3 people are required for performing the intravenous injection. Weigh the animal and determine the volume of isotope solution to be injected (0.1 mL/10 g body weight); record both on worksheet. For the example used here, a 31 g mouse would receive 0.31 mL. Draw this volume into the syringe through the injection line ensuring that there are no air bubbles. Make a small slit in the center of the cheese cloth and thread the mouse tail through it. Fold the cheese cloth over the mouse and roll the mouse up so that it is totally restrained in the cloth with only the tail protruding (see Note 14). Holding the body of the mouse, dip the entire tail in the warm water for 90 s to dilate the tail veins. Lay the mouse on the bench on its side with the tail protruding under a bright light. One person should hold the mouse in this position and simultaneously hold the base of the tail firmly to prevent the mouse from jerking it during the injection. While also holding the tail, a second person inserts the needle into a lateral tail vein about half way along its length. The needle should be held in line with the vein and enter the skin at as flat an angle as possible, bevel side up. As soon as the needle looks like it is in the vein, a little suction is applied from the syringe by the third person, and some blood should be visible in the tubing if the needle is in the vein. If no blood returns, the needle is not in the vein; in this case remove the needle and attempt again

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proximal to the previous attempt (see Note 15). Once the needle is in the vein, the third person injects the solution over approximately 10 s (proportionally less or more time for smaller or bigger volumes), while the second person holds the tail and the needle, and the first holds the mouse and the tail. A timer is started (counting up) when half the solution has been injected. Once the tail is warmed, the injection can be completed in 30–45 s (see Note 16). After the entire volume is injected, the needle is removed, and pressure applied to the tail until it stops bleeding. The mouse is unwrapped and returned to its cage (ideally in a separate room). 4. Approximately 10 min later, the mouse is again taken from the cage, and using a 20G hypodermic needle, the opposite lateral tail vein to the one injected is punctured and 25 ML of blood is collected with a pipette and immediately mixed with 0.25 mL of 0.2M chilled PCA, vortexed, and set on ice. The exact amount of time elapsed at the time the blood was collected is recorded on the worksheet (T1). The tail bleeding is stopped by applying pressure and the mouse is returned to its cage (see Note 17). 5. After 29.75 min from the injection time, the animal is removed from the cage, and at exactly 30 min, the head of the mouse is excised (see Note 18) and the time is noted (T2). From the head, 50 ML of blood is collected and mixed with 0.5 mL chilled 0.2M PCA, vortexed well and placed on ice. The trunk blood is collected into another tube and kept on ice to clot; this is used for measuring hormones, metabolites, etc., and not pertinent to the protein synthesis measurement itself. Twenty to 30 s after the decapitation, both hind limbs are severed at the joint between the femur and the pelvis using scissors, the skin is peeled off, and the limbs are placed in the prechilled foil pouch, wrapped up, and buried in the ice to chill. This is recorded as the “chill time” (T3) for the hind limb muscles and represents the duration of the protein labeling period; no more than 40–50 s should elapse from decapitation to chilling. If the diaphragm is to be measured, the skin and abdominal wall muscles are cut, the organs below the diaphragm are rapidly removed and the carcass is submerged in the chilled water; the time of submersion is recorded (as T3 for the diaphragm). Similarly, if front limb muscles are needed, the arms are detached, skinned, and chilled as described for the hind limbs (see Note 19-very important). 6. One minute after the limbs are chilled, they are individually removed from the pouch (they should not be wet), and pinned out on the chilled and dried dissection dish in the ice tray. The muscles needed for measuring protein FSR are rapidly and quantitatively dissected using dry, chilled dissection instruments, and

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then dropped on the foil boats floating on the liquid nitrogen where they freeze instantaneously (see Note 20). After one limb is completely dissected, the second is removed from the pouch and the process repeated. The carcass is pinned out and the diaphragm dissected quantitatively, mopped rapidly with a Kimwipe® to remove surface water, and frozen. The muscles should not be on the chilled dissection dish for more than 2–3 min. The bones that subtend the muscles can be reserved and their lengths determined with Vernier calipers when time permits. 7. Once all muscles have been dissected and frozen, they are rapidly weighed on the analytical balance, transferred to chilled storage vials, and held in liquid nitrogen or dry ice until they can be stored at −80°C. After sitting on ice for at least 10 min, the PCA-treated blood samples are microfuged at 13,200 × g for 3 min and the clear supernatant is transferred to a microfuge tube, and stored frozen at −80°C. The whole blood is allowed to clot for >30–40 min, centrifuged, and the serum is recovered and stored frozen at −80°C. 8. To avoid cross-contamination, all instruments and dishes are washed and rechilled between mice. The needle on the injection line is replaced, and the process is repeated for the next animal. With experience and three people, the process from the time the mouse is euthanized to completion can be accomplished in less than 10 min. 3.2. Preparation of Tissue and Blood for Measurement of L-[43H]-Phenylalanine Specific Radioactivity

Keep in mind that all samples and waste are radioactive and need to be handled appropriately. 1. Blood: Thaw out blood PCA-supernatants on ice. Lay out a strip of pH paper on the bench. To each sample, add 10–20 ML chilled 4M KOH, vortex, and test pH. Repeat this process until pH 7 is attained (see Note 21). At neutral pH, potassium perchlorate is insoluble and precipitates out as white crystals. Repeat for all samples. Once samples have all been on ice for at least 30 min, centrifuge at 8,000 × g for 20 min at 2–4°C. Carefully remove supernatant and place in a clean microfuge tube and discard the precipitate. Place samples in the Speedvac to dry overnight. Cap and store at 4°C until they are purified over the ion-exchange column. 2. Purification of blood and muscle free precursor pool (see Note 22): Prepare ion exchange resin in H+ form as follows: add 1.25 mL of Dowex®-50 resin (prepared as in Note 11) to each column, and wash resin with 100 mL mQ water; then use pH paper to check that the pH of the eluant from the column is the same as the water. Add 5.0 mL of 1M HCl to each column; use pH paper to check that the final column eluant is acidic. Wash each column with ~75 mL mQ water and check that the

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eluant pH is the same as the water. Now columns are ready to use (see Note 23). Dissolve dried blood (from Subheading 3.2, step 1) or muscle free pools (from Subheading 3.2, step 4) in 1.25 mL of 1M acetic acid; vortex to dissolve and apply to prepared Dowex®-50 column. Wash the sample vial with 1 mL 1M acetic acid and apply to the same column; repeat this 3× so that a total of 5.25 mL is applied to the column. Check that the pH of the eluant is acidic using pH paper. Rinse Dowex®50 with 75 mL mQ water; then check that the pH of the eluant from the column is the same as water using pH paper. Put a clean (Chromerge®-washed), labeled glass vial under each column and elute amino acids with 3 mL of 3M NH4OH. Apply a further 3 mL of water to the column and collect eluant into vial. Place vials in the evaporator with the heating block at 65°C; direct nitrogen gas into each vial using the needle manifold assembly and reduce the volume by half at which point no ammonia smell should be detectable. Then transfer the vials to the Savant Speedvac to complete the drying. Dissolve each sample in 1.0 mL of mQ water and then filter into a microfuge tube using a syringe and syringe filter. Dry the filtered samples in the Savant Speedvac and reconstitute in a volume of water or reagent as required by the technique that will be used to isolate the phenylalanine and determine the radioactivity or stable isotope enrichment associated with it (see Note 24). 3. Muscle homogenization: Using the steel pestle and mortar chilled in liquid nitrogen, powder all muscles (see Note 25). Very rapidly weigh out approximately 50 mg of powdered muscle into a chilled, labeled, polypropylene tube. Record the exact weight on a worksheet. With the tube on the balance, tare, and add 1 mL of chilled mQ water. Record weight of water on worksheet and return the tube immediately to ice. Dry the chilled dispersion probe on the homogenizer by running it briefly in air while wiping with a Kimwipe®. Holding the sample tube in an ice jacket, homogenize the muscle for 3 × 30 s with 15 s rests in between. Cap the tube and place it on ice. Wash, chill, and dry the dispersion probe thoroughly between samples. Homogenize all samples before proceeding to the next step. 4. Analyzing for TP: Gently invert each tube several times to mix the homogenate, take an aliquot and make a 1:200 dilution with 0.1M NaOH for measuring TP using the BCA assay (see Note 26). Remove 400 ML of homogenate into another prechilled polypropylene tube; add 66 ML chilled 1.2M PCA, vortex immediately, and incubate on ice for >20 min. Centrifuge at 8,000 × g at 4°C for 15 min. Transfer the supernatant which contains the muscle free precursor pool into a 1.5 mL microfuge tube; cap and freeze at −20°C. These samples will then be

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processed in the same way as the blood free pool described in Subheading 3.2, step 1. Wash the remaining precipitate with 1 mL 0.2M PCA using a glass rod to help break up the pellet; vortex thoroughly. Incubate on ice for >15 min before centrifuging at 8,000 × g at 4°C for 15 min. Discard the supernatant. Repeat this wash step 4 more times (see Note 27). Before the last centrifugation, transfer the protein slurry quantitatively to a KIMAX® hydrolysis tube, using additional 0.2M PCA to rinse and ensure complete transfer. Centrifuge the tube at 1,500 × g at 4°C for 30 min. Discard the supernatant and invert the tube to drain off any remaining PCA. Then cap (with Teflon® insert) and store at −20°C until the hydrolysis is performed (see Subheading 3.2, step 6). 5. Isolation of myofibrillar proteins (MP, see Note 28): After taking the TP homogenate aliquot, add an equal volume of the 2× low salt/sucrose buffer to the remainder of the homogenate, cap, mix well, and place the tubes on the shaker in the cold room for 60 min to solubilize membranes. Then, centrifuge the homogenate at 1,500 × g at 4°C for 10 min to separate the soluble proteins (sarcoplasmic, hemoglobin, plasma proteins, and soluble extracellular proteins) from the insoluble MP pellet. Place the tubes on ice after centrifugation being careful not to shake them as the pellet is fragile. Carefully remove and discard the supernatant. Wash the MP pellet with 1 mL of the 1× low salt/sucrose buffer. Vortex well, and then centrifuge as in the previous step. Discard the supernatant and keep the pellet. Repeat the MP pellet wash in 1× low salt/ sucrose buffer. To the resulting MP pellet, add 1 mL of ice cold mQ water and vortex well to resuspend myofibrils. Centrifuge the suspension at 1,500 × g at 4°C for 10 min. Discard the supernatant and repeat the wash in ice cold mQ water. Resuspend the MP pellet in 1 mL of cold mQ water (vortex thoroughly) and quantitatively transfer the slurry to a large prechilled, polypropylene tube. Centrifuge at 8,000 × g for 15 min at 4°C. Gently remove supernatant and discard (see Note 29). To the MP pellet add 2.0 mL of 0.3M NaOH; vortex well and incubate in a water bath at 37°C for exactly 60 min, vortexing every 15 min to assist solubilization of MPs. This step separates the alkali-soluble MPs from the alkali-insoluble collagen. Repeat the centrifugation and then transfer the supernatant to another hydrolysis tube, and chill on ice. To the hydroxide MP solution, add 2/3 of its volume of chilled 1.2M PCA and incubate on ice for 30 min. Centrifuge at 1,500 × g at 4°C for 30 min and discard the supernatant. Wash the pellet with 1 mL chilled 0.2M PCA; vortex hard to disperse and centrifuge as before. Pour off the supernatant and leave tubes inverted at room temperature to drain; cap (with Teflon® liners) and store at −20°C.

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6. Acid hydrolysis of protein precipitates: The TP and MP acid precipitates should be in KIMAX® tubes at −20°C. Allow samples to thaw at room temperature; working in a fume hood, add 6M HCl to each tube (see Note 30). Use an acid-washed glass rod or glass Pasteur-pipette to completely disperse the pellet in the acid. Place the tubes in the evaporator heating blocks set at 110°C. While the samples are warming up, very gently blow nitrogen gas into each tube taking extreme care not to splash the acid and to avoid cross-contamination. The purpose of this is to displace as much oxygen as possible from the tubes and minimize sample oxidation. After each tube is gassed, loosely screw on the cap. When the samples start to bubble, repeat the nitrogen gassing and this time screw the cap on very tightly. When all samples are done, transfer the tubes to a metal rack and place them in an oven at 110°C to hydrolyze (see Note 31). After exactly 24 h remove the samples from the oven, cool, and place them in the Savant Speedvac to dry. Once dried, add 4 mL mQ water to each tube and leave to soften at room temperature for 30 min. Vortex at length to redissolve the pellet and then dry the sample down again. Repeat previous wash step two more times, for a total of three washes, and then resuspend the final pellet in 1 mL mQ water. Use a 3 mL syringe to filter the hydrolysate through a 13 mm, 0.4 Mm syringe filter into a 1.5 mL microfuge tube. Dry the sample and resuspend it in a volume of water or other diluent as required by the technique that will be used to isolate the phenylalanine and to determine the radioactivity or stable isotope enrichment associated with it (see Note 32). 7. The specific radioactivity of the phenylalanine in the blood and tissue free amino acid pools and in the protein hydrolysates is measured by HPLC (see Note 33) and scintillation counting (for radioactive tracers or mass spectrometry for stable isotope tracers). Values (dpm/nmol) are calculated for the specific radioactivity (or MPE if a stable isotope tracer was used) of the blood phenylalanine (SABLD) at T1 and T2, the tissue free pool (SAFP), and the protein (TP and MP) bound tracer (SATP or SAMP). 3.3. Calculations for Estimating Protein Synthesis Rates

Before performing the final calculations, determine the ratio of tissue SAFP at T3 to the SABLD at T2 for each animal. If values are <0.95 or >1, it indicates that there likely was a problem (several possible causes), and it is unlikely that accurate, valid FSR values will be obtained. To determine the average SAFP ( SA FP ) for the entire labeling period (see Notes 34 and 35), calculate the rate of change of SABLD over the labeling time: ' BLD [(dpm/nmol) / min] [(SA BLD at T1)  (SA BLD atT2)] / (T2  T1).

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Then use this value, to calculate SA FP : SA FP

SA FP at T3  [' BLD u (T3 / 2)].

Calculate the FSR (in %/day) values for TP and MP: FSR TP

(SA TP / SA FP ) u (1, 440 / T3 ) u 100,

FSR MP

(SA MP / SA FP ) u (1, 440 / T3 ) u 100.

The ASRTP (mg protein/day) is estimated as: ASR TP

(FSR TP / 100) u TP mass of the muscle.

The TP mass of a muscle is the product of the total wet weight of that muscle and the protein concentration determined from the sample of homogenate taken in Subheading 3.2, step 4. 3.4. Approach for Estimating Rates of Protein Degradation

Estimates of protein degradation averaged over relatively chronic periods (days) during which there are measureable changes in protein mass can be calculated from the protein synthesis and protein accretion rates (for example see refs. (13, 14)). To derive these data, the change in the protein mass (as mg/day) of the muscle of interest over an interval is measured by collecting and quantifying protein mass at the beginning and the end of the interval. The average protein synthesis rate over this time is then determined; a single measurement can be made at the midpoint of the interval, or the weighted average of a number of measurements is taken. Alternatively, if several values are obtained so that the change in muscle mass over time can be described with an equation, the velocity at the time points when protein synthesis is measured can be estimated by differentiation. The difference between protein synthesis and accretion rates will represent the average degradation rate.

4. Notes 1. The tracer should be administered in the smallest feasible volume. This is dictated by the solubility of phenylalanine, which is close to 150 mM. Thus, administration of 0.1 mL/10 g body weight of a 150 mM solution (15 Mmol/10 g body weight) will constitute a flooding dose. Alternatively, a greater volume of a more dilute solution can be administered (e.g., 0.2 mL/10 g body weight of a 75 mM solution). This may be necessary if a final specific radioactivity activity higher than 6.67 mCi/mmol phenylalanine is needed.

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2. The amount of radioactive tracer to add is a compromise between the cost of the isotope-labeled tracer and the counting precision for the incorporated 3H-phenylalanine. 14C-ring labeled phenylalanine could be used but is much more expensive. It is useful to keep the following facts in mind in estimating how much tracer to use: (a) on average phenylalanine constitutes 6% of body proteins by weight; (b) when using a radioactive 3H-labeled tracer, the largest source of error usually comes from the uncertainty in counting the 3H associated with the phenylalanine incorporated into the protein(s); the error is the square root of the total net counts accumulated. For example, the uncertainty in a net (total – background) value of 400 counts is ±20 counts, i.e. ±5%. Thus, administering more tracer to the animal, increasing the amount of sample analyzed, reducing background activity, increasing the counting time of the samples (e.g., if the 400 counts were collected over 20 min, increasing this to 60 min would reduce the error to ±3%), and factors that affect the efficiency of counting 3H (the scintillation counter itself, the scintillation fluid, the composition and preparation of the sample analyzed) all factor into the final decision of the specific radioactivity of the injection solution. Ideally, one should aim for a counting error of the incorporated 3H-phenylalanine of less than ±2–3%. 3. It is important that the “cold” phenylalanine used is as close to 100% pure L-phenylalanine. It should be stored desiccated. 4. The solution should be certified as being >98% pure L-phenylalanine. Unless one has a chiral column to independently verify the R-phenylalanine content, one has to take the manufacturer’s word for it. Ideally, the solution should be run over an HPLC column and counted to verify that all the 3H counts are in phenylalanine and that the amount specified is present (we find that it is usually approximately 93–95%). However, this is not the case with every batch, in which case it should be either returned for another batch, purified in-house, or take into account the activity present, provided it is not also present in another amino acid (such as tyrosine, as is often the case). The amount of “cold” carrier phenylalanine in the solution is negligible and can be ignored in the calculations. The stock tracer solution is vulnerable to radiolysis and should be stored at 4°C for periods of <1–2 months or frozen for longer periods. 5. The phenylalanine takes a while to dissolve (hours). To facilitate the process, warm the solution in a water bath at 37°C before putting the vial on the stir plate. Spin the stir bar as fast as possible (tape the vial to the plate so it does not fall over). The solution must be completely clear.

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6. This procedure is for the acute intravenous administration of the amino acid tracer solution. If the animal has a catheter already implanted, it can be used to administer the tracer. To remove the hub from the needle, hold the needle with a pair of forceps and flame the hub with a Bunsen burner (5–10 s), enough to soften the plastic material. Use hemostats to pull the hub off the needle. This should leave the “hub-end” of the needle with a white coating and slightly flared at the point where the needle emerged from the hub. If the white coating is removed, you cannot use the needle as it will be too small for the tubing. Prepare one needle per animal. 7. If the animal does not have an indwelling catheter to permit the rapid administration of a large dose of anesthesia, decapitation without the use of anesthesia is used to euthanize rodents and collect trunk blood. This requires detailed justification and special approval from the IACUC committee. Its use is warranted because the prior use of anesthesia or sedation will alter the rate of protein synthesis to variable and not always predictable extents (24). See Note 18 for alternatives. 8. Make these from heavy duty aluminum foil. 9. PCA is hazardous and should always be used with caution. 10. To prepare the Dowex® resin, suspend the entire bottle of resin in mQ water in a 2 L beaker. Mix with a spatula; never use a magnetic stirrer. Pour off water and repeat the wash 3×. Suspend the resin in mQ water again, and allow it to sit for 30 min with occasional stirring. Decant the water and repeat this last step 6×. Suspend the resin in 1M NaOH and allow it to sit overnight with occasional stirring. Decant and resuspend in the resin in 0.4 L of 1M NaOH. Allow it to sit for 4 h with occasional stirring. Repeat the last step 6×, leaving overnight when necessary. Decant and resuspend the resin in 2 volumes of 1M NaOH (verify that it is very basic) and store at 4°C. 11. This is expensive; only make up as much as you need. Prepare in the fume hood. 12. As concentrated HCl will be evaporated, the system must have an acid trap, and the refrigeration component must attain −104°C to prevent HCl fumes from reaching and destroying the vacuum pump. 13. If the fasting and refeeding approach is taken to ensure that postprandial FSR values are obtained, the animal should be weighed before and after refeeding, and/or stomach contents recorded to ascertain that the animal had eaten. 14. This is often named the “mouse burrito” because of the similar appearance. If done correctly, the mouse is lightly restrained and stays quite still.

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15. Before inserting the needle, always make sure that the isotope solution is all the way to the tip and that no air bubbles have been inadvertently introduced. 16. With experience, it is possible for the same person to place the needle and do the injection. It is possible to use this approach to reliably inject mice as small as 10 g. For mice smaller than 10 g, or if the tail vein injection cannot be performed, the isotope tracer can be administered intraperitoneally using a 30G needle. This has the disadvantage that one cannot be sure until after the analysis if any of the isotope solution was injected into the intestine, and the equilibration time within the precursor free pool is not immediate as when the dose is administered intravenously (25, 26). Additionally, some of the isotope solution may leak out of the hole left by the needle, especially in smaller animals. It is important to keep precise notes on each animal to establish if any technical issues encountered are associated with FSR values that deviate significantly from others in the same treatment group and, therefore, are most likely erroneous. 17. If the total duration of the labeling period is shortened to 15 min or less, the intermediate time point blood sample can be omitted. 18. If the animal has an indwelling catheter, an overdose of anesthetic can be used to rapidly euthanize it (<15–30 s). Once the animal is unconscious, it is decapitated, and the same procedures followed. 19. Once the tissue has been chilled, it must NEVER be allowed to warm up until the isolated protein has been treated with PCA. This is important to prevent proteolysis. Proteolysis will result in the dilution of the precursor pool with unlabeled phenylalanine and, thus, the specific radioactivity values for the precursor pool will be inaccurate (too low), and the estimate of FSR will be wrong (overestimated). 20. An alternative approach is to use one limb to take pieces of muscle rapidly (not quantitatively) and freeze for measuring protein FSR. The second leg is then used to quantitatively dissect out the muscles with less concern about them warming up. This is possible with larger animals/muscles where sufficient muscle for performing all the analyses can be obtained from one side. 21. Care should be taken to ensure that the KOH does not become contaminated. If the samples become too alkaline, they can be back titrated with PCA. 22. Purification of the free pool amino acids over the ion exchange column is necessary to obtain a “clean” phenylalanine peak when the samples are subjected to HPLC using an anion exchange

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column (PA-1 column, DIONEX). If an alternative technique is used, the need for this clean up step may be unnecessary. We have found that it is rarely necessary to perform this step on samples from rats and pigs, but essential for mouse samples. 23. The columns can be prepared ahead of time and then capped with mQ water and stored in the cold room (for a day or so). The water is drained from the columns before the sample is applied. It is not difficult to process up to 24 samples at once. 24. The amount of free tracee amino acid in the sample can be roughly estimated. For example, in the present protocol the mice are administered 1.5 mmol phenylalanine/kg body weight which equilibrates in the body fluid compartment. If we assume the latter to be approximately 60% of body weight, the concentration will be approximately 2.5 Mmol/mL water, or 40 nmol in 50 ML of blood, or 35 nmol in a 400 ML aliquot of the 1:20 muscle homogenate. The sensitivity of the detection system will dictate how much the samples should be diluted, and the appropriate sample size that should be analyzed in the first place. 25. It is best to powder all samples before proceeding to the analysis step. To avoid cross-contamination of samples, the mortar and pestle should be warmed to room temperature, washed, and chilled in liquid nitrogen between every sample. 26. For an adult muscle, this will yield a solution of approximately 50 Mg protein/mL. After incubating the samples at 37°C for 1 h to solubilize the proteins, the exact protein content is determined using a protein assay with the protocol optimized for this concentration. 27. All the washes are very important to ensure that the free phenylalanine tracer is completely removed. The specific radioactivity of the precursor pool can be several hundred times higher than that of the protein. Thus, even minute levels of contamination can lead to overestimation of the final FSR values. 28. The isolation of the MPs is not quantitative in this procedure. Thus, only FSR of the MPs will be obtained. An independent quantitative measurement of the MP concentration in the muscle is needed to estimate ASR. 29. At this point an aliquot of the MP fraction can be reserved and used to isolate individual MP proteins, e.g., by SDS/PAGE electrophoresis with or without prior immunoprecipitation of the protein of interest. The purified protein is hydrolyzed as described in Subheading 3.2, step 6, and its phenylalanine specific radioactivity is then determined. The FSR of that protein is estimated using the same precursor pool values as for the TP and MP. If these analyses are to be performed, it may be necessary to begin with a larger sample size.

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30. The volume of HCl added depends on how much protein is to be hydrolyzed; this should be at least 1 mL/2 mg of protein. For example, the TP pellet from a 400 ML aliquot of the 1:20 muscle homogenate will contain roughly 4 mg of protein (assumes a muscle protein concentration of 200 mg/g muscle) and the corresponding MP pellet will be 3 mg (assumes MPs to be 50% of TP). Thus, 2 and 1.5 mL of 6M HCl will be added to the TP and MP pellets, respectively. As the exact amount of sample used is known and the protein content of the muscle is measured, the actual amount of protein can be estimated more accurately; however, this is not really necessary unless the muscle protein concentration is expected to be very different, e.g., in immature muscles. 31. The temperature is critical. At lower temperatures, the hydrolysis of the proteins may not be complete in 24 h. At higher temperatures, the H isotope atom label on the C-4 position of the phenylalanine ring can exchange with H or OH and must be avoided. It is recommended that the heating block and oven are monitored to ensure that they do not go above 110°C before the samples are placed in them. 32. Again one can make some rough calculations based on the initial amount of protein and assuming that protein is 6% phenylalanine. Thus, a 4 mg protein pellet will contain approximately 0.24 mg or 1.4 Mmol of phenylalanine. 33. There are some reports where the radioactivity of the samples (blood and tissue free pools, and protein bound) are counted directly, and the concentration of the phenylalanine is measured on a separate aliquot of the sample rather than by isolating the phenylalanine and determining its radioactivity. The former approach has the advantage that it is faster and less tracer can be used for labeling the animal in vivo because larger amounts of sample can be counted. However, it has the major drawback that it assumes that all the activity in the sample is associated with the phenylalanine; this probably is not the case and would introduce an unknown amount of error. 34. A number of assumptions go into the estimation of the average specific radioactivity or enrichment of the precursor pool. The first is that the specific radioactivity of the aminoacyl-tRNA, the true precursor, and muscle free amino acid pools (a surrogate that is measured in this protocol) are equilibrated throughout the labeling period. There has been extensive debate in the literature, because the routine measurement of phenylalaninetRNA specific radioactivity or enrichment is not as practical as measuring that of the tissue free pool. Under the conditions described in this protocol, we have verified that the assumption is valid (16). If the isotope solution has been administered intraperitoneally, a correction to the precursor pool value is

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recommended to account for the slower equilibration of the injected tracer (25, 26). In the present calculations, we also assume that any decrease in specific radioactivity over the 30 min labeling period is linear. It is more likely to be an exponential decrease, but the error introduced by assuming it is linear is negligible. 35. If a number of muscles from the same animal are analyzed separately, it is not necessary to measure the SAFP for each muscle. We have evaluated this numerous times, and determined that the variation in SAFP among muscles is not greater than the analytical error for any one muscle. This assumption is not necessarily valid for different tissues; e.g., it is not advisable to use a muscle SAFP value for calculating liver FSR. References 1. Davis TA, Fiorotto ML (2009) Regulation of muscle growth in neonates. Curr Opin Clin Nutr Metab Care 12:78–85 2. Davis TA, Fiorotto ML (2005) Regulation of skeletal muscle protein metabolism in growing animals. In: Burrin DG, Mersmann HJ (eds). Biology of Metabolism of Growing Animals. Elsevier, The Netherlands, pp. 37–68 3. Rooyackers OE, Nair KS (1997) Hormonal regulation of human muscle protein metabolism. Annu Rev Nutr 17:457–485 4. Davis TA, Fiorotto ML, Burrin DG, et al (1999) Protein synthesis in organs and tissues: quantitative methods in laboratory animals. In: El-Khoury AE (ed), Methods for Investigation of Amino Acid and Protein Metabolism. CRC Press, Boca Raton, 49–68 5. Reeds PJ, Davis TA (1999) Of flux and flooding: the advantages and problems of different isotopic methods for quantifying protein turnover in vivo: I. Methods based on the dilution of a tracer. Curr Opin Clin Nutr Metab Care 2:23–28 6. Davis TA, Reeds PJ (2001) Of flux and flooding: the advantages and problems of different isotopic methods for quantifying protein turnover in vivo: II. Methods based on the incorporation of a tracer. Curr Opin Clin Nutr Metab Care 4:51–56 7. Garlick PJ, McNurlan MA, Essen P, et al (1994) Measurement of tissue protein synthesis rates in vivo: a critical analysis of contrasting methods. Am J Physiol Endocrinol Metab 266:E287–E297 8. Wagenmakers AJ (1999) Tracers to investigate protein and amino acid metabolism in human subjects. Proc Nutr Soc 58:987–1000

9. Chinkes DL (2005) Methods for measuring tissue protein breakdown rate in vivo. Curr Opin Clin Nutr Metab Care 8:534–537 10. Bergen WG (2008) Measuring in vivo intracellular protein degradation rates in animal systems. J Anim Sci 86:E3–12 11. Vissers YL, von Meyenfeldt MF, Braulio VB, et al (2003) Measuring whole-body actin/ myosin protein breakdown in mice using a primed constant stable isotope-infusion protocol. Clin Sci (Lond) 104:585–590 12. Emery PW, Preedy VR (2003) Measuring muscle protein turnover in vivo: what can 3-methylhistidine production tell us? Clin Sci (Lond) 104:557–558 13. Johnson JD, Dunham T, Skipper BJ, et al (1986) Protein turnover in tissues of the rat fetus following maternal starvation. Pediatr Res 20:1252–1257 14. Fiorotto ML, Davis TA, Reeds PJ (2000) Regulation of myofibrillar protein turnover during maturation in normal and undernourished rat pups. Am J Physiol Regul Integr Comp Physiol 278:R845–R854 15. Garlick PJ, McNurlan MA, Preedy VR (1980) A rapid and convenient technique for measuring the rate of protein synthesis in tissues by injection of [3H]phenylalanine. Biochem J 192:719–723 16. Davis TA, Fiorotto ML, Nguyen HV, et al (1999) Aminoacyl-tRNA and tissue free amino acid pools are equilibrated after a flooding dose of phenylalanine. Am J Physiol Am J Physiol Endocrinol Metab 277:E103–E109 17. Louis M, Poortmans JR, Francaux M, et al (2003) No effect of creatine supplementation on human myofibrillar and sarcoplasmic

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19.

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M.L. Fiorotto et al. protein synthesis after resistance exercise. Am J Physiol Endocrinol Metab 266: E287–E297 Balagopal P, Ljungqvist O, Nair KS (1997) Skeletal muscle myosin heavy-chain synthesis rate in healthy humans. Am J Physiol Endocrinol Metab 272:E45–E50 Rooyackers O, Adey D, Ades P, et al (1996) Effect of age on in vivo rates of mitochodrial protein synthesis in human skeletal muscle. Proc Nat Acad Sci USA 93: 15364–15369 Welle S, Thornton C, Jozefowicz R, et al (1993) Myofibrillar protein synthesis in young and old men. Am J Physiol Endocrinol Metab 264:E693–E698 Rennie MJ, Smith K, Watt PW (1994) Measurement of human tissue protein synthesis: an optimal approach. Am J Physiol Endocrinol Metab 266:E298–E307

22. Smith PK, Krohn RI, Hermanson GT, et al (1985) Measurement of protein using bicinchoninic acid. Anal Biochem 150:76–85 23. Davis TA, Fiorotto ML, Nguyen HV, et al (1991) Response of muscle protein synthesis to fasting in suckling and weaned rats. Am J Physiol Regul Integr Comp Physiol 261:R1373–R1380 24. Heys SD, Norton AC, Dundas CR, et al (1989) Anaesthetic agents and their effect on tissue protein synthesis in the rat. Clin Sci (Lond) 77:651–655 25. Jepson MM, Pell JM, Bates PC, et al (1986) The effects of endotoxaemia on protein metabolism in skeletal muscle and liver of fed and fasted rats. Biochem J 235:329–336 26. Bregendahl K, Liu L, Cant JP, et al (2004) Fractional protein synthesis rates measured by an intraperitoneal injection of a flooding dose of L-[ring-2H5]phenylalanine in pigs. J Nutr 134:2722–2728

Part IV Generation of Viral Vectors and Transgenic Mice

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Chapter 15 Recombinant Adeno-Associated Viral Vector Production and Purification Jin-Hong Shin, Yongping Yue, and Dongsheng Duan Abstract Gene delivery vectors based on recombinant adeno-associated virus (AAV) are powerful tools for studying myogenesis in normal and diseased conditions. Strategies have been developed to use AAV to increase, down-regulate, or modify expression of a particular muscle gene in a specific muscle, muscle group(s), or all muscles in the body. AAV-based muscle gene therapy has been shown to cure several inherited muscle diseases in animal models. Early clinical trials have also yielded promising results. In general, AAV vectors lead to robust, long-term in vivo transduction in rodents, dogs, and non-human primates. To meet specific research needs, investigators have developed numerous AAV variants by engineering viral capsid and/or genome. Here we outline a generic AAV production and purification protocol. Techniques described here are applicable to any AAV variant. Key words: AAV, Adeno-associated virus, Muscle, Gene therapy, Gene transfer/delivery, Serotype, Muscular dystrophy, Dystrophin, Alkaline phosphatase

1. Introduction Adeno-associated virus (AAV) is a single-stranded DNA virus discovered in 1965 (1). It belongs to the Dependovirus genus of the Parvoviridae family. The 4.7 kb wild type AAV genome encodes two major open reading frames. The rep gene expresses viral replication proteins and the cap gene expresses viral capsid proteins. At the ends of the AAV genome are the inverted terminal repeats (ITRs). The ITR forms a T-shaped hairpin structure. It is essential for AAV replication and packaging. The mature virion is a ~20 nm nonenveloped icosahedral particle containing either a plus- or minus-strand genome. Although mature AAV virion is infectious in mammalian cells, the replicative AAV life cycle requires helper

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function from adenovirus or herpesvirus (2). In the absence of helper virus coinfection, AAV genome is either integrated in the host genome or maintained as double stranded circular episomes (3–5). Recombinant AAV vector is generated by replacing the wild type AAV open reading frames with a target (therapeutic or marker) gene expression cassette. Since initial cloning of the AAV genome into a plasmid format in early 1980s (6, 7), tremendous progress has been made in developing AAV into a versatile and effective gene delivery vehicle (8). Recent clinical success in AAV gene therapy for inherited diseases further raises the enthusiasm of applying AAV technology in translational medicine (9–12). AAV vector is one of the most attractive gene transfer tools in studying basic muscle biology and in developing novel genetic therapies for muscle diseases. Direct local muscle injection and systemic (intravascular or intraperitoneal) AAV administration have been used to achieve single muscle, muscle group, and even whole body muscle transduction. These preclinical studies have revealed robust and persistent (in months and years) transgene expression in normal and diseased muscles in various animal models including non-human primates. Besides gene addition/replacement/overexpression, AAV has also been used to down-regulate gene expression (e.g., via RNA interference) or to modulate RNA processing (e.g., via exon skipping). In many cases, AAV-mediated muscle gene transfer has helped investigators to obtain critical information that may otherwise take years to get if a conventional approach is used (such as transgenic modeling in mice). Most of the earlier AAV gene transfer studies used AAV serotype-2 (AAV-2). To further improve the efficiency and specificity of AAV-mediated gene transfer, investigators have developed numerous AAV variants by viral genome engineering and/or capsid modification. The single-stranded nature of the AAV genome impedes immediate transcription. To circumvent this problem, self-complementary AAV (scAAV) vectors are developed by removing the terminal resolution site in one of the ITRs (13). By deleting the d-sequence in one of the ITRs, single polarity AAV vector can also be generated containing either the plus or the minus strand only (14, 15). A major limitation of AAV vector is its small packaging capacity. A series of dual vector strategies including cis-activation, trans-splicing, overlapping, and hybrid are now available to effectively overcome this constraint (16). Several rate-limiting steps of AAV transduction (such as the uptake, intracellular processing, and uncoating) are determined by the capsid. Besides naturally occurring AAV serotypes, many creative strategies have been explored to generate novel viral particles with distinctive phenotypes. These include proviral sequence rescue from mammalian tissues, rational design based on known AAV structure/biology (such as peptide ligand insertion, mosaic

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capsid reconstitution, and tyrosine mutation), and direct evolution by error-prone PCR/DNA shuffling (reviewed in refs. (17– 20)). Collectively, these newly engineered AAV vectors offer a broad range of selection to meet different experimental needs. With the expansion of our knowledge on AAV transduction biology and AAV vector engineering, methods for AAV production and purification have also gone through revolutionary changes (reviewed in refs. (21–27)). The original method requires three components including a cis plasmid carrying an ITR-flanked target gene expression cassette, a trans plasmid supplying AAV replication and structural proteins, and a wild type adenovirus as the helper. A complicated procedure involving plasmid cotransfection and adenovirus coinfection is carried out to generate crude AAV lysate. AAV vector is then extracted from the crude lysate through one round of cesium chloride (CsCl) gradient banding. Besides being cumbersome and low yield, high level adenovirus carryover often skews experimental results. The newly developed transient plasmid cotransfection method has essentially solved the issue of adenovirus contamination. To meet the need of large animal study and clinical trial, new platforms have also been developed using producer cell lines, baculovirus system, and column chromatography purification for large-scale production. In this protocol, we outline a procedure based on transient plasmid cotransfection and CsCl isopycnic ultracentrifugation. This method can be used to generate high quality vector stock of any AAV variant.

2. Materials 2.1. Recombinant AAV Vector Production

1. 293 Cells (American Type Culture Collection). This is an adenovirus transformed human fetal kidney cell line (28). A 4.3 kb adenoviral DNA (nucleotide 1–4,344) is inserted in chromosome 19 in these cells. They constitutively express adenoviral E1a and E1b gene (29) (see Note 1; Fig. 1). 2. Dulbecco’s modified Eagle’s medium (DMEM) containing high glucose and l-glutamine. Store at 4°C. 3. Fetal bovine serum (FBS). Store at −20°C. 4. 100× Penicillin G (10,000 Unit/mL) – Streptomycin (10 mg/ mL). Store at −20°C. 5. Adenoviral helper plasmid (pHelper) (Stratagene) (see Notes 2 and 3). 6. AAV helper plasmid (see Notes 2 and 4). 7. The cis plasmid carrying the vector genome (ITR-promotertarget gene-pA-ITR) (see Note 2).

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Fig. 1. 293 Cell propagation and transfection. (a) Schematic outline of 293 cell propagation. A freshly thawed stock vial may take up to 3 days to reach confluency. Split it 1:3. Forty-eight hours later, split again to 10 × 150 mm plates. After another 48 h, split to 30 × 150 mm plates. At this stage you may split 1:3 for the number of plates needed for your adeno-associated virus (AAV) preparation and freeze the remaining as stock. If you are doing a large preparation, you may split all 30 plates to 90 plates. Cells are usually ready for transfection at ~48 h after the last split. (b) Representative photomicrographs showing 293 cells at ~60% (left ), 70% (middle) and 80% (right ) confluency. Transfection at 70% confluency gives the highest AAV yield. (c, d) Representative photomicrographs showing transfection efficiency at 50 h after calcium phosphate transfection. (c) Shows a good transfection and (d) shows a poor transfection. In case of poor transfection, one should stop AAV preparation and re-start. The viral yield is usually several log folds lower from poorly transfected cells.

8. 2.5 M CaCl2. Sterilize by filtration and store at −20°C. 9. 2× HBS buffer: 300 mM NaCl, 1.5 mM Na2HPO4, and 40 mM HEPES, pH 7.05 ± 0.05. Sterilize by filtration and store at −20°C (see Note 5). 10. 150 mL Corning Pyrex fleaker. 11. Cell lifter (Corning). 12. 150 mm Cell culture plates. 13. IEC Centra CL3R refrigerated table top centrifuge. 14. 5, 10, and 25 mL Sterile Costar Stripette disposable pipettes. 15. 250 mL Sterile polypropylene centrifuge bottle. 16. 10 mM Tris–HCl (pH 8.0). 17. 15 mL Sterile polypropylene centrifuge tube.

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1. Dry ice/ethanol bath. 2. 37°C Water bath. 3. 15 and 50 mL Sterile polypropylene centrifuge tubes. 4. Misonic Sonicator S3000. 5. DNase I (11 mg protein/vial, total 33 K [kunitz] units) (see Note 6). 6. 10% Sodium deoxycholate. Store at room temperature. 7. 0.25% Trypsin-EDTA. Store at 4°C. 8. Optical grade CsCl. 9. Eppendorf 5810R bench-top refrigerated centrifuge. 10. Beckman-Coulter Optima XL-80 ultracentrifuge (see Note 7). 11. Beckman swinging bucket 55 titanium (SW55 Ti) rotor (see Note 7). 12. 1 in. 20G Needle. 13. 1.5 in. 25G Needle. 14. 5.1 mL (13 × 51 mm) Beckman polyallomer ultracentrifuge tube (see Note 8). 15. 10,000 Molecular weight cutoff (MWCO) Slide-A-Lyze dialysis cassette (Pierce) (see Note 9). 16. AAV Dialysis buffer: 150 mM NaCl, 20 mM HEPES, pH 7.4. Autoclave. Cool to 4°C before use.

2.3. Titer Determination and Quality Control

1. AAV slot blot digestion buffer: 400 mM NaOH, 20 mM EDTA. Freshly made before use. 2. Slot blot hybridization solution: 5× SSC, 5× Denhardts’s solution, 1% sodium dodecyl sulfate (SDS), and 50% formamide, add 100 Mg/mL denatured salmon sperm DNA just before use. 3. Bio-Dot SF manifold microfiltration apparatus (Bio-Rad). 4. Hybond-N plus membrane (GE Healthcare). 5. Techne HB-1D roller bottle hybridization oven. 6. AAV PCR alkaline digestion buffer: 25 mM NaOH, 0.2 mM EDTA. 7. AAV PCR neutralization buffer: 40 mM Tris–HCl, pH 5.0. 8. Primers for quantitative PCR determination of AAV titer (see Table 1). 9. ABI 7900HT fast real-time PCR system. 10. Fast SYBR green PCR master mixture. 11. MicroAmp optical 96-well reaction plate. 12. MicroAmp optical adhesive film. 13. AAV copy number controls (see Note 10).

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Table 1 Primers for quantitative PCR Target region

Primer sequence

Product size (bp)

RSV promoter

Forward (DL553): GGTTGTACGCGGTTAGGAGT Reverse (DL554): GGCATGTTGCTAACTCATCG

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CMV/CAG enhancer

Forward (DL560): TTACGGTAAACTGCCCACTTG Reverse (DL561): CATAAGGTCATGTACTGGGCATAA

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Human DMD ex 69/70

Forward (DL1294): TTTTCTGGTCGAGTTGCAAAAG Reverse (DL1295): CCATGTTGTCCCCCTCTAAGAC

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14. JEOL JEM-1400 transmission electron microscope. 15. Holey carbon coated copper 200 mesh grid (Polysciences). 16. 98% Uranyl acetate (see Note 11). 17. EndoSafe Limulus Amebocyte Lysate (LAL) gel clot test kit (Charles Rivers Laboratory).

3. Methods 3.1. Recombinant AAV Vector Production

1. Propagate 293 cells in high-glucose DMEM containing 10% FBS and 1× penicillin G-streptomycin. Approximately 1½ week prior to viral production, thaw a vial of stock 293 cells in one 150 mm culture plate (see Fig. 1). When cells reach 70% confluency (24–72 h), split cells to three 150 mm culture plates. Forty-eight hours later, split cells to 10 × 150 mm plates. After another 48 h, split again to 30 × 150 mm plates. At this stage, either split cells to 90 × 150 mm plates and use them for AAV production, or split to the number of the plates as needed and save the remaining in −80°C (e.g., split 10 plates to 30 plates for AAV production and freeze 20 plates for future use) (see Fig. 1a). Usually, 48 h after the last split, cells should be ready for transfection. Change to fresh culture media about 1–2 h before transfection (see Note 12). 2. Prepare DNA-calcium-phosphate precipitate for cotransfection of the cis plasmid, AAV helper plasmid, and the adenoviral helper plasmid. For a 15 × 150 mm plate preparation, use 187.5 Mg of the cis plasmid, 562.5 Mg of the AAV helper plasmid and 562.5 Mg pHelper. Warm up 2.5 M CaCl2 and 2× HBS buffer at 37°C for 20 min. Mix all plasmids thoroughly in 15.2 mL H2O. Incubate at 37°C for 10 min. Add 1.68 mL of

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2.5 M CaCl2 to the plasmid mixture (a final CaCl2 concentration of 250 mM). Mix well and incubate at 37°C for 5 min. Add 16.8 mL of 2× HBS to a 150 mL Corning fleaker. Slowly drop the DNA/CaCl2 mixture to 2× HBS to generate DNAcalcium-phosphate precipitate. Gently swirl the 2× HBS buffer while dropping the DNA/CaCl2 mixture. Incubate at room temperature for 15 min (see Notes 13 and 14). 3. Gently apply the DNA-calcium-phosphate precipitate (~2.2 mL/150 mm plate) to 293 cells drop-by-drop evenly to the entire plate while swirling the culture plate. 4. Around 60 h after transfection, scrape cells from 150 mm plates with a cell lifter. Split crude lysate to two 250 mL Corning centrifuge bottles. Carefully rinse off all cells from plates to the centrifuge bottles (see Note 15). 5. Spin at 1,500 × g for 5 min at 4°C in an IEC Centra CL3R centrifuge. Resuspend cell pellet into 10 mM Tris–HCl at 5 mL/centrifuge bottle. Rinse each centrifuge bottle with 2 mL of 10 mM Tris–HCl. Combine cell lysate into two 15 mL centrifuge tubes (7 mL/tube). Store the crude lysate in −80°C until purification. 3.2. Recombinant AAV Purification

1. Freeze/thaw the crude lysate (in 15 mL tubes) 8 times by rotating through a dry ice/ethanol bath (7 min/round) and a 37°C water bath (7 min/round). 2. Combine the crude lysate to a 50 mL tube and bring the final volume to ~21 mL with 10 mM Tris–HCl. 3. Sonicate the crude lysate on ice using the Misonic Sonicator at the power output of 2 for 7 min (see Note 16). 4. Add 2 mL of reconstituted DNase I and incubate at 37°C for 30 min (see Note 6). 5. Sonicate the crude lysate again under the same setting (power output 2 for 7 min). 6. Add 2.5 mL of 10% sodium deoxycholate and 2.1 mL of 0.25% trypsin-EDTA. Mix well. Incubate at 37°C for 30 min and then chill on ice for 20 min (see Note 17). 7. Add 16.9 g CsCl. Mix well. Incubate at 37°C for 20 min. Periodically shake the tube to assist CsCl dissolving (see Note 18). 8. Centrifuge at 3,000 × g for 30 min at 4°C in an Eppendorf 5810R centrifuge. Carefully transfer the clear lysate to six 5.1 mL Beckman polyallomer ultracentrifuge tubes (~5 mL/ tube) (see Note 19) (see Fig. 2a). 9. Load the lysate to a SW55 Ti rotor and spin at 266,400 × g (53,000 rpm) for 30 h at 4°C in a Beckman-Coulter Optimal XL-80 ultracentrifuge (see Notes 7 and 20).

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Fig. 2 Self-made needle stylet and the position of needle penetration when collecting AAV fractions. (a) The solution appears turbid after CsCl is completely dissolved (left ). Cell debris forms a thin lipid-like layer at the top after spinning (right). (b) Left, a 1 in. 20G needle and a 1.5 in. 25G needle; Right, assembled needle stylet. (c) A horizontal line is visible at ~0.5 cm above the bottom of a 5.1 mL Beckman polyallomer ultracentrifuge tube. When pulling fractions, insert the needle stylet horizontally into the centrifuge tube at the level of this line. Stop when the needle tip reaches at the center. Face the needle tip opening upward. Start collecting fractions.

10. Assemble a self-made needle stylet by inserting a 1.5 in. 25G needle into the lumen of a 1 in. 20G needle (see Fig. 2b). Collect fractions from the bottom of the tube with the needle stylet (see Fig. 2c) (see Note 21). Identify the viral containing fractions by slot blot or quantitative PCR (see Subheading 3.3) (see Note 22). 11. Combine AAV-containing fractions into a new 5.1 mL Beckman polyallomer ultracentrifuge tube (see Note 21). Repeat ultracentrifugation under the same setting. 12. Combine fractions with the highest AAV titer from each Beckman ultracentrifuge tube and perform the third round of ultracentrifugation under the same setting (see Note 23). 13. Combine fractions with the highest AAV titer. Dialyze virus in three changes of AAV dialysis buffer at 4°C (4 L buffer/ change, 12–16 h/change) (see Note 24). 3.3. Titer Determination and Quality Control

1. Determine AAV titer by slot blot. Denature samples (1, 5, and 10 ML in duplicates) and plasmid copy number controls (107–1011 copies) in 50 ML AAV digestion buffer at 100°C for 10 min. Immediately chill on ice and bring up volume to 400 ML with the digestion buffer. Load onto a Hybond-N plus

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membrane with a Bio-Dot SF manifold microfiltration apparatus. After blotting, crosslink DNA to the membrane with UV irradiation. Prehybridize the membrane in 10 mL slot blot hybridization solution in a Techne roller bottle hybridization oven at 37°C for 2 h. Hybridize the membrane with a 32 P-labeled transgene-specific probe in the slot blot hybridization solution at 37°C for 5 h. Wash the membrane at 37°C in 2× SSC/1% SDS (2 × 15 min). Wash the membrane in 0.5× SSC/1% SDS (2 × 15 min). Wash the membrane in 0.1× SSC for 15 min. Expose the membrane in an X-ray film for 2 h (or overnight). Develop the film and determine the viral genome particle titer by comparing the intensity of the viral sample bands to those of the copy number controls. 2. Determine AAV titer by quantitative PCR. Denature viral samples (2 ML in triplicates) and plasmid copy number controls (106–1011 copies/ML) in 50 ML of AAV PCR alkaline digestion buffer at 100°C for 10 min. Immediately chill on ice and add 50 ML of AAV PCR neutralization buffer. Mix well and use as the PCR template. Prepare the PCR reaction mixture on ice in a 96-well reaction plate. Each reaction mixture (20 ML/well) contains 10 ML of Fast SYBR green PCR master mixture, 0.3 ML of 10 MM forward primer, 0.3 ML of 10 MM reverse primer (see Table 1), 1 ML of the PCR template, and 8.4 ML of PCR quality water. Warm up the ABI 7900HT real-time PCR machine for 5 min. Select absolute quantification for the study type and SYBR for the detector. Designate the sample, standard (copy number control), and control (no template) wells. Turn on the ROX passive reference. Set the thermal cycler condition (see Table 2). Load the 96-well plate. Start the run. Obtain the quantity mean from the on-screen result table and calculate the viral genome copy number titer (see Note 25).

Table 2 Conditions for quantitative PCR Stage 1 (initial denaturation)

Stage 2 (amplification reaction)

Stage 3 (dissociation curve; optional)

Thermal profile

95°C, 20 s

95°C, 5 s l 60°C, 20 s; ×40

95°C, 15 s l 60°C, 15 sl 95°C, 15 s

Autoincrement

+0, +0

+0, +0

+0, +0

Ramp rate

100%

100%

Final 95°C is 2%; the others 100%

Data collection

None

At 60°C step

At slope between 60 and 95°C

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Fig. 3. AAV vector genome titer determination and viral quality examination. (a) A representative slot blot from a first round CsCl ultracentrifugation. Fractions (Fr.) 5–8 usually contain most of the virus. (b) Representative results from quantitative PCR. The top is the quantification curve. The bottom is the amplification curve. Fractions are marked in numbers. (c) Representative electron microscopic images of AAV-2 (left) and AAV-5 (right). Scale bar applies to both images. Arrowhead indicates a partially packaged virus.

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3. Determine AAV quality by electron microscope. Place one drop of purified AAV on a 200 mesh carbon-coated copper grid for 5 min. Gently wash in ultra pure water for four times. Apply one drop of 1% uranyl acetate on the sample. Air dry for 5 min. Visualize viral particles using a JEOL JEM-1400 transmission electron microscope (see Fig. 3). 4. Determine the endotoxin level with the LAL assay. Reconstitute control standard endotoxin (CSE) with LAL reagent water provided with the EndoSafe LAL gel clot test kit. The reconstituted CSE is stable for 4 weeks at 4°C. Place 200 ML sample (or control) to each LAL gel clot reaction tube. Vortex briefly. Incubate at a 37°C water bath for 60 min. A positive result is defined as the formation of a clot that retains its integrity (either at the bottom or slide down to the side of the tube) when the tube is inverted 180°. The formation of a viscous solution which breaks apart and slides down the side of the tube is considered negative (see Note 26).

4. Notes 1. Latent infection of 293 cells by wild type AAV may result in wild type AAV contamination in purified vector stocks (30). We suggest checking wild type AAV contamination periodically in 293 cells (31). Although 293 cells remain the mainstay for laboratory-scale AAV production, different producer cell lines have been developed for industrial-scale manufacturing. These cells contain integrated AAV rep and cap genes. Some also carry the vector genome. AAV production is initiated with a helper virus (such as adenovirus and herpes simplex virus) infection (reviewed in refs. (22–24, 26)). 2. All the plasmids (including adenovirus helper plasmid, AAV helper plasmid, and cis plasmid) are prepared by the triton-lysis/ CsCl-ethidium bromide density gradient centrifugation method. We have found that plasmids prepared this way give the highest AAV yield (32). The vast majority of AAV variants are based on the AAV-2 ITR. Because of the high recombination nature of the ITR, we strongly suggest to propagate the cis plasmid in the SURE cells (Stratagene) or Stbl2 cells (Gibco-Invitrogen). 3. Adenovirus contamination has been a major concern of AAV stock. This hurdle is now overcome with the development of helper virus-free AAV production system. In this system, a helper plasmid is used to express adenoviral virus-associated RNA (VA RNA), E2a, and E4 genes. Since 293 cells express adenoviral E1a and E1b genes, all adenoviral helper function is now reconstituted. This technology has completely eliminated

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the need of adenovirus coinfection in AAV preparation. We have used the Stratagene pHelper plasmid, which expresses adenoviral E2a, E4, and VA RNA (33). Several similar adenoviral helper plasmids have also been published including pXX6 (available at the UNC Vector Core Facility, University of North Carolina at Chapel Hill, NC) and pDG (PlasmidFactory, Heidelberg, Germany) (34–36). Besides adenoviral helper genes, pDG also carries AAV-2 rep and cap genes (35). 4. For most serotypes (or AAV variants), a single AAV helper plasmid is used to express both cap and rep genes. The selection of the cap gene is determined by the intended serotype. However, most AAV helper plasmids express the AAV-2 rep gene (this is because the AAV-2 ITR is often used as the replication/packaging signal). Two exceptions are AAV-5 and 6. In these cases, two AAV helper plasmids are used. For AAV-5, one helper expresses the AAV-2 rep gene and the other expresses the AAV-5 rep and cap genes (37, 38). For AAV-6, one helper expresses the AAV-2 rep gene under the mouse metallothionein (MT) promoter and the other expresses AAV-6 cap gene under the cytomegalovirus (CMV) promoter (39). The AAV-2 helper plasmid (pAAV-RC) can be purchased from Stratagene. The helper plasmids for AAV-1 to 8 can be purchased from PlasmidFactory. We have obtained AAV-5 helper plasmids (pAV5-Trans and pAV2-Rep) from Dr. John F. Engelhardt (University of Iowa, Iowa City, IA). We have obtained AAV-6 helper plasmids (pMT-Rep2 and pCMV-Cap6) from Dr. A. Dusty Miller (Fred Hutchinson Cancer Research Institute, Seattle, WA). We have obtained AAV1, 7, 8, and 9 helper plasmids from Dr. James M. Wilson (University of Pennsylvania, Philadelphia, PA). 5. Since pH affects transduction efficiency, it is highly suggested to double check pH of 2× HBS buffer before each usage. After thawing the buffer, make sure to mix well by inverting the tube several times. 6. DNase I (from the bovine pancrease) is used to release AAV particles from the nucleus. Reconstitute DNase I by adding 4 mL of double distilled water to one vial of lyophilized powder. Mix well before use. Besides DNase I, one can also use Benzonase (25,000 Unit). A recent study suggests that for AAV serotype-1, 8, and 9, mature viral particles are also released to the culture medium. For these serotypes, high quantity of biologically active AAV virions can be directly harvested from the culture medium (40). 7. We have also used the Beckman L-60, Beckman L8-70, and Sorvall Discovery SE100 ultracentrifuges and the Beckman SW50.1 and Sorvall AH650 rotors. It is important to match the centrifuge speed with the rotor.

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8. Others have used the Beckman ultraclear ultracentrifuge tube. This type of tube may lead to better visualization of the virus band (e.g., during adenovirus preparation). However, we find the ultraclear ultracentrifuge tube is difficult to penetrate with a needle. Since the AAV band is often not detectable by eyeballing, we recommend use of the polyallomer tube. 9. Besides Pierce’s Slide-A-Lyze dialysis cassette, we have also obtained excellent results with self-prepared 12–14,000 MWCO dialysis tubing. Briefly, boil dialysis tubing (6.4 mm) in a solution containing 238 mM NaHCO3 and 1 MM EDTA for 1 h. Wash extensively with tap water until pH of the washout reaches that of tap water. Store the dialysis tubing at 4°C in a dark bottle containing 0.04% sodium azide. 10. We use 107–1011 copies and 106–1011 copy/ML of the vector genome in slot blot and quantitative PCR, respectively. The copy number control is made with the cis plasmid according to the length (in bp) and the concentration (in ng/ML) of the plasmid. The formula for calculating single-stranded AAV copy number is: [Plasmid concentration × 1.2 × 1015]/[(plasmid DNA length × 607.4) + 157.9]. Store the copy number control in −20°C in 50 ML aliquots. Avoid repeated freeze/thaw. 11. Uranyl acetate is highly toxic and radioactive. Handle with care. Dilute the stock with double-distilled water to 1% working solution and filter through a 0.45 Mm filter. Store in dark at 4°C. 12. It is critical to double check cell confluency prior to plasmid transfection (see Fig. 1b). We perform transfection at 70% confluency. Differences in cell confluency (e.g., 60 or 80%) may result in suboptimal transfection and low AAV yield. 13. The type/number of the plasmids may vary. For AAV-1, 2, 7, 8 and 9, we use triple plasmid transfection (the cis plasmid at 12.5 Mg/150 mm plate, the pHelper at 37.5 Mg/150 mm plate, and a rep-cap containing AAV helper plasmid at 37.5 Mg/150 mm plate). Four plasmids are used in AAV-5 and 6 preparations. For AAV-5, it includes the cis plasmid at 12.5 Mg/150 mm plate, the pHelper at 37.5 Mg/150 mm plate, an AAV-5 rep-cap plasmid at 37.5 Mg/150 mm plate and an AAV-2 rep plasmid at 12.5 Mg/150 mm plate. For AAV-6, it includes the cis plasmid at 12.5 Mg/150 mm plate, the pHelper at 37.5 Mg/150 mm plate, the pMT-Rep2 at 12.5 Mg/150 mm plate and the pCMACap6 at 37.5 Mg/150 mm plate. If adenoviral helper function and AAV helper function are combined in one plasmid (such as in pDG), only two plasmids will be needed (the pDG and a cis plasmid) for transfection (35). We have consistently observed similar transfection efficiency using either three or four plasmids.

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14. Here, we described the calcium phosphate coprecipitation method. Under optimal condition, transfection efficiency reaches 90% (see Fig. 1c). However, depending on the conditions used (such as the pH of the solution, the size of the precipitates), transfection efficiency can be dramatically reduced (see Fig. 1d). We recommend routinely monitoring calcium phosphate precipitate on a coverslip using a phase contrast microscope. Under optimal condition, one should see uniform fine precipitates. Large aggregates often lead to poor transfection and low AAV yield. We also recommend monitoring the quality of the transfection reagents (such as CaCl2 and 2× HBS) with a pilot test using a reporter gene plasmid. Alternatively, a separate 35 mm plate of 293 cells (from the same split of 150 mm plates) should be transfected with the same transfection cocktail and examined for transfection efficiency (e.g., by histochemical staining for the LacZ and AP genes, or by immunofluorescence staining for a particular target gene). Some investigators have also spiked one to 20–100th of a GFP plasmid that does not contain AAV ITR as internal control to monitor transfection efficiency. The protocol described here has been optimized for 15 × 150 mm plate transfection. Besides calcium phosphate coprecipitation method, others have also used polyethylenimine coprecipitation and cationic lipid transfection methods (reviewed in ref. (27)). 15. From 15 × 150 mm plates, we usually get ~350 mL crude lysate. Cells usually pellet at the bottom of the 250 mL Corning centrifuge bottle. We rinse off the remaining cells from the plates with the supernatant from the centrifuge bottle. 16. Clean the sonicator probe with 70% ethanol followed by rinse with 10 mM Tris–HCl (pH 8.0) before each use. Make sure to submerge the probe into the crude lysate so that the tip of the probe is ~2.5 cm above the bottom of the 50 mL tube. 17. Final volume will be approximately 27.6 mL prior to the addition of CsCl. 18. The buoyant density of AAV is between 1.32 and 1.47 g/mL in CsCl gradient (41). The lighter particles (<1.39 g/mL) represent empty or partially packaged virions (42). The fully packaged AAV particles have a density of 1.39–1.42 g/mL. The most infectious AAV particles are found at the density of 1.41 g/mL. Heavy particles (>1.42 g/mL) seem to associate with an unknown high molecular weight protein and their infectivity is significantly reduced (43, 44). Adding 16.9 g CsCl to 27.6 mL crude lysate (0.613 g/mL) results in a final CsCl contraction of 1.41 g/mL prior to ultracentrifugation. 19. After centrifugation, cell debris forms a thin lipid-like layer at the top (see Fig. 2a). If not proceed to ultracentrifugation right away, the clear lysate can be stored at 4°C for up to 1 week.

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20. For Beckman SW50.1 and Sorval AH650 rotors, we suggest spinning at 198,000 × g (46,000 rpm) for 40 h. 21. We usually insert the self-made needle stylet at the level of the horizontal line on the Beckman ultracentrifuge tube (see Fig. 2c). To get consistent results, it is important to always enter at the same level. Position the needle tip opening to face up. Remove the 25G needle. Discard the first 14 drops and then collect 12 drops (~750 ML) /fraction. From each 5.1 mL Beckman ultracentrifuge tube, we usually get 12 fractions. AAV often appears in fractions 5–8. 22. For the first and the second rounds of ultracentrifugation, it is not necessary to include the copy number controls in slot blot or quantitative PCR. 23. Prior to the third round of ultracentrifugation, we usually add 200 mg CsCl to 5 mL of combined viral fractions collected from the second round of ultracentrifugation. Addition of CsCl is necessary to maintain the isopycnic gradient during the third round of ultracentrifugation. From 15 to 150 mm plates, we get ~30 mL lysate for the first round of ultracentrifugation. From the first round of ultracentrifugation, we collect ~15 mL of AAV containing fraction (enough to fill three 5.1 mL Beckman ultracentrifuge tubes). After the second round of ultracentrifugation, we usually get ~3 mL of AAV containing fraction. To have enough volume for the third round of ultracentrifugation, we usually start with 150 × 150 mm plates AAV preparation. This will generate ~30 mL of AAV containing fraction after two rounds of ultracentrifugation. However, if the preparation started with 15 × 150 mm plates, the volume from the second round of ultracentrifugation will be less than 5 mL and not enough to fill up a single 5 mL ultracentrifuge tube. In this case, we suggest using CsCl/10 mM Tris–HCl (pH 8.0) (0.613 g/mL) to bring up the final volume to 5 mL. 24. Double check to make sure the dialysis tubing is not leaking. Remember to place a magnetic stir bar to gently agitate the dialysis buffer. We usually get a yield of 5 × 1012 to 1 × 1013 viral genome particles/mL of AAV vectors. 25. If PCR templates are not used immediately, they can be stored in 20 ML aliquots at −20°C. Each aliquot is only good for one use. Make sure to tightly seal the wells with an adhesive film applicator and do not taint the surface of the adhesive film. After running the PCR reaction, check the standard curve (the R2 should be higher than 0.95) and the dissociation curve (all reactions should point to a single peak). Ideally, the standard deviation of the viral samples should be less than one tenth of the titer. 26. It is critical to use depyronated pipette tips and dilution tubes provided with the EndoSafe LAL gel clot test kit. Prior to the use of each new lot of the kit, validate the endpoint control to

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confirm the listed sensitivity. For each new type of biological sample, perform a positive product inhibition control to determine whether there are proteins/chemicals in the sample that may inhibit the gel clot reaction. Always include a positive and a negative control. The LAL assay requires 200 ML sample volume for each sample. To avoid wasting AAV virus, viral stock can be diluted with the LAL reagent water prior to the assay. After removing the tube from the 37°C water bath, examine the clot formation immediately (within 2 min). Waiting too long will lead to false readings.

Acknowledgments The protocols were developed with the grant support from the National Institutes of Health (AR-49419 and HL-91883 to DD), the Muscular Dystrophy Association (DD), and the Parent Project for Muscular Dystrophy. We thank Duan lab members for helpful discussion. References 1. Atchison, R. W., Casto, B. C., and Hammon, W. M. (1965) Adenovirus-Associated Defective Virus Particles. Science 149, 754–6. 2. Flotte, T. R., and Berns, K. I. (2005) Adenoassociated virus: a ubiquitous commensal of mammals. Hum Gene Ther 16, 401–7. 3. Schnepp, B. C., Jensen, R. L., Chen, C. L., Johnson, P. R., and Clark, K. R. (2005) Characterization of adeno-associated virus genomes isolated from human tissues. J Virol 79, 14793–803. 4. Duan, D., Sharma, P., Yang, J., Yue, Y., Dudus, L., Zhang, Y., Fisher, K. J., and Engelhardt, J. F. (1998) Circular Intermediates of Recombinant Adeno–Associated Virus have Defined Structural Characteristics Responsible for Long Term Episomal Persistence in Muscle. J Virol 72, 8568–77. 5. Huser, D., Gogol-Doring, A., Lutter, T., Weger, S., Winter, K., Hammer, E. M., Cathomen, T., Reinert, K., and Heilbronn, R. (2010) Integration preferences of wildtype AAV-2 for consensus rep-binding sites at numerous loci in the human genome. PLoS Pathog 6, e1000985. 6. Senapathy, P., and Carter, B. J. (1984) Molecular cloning of adeno-associated virus variant genomes and generation of infectious virus by recombination in mammalian cells. J Biol Chem 259, 4661–6.

7. Samulski, R. J., Srivastava, A., Berns, K. I., and Muzyczka, N. (1983) Rescue of adeno-associated virus from recombinant plasmids: gene correction within the terminal repeats of AAV. Cell 33, 135–43. 8. Carter, B. J. (2004) Adeno-associated virus and the development of adeno-associated virus vectors: a historical perspective. Mol Ther 10, 981–9. 9. Mendell, J. R., Rodino-Klapac, L. R., RosalesQuintero, X., Kota, J., Coley, B. D., Galloway, G., Craenen, J. M., Lewis, S., Malik, V., Shilling, C., Byrne, B. J., Conlon, T., Campbell, K. J., Bremer, W. G., Viollet, L., Walker, C. M., Sahenk, Z., and Clark, K. R. (2009) Limbgirdle muscular dystrophy type 2D gene therapy restores alpha-sarcoglycan and associated proteins. Ann Neurol 66, 290–7. 10. Maguire, A. M., Simonelli, F., Pierce, E. A., Pugh, E. N., Jr., Mingozzi, F., Bennicelli, J., Banfi, S., Marshall, K. A., Testa, F., Surace, E. M., Rossi, S., Lyubarsky, A., Arruda, V. R., Konkle, B., Stone, E., Sun, J., Jacobs, J., Dell’Osso, L., Hertle, R., Ma, J. X., Redmond, T. M., Zhu, X., Hauck, B., Zelenaia, O., Shindler, K. S., Maguire, M. G., Wright, J. F., Volpe, N. J., McDonnell, J. W., Auricchio, A., High, K. A., and Bennett, J. (2008) Safety and efficacy of gene transfer for Leber’s congenital amaurosis. N Engl J Med 358, 2240–8.

15 11. Cideciyan, A. V., Hauswirth, W. W., Aleman, T. S., Kaushal, S., Schwartz, S. B., Boye, S. L., Windsor, E. A., Conlon, T. J., Sumaroka, A., Roman, A. J., Byrne, B. J., and Jacobson, S. G. (2009) Vision 1 year after gene therapy for Leber’s congenital amaurosis. N Engl J Med 361, 725–7. 12. Bainbridge, J. W., Smith, A. J., Barker, S. S., Robbie, S., Henderson, R., Balaggan, K., Viswanathan, A., Holder, G. E., Stockman, A., Tyler, N., Petersen-Jones, S., Bhattacharya, S. S., Thrasher, A. J., Fitzke, F. W., Carter, B. J., Rubin, G. S., Moore, A. T., and Ali, R. R. (2008) Effect of gene therapy on visual function in Leber’s congenital amaurosis. N Engl J Med 358, 2231–9. 13. McCarty, D. M. (2008) Self-complementary AAV vectors; advances and applications. Mol Ther 16, 1648–56. 14. Zhou, X., Zeng, X., Fan, Z., Li, C., McCown, T., Samulski, R. J., and Xiao, X. (2008) Adenoassociated virus of a single-polarity DNA genome is capable of transduction in vivo. Mol Ther 16, 494–9. 15. Zhong, L., Zhou, X., Li, Y., Qing, K., Xiao, X., Samulski, R. J., and Srivastava, A. (2008) Single-polarity Recombinant Adeno-associated Virus 2 Vector-mediated Transgene Expression In Vitro and In Vivo: Mechanism of Transduction. Mol Ther 16, 290–95. 16. Ghosh, A., Yue, Y., Lai, Y., and Duan, D. (2008) A hybrid vector system expands adenassociated viral vector packaging capacity in a transgene independent manner. Mol Ther 16, 124–30. 17. Kwon, I., and Schaffer, D. V. (2008) Designer gene delivery vectors: molecular engineering and evolution of adeno-associated viral vectors for enhanced gene transfer. Pharm Res 25, 489–99. 18. Vandenberghe, L. H., Wilson, J. M., and Gao, G. (2009) Tailoring the AAV vector capsid for gene therapy. Gene Ther 16, 311–9. 19. Wu, Z., Asokan, A., and Samulski, R. J. (2006) Adeno-associated Virus Serotypes: Vector Toolkit for Human Gene Therapy. Mol Ther 14, 316–27. 20. Gao, G., Vandenberghe, L. H., and Wilson, J. M. (2005) New recombinant serotypes of AAV vectors. Curr Gene Ther 5, 285–97. 21. Virag, T., Cecchini, S., and Kotin, R. M. (2009) Producing recombinant adeno-associated virus in foster cells: overcoming production limitations using a baculovirus-insect cell expression strategy. Hum Gene Ther 20, 807–17.

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22. Clement, N., Knop, D. R., and Byrne, B. J. (2009) Large-scale adeno-associated viral vector production using a herpesvirus-based system enables manufacturing for clinical studies. Hum Gene Ther 20, 796–806. 23. Zhang, H., Xie, J., Xie, Q., Wilson, J. M., and Gao, G. (2009) Adenovirus-adeno-associated virus hybrid for large-scale recombinant adenoassociated virus production. Hum Gene Ther 20, 922–9. 24. Zolotukhin, S. (2005) Production of recombinant adeno-associated virus vectors. Hum Gene Ther 16, 551–7. 25. Cecchini, S., Negrete, A., and Kotin, R. M. (2008) Toward exascale production of recombinant adeno-associated virus for gene transfer applications. Gene Ther 15, 823–30. 26. Thorne, B. A., Takeya, R. K., and Peluso, R. W. (2009) Manufacturing recombinant adenoassociated viral vectors from producer cell clones. Hum Gene Ther 20, 707–14. 27. Wright, J. F. (2009) Transient transfection methods for clinical adeno-associated viral vector production. Hum Gene Ther 20, 698–706. 28. Graham, F. L., Smiley, J., Russell, W. C., and Nairn, R. (1977) Characteristics of a human cell line transformed by DNA from human adenovirus type 5. J Gen Virol 36, 59–74. 29. Louis, N., Evelegh, C., and Graham, F. L. (1997) Cloning and sequencing of the cellularviral junctions from the human adenovirus type 5 transformed 293 cell line. Virology 233, 423–9. 30. Duan, D., Fisher, K. J., Burda, J. F., and Engelhardt, J. F. (1997) Structural and functional heterogeneity of integrated recombinant AAV genomes. Virus Res 48, 41–56. 31. Katano, H., Afione, S., Schmidt, M., and Chiorini, J. A. (2004) Identification of adenoassociated virus contamination in cell and virus stocks by PCR. Biotechniques 36, 676–80. 32. Heilig, J. S., Elbing, K. L., and Brent, R. (2001) Large-scale preparation of plasmid DNA. Curr Protoc Mol Biol Chapter 1, Unit1 7. 33. Matsushita, T., Elliger, S., Elliger, C., Podsakoff, G., Villarreal, L., Kurtzman, G. J., Iwaki, Y., and Colosi, P. (1998) Adenoassociated virus vectors can be efficiently produced without helper virus. Gene Ther 5, 938–45. 34. Grimm, D., Kay, M. A., and Kleinschmidt, J. A. (2003) Helper virus-free, optically controllable, and two-plasmid-based production of adeno-associated virus vectors of serotypes 1 to 6. Mol Ther 7, 839–50.

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35. Grimm, D., Kern, A., Rittner, K., and Kleinschmidt, J. A. (1998) Novel tools for production and purification of recombinant adenoassociated virus vectors. Hum Gene Ther 9, 2745–60. 36. Xiao, X., Li, J., and Samulski, R. J. (1998) Production of high-titer recombinant adenoassociated virus vectors in the absence of helper adenovirus. J Virol 72, 2224–32. 37. Duan, D., Yan, Z., Yue, Y., Ding, W., and Engelhardt, J. F. (2001) Enhancement Of Muscle Gene Delivery With Pseudotyped AAV-5 Correlates With Myoblast Differentiation. J Virol 75, 7662–71. 38. Yan, Z., Zak, R., Luxton, G. W., Ritchie, T. C., Bantel-Schaal, U., and Engelhardt, J. F. (2002) Ubiquitination of both adeno-associated virus type 2 and 5 capsid proteins affects the transduction efficiency of recombinant vectors. J Virol 76, 2043–53. 39. Allen, J. M., Halbert, C. L., and Miller, A. D. (2000) Improved adeno-associated virus

vector production with transfection of a single helper adenovirus gene, E4orf6. Mol Ther 1, 88–95. 40. Vandenberghe, L. H., Xiao, R., Lock, M., Lin, J., Korn, M., and Wilson, J. M. (2010) Efficient serotype-dependent release of functional vector into the culture medium during AAV manufacturing. Hum Gene Ther 21, 1251–7. 41. de la Maza, L. M., and Carter, B. J. (1980) Molecular structure of adeno-associated virus variant DNA. J Biol Chem 255, 3194–203. 42. Torikai, K., Ito, M., Jordan, L. E., and Mayor, H. D. (1970) Properties of light particles produced during growth of Type 4 adeno-associated satellite virus. J Virol 6, 363–9. 43. Lipps, B. V., and Mayor, H. D. (1982) Characterization of heavy particles of adenoassociated virus type 1. J Gen Virol 58 Pt 1, 63–72. 44. de la Maza, L. M., and Carter, B. J. (1980) Heavy and light particles of adeno-associated virus. J Virol 33, 1129–37.

Chapter 16 Generation of Lentiviral Vectors for Use in Skeletal Muscle Research Christophe Pichavant and Jacques P. Tremblay Abstract Gene therapy is a promising approach for the treatment of a variety of disorders including genetic diseases and cancer. Among the viral vectors used in gene therapy, the lentiviral vector, based on HIV-1, is the only integrative vector able to transduce nondividing cells. The first generation of lentiviral vector was established in 1996. Since then, other generations of lentiviral vector packaging systems were developed to improve this first vector. In this chapter, we describe these different packaging systems, the generation of lentiviral vector from productive cells, the 293T cell line, and the transduction of myogenic cells with a lentiviral vector as well. Key words: Lentiviral vector, Virus production, Transfection, Transduction, Myoblast

1. Introduction The interest and the use of lentiviral vectors started in 1996 when Naldini et al. showed that vectors based on human immunodeficiency virus type-1 (HIV-1) were able to transduce quiescent cells (1). Lentivirus, similarly to retrovirus, is part of the Retroviridae family. Its viral capsid is enveloped and its diameter varies between 80 and 100 nm. It contains two copies of positive-sense single-stranded RNA as well as integrase (IN), protease (PRO), and reverse transcriptase (RT) necessary for its transduction. Lentivirus integration is random but tends to occur in chromosomal regions that are more accessible (2) and more likely in transcriptional active sites (3). The HIV-1 genome is composed of three structural genes: Gag, Pol, and Env. Adding to these structural genes, there are 2 regulatory genes: Tat and Rev, and 4 accessory genes: Vif, Vpr, Vpu, and Nef (4). There are also specific sequences that are present in the lentiviral genome such as long terminal repeats (LTRs), Joseph X. DiMario (ed.), Myogenesis: Methods and Protocols, Methods in Molecular Biology, vol. 798, DOI 10.1007/978-1-61779-343-1_16, © Springer Science+Business Media, LLC 2012

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primer binding site (PBS), \, REV response element (RRE), and central PolyPurine Track (cPPT). Regarding the structural genes, the GAG product is cleaved into different protein subunits: the MA (matrix), which is essential for virion assembly and for quiescent cell transduction; the CA (capsid), which is also important for the assembly and maturation of the virion; the NC (nucleocapsid), which provides a strong support for viral RNA within the viral particle; and several polypeptides of small size such as p1, p2, or p6, which roles are still not completely understood. Pol encodes for three enzymes involved in viral replication: IN, PRO, and RT. Env allows the expression of the glycoprotein gp120 and gp41 at the virion surface creating the viral envelope. Gp120 is required to bind to cellular receptors on the host cell, and gp41 is involved in the lentivirus fusion with the cellular membrane (5). Concerning the regulatory genes, Rev encodes for the REV protein implicated in viral RNA exportation. On the one hand, this protein is bound to the viral genome via the RRE. On the other hand, it interacts with the nucleoporins of the nucleus. Linked in such manner to the viral mRNA, REV allows its exportation from the nucleus to the cytoplasm (6). The other regulatory gene Tat encodes for a protein, which is characterized as a strong transcription activator. Indeed, when TAT is bound to the trans-activating response (TAR) present in the 5c of the viral genome, high levels of mRNA transcription are found (7). Vif, Vpr, Vpu, and Nef are genes involved in HIV-1 virulence and are named “accessory” since they are not essential for viral replication in cell culture (8). Sequences, other than the RRE described earlier, are important as well for the HIV-1 (9). LTRs are sequences localized at each extremity of a viral genome transcription. These sequences have the role of a promoter in 5c and of a polyadenylation signal in 3c. Even though this promoter is weak, its transcriptional activity is enhanced by the presence of TAT (7). The LTR in 5c is followed by a PBS needed for transcription initiation and by a \ sequence required for viral RNA encapsidation. Another important sequence is the cPPT. This sequence gives the viral DNA a specific conformation allowing its access to the nucleus even when the host cell is not in a mitosis phase (10). To generate lentivirus particles, a three-plasmid expression system was developed by transient transfection in 1996 (1). The first plasmid carried viral genes except for Env and Vpu (the packaging plasmid). The second one coded for an envelope, and the last one contained the gene of interest to transfer. This first generation of lentiviral vector packaging system (LVPS) was able to transduce cells, but the produced virions contained VIF, VPR, and NEF proteins, which are implicated in HIV-1 toxicity. Thus, a second generation was created by removing these genes from the packaging plasmid (11). To reduce the number of viral genes even more, a third generation expression system was designed (12). This

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Fig. 1. Schematic representations of the three generations of lentiviral vector packaging systems. (a–c) Represent respectively the first, the second, and the third generation of LVPS. The first and the second generations are based on a threeplasmid expression system whereas the third generation is based on a four-plasmid expression system.

generation is based on a four-plasmid expression system. The first plasmid codes for the envelope, the second for GAG/POL, the third for REV and the last one contains the gene of interest. These three generations are schematized in Fig. 1. To enhance the biosafety profile, the LTR sequences can be also mutated; this modification was named Self-INactivating (SIN) (13). The maximum size of the transgene placed between the two LTRs with its own promoter is about 8–9 kb. To increase the transgene expression, a Woodchuck hepatitis virus Posttranscriptional Regulatory Element (WPRE) sequence can be placed at the 3c region of the transgene (14). As mentioned earlier, lentiviral vectors are obtained by transient transfection of different plasmids. The first one contains the transgene to be expressed. Another one carries the envelope gene. Since the most used envelope gene is vesicular stomatitis virus-G glycoprotein (VSV-G) due to its broad tropism (15), VSV-G will be used as the reference envelope protein throughout this chapter. Afterwards, depending on the generation of LVPS, different packaging plasmids have been used. To produce a lentiviral vector of second generation, a third plasmid containing Gag/Pol, Rev, and Tat is added, whereas for a lentiviral vector of third generation, two more plasmids are necessary. The first one codes for GAG/POL and the second for REV. In addition, the plasmid containing the gene of interest can have the SIN modification. The SIN-vector plasmid can be packaged into both, second and third generation of LVPS. These three or four plasmids are transfected in HEK 293T cells (human embryonic immortalized renal cells) since they can be easily transfected. Pol allows the RT expression, and therefore the viral RNA can be retrieved. The capsid and envelope proteins are provided through Gag and VSV-G. Rev exports quickly the viral mRNA toward the cytoplasm through the RRE sequence where it is encapsided by the \ sequence. It must be mentioned that RT, PRO, and IN are also present in the viral capsid. When the assembly of viral particles is completed, they bud from the cell membranes and are released in the supernatant. The obtained titers are about 106–107 viral particles per mL and can be increased after ultrafiltration or ultracentrifugation. The produced lentivirus contains the necessary sequences for its transgene integration in the host genome while still being nonreplicative.

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The quality of a lentiviral vector depends on different factors. The first one is the design of the lentiviral plasmid backbone. It is important to select a strong and appropriate promoter to drive transgene expression. An antibiotic resistance gene can also be incorporated in the backbone. When the lentiviral plasmid backbone is obtained, the next step is to produce the virus. The lentivirus production protocol has to be optimized to obtain an efficient virus as well as the lentivirus transduction protocol to achieve a high level of transduction. These different factors are discussed later.

2. Materials Prepare all solutions using distilled water. Culture media and reagents must be sterile and stored at 4°C. Plasmids are stored at −20°C as well as the polybrene solution. All cells are maintained at 37°C under 5% CO2. 2.1. Cell Culture

1. HEK 293T cells (see Note 1). 2. Medium A: Dulbecco’s modified eagle medium – high glucose (DMEM-HG) supplemented with 10% heat-inactivated fetal bovine serum (FBS) and 1% penicillin–streptomycin (PS). 3. Myoblasts. 4. Medium B: MB-1 medium (Hyclone) supplemented with 15% heat-inactivated FBS, 1% PS, and 10 Pg/L of basic fibroblast growth factor (Feldan). 5. Medium C: Medium B with 2% heat-inactivated FBS instead of 15%.

2.2. Lentivirus Production

1. 100 mm petri dishes (Sarstedt). 2. Medium D: Medium A supplemented with 0.25% of 1 M 4-(2-hydroxyethyl)-1-piperazineethanesulfonic acid (HEPES). 3. Distilled water buffered with 2.5 mM HEPES. 4. 2 M CaCl2. 5. 2× HEPES-buffered saline (HeBS): 50 mM HEPES, 280 mM NaCl, 1.5 mM Na2HPO4, pH 7.20. 6. Mechanical pipettor and Pasteur pipettes. 7. 15- and 1.5-mL conical tubes. 8. Plasmids coding for your transgene, VSV-G and Gag/ Pol-Rev-Tat (when using second generation of LVPS). 9. Plasmids coding for your transgene, VSV-G, Gag/Pol, and Rev (when using third generation of LVPS). 10. 0.22-Pm Syringe filters.

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1. Six-well plates. 2. FBS. 3. Medium C for myoblast transduction. 4. Medium D for myoblast differentiation. 5. 8 Pg/mL polybrene. 6. Hank’s buffered salt solution (HBSS).

3. Methods Carry out procedures at room temperature in a sterile environment. 3.1. Production of Lentivirus by Calcium Chloride Transfection (see Note 2)

1. Day 0: late in the afternoon, plate 293T cells (see Note 3) at the concentration of 2.5 × 106 cells per 100 mm petri dish (see Note 4) with 8 mL medium A (see Note 5). 2. Day 1: early in the morning, aspirate the medium and add 8 mL warm medium D (see Notes 5 and 6). 3. Day 1: late in the afternoon, prepare as many 1.5 and 15 mL conical tubes as dishes to transfect (see Note 7). 4. Depending on which generation of LVPS is produced, the plasmids chosen to transfect are different (see Notes 8 and 9) (Fig. 1). 5. If producing a lentivirus from the second generation of LVPS, place in each 1.5-mL tube: (a) 15 Pg of the plasmid containing your transgene of interest (see Note 10). (b) 15 Pg of the Gag/Pol-Rev-Tat plasmid (i.e., psPAX2, pCMV-śR8.91). (c) 5 Pg of the VSV-G plasmid (i.e., pMD2.G, pCMV-VSV-G). 6. If producing a lentivirus from the third generation of LVPS, place in each 1.5-mL tube: (a) 15 Pg of the plasmid containing your transgene of interest (see Note 10). (b) 2.5 Pg of the Rev plasmid. (c) 6.5 Pg of the Gag/Pol plasmid. (d) 2.5 Pg of the VSV-G plasmid. 7. Add 62.5 PL of 2 M CaCl2 to each tube (see Note 11). 8. Fill up to a total volume of 500 PL with distilled water buffered with 2.5 mM HEPES (see Note 12). 9. Mix gently by pipetting. 10. Place in each 15-mL conical tube 500 PL of 2× HeBS, pH 7.20 (see Note 13).

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11. Bubble the 2× HeBS using a mechanical pipettor attached to a filtered Pasteur pipette. 12. While bubbling the HeBS, add the volume (500 PL) of the 1.5 mL tube dropwise with a 1-mL tip. 13. Mix by pipetting. 14. Wait 20 min at room temperature to allow precipitate formation. 15. Add dropwise the precipitated DNA (1 mL) over the cells in the culture dish and rotate gently the dish to mix the precipitate and the medium. 16. Day 2: early in the morning, aspirate the medium and add 6 mL warm medium A (see Notes 5 and 6). 17. Day 2: late in the afternoon, collect the supernatant and add 6 mL warm medium A (see Notes 5 and 6). 18. Centrifuge 5 min at 500 × g to remove cell debris and filtrate at 0.22 Pm. 19. Pool supernatants and this first harvest of virus is named Passage 1. 20. Supernatants containing the lentivirus can be used directly, stored at 4°C for 2–3 days, at −80°C or concentrated if needed (see Note 14). 21. Day 3: in the morning, collect the supernatant and add 6 mL warm medium A (see Notes 5, 6, and 15). 22. Treat supernatants in the same way as Day 2. This second harvest of virus is named Passage 2. 23. Day 4: in the morning, collect the supernatant and treat them in the same way as Day 2, this last harvest of virus is named Passage 3. 3.2. Myoblast Transduction

1. Day 0: plate 1.5 × 105 myoblasts in a “6 well plate” with 2 mL warm medium B (see Note 16). 2. Day 1: Aspirate the medium and add: (a) x mL of virus (see Notes 17 and 18). (b) 5% FBS. (c) 24 Pg of polybrene (3 PL of the solution at 8 Pg/mL). (d) Fill up to a total volume of 3 mL with warm medium B. 3. Wait 15 min at 37°C. 4. Centrifuge 30 min at 750 × g (see Note 19). 5. Replace cells in incubator. 6. Day 2: aspirate the medium, wash gently with HBSS, and add 2 mL warm medium B. 7. Day 3: transduced cells are ready to be used in your experiment (see Notes 20–23).

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4. Notes 1. This cell line is known to not attach well to the plate. If some adhesion problems are observed, changing simply the manufacturer of the plates/dishes can sometimes resolve these problems. 2. Lipofectamine (Invitrogen) can also be used to transfect 293T cells instead of CaCl2 but their transfection efficiencies are identical in these cells. Therefore, the CaCl2 transfection was preferred since its price is lower than Lipofectamine. 3. 293T cells are best transfected when passaged often. 293FT cells can also be used to produce lentivirus but in our experiments, no significant difference in the lentiviral vector production was noticed. Thus, we recommend to use the 293T cells since it is easier to work with this cell line than that of 293FT cells. 4. In plating 4 × 106 cells instead of 2.5 × 106, no difference has been noticed in the virus production. Thus, the protocol using the least amount of cells (2.5 × 106) was kept. 5. Add 1% L-glutamine (Gibco) to the medium if it was prepared more than a week ago. 6. As already mentioned, 293T cells come off the plate easily. Therefore, medium should be changed gently and carefully. Even if you are manipulating carefully, is it possible that cells begin to come off the plate on the side. If the cell detachment is less than 10% of the total cells, it will not influence the lentivirus production. Nevertheless, you must be careful since cells come off the plate more often with the higher number of medium replacement. 7. When you are going to produce your lentivirus coding for your transgene of interest, we suggest producing simultaneously a lentivirus coding for a reporter gene (such as GFP under a ubiquitous promoter) so that you can monitor the effectiveness of the lentivirus batch. The transfection of a GFP-lentiviral plasmid backbone can be considered as a positive control since you will be able to check under fluorescence light 1 or 2 days after your transfection. Normally, at least 95% of GFPtransfected 293T cells express this transgene when using our protocol as shown in Fig. 2. GFP is also a good marker to follow the transduction efficiency. 8. Non-SIN lentiviruses have to be produced with the second generation of LVPS since Tat is not present in the third generation.

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Fig. 2. GFP expression in GFP-transfected 293T cells. This figure shows 293T cells transfected with a lentiviral plasmid backbone coding for GFP. Two days after the calcium chloride transfection of this plasmid, at least 95% of transfected cells expressed GFP.

9. The lentivirus productions coming from the second and the third generation of LVPS were compared. Using a SINlentiviral plasmid backbone coding for the GFP, we obtained 20% more of GFP positive cells after transducing myoblasts with lentiviruses produced with the three-plasmid expression system than with the four-plasmid system. 10. In the second and third generation of LVPS, 15 Pg of the plasmid containing the transgene of interest is used. In some specific cases, we have observed better transduction efficiency when the plasmid quantity was reduced during the transfection (i.e., lentiviral vector coding for H2Dd (6 Pg)). In other cases, similar results are found for a reduced quantity of plasmid (i.e., lentiviral vector coding for GFP (7.5–15 Pg)) or for an increased quantity (i.e., lentiviral vector coding for GFP (7.5–20 Pg) or for micro-dystrophin (15–20 Pg)). Generally, a larger plasmid will require a higher quantity of plasmid. Nevertheless, the best results are usually obtained in using 15 Pg of the plasmid containing your transgene of interest. 11. It is important to add the CaCl2 solution at the end to obtain a better DNA precipitate. Other concentrations of CaCl2 can also be used but it is important to have a final concentration of 250 mM (in 500 PL). 12. In some lentivirus production protocols, 1× TE is added before mixing the DNA precipitate with HeBS. Therefore, we also tried this method by replacing 250 PL of distilled water buffered with 2.5 mM HEPES (used in our current protocol) with 250 PL 1× TE. Thus, lentivirus productions (coding for GFP) coming from protocols with or without 1× TE were compared.

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No difference in the percentage of transduced myoblasts was observed between these two lentivirus protocols. Thus, we decided not to use 1× TE in our protocol. 13. The pH of the HeBS solution is critical to obtain good cell transfection. Indeed, we have tested different pH for HeBS solutions: 7.05, 7.10, 7.15, 7.20, and 7.25 to produce lentiviruses coding for GFP. Only a few myoblasts transduced by the lentiviruses produced with the HeBS solutions at pH 7.05 and 7.10 were able to express GFP whereas those transduced by the lentiviruses produced with the other pH solutions were all positive for this transgene. Although the pH 7.15, 7.20, and 7.25 provided good results, we suggest the use of HeBS solution pH 7.20 since it is the most used pH. 14. Since each lentivirus production is different, we suggest to store and test separately a small aliquot (1 mL is enough) for each lentivirus produced (if your experimental conditions allow it). To store lentivirus, freeze directly the aliquots in liquid nitrogen and stored them at −80°C. 15. If you produced a GFP-lentivirus containing a gene under a strong ubiquitous promoter such as CMV, you can verify that almost all the cells transfected with this vector are positive for that gene. If this is not the case, there is a problem with your transfection. This can be due to the following: –

The design or the production of the lentiviral plasmid backbone.

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The transfection protocol: concentration or pH of the solutions used (CaCl2, HeBS, and HEPES).

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293T cells: verify their proliferation capacity and their shape.

16. This medium is adequate for the proliferation of human, monkey, and dog myoblasts. For the proliferation of mouse myoblasts, use the medium A. 17. Use a small quantity of your virus (i.e., 1 mL or less) to test your lentivirus. We recommend to test each passage (1–3) of your lentivirus production. We have generally noticed that the best production was Passage 2. We suggest to transduce simultaneously the same cell line with a lentivirus containing the transgene of interest and with a lentivirus coding for a reporter gene, such as GFP, or with a previously tested lentivirus. Cells transduced with these viruses can be used as a positive control of the transduction. 18. There is no direct correlation between the quantity of virus used and the percentage of cells being transduced. Indeed, a large quantity of lentivirus can be toxic for cells. Moreover, when myoblasts are transduced, their proliferation medium (MB-1) is different than that of virus production. In other words, when a volume of virus is added to myoblasts, there is

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a mix between the DMEM-HG and MB-1 media. We suggest that the final volume of transduction medium should be composed of at least 50% of MB-1 medium. This recommendation is also suitable when the medium of the cells to transduce is different from that used for virus production. This problem can be solved if you have concentrated your virus by ultracentrifugation or ultrafiltration. 19. After transducing myoblasts with a lentivirus coding for GFP or for micro-dystrophin, the centrifugation of these cells (already attached to the plate) allowed us to obtain a 20% increase of the cells expressing these transgenes. 20. If your transgene is under a strong ubiquitous promoter, transduced myoblasts should already express it. If your transduced cells do not express the transgene whereas the transfected 293T expressed it, there is a problem with your packaging plasmids. 21. If your transgene is under a muscle specific promoter such as muscle creatine kinase (MCK) promoter, transduced myoblasts have to be differentiated in a specific medium (medium C) to express the transgene. Generally, 2 days of differentiation are sufficient to obtain myotubes. If you are planning to introduce a truncated version of dystrophin in myoblasts, we suggest the use of a muscle specific promoter to drive this transgene since dystrophin is normally just expressed in myotubes or fibers and its expression seems toxic for myoblasts. 22. If you have an antibiotic resistance gene (under a ubiquitous promoter such as SV40) included in your lentiviral plasmid backbone, you can start the selection 2 days after the transduction. We recommend the use of puromycin instead of hygromycin since the selection is more rapid than with hydromycin. Puromycin at 1 Pg/PL during 2 days is normally sufficient to kill the untransduced cells. Even if your transduced cells expressed the puromycin resistance gene, do not extend the treatment more than 2–3 days since it is weakly toxic for eukaryotic cells. 23. Generally, the efficacy of a lentivirus is inversely proportional to the transgene size. This is due to the lentiviral vector encapsidation limit (±8–9 kb). References 1. Naldini L, Blomer U, Gallay P, Ory D, Mulligan R, Gage FH, Verma IM, Trono D (1996) In vivo gene delivery and stable transduction of nondividing cells by a lentiviral vector. Science 272:263–267 2. Taganov KD, Cuesta I, Daniel R, Cirillo LA, Katz RA, Zaret KS, Skalka AM (2004) Integrase-specific enhancement and suppression of retroviral DNA integration by com-

pacted chromatin structure in vitro. J Virol 78:5848–5855 3. Scherdin U, Rhodes K, Breindl M (1990) Transcriptionally active genome regions are preferred targets for retrovirus integration. J Virol 64:907–912 4. Luciw PA (1996) Human immunodeficiency viruses and their replication, in Fields Virology (Fields, B. N., Knipe, D. M., Howley, P. M.,

16 Chanock, R. M., Melnick, J. L., Monath, T. P., Roizman, B., and Straus, S. E.) 3rd ed., pp 1881–1952, Lippincott-Raven Publishers, Philadelphia, PA 5. Chan DC, Kim, PS (1998) HIV entry and its inhibition. Cell 93:681–684 6. Malim MH, Hauber J, Le SY, Maizel JV, Cullen BR (1989) The HIV-1 rev trans-activator acts through a structured target sequence to activate nuclear export of unspliced viral mRNA. Nature 338:254–257 7. Cullen BR (1986) Trans-activation of human immunodeficiency virus occurs via a bimodal mechanism. Cell 46:973–982 8. Trono D (1995) HIV accessory proteins: leading roles for the supporting cast. Cell 82: 189–192 9. Klimatcheva E, Rosenblatt JD, Planelles V (1999) Lentiviral vectors and gene therapy. Front Biosci 4:D481–496 10. Zennou V, Petit C, Guetard D, Nerhbass U, Montagnier L, Charneau P (2000) HIV-1

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genome nuclear import is mediated by a central DNA flap. Cell 101:173–185 11. Zufferey R, Nagy D, Mandel RJ, Naldini L, Trono D (1997) Multiply attenuated lentiviral vector achieves efficient gene delivery in vivo. Nat Biotechnol 15:871–875 12. Dull T, Zufferey R, Kelly M, Mandel RJ, Nguyen M, Trono D, Naldini L (1998) A thirdgeneration lentivirus vector with a conditional packaging system. J Virol 72:8463–8471 13. Zufferey R, Dull T, Mandel RJ, Bukovsky A, Quiroz D, Naldini L, Trono D (1998) Selfinactivating lentivirus vector for safe and efficient in vivo gene delivery. J Virol 72:9873–9880 14. Zufferey R, Donello JE, Trono D, Hope TJ (1999) Woodchuck hepatitis virus posttranscriptional regulatory element enhances expression of transgenes delivered by retroviral vectors. J Virol 73:2886–2892 15. Cronin J, Zhang XY, Reiser J (2005) Altering the tropism of lentiviral vectors through pseudotyping. Curr Gene Ther 5:387–398

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Chapter 17 Generating Tamoxifen-Inducible Cre Alleles to Investigate Myogenesis in Mice Christoph Lepper and Chen-Ming Fan Abstract Gene inactivation has become the gold standard for determining gene function in the mouse. Many genes inactivated in the germ line cause early lethality that precludes phenotypic assessment at a later time point. Conditional gene inactivation using Cre recombinase expressed via a tissue specific promoter/enhancer allows phenotypic analyses of selected tissues, but lacks temporal control. Recent development of the tamoxifen-inducible Cre-ERT2 offers both cell type-specific and temporal control of conditional gene inactivation. As an example, we describe the design and step-wise construction of a Cre-ERT2 knock-in allele at the Pax7 locus using the recombineering method – Pax7 is selectively expressed in embryonic muscle progenitors and adult muscle stem cells. The resulting Pax7-Cre-ERT2 (Pax7CE) allele has been successfully applied to embryos and adults for tamoxifen-regulated myogenic lineage tracing and gene inactivation (Nature 460:627–631, 2009; Genesis 48:424–436, 2010). Key words: Pax7, Myogenic progenitor, Satellite cell, Lineage tracing, Tamoxifen, Cre, Cre-ERT2, Homologous recombination, Recombineering, Knock-out, Knock-in

1. Introduction In recent years, Cre-mediated recombination between loxP sites to activate reporter gene expression has been intensely used for lineage tracing in the mouse (1). The advantage of the Cre system is that it also provides versatility for concomitant gene inactivation of conditional alleles with loxP sites. However, conventional Cre is constitutively active and thus, its application to reporter gene activation and/or conditional gene inactivation is a cumulative assay for all cells at any one time expressing Cre. The advanced Cre technology utilizes a tamoxifen inducible Cre-estrogen receptor (ER) fusion protein (2, 3). For this, Cre is

Joseph X. DiMario (ed.), Myogenesis: Methods and Protocols, Methods in Molecular Biology, vol. 798, DOI 10.1007/978-1-61779-343-1_17, © Springer Science+Business Media, LLC 2012

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fused with the ligand-binding domain of a mutated ligand-binding domain of ER, which does not bind to estrogen but binds to tamoxifen with high affinity. Cre-ER proteins are sequestered in the cytoplasm via association with the HSP90 chaperone. Upon addition of tamoxifen, tamoxifen-bound Cre-ER dissociates from HSP90, translocates into the nucleus, and carries out site-specific recombination between flanking loxP sites. A more advanced version is Cre-ERT2, in which additional mutations have been engineered into the ER ligand-binding domain to bind tamoxifen with a higher affinity (4). Therefore, lower doses of tamoxifen can be used, reducing unwanted side effects – tamoxifen is also an inhibitor to protein kinase C (5). For manipulating adult muscle stem cells, two Cre-ER knockin (KI) alleles at the Pax7 locus have been reported (6–8). Below we detail the construction of the KI/knock-out (KO) allele of the Pax7-Cre-ERT2 allele (6, 7). We contrast our design to an earlier version of the Pax7-IRES-Cre-ERT allele (8) in both the main text and the note section. All steps described here are generally applicable to place Cre-ERT2 into any gene of interest.

2. Materials There are two major steps in making a Pax7-Cre-ERT2 homologous recombination vector: (1) Sub-clone a genomic fragment of Pax7 from a bacterial artificial chromosome (BAC); (2) Place the Cre-ERT2 and a companion positive selection cassette into the desired position (see later) of the vector containing the genomic fragment of Pax7. 1. BAC with ample sequences flanking the location (~10–20 kb on each side) where you plan to place Cre-ERT2 into your favorite gene. We screened a 129sv genomic library for Pax7-containing BACs (Invitrogen). Now the Ensembl genome browser can be searched for 129sv BACs made from AB2.2 ES cells to obtain the target Pax7 genomic fragment. This BAC library has been end-sequenced and is publicly available (9). Instructions for searching the database and ordering end-sequenced BACs are detailed in the referenced paper (see Note 1). 2. Recombineering reagents: We recommend you familiarize yourself with the recombineering method (10, 11) for making homologous recombination or transgenic vectors. The protocols and reagents are described in the above two references. Login to the following website: http://web.ncifcrf.gov/ research/brb/reagents/recombineeringReagent.aspx to request reagents and download protocols. Recombineering is ideally suited for this type of cloning as it does not depend on restriction

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enzyme sites to place Cre-ERT2 into a desired location (see Note 2). The basic components needed are as follows: (1) SW102 bacteria strain, which contains heat-shock inducible L phage genes exo, bet, and gam for recombination (see Note 3). (2) pL253 plasmid, which contains a tk cassette for negative selection in ES cells. 3. A Cre-ERT2-IRES-DsRed-Frt-neo-Frt (6) or a Cre-ERT2-Frt-neoFrt cassette. Both cassettes are available from our lab. You may find/order more advanced versions from Addgene (see Note 4). 4. Electroporator. 5. Gene-specific oligonucleotides for recombineering (see Method for their design). 6. T4 polynucleotide kinase. 7. Agarose. 8. Ethidium bromide, stock at 10 mg/mL. 9. Qiagen large construct kit. 10. Zymogen gel recovery kit. 11. Luria Broth (LB): 10 g tryptone, 5 g yeast extract, 10 g NaCl per liter of H2O. 12. LB-chloramphenicol (LB-chlor, 12.5 Mg/mL chloramphenicol) plates. 13. LB-ampicillin (LB-amp, 50 Mg/mL ampicillin) plates. 14. LB-kanamycin (LB-kan, 25 Mg/mL kanamycin) plates. 15. 32°C incubator. 16. 32°C shaker. 17. 37°C incubator. 18. 42°C water bath.

3. Methods 3.1. Confirm the Ordered BAC

Based on end sequence information, an in silico prediction of the restriction enzyme map of the BAC is necessary for two reasons: It helps to confirm the BAC to be correct when it arrives; it helps to design restriction enzyme digests and probes for Southern analysis of potential ES cell clones. 1. BAC clones arrive as bacterial stabs. Streak out the bacteria on a LB-chlor plate. The BAC library referenced is constructed in the BACe3.6 vector (chloramphenicol resistant).

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2. Inoculate five colonies for mini-prep using standard alkaline lysis method (12). Digest ½ of the miniprep DNA with a restriction enzyme chosen based on the in silico predicted map and run the digested DNA on a 1% agarose gel. 3. Stain the gel with ethidium bromide (5 Mg/mL) and check digested patterns for two things: First, all or most of the five clones give you the same pattern (i.e., the BAC is not a mixture); Second, the digest pattern is as predicted based on the in silico analysis (i.e., no gross rearrangements). 4. Inoculate a clone with the predicted digest pattern for a 500 mL prep. The Qiagen large-construct kit can be used. Intact clean BAC DNA is needed for electroporation into SW102 bacteria for recombineering. To be certain that the BAC is as ordered, resequence BAC ends with T7 and Sp6 primers. 3.2. Transfer the BAC into SW102

1. Inoculate a single colony of SW102 in 5 mL of LB in a 14 mL snap cap tube. Grow at 32°C until OD600 = 0.4–0.6. Chill on ice for 5 min. Keep cold during the following steps. 2. Spin down the bacteria in a prechilled centrifuge (4°C, 4,000 rpm (Sorvall RC-3B centrifuge), 5 min), and wash with 5 mL of ice-cold deionized water. Repeat the spin-wash step 3 times. After the third wash, resuspend bacteria in 50 ML of deionized water. 3. Electroporate 100 ng of the BAC (from Subheading 3.1) into SW102. Any electroporator designed for bacteria transformation should work for this step. Dilute the electroporated bacteria into 1 mL of LB, incubate at 32°C for 1 h in a shaker. 4. Plate 100 ML onto one LB-chlor plate, and the remainder onto an additional plate. Incubate at 32°C for 24 h. 5. Pick three colonies for mini-prep and perform restriction enzyme digest as above in Subheading 3.1 to be sure that the transferred BAC remains intact. 6. Make a glycerol stock of the BAC clone in SW102 for longterm storage, or proceed to the next step.

3.3. Prepare Oligos and pL253 Vector to Capture Desired Genomic Region from the BAC

The Pax7-Cre-ERT2 allele that we generated is a KI allele as well as a KO allele. We replaced exon 1 of Pax7 with Cre-ERT2 (see Fig. 1). For this, we retrieved a genomic fragment containing ~18 kb 5c arm and ~2 kb 3c arm (see Note 5). If you do not wish to inactivate the gene function, place the Cre-ERT2 cassette in the 3c UTR for bicistronic expression via IRES, e.g., the Pax7-IRES-Cre-ERT allele (8) (see Note 6). pL253 contains a polylinker for linearizing the final vector prior to transfection into ES cells. It also contains a tk negative selection cassette to reduce the number of ES cell colonies containing random insertions.

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Fig. 1. Flow chart of creating a Cre-ERT2 knock-in targeting vector. The two recombineering steps are outlined. Vectors, cassettes, and pertinent elements of the constructs are as depicted and labeled; oligos are short lines; squares, restriction enzyme site overhangs; p, phosphorylated ends. We recommend you draw out the recombination events so that the oligonecleotide sequences/orientations and the overhangs are placed in the correct order for ligation and recombineering. At the bottom, a simpler Cre-ERT2 cassette is diagrammed (see Note 8) and can be used as an alternative.

1. Make (order) four oligonucleotides (see Fig. 1 for configuration): The two 5c oligos are complementary but with a NotI overhang for ligation to NotI-cut pL253. The two 3c oligos are to be ligated to BamHI-cut pL253. For both, we designed 65 bp oligos identical to the 5c and 3c ends of Pax7 genomic fragment to be retrieved from the BAC (see Note 7). 2. Resuspend each oligo at 1 Mg/ML in water. For each pair, set up kinase reaction (using T4 polynucleotide kinase) with 5 Mg of the oligo with the restriction enzyme site overhang using standard conditions (12) in a 45 ML reaction.

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3. After the kinase reaction, add 5 Mg of the corresponding complementary oligo. Immediately boil them together for 5 min. Let them gradually cool to room temp to anneal. Now, they are ready to use at 200 ng/ML. 4. Cut 3 Mg of pL253 with NotI and BamHI. Run a 1% agarose gel and gel purify the linearized vector using the Zymogen gel recovery kit and elute DNA in 10 ML of water. Save 1/10 of this for step 6 below. 5. Ligate 9 ML of the linearized vector with 5 ML (1 Mg) each of the 5c and 3c annealed oligonucleotides in a 50 ML reaction, overnight at 16°C. 6. Run out the ligation mix on a 1% agarose gel. Run the saved unligated linearized vector (from step 3) side by side as control. There are many bands when the ligation products are run out on the gel, including the ligated vector multi-mers and oligo dimers. 7. Gel isolate the ligated vector, which is only slightly larger than the unligated vector control, using the Zymogen kit and elute DNA in 6 ML of water. 8. Determine the DNA concentration. Minimally 100 ng/ML is needed. The ligated vector is now ready for electroporation into SW102 containing the BAC (from Subheading 3.2). 3.4. Capture the Genomic Fragment from BAC by Recombineering

1. Grow up 5 mL of SW102 bacteria with the Pax7 containing BAC in LB-chlor (12.5 Mg/mL) at 32°C until OD600 = 0.4–0.6. 2. Heat shock in 42°C water bath (to induce L phage recombination system) for 15 min with occasional swirling. 3. Immediately chill on ice and proceed to make competent cells as in Subheading 3.2, Step 2. 4. Electroporate 1–4 ML of purified oligo-ligated vector (preferably t 400 ng) into competent cells. 5. Transfer electroporated cells into 1 mL of LB and culture at 32°C for 1 h. 6. Plate the bacteria onto two LB-amp plates, one with 100 ML, and the other with the rest. Incubate at 32°C for at least 24 h.

3.5. Screen for Recombinants

1. Inoculate 5–10 of the smallest colonies in LB-amp and grow them at 32°C until saturation. Bacteria carrying recombined plasmids grow very slowly compared with those carrying recircularized pL253. 2. Isolate DNA by standard alkaline lysis miniprep (12). 3. Without enzyme digest, run 1/5 of miniprep on a 1% agarose gel with size markers. The potential recombinants have only one band of large size (>10 kb). Those that have a prominent 4–5 kb band are clones with recircularized pL253.

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4. Analyze potential recombinants by restriction enzyme digest to determine whether they have the predicted digest pattern. To be sure, make primers flanking the polylinker of pL253 for sequencing the recombined junction. 5. Make a glycerol stock of the SW102 bacteria harboring the recombined plasmid. The bacteria can be used directly for the following step. 3.6. Replace Exon 1 of Pax7 with a Cre-ERT2Frt-neo-Frt Cassette

The Cre-ERT2 cassette developed by the Chambon lab (4) has a rabbit globin intron and a SV40 early polyA signal. Both are important for efficient nuclear export of the transcript. We placed an IRESDsRed between Cre-ERT2 and the polyA signal for visualizing Pax7expressing cells. For ES cell selection, we included an Frt-neo-Frt cassette (derived from the pL452 plasmid (10)) 3c to the polyA signal to make it as one cassette (6). The final cassette has 5c SpeI and 3c SalI sites for oligo ligation/recombineering (see Note 8). 1. Similar to capturing the genomic fragment (same principle as in Subheading 3.3), make four oligonucleotides. The 5c pair of oligos is complementary with a SpeI overhang. The 3c pair of oligos has a SalI overhang (see Note 8). 2. Resuspend, kinase, and anneal the oligos as in Subheading 3.3. 3. Digest 3 Mg of Cre-ERT2-IRES-DsRed-Frt-neo-Frt (6) cassette with SpeI and SalI, and isolate the digested cassette using the Zymogen kit. 4. Ligate oligos to the cassette. 5. Gel purify the ligated cassette using the Zymogen kit, and elute in 6 ML of water. 6. Prepare electrocompetent cells from heat-shocked SW102 containing the captured Pax7 genomic fragment in pL253 (from Subheading 3.5) using the same method described in Subheading 3.4. 7. Electroporate the ligated cassette into competent SW102, add 1 mL of LB and incubate for 1 h at 32°C. 8. Plate at two dilutions, 100 and 900 ML, onto LB-kan plates (the neo gene provides kan resistance). Incubate for at least 24 h at 32°C.

3.7. Screen for Recombinants

1. Pick 5–10 of the smallest colonies for culture in LB-kan at 32°C. 2. Miniprep and run 1/5 of DNA on a 1% agarose gel without enzyme digest. 3. For those with only one large band (potential recombinants), dilute 1 ML of DNA in 100 ML of water.

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4. Use 1 ML of each diluted DNA of potential recombinants (3–5 are usually sufficient) for transformation into regular chemically competent cells (e.g., TOP10 or DH5A). Plate on LB-kan plates, and incubate overnight at 37°C. 5. Pick one colony from each transformation for culture and miniprep. Digest the DNA with the selected restriction enzyme(s) to check for correct recombination. Unlike the BAC, pL253 is a multi-copy plasmid and presumably only one of the multiple plasmids goes through recombination. Thus, the above recombineering step results in mixed plasmids in the bacteria. Retransformation is necessary to remove unrecombined plasmids (not kan-resistant) before analysis by restriction enzyme digest pattern can be carried out accurately. 3.8. Sequence the Recombined Construct

To be sure the construct is the correct recombinant before electroporating it into ES cells, sequence the targeting vector. 1. Sequence the junctions between pL253 and the genomic fragment. 2. Sequence the junctions between Cre-ERT2-IRES-DsRed-Frtneo-Frt and its flanking regions. If all junction sequences and restriction enzyme digest patterns are as predicted, then the final vector is done.

3.9. Additional Steps

To obtain the mouse line, you need to carry out additional steps. As these steps are routine now, we only point out some main points needing attention. Your transgenic facility manager should also be able to help you out with the steps below. 1. Make primers to PCR across the short arm to screen for homologous recombination events in ES cell clones : one inside in the Cre-ERT2-IRES-DsRed-Frt-neo-Frt cassette, and the other outside of the homology arm. 2. Design Southern probes located outside of homology arms to determine true homologous recombination. PCR across the short arm is only for quick screening. Southern analysis should be done for confirmation, preferably on both ends. 3. Remove the Frt-neo-Frt cassette either by transfecting a FLPe expressing plasmid (available from Addgene) into the targeted ES cells (and select for clones that have the neo cassette deleted) prior to blastocyst injection or by crossing to the Actin-FLPe mice (available from Jackson Lab) after germ line transmission of the allele. 4. Cross the Cre-ERT2 allele to a Cre reporter line, e.g., Rosa26RLacZ or Rosa26RYFP (available from Jackson Lab) to test its efficacy (see Note 9).

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5. Preparing tamoxifen: Tamoxifen can be made in corn oil at 20 mg/mL, aliquoted, and stored in −80°C. Thawed tamoxifen can be stored at 4°C and used up to 2 weeks with full activity (see Note 10). 6. Administrating tamoxifen: The dosage of tamoxifen needs to be titrated pending on your experimental needs. For the Pax7Cre-ERT2 allele, we used 1.5–3 mg/40 g of mouse body weight. Intraperitoneal injection of tamoxifen is the easiest and requires only minimal training. Oral gavages are less harmful to the animal but require a bit more training. Food supplementation is also an option – contact Harland Tek for more information on manufacturing customized food.

4. Notes 1. Isogenic DNA is preferred for homologous recombination in ES cells. If your transgenic facility uses a 129sv derived ES cell line, buy the BAC from the AB2.2 BAC library. C57/BL6 ES cells are becoming more widely used. There are several C57/ BL6 BAC libraries listed on the CHORI (Children’s Hospital Oakland Research Institute) website. You can use the UC Santa Cruz Genome Browser website to search for CHORI BACs containing your genes of interest. The website has a direct link to paired-end sequenced BAC clones for purchase. 2. There are newer versions of recombineering methods and materials from other labs. They have become more sophisticated and easier to use. For example, the method developed by the Capecchi lab (13) may be easier to use for certain kinds of targeting vectors. 3. When we did recombineering for the Pax7-Cre-ERT2 vector, NCI provided the DY380 bacteria (discontinued) for recombineering. It has been substituted by SW102, hence we use SW102 in the text. There are also other updated changes incorporated into this protocol, which are not always explicitly stated. 4. Addgene has many published vectors deposited for public distribution. We find it easier to get plasmids from Addgene than to ask from investigators. 5. The long and short of the homology arms: We now design all recombination constructs with a very long arm and a less than 2 kb short arm. Arms less than 2 kb are consistently more reliable for PCR screening of the homologous recombinant. If your lab is proficient in using Southern hybridization for screening ES cells, then you can make both arms as long as you wish (note that high copy number plasmids have an upper size

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limit around 27–30 kb for propagation). We cannot emphasize enough that a Southern is essential to be certain of correct homologous recombination. Because the particular place you “have to” place the Cre-ERT2 cassette, you may not have many choices for restriction enzymes. Hence, a careful in silico investigation prior to starting a project is crucial. 6. To KO or not to KO when creating a KI: You can insert the Cre-ERT2 cassette after the stop codon and employ the IRES to express Cre-ERT2, which is followed by the gene’s own 3c UTR. A Frt-neo-Frt cassette can be placed behind the Cre-ERT2 cassette for positive selection in ES cells. The advantages of this approach are that the function of the gene is retained and that the endogenous 3c UTR is utilized. As microRNA regulation is prevalent, preservation of the natural 3c UTR offers an additional layer of control for regulated expression. On the flip side, the use of IRES often leads to a lower level of expression of the downstream gene (the Cre-ERT2). For this to work as intended, you should not put an artificial polyA signal after IRES-Cre-ERT2 and you should remove the Frt-neo-Frt cassette. The Pax7-Cre-ERT2 allele described here is also a KO allele (6, 7): The advantage is that the Cre-ERT2 is likely expressed at a higher level than that of utilizing IRES. The disadvantages are that the mouse is heterozygous for Pax7 and that the 3c UTR is artificial. 7. The design principle of the oligonucleotides: Although 45 bp of homology is said to be sufficient for recombineering, we find that in practice 65 bp in length is better. If you experience difficulty to obtain recombinants using oligonucleotides, then use PCR to generate larger homology fragments. We had to do so for the second step. 300–400 bp fragments for both sides can increase efficiency dramatically. Incorporate the restriction sites at the same end as the oligonucleotides. In this case, cut the PCR fragment with restriction enzymes so that they can be ligated to the cassette. We recommend you draw out each recombination step with your designed oligonucleotides and target sequences to make sure everything is in the right order. If you place the overhang at the wrong end of the oligonucleotides, you will not get the recombination events that you plan. 8. In our Pax7-Cre-ERT2 mice, Ds-Red was not detected, likely due to inefficiency from IRES-mediated bicistronic expression. Thus, “over-design” does not always pay off. There are other cassettes deposited at Addgene and you should check for updated versions that suit your need. We recently made a simpler cassette with two polyA signal sequences (one SV40 late polyA followed by SV40 early

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polyA), which hypothetically should increase the efficiency of polyA addition. This new cassette has 5c NotI and 3c SalI sites for ligation/recombineering. Both cassettes are diagrammed in Fig. 1 and are available upon request. 9. The keys to characterize a new Cre-ERT2 line are the following: (a) Set up crosses to obtain mice carrying both your Cre-ERT2 and a reporter allele. Without administrating tamoxifen, there should be no reporter gene expression, i.e., no leaky Cre activity. For example, the Pax7-IRES-Cre-ERT mouse is reported to have spontaneous activity, though minimal (8). (b) Choose the Rosa26 reporter allele: there are many alleles with different reporters available from the Jackson Lab. The original Rosa26RLacZ allele is still popular because of the simplicity of detecting reporter gene expression by X-gal staining. For those who are interested in cellular morphology or live imaging, fluorescent reporter alleles may be favored. 10. You should titrate the dosage of tamoxifen for your experiment. Because each Cre-ERT2 line expresses different levels of Cre in different cell types, different tamoxifen dosages will likely be required to achieve your goal. For example, a single low dosage of tamoxifen may be preferred for clonal lineage tracing. On the contrary, multiple injections are likely needed to efficiently inactivate conditional alleles. References 1. Soriano P (1999) Generalized lacZ expression with the ROSA26 Cre reporter strain, Nat Genet 21, 70–71. 2. Metzger D, Clifford J, Chiba H, Chambon P (1995) Conditional site-specific recombination in mammalian cells using a ligand-dependent chimeric Cre recombinase, Proc Natl Acad Sci USA 92, 6991–6995. 3. Indra AK, Warot X, Brocard J, Bornert JM, Xiao J.H, Chambon P, Metzger D (1999) Temporally-controlled site-specific mutagenesis in the basal layer of the epidermis: comparison of the recombinase activity of the tamoxifeninducible Cre-ER(T) and Cre-ER(T2) recombinases, Nucleic Acids Res 27, 4324–4327. 4. Feil R, Wagner J, Metzger D, Chambon P (1997) Regulation of Cre recombinase activity by mutated estrogen receptor ligand-binding domains, Biochem Biophys Res Commun 237, 752–757. 5. O’Brian CA, Liskamp RM, Solomon DH, Weinstein IB (1985) Inhibition of protein

kinase C by tamoxifen, Cancer Res 45, 2462–2465. 6. Lepper C, Conway SJ, Fan, CM (2009) Adult satellite cells and embryonic muscle progenitors have distinct genetic requirements, Nature 460, 627–631. 7. Lepper C, Fan CM (2010) Inducible lineage tracing of Pax7-descendant cells reveals embryonic origin of adult satellite cells, Genesis 48, 424–436. 8. Nishijo K, Hosoyama T, Bjornson CR, Schaffer BS, Prajapati SI, Bahadur AN, Hansen MS, Blandford MC, McCleish AT, Rubin BP, Epstein JA, Rando TA, Capecchi MR, Keller C (2009) Biomarker system for studying muscle, stem cells, and cancer in vivo, FASEB J 23, 2681–2690. 9. Adams DJ, Quail MA, Cox T, van der Weyden L, Gorick BD, Su Q, Chan WI, Davies R, Bonfield JK, Law F, Humphray S, Plumb B, Liu P, Rogers J, Bradley A (2005) A genomewide, end-sequenced 129Sv BAC library

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resource for targeting vector construction, Genomics 86, 753–758. 10. Lee EC, Yu D, Martinez de Velasco J, Tessarollo L, Swing DA, Court DL, Jenkins NA, Copeland NG (2001) A highly efficient Escherichia coli-based chromosome engineering system adapted for recombinogenic targeting and subcloning of BAC DNA, Genomics 73, 56–65. 11. Yu D, Ellis HM, Lee EC, Jenkins NA, Copeland NG, Court DL (2000) An efficient recombina-

tion system for chromosome engineering in Escherichia coli, Proc Natl Acad Sci USA 97, 5978–5983. 12. Sambrook J, Fritsch E., Maniatis T (1989) Molecular Cloning: A Laboratory Manual, 2nd ed., Cold Spring Harbor Press, Cold Spring Harbor, New York. 13. Wu S, Ying G, Wu Q, Capecchi MR (2008) A protocol for constructing gene targeting vectors: generating knockout mice for the cadherin family and beyond, Nat Protoc 3, 1056–1076.

Part V Muscle Profiling

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Chapter 18 Gene Profiling Studies in Skeletal Muscle by Quantitative Real-Time Polymerase Chain Reaction Assay Shephali Bhatnagar, Siva K. Panguluri, and Ashok Kumar Abstract Gene profiling is an excellent tool to identify the genetic mechanisms, networks, and molecular pathways involved in skeletal muscle development and muscular disorders. Oligonucleotide or cDNA microarray can be the first step to identify the global gene expression in the study of interest. As microarray techniques provide a large set of differentially expressed genes in a given comparison, the expression profile can be narrowed down by taking various parameters into consideration such as fold values, p-values, and their relevance to the study. Every technique has its own limitations. Therefore, further validation of the results with a different technique is always necessary. Quantitative real-time reverse-transcriptase polymerase chain reaction (qRT-PCR) is the most common technique to validate microarray data and to study the relative expression of specific genes in any experimental set-up. Here, we describe, the qRT-PCR technique, in detail, for successful gene expression studies in skeletal muscle cells and tissues. Key words: Skeletal muscle, Quantitative real-time PCR, C2C12 myoblasts, RNA isolation, RNA analysis, Gene expression

1. Introduction Skeletal muscle constitutes approximately 40% of human body mass and is required for basic functions such as locomotion, metabolism, and respiration. Myogenesis is a multistep developmental program that generates skeletal muscle in embryos (1, 2). This process is also required for postnatal growth and the regeneration of myofibers after injury (1). Adult skeletal muscle exhibits a very high level of plasticity. Resistance exercise and nutritional uptake lead to skeletal muscle hypertrophy (3). Conversely, in a wide variety of disease states and conditions, skeletal muscle undergoes atrophy or wasting, which is a critical determinant of human morbidity and mortality (4–6). Furthermore, mutations in many Joseph X. DiMario (ed.), Myogenesis: Methods and Protocols, Methods in Molecular Biology, vol. 798, DOI 10.1007/978-1-61779-343-1_18, © Springer Science+Business Media, LLC 2012

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muscle structural genes lead to muscular dystrophy, a group of fatal genetic disorders involving striated muscle (7). Skeletal muscle development as well as pathology is governed by coordinated changes in the activity and expression of several genes (1, 8). Different physiological and pathophysiological stimuli modulate the activity of various signaling pathways and transcription factors leading to expression of a specific set of genes in nuclei. A number of gene profiling studies using microarray approaches have been performed leading to the identification of sets of genes involved in skeletal muscle development, plasticity, and muscular disorders such as muscular dystrophy and inflammatory myopathies (9–14). Although microarray is a powerful approach to simultaneously study the expression of a large number of genes, the reliability of microarray results depends on several factors such as array production, RNA extraction, probe labeling, hybridization conditions, and image analysis. Therefore, the genes identified as differentially expressed by microarray approaches must be confirmed by another method. Quantitative real-time PCR (qRT-PCR) is a highly sensitive and rapid technique, which requires 1,000-fold less RNA compared to conventional assays such as Northern blot for gene expression studies (15). With the advent of multiplexing technology, it has now become possible to study the expression of multiple genes in a single qRT-PCR reaction. Furthermore, the qRT-PCR is an ideal technique to study relative expression of known muscle genes in experimental or clinical settings. In addition to animal models, the major steps of muscle development and plasticity can be recapitulated using C2C12 myoblasts/myotubes in vitro. Indeed, C2C12 myoblasts are the most commonly used cells for studying skeletal muscle in culture. Here, we provide the detailed methodology for RNA isolation and analysis and qRT-PCR technique for gene expression studies in skeletal muscle cells and tissues. Though most of the procedures given in this chapter are similar across all types of real-time PCR machines, we have provided details of qRT-PCR assays using 7,300 Sequence Detection System (Applied Biosystems) and SYBR Green dye.

2. Materials Prepare all solutions using nuclease-free water and under nucleasefree conditions (see Notes 1–3). 2.1. Cell Culture

Perform all the following steps aseptically under the hood. All the cell culture reagents are from Invitrogen unless specified. 1. C2C12 myoblasts (American Type Culture Collection). 2. Growth Medium: To 500 mL of DMEM high glucose 1×, add 5 mL of HEPES from 100× stock, 5 mL of Penicillin/

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streptomycin from 100× stock, and 50 mL heat-inactivated fetal bovine serum. 3. 0.25% Trypsin solution. 4. Differentiation medium: To 500 mL of DMEM high glucose 1× add: 5 mL of HEPES from 100× stock, 5 mL of Penicillin/ streptomycin from 100× stock, and 10 mL horse serum (Sigma). 2.2. Quantitative Real-Time RT-PCR 2.2.1. Total RNA Isolation

1. TRIzol reagent: Store the reagent at 4°C in amber color bottle (see Note 4). 2. Diethyl pyrocarbonate (DEPC)-treated water: Add 1 mL of 0.1% DEPC to sterile water and mix well. Alternatively, water can be autoclaved after adding DEPC (see Note 5). 3. 75% Ethanol: Add 25 mL of DEPC-treated water to 75 mL absolute ethanol. Mix well and refrigerate until further use. 4. RNeasy kit. 5. Chloroform (Molecular Biology grade).

2.2.2. cDNA Synthesis

1. High Capacity cDNA Reverse Transcription Kit (Applied Biosystems). The kit consists of the following components: 10× reverse transcription (RT) buffer, 100 mM dNTP mix, 10× RT random primers, multiscribe reverse transcriptase 50 U/PL. 2. DNase/RNase free water.

2.2.3. RNA Quality Analysis Using Agilent 2100 Bioanalyzer

1. RNA samples (up to 12 per chip). 2. RNA 6,000 ladder (Ambion). 3. Bioanalyzer RNA Nano Kit, includes chips and reagents. 4. RNaseZap.

2.2.4. SYBR Green qRT-PCR

All the components are from Applied Biosystems unless specified. 1. cDNA made previously (see Subheading 2.2.2). 2. DNase/RNase free water. 3. 20 PM Primers, forward and reverse from your favorite vendor. 4. Power SYBR Green PCR Master Mix. 5. 96-Well Optical Reaction Plate with barcode 128. 6. MicroAmp optical Adhesive Film, PCR compatible, DNA/ RNA/RNase free.

2.2.5. Primers

We design our primers using Vector NTI software (Invitrogen). Using this software, we get >95% success rate that the primers will work in the qRT-PCR assays.

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3. Methods 3.1. Culturing C2C12 Myoblasts and Their Differentiation into Myotubes

1. Culture C2C12 myoblasts in growth medium at 37°C and 5% CO2 in an incubator until they reach 70–80% confluency. 2. Split the myoblasts in 1:3 ratio to maintain the undifferentiated myoblast population. 3. Differentiate C2C12 myoblast cultures by replacing the growth medium with differentiation medium (DM). 4. Change differentiation medium every 48 h to maintain healthy myotubes.

3.2. Extraction of Total RNA from Cultured Myoblasts/Myotubes

Perform all centrifugation steps in an Eppendorf microcentrifuge. 1. Depending on the study the treated myoblasts/myotubes were harvested by trypsinizing the cells using 0.25% trypsin solution. 2. Collect the cells in microcentrifuge tubes and pellet them by centrifuging at 8,000 rpm for 2 min at 4°C. 3. Discard the supernatant and add 1 mL TRIzol reagent. 4. Pipette the cell pellet in TRIzol 5–6 times to ensure proper lysis. 5. To the homogenate, add 200 PL chloroform and mix well by vortexing the tube for 1 min. 6. Incubate at room temperature for 5 min. 7. Centrifuge the tubes at 12,000 rpm in refrigerated centrifuge for 15 min. 8. Carefully transfer the top aqueous phase (approximately 600 PL) into a new eppendorf tube, add an equal volume of 75% ethanol, and vortex vigorously for 5 s. 9. Add the mixture to the RNeasy spin column from the RNeasy kit. 10. Centrifuge the tubes at 15,000 rpm for 20 s and discard the flow-through. 11. Add 100 PL of buffer RW1 (from RNeasy kit) and spin at 15,000 rpm for 20 s. 12. Discard the flow-through and the collection tubes and put columns into new collection tubes. 13. Add 500 PL of buffer RPE (from RNeasy kit) and centrifuge at 15,000 rpm for 30 s. 14. Discard the flow-through and repeat step 13. 15. Discard the flow-through and centrifuge again at 15,000 rpm for 30 s without buffer. 16. Discard the flow-through and collection tubes and put columns into new sterile 1.5 mL eppendorf tubes.

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17. Add 30 PL of RNase-free water in the center of the tube. 18. Incubate at room temperature for 5 min and centrifuge at 15,000 rpm for 1 min. 19. Measure the concentration of RNA by spectrophotometer or any other standard methods. 3.3. Total RNA Isolation from Skeletal Muscle Tissues

1. Place approximately 30–40 mg skeletal muscle tissue in a prelabeled microcentrifuge tubes placed on ice. 2. Chop tissues using a sterile razor blade in a dish (kept on ice) in a drop of TRIzol reagent. 3. Transfer chopped tissue to a glass mortar and pestle pretreated with DEPC water for 2 h. 4. Add 1 mL of TRIzol reagent to it and homogenize well (see Note 6). 5. Transfer the homogenate into a 1.5-mL eppendorf tube. 6. For rest of the process, follow steps 5–19 as described above for isolation of total RNA from cultured myoblasts.

3.4. Evaluating RNA Quality

We use Agilent 2100 Bioanalyzer to check the quality of RNA (see Note 7). The complete protocol for analyzing RNA quality is described as follows: 1. Clean the electrodes by placing a washing chip filled with 350 PL RNAseZap in the Bioanalyzer for 1 min. Place a washing chip filled with 350 PL DEPC water in the Bioanalyzer for 30 s. 2. Leave the lid open for 10 s to allow the electrodes to dry. 3. Place 400 PL of RNA gel matrix into the top receptacle of a spin filter (included in RNA Nano kit), place the spin filter in a microcentrifuge and spin at 4,000 rpm for 10 min. Filtered gel must be used within 4 weeks. 4. Put 130 PL of the filtered RNA gel matrix into an RNAase-free eppendorf tube and add 2 PL of RNA dye concentrate provided in the RNA Nano kit. 5. Vortex vigorously to ensure proper mixing of gel and dye (see Note 8). 6. Take a new RNA chip out of its sealed bag and place it on the chip priming station. 7. Take 9.0 PL of the gel-dye mix with a pipette and dispense this in the well marked with a black circle “G.” 8. Ensure that the plunger is at 1 mL. Then close the chip priming station. Press the plunger until it is held by the syringe clip. 9. Wait for 30 s. Slip plunger off clip and wait 5 s and then slowly pull the plunger back to the 1 mL mark. Open chip priming station.

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10. Pipette 9 PL of the gel-dye mix in each of the wells marked with an uncircled “G” (see Notes 9 and 10). 11. Pipette 5 PL of the RNA 6000 Nano Marker and put it in the well marked with the ladder symbol. 12. Dispense 5 PL of the RNA 6000 Nano Marker into each of the 12 sample wells. 13. Dilute RNA samples at 1:10 and 1:100 using RNAase-free water. Also put 2 PL RNA ladder in a tube. 14. Heat the tubes at 70°C for 2 min to denature RNA samples followed by snap cooling by putting the tubes on ice. 15. Pipette 1 PL of RNA 6000 ladder and put it in the well with the ladder symbol. 16. Pipette 1 PL of each sample and put it in each sample well (see Note 11). 17. Pipette 1 PL of RNA Nano Marker in each unused well (see Note 12). 18. Place the chip in the Agilent 2100 Bioanalyzer scanner and follow the user manual protocol for operating the machine and extracting the analyzed data. 19. Finally print results. A typical gel picture and results of one RNA sample of good quality are shown in Fig. 1 (see Note 13).

Fig. 1. Analysis of RNA quality by using Agilent 2100 Bioanalyzer. (a) A representative gel-like picture after running the total RNA samples on Agilent 2100 Bioanalyzer is presented here. Six different C2C12 RNA preparations were analyzed with the Agilent 2100 Bioanalyzer. (b) The electropherograms of RNA sample #1 showing the presence of ribosomal RNA 28S and 18S peaks is shown here.

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1. Use 2 Pg of RNA in a final volume made to 10 PL with DNase/ RNase-free water in a 0.2-mL PCR tube (see Note 14). 2. Make PCR master-mix using the High Capacity cDNA Reverse Transcription Kit as follows (see Note 15): Reagents

Volume (mL)

10× RT buffer

2.0

20× dNTP mix (100 mM)

0.8

10× RT random primers

2.0

Multiscribe reverse transcriptase

1.0

Nuclease free water

4.2

Total

10.0

3. Add 10 PL of PCR master-mix to RNA sample (i.e., from step 1 above). Total volume is now 20 PL. 4. Briefly vortex and centrifuge the tubes. 5. Program a PCR machine as follows: Conditions

Step 1

Step 2

Step 3

Step 4

Temperature (°C)

25

37

8

4

Time

10 min

120 min

5s

Forever

6. Put the tubes on PCR heating block and run PCR reaction. Store the tubes at −20°C after the PCR reaction is finished. 3.6. Quantitative Real-time RT-PCR

The Quantitative real-time RT-PCR consists of two steps: (1) designing primers and (2) setting up the reaction.

3.6.1. Designing Primers

We design primers using Vector NTI software (see Note 16). Basic steps for designing good quality primers using Vector NTI software are as follows: 1. Find the cDNA or mRNA sequence for the gene of interest in PubMed nucleotide database and copy the gene’s unique gene ID number. 2. Open Vector NTI software followed by clicking on Tools in the main menu, and then click on open link GID. 3. Paste the gene ID in the dialog box and click OK. The program will download the nucleotide sequence on your computer. 4. Select a region of 300–400 base pairs (bp) within the cDNA sequence. 5. Go to Analyses in main menu and click on Primer Design. A dialog box will appear.

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6. Make the following entry in only specific fields. The others are default; do not change the values in them. Product length: Min:100 bp, Max: 200 bp. Maximum number of output options: 50. Tm (C): t55 and d60. %GC: t55 and d60. Length: t20 and d25. 7. Click “Apply.” A window will open on upper left corner containing sequence of 50 primer sets. 8. Look at each forward and reverse primer sequence individually. The GC difference should be 0°C. Tm difference should not be more than ±1°C between two primers. 9. Click on the first primer meeting the required parameters, and then select Analyze. A new window will open containing the selected primer information. 10. Check for palindrome and repeats. Palindrome and repeats should be 0. If there is any number of Palindrome or Repeats in either forward or reverse primers, do not use this pair. Perform the same on other set of primers. 11. Next click on “Dimers and Hairpin Loops” icon. A new window will open providing separately the number of hairpin loops and dimers in the selected primer. The ideal situation is that we should have no hairpin loop and dimers. However, it is rare for most of the primers. The following criteria can be used to pick the good primers even with those having hair-pin loops and primer dimers. 12. Make sure that the primer does not have more than eight hairpin loops or dimers. Minimum is better but up to eight are acceptable. 13. Check the dimer dG and hairpin dG for each dimer and hairpin loop, respectively, by clicking button on the window. 14. The best value for dG should be 0 kcal/mol. However, dG values between −1.8 and +1.8 kcal/mol can be acceptable. If any of the two primers in the pair has a dG value outside this range, then do not use this pair and analyze other primer sets (see Note 17). 15. Once the right primer set is found, copy primer sequence and send for primer synthesis (see Note 18). 16. Test run qRT-PCR using these primers with a few samples. The primer set which shows a good dissociation curve should be used for qRT-PCR. 17. Figure 2a shows a typical dissociation curve for a good primer set with no primer dimers. Primer sets having primer dimers are shown in Fig. 2b. The dissociation curve of a good qRTPCR assay should look similar to Fig. 2a (see Note 19).

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Fig. 2. Dissociation curves in qRT-PCR assay. (a) A typical dissociation curve with no primer dimer after running a qRT-PCR assay is shown here. (b) The left peak represents a primer dimer in this dissociation curve.

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3.6.2. Setting Up the Reaction

After synthesizing the cDNA and obtaining primers, the real time PCR reaction is set up as follows: Components

Volume (mL)

cDNA

1

20 Pm Stock primer 1

1

20 PM Stock primer 2

1

RNase-free water

7

2× SYBER Green Master Mix

10

Total

20

1. Prepare the master-mix with all the ingredients except cDNA. 2. Dispense 1 PL of cDNA in the individual wells of 96-well Optical Reaction Plate with barcode 128 and then add 19 PL of master mix to each well. All reactions should be carried out in duplicate or triplicate to reduce variation. 3. Data normalization is accomplished using the endogenous control such glyceraldehyde 3-phosphate dehydrogenase (GAPDH) or E-actin (see Note 20). 4. Seal the plate using MicroAmp optical adhesive film and spin the plate in PCR plate centrifuge. 5. Insert the plate into the 7300 Sequence Detection System. 6. Set thermal conditions for qRT-PCR using 7300 system SDS software as follows: (a) Denaturation at 95°C for 10 min (b) 40 Cycles of denaturation at 95°C for 15 s, annealing and extension at 60°C for 1 min (c) Finally, a melting curve of 95°C for 15 s, 60°C for 15 s, and 95°C for 15 s. 7. Click on the 7300 system SDS software icon on the computer attached with 7300 Sequence Detection System. Click on Create New Document tab. 8. A new window will appear, select ddCt (Relative Quantitation) Plate in the Assay pull down menu. 9. Click “Next” and enter the name of the primes to be used in left side window. Then select the primer in left window and click Add button. After adding all primer names, click Next tab. 10. Enter the sample information for each well and save the file as .sds document. 11. In the same window, click Instrument tab and then click on Add Dissociation Stage tab. 12. Finally, click on Start tab. This will start the program. Do not disturb the program until the run is finished.

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1. Open 7300 system SDS software on the computer. 2. Click on Create New Document. 3. Select ddCt (Relative Quantitation) Study from the pull-down menu of the Assay tab. 4. Click on Next tab. A new window will appear. Click on Add plates tab. 5. Select the desired .sds file saved at the time of setting-up the qRT-PCR assay. Click open. The file will appear. Click Finish tab in the dialog box. 6. A new window will open. Select all the fields in the upper left box and click green arrow in the main menu. 7. Again select all the fields in upper left window. The corresponding Ct values will appear in bottom left window. 8. Go to main menu and save file as .sdm (SDS Multi-plate documents). 9. Click on main menu File tab, and sequentially click on Export, Results, and Both. Save the file as .csv file. 10. Close the application and proceed for analysis part using .csv file. 11. Open .csv file using Microsoft Excel program, calculate the averages for the duplicates/triplicates of each sample and normalizing gene. This gives us śCT values. 12. Deduct the $CT values of the normalizing gene from the corresponding śCT values of the samples. 13. Calculate the final average by taking the average of all control śCT values. 14. Subtracting the $CT values from the final average gives us the śśCt values. 15. The corresponding fold change is calculated as two to the power of śśCt values. This gives us the fold change in the samples when compared with control which can be plotted on a graph.

4. Notes 1. Prepare and store all cDNA synthesis and qRT-PCR reagents at −20°C (unless indicated otherwise). Cell culture medium should be stored at 4°C. 2. Diligently follow all waste disposal regulations (especially phenol based reagents used during RNA isolation) according to the material safety data sheet provided by the manufacturer. Store all other reagents at room temperature. 3. Always use sterile gloves while working with RNA. Avoid contact with skin or clothing. All the procedures should be done

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in RNAse/DNase free environment. Autoclave all the tips and eppendorf tubes properly before using. Clean the working area, pipettes, and other equipment used for RNA extraction with RNAZap and/or with 70% ethanol to ensure RNase free conditions. 4. TRIzol reagent may be corrosive and cause irritation. Therefore, avoid contact with skin directly. Use gloves and lab coats while working with TRIzol reagent (Read MSDS of the product for detailed protection and safety). 5. DEPC is not miscible with water; shake vigorously after adding DEPC to water for proper mixing. Also DEPC should not be stored more than 24 months (see MSDS of the product for full details). Follow the hazardous waste disposal methods for proper disposal of expired DEPC. 6. For good yield of RNA, muscle tissue should be properly homogenized in TRIzol reagent. When tissue is homogenized, the solution becomes turbid with no major tissue clumps visible. 7. It is always a good practice to check the quality of RNA before using it for qRT-PCR assay especially when RNA samples were stored at −80°C for several days. RNA samples can also be stored in small aliquots to avoid repeated freeze thaw. 8. To protect the gel–dye mix from light, cover the tube with foil. Remember to return the reagents to the cold room when you are finished. 9. Loading the gel–dye mix could be difficult. If you do not get the gel distributed evenly and without bubbles throughout the channels, you will not get good results. Look at the back of the chip for bubbles. 10. Use the chip within 5 min of preparation to prevent evaporation. Alternatively, cover the chip if it will be left standing for any length of time. 11. Sample concentrations should ideally be 100–200 ng/PL, though a concentration as low as 50 ng/PL can be used. 12. You must put the nano marker in every sample and the ladder well. Add water or nano marker to unused wells to bring the volume up to 6 PL. 13. Genomic DNA contamination of RNA samples can produce stray bands or clog the capillary. To check for genomic DNA, treat the samples with DNase. Run a DNase-treated sample next to an untreated sample. 14. Accurate pipetting is very important. Therefore, use properly calibrated pipettes. Place the tip at the bottom and center of each well when dispensing. Do not try to push pass the first resistance point on the pipette to avoid bubbles. You may pipette up and down gently to mix samples in the wells.

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15. To avoid multiple pipetting, it is better to make a master mix and then add master mix to the tubes containing cDNA. 16. Having good primer sets is critical for the success of qRT-PCR assay. 17. It is quite possible that you will not get any good quality primers in a selected region of 300–400 bp. If the primers are not good, move across the sequence (by shifting starting point 200 bp downstream) and perform the same search. 18. If the gene of interest is not giving good primer sets, we order two to three best possible sets of primers and test them in qRTPCR assays. 19. After finishing the run, it is a good idea to run the PCR products on agarose gel electrophoresis. This gel should show a single PCR product and no primer dimer. 20. In pathological conditions, skeletal muscles are infiltrated by other cells such as immunocytes and fibroblasts. Since GAPDH and E-actin are expressed by all cell types, using these genes as normalizing controls, it is not possible to distinguish whether the observed change in expression of a specific gene is because of its altered expression in skeletal muscle or other cell type. To determine the changes in expression level only in skeletal muscle, we suggest using some skeletal muscle specific endogenous gene for normalization purposes. Myosin heavy chain four or muscle creatine kinase could serve as good normalizing controls.

Acknowledgment This work was supported by National Institute of Health grants (AG029623 and AR059810) to Ashok Kumar. References 1. Charge SB, Rudnicki MA (2004) Cellular and molecular regulation of muscle regeneration. Physiol Rev 84:209–238 2. Perry RL, Rudnick MA (2000) Molecular mechanisms regulating myogenic determination and differentiation. Front Biosci 5:D750–D767 3. Glass DJ (2005) Skeletal muscle hypertrophy and atrophy signaling pathways. Int J Biochem Cell Biol 37:1974–1984 4. Jackman RW, Kandarian SC (2004) The molecular basis of skeletal muscle atrophy. Am J Physiol Cell Physiol 287:C834–C843 5. Li H, Malhotra S, Kumar A (2008) Nuclear factor-kappa B signaling in skeletal muscle atrophy. J Mol Med 86:1113–1126

6. Kandarian SC, Stevenson EJ (2002) Molecular events in skeletal muscle during disuse atrophy. Exerc Sport Sci Rev 30:111–1116 7. Emery AE (2002) The muscular dystrophies. Lancet 359:687–695 8. Cao PR, Kim HJ, Lecker SH (2005) Ubiquitinprotein ligases in muscle wasting. Int J Biochem Cell Biol 37:2088–2097 9. Bodine SC, Latres E, Baumhueter S, Lai VK, Nunez L, Clarke BA, Poueymirou WT, Panaro FJ, Na E, Dharmarajan K, Pan ZQ, Valenzuela DM, DeChiara TM, Stitt TN, Yancopoulos GD, Glass DJ (2001) Identification of ubiquitin ligases required for skeletal muscle atrophy. Science 294:1704–1708

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10. Giresi PG, Stevenson EJ, Theilhaber J, Koncarevic A, Parkington J, Fielding RA, Kandarian SC (2005) Identification of a molecular signature of sarcopenia. Physiol Genomics 21:253–263 11. Haslett JN, Sanoudou D, Kho AT, Bennett RR, Greenberg SA, Kohane IS, Beggs AH, Kunkel LM (2002) Gene expression comparison of biopsies from Duchenne muscular dystrophy (DMD) and normal skeletal muscle. Proc Natl Acad Sci USA 99:15000–15005 12. Panguluri SK, Bhatnagar S, Kumar A, McCarthy JJ, Srivastava AK, Cooper NG, Lundy RF, Kumar A (2010) Genomic profiling of messenger RNAs and microRNAs reveals potential

mechanisms of TWEAK-induced skeletal muscle wasting in mice. PLoS One 5:e8760 13. Stevenson EJ, Giresi PG, Koncarevic A, Kandarian SC (2003) Global analysis of gene expression patterns during disuse atrophy in rat skeletal muscle. J Physiol 551:33–48 14. Delgado I, Huang X, Jones S, Zhang L, Hatcher R, Gao B, Zhang P (2003) Dynamic gene expression during the onset of myoblast differentiation in vitro. Genomics 82:109–121 15. Rajeevan MS, Ranamukhaarachchi DG, Vernon SD, Unger ER (2001) Use of real-time quantitative PCR to validate the results of cDNA array and differential display PCR technologies. Methods 25:443–451

Chapter 19 Analysis of Lipid Profiles in Skeletal Muscles Vassilis Mougios and Anatoli Petridou Abstract The lipidome of skeletal muscles is a worthwhile target of research, as it affects a multitude of biological functions, and is, in turn, affected by factors such as diet, physical activity, and development. We present two methods for the analysis of the main lipid classes in skeletal muscles of humans and other animals, that is, triacylglycerols and phospholipids. The methods differ in that the former concerns total phospholipids, while the latter concerns individual phospholipids. In both methods, lipids are extracted from muscle after the addition of internal standards, and they are separated by one-dimensional (1D) thin-layer chromatography (TLC). This is sufficient for the separation of triacylglycerols and total phospholipids. In the first method, the two classes are subsequently subjected to methanolysis to produce methyl esters of fatty acids (and, to a lesser extent, dimethyl acetals of fatty aldehydes derived from plasmalogens), which are analyzed by gas chromatography (GC). Quantitation is achieved on the basis of the internal standards. In the second method, 1D TLC is used for the analysis of triacylglycerols only, whereas individual phospholipids are separated by two-dimensional TLC. This results in the isolation of phosphatidyl choline, lysophosphatidyl choline, phosphatidyl ethanolamine, phosphatidyl serine, phosphatidyl inositol, cardiolipin, and sphingomyelin. Methanolysis and subsequent analysis by GC results in the determination of the fatty acid and aldehyde profiles of the individual muscle phospholipids. Key words: Dimethyl acetals, Fatty acids, Gas chromatography, Lipids, Methyl esters, Phospholipids, Plasmalogens, Skeletal muscle, Thin-layer chromatography, Triacylglycerols

1. Introduction Lipids are integral components of all cells. The plasma membrane forming the boundary of a cell with its surroundings is basically lipid (in particular, phospholipid) in nature. The same holds true for all membranes delimiting the subcellular organelles of eukaryotic cells (the nucleus, mitochondria, endoplasmic reticulum, Golgi apparatus, lysosomes, peroxisomes, etc.) and specialized structures like the sarcoplasmic reticulum of muscle fibers and the disks of the

Joseph X. DiMario (ed.), Myogenesis: Methods and Protocols, Methods in Molecular Biology, vol. 798, DOI 10.1007/978-1-61779-343-1_19, © Springer Science+Business Media, LLC 2012

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rod outer segment. Lipids, mainly in the form of triacylglycerols, are stored in cells of several types, including muscle fibers, hepatocytes, and adipocytes, the latter consisting of triacylglycerols by about 80%. In addition, lipids are present in biological fluids. For example, the blood plasma hosts fatty acids, triacylglycerols, phospholipids, and sterols, transporting them to tissues. Lipid analysis presents a challenge emanating from the great heterogeneity of a class of biological compounds united only by their poor solubility in water. Indeed, there is little chemical similarity between, say, a triacylglycerol and cholesterol. (By contrast, a protein will always consist of amino acid residues that are variations on a theme, and nucleic acids will always consist of nucleotide residues that are variations on another theme). Hence, one needs a variety of methods to study the full lipid complement in a biological sample. The aim of this chapter is to provide detailed methods for the analysis of the main lipid classes in the skeletal muscles of humans and other animals, that is, triacylglycerols and phospholipids, both of which are affected by factors such as diet (1), physical exercise (2), and development (3, 4), and both of which, in turn, affect a multitude of biological functions, including ion homeostasis, gene expression, and signal transduction (5, 6) (see Note 1). Two alternative methods are described below (see Fig. 1). In both, lipids are extracted from muscle with a powerful organic solvent (chloroform-methanol, 2:1), followed by the addition of water that results in the formation of two phases, one containing the lipids Analysis of triacylglycerols and total phospholipids

Analysis of triacylglycerols and individual phospholipids

Lipid extraction from tissue

Lipid extraction from tissue

1D TLC

1D TLC

2D TLC

Isolation of triacylglycerols and phospholipids

Isolation of triacylglycerols

Isolation of individual phospholipids

Methanolysis

Methanolysis

Methanolysis

Analysis of fatty acid methyl esters by GC

Analysis of fatty acid methyl esters by GC

Analysis of fatty acid methyl esters by GC

Fig. 1. Major steps in the two methods of analysis of lipid profiles in skeletal muscles described in this chapter. 1D one-dimensional; 2D two-dimensional; GC gas chromatography; TLC thin-layer chromatography.

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and another containing nonlipid constituents (7). Then, in the first method, triacylglycerols (represented schematically in Fig. 2a) and phospholipids (see Fig. 2b–j) as a whole are separated from each other and from minor lipid constituents of skeletal muscles by one-dimensional (1D) thin-layer chromatography (TLC). The two a

b Acyl group Acyl group

Acyl group

c

Glycerol unit

Acyl group

Glycerol unit

Acyl group

Phosphate

Choline

group

unit

Phosphate

Choline

group

unit

d 1-Alkenyl group Phosphate

Choline

group

unit

Acyl group

e

Glycerol unit

Glycerol unit

Acyl group

f 1-Alkenyl group Phosphate

Ethanolamine

group

unit

g

Acyl group

Glycerol unit

Acyl group

Glycerol unit

Acyl group

Phosphate

Ethanolamine

group

unit

h Acyl group Phosphate group

Acyl group

Serine unit

Glycerol unit

Acyl group

Glycerol unit

Acyl group

Phosphate

Inositol

group

unit

i

group

Phosphate group

Glycerol unit

Phosphate

Glycerol unit

Acyl group

Glycerol unit

Acyl group

Acyl group Acyl group

j Sphingosine unit

Acyl group

Phosphate

Choline

group

unit

Fig. 2. Schematic representation of skeletal muscle lipids determined by the methods described in this chapter. (a) Triacylglycerol, (b) phosphatidyl choline, (c) lysophosphatidyl choline, (d) phosphatidal choline (a plasmalogen), (e) phosphatidyl ethanolamine, (f) phosphatidal ethanolamine (a plasmalogen), (g) phosphatidyl serine, (h) phosphatidyl inositol, (i) cardiolipin, (j) sphingomyelin.

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fractions are subsequently subjected to methanolysis to produce methyl esters (MEs) of fatty acids (and, to a lesser extent, dimethyl acetals, DMAs, of fatty aldehydes derived from plasmalogens), which are analyzed by gas chromatography (GC). Internal standards added at the beginning of the method compensate for losses during the entire procedure. As internal standards, we use a triacylglycerol and a phospholipid with acyl groups (hepatadecanoyl, abbreviated 17:0, see Note 2) that are absent from the natural muscle triacylglycerols and phospholipids. The internal standards comigrate with the natural lipids of the same class during TLC and, upon methanolysis, yield methyl heptadecanoate (17:0 ME), which serves as reference for calculating the amounts of the endogenous MEs and DMAs after they are separated by GC. The method outlined above is useful if one is content with determining the fatty acid composition of total phospholipids. However, an added complexity of the lipidome is diversity within the class of phospholipids. This diversity stems from: 1. The presence of either of two possible alcohols, namely, glycerol (see Fig. 2b–i) and sphingosine (see Fig. 2j), as the backbones to which the acyl groups and phosphate are linked. 2. The presence of either of several alcohols, such as choline (see Fig. 2b–d, j), ethanolamine (see Fig. 2e, f), serine (see Fig. 2g), and inositol (see Fig. 2h), at the end of the polar head group. 3. The unusual structure of cardiolipin (CL) (see Fig. 2i), being almost a diphosphatidyl glycerol. 4. The presence of lysophospholipids (see Fig. 2c), that is, phospholipids lacking one acyl group. 5. The presence of ether, rather than ester, linkages in certain phospholipids (see Fig. 2d, f). If one is then interested in separating all of the individual phospholipids that are present in skeletal muscles in appreciable amounts, one has to resort to two-dimensional (2D) TLC, followed by methanolysis of each phospholipid and by GC of the resulting MEs and DMAs. These procedures constitute the second method. Description of the two methods is preceded by instructions on how to establish the method for the gas chromatographic analysis of MEs and DMAs, which is a prerequisite for both.

2. Materials Use analytical grade reagents. To prevent contamination of reagents and samples by lipids present on the skin and to protect hands from hazardous organic solvents, wear gloves throughout all steps

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of the analysis. Avoid inhaling vapors of organic solvents. Observe the regulations for the safe disposal of waste materials. 2.1. Gas Chromatography

1. Gas chromatograph equipped with flame ionization detector. Carrier gas may be helium or hydrogen. 2. Column. Many commercially available capillary GC columns that are suitable for analysis of fatty acid MEs will do. We recommend a length of 30 m, an internal diameter of 0.25 mm, and a film thickness of 0.25 μm. 3. Analytical balance displaying four decimal points of the gram. 4. Glass screw top vials, 2 and 4 mL, along with perforated (open top) screw caps and PTFE/silicone septa. 5. Hexane. 6. ME standards. We have found 22 MEs to be present at appreciable amounts (that is, at least 0.1% of total) in skeletal muscle triacylglycerols and phospholipids. These and 17:0 ME (derived from methanolysis of the internal standards) are listed in Table 1, along with their relative molecular masses (Mr) to aid you in the calculations (see Subheadings 3.2.3 and 3.3.4). Obtain all MEs at the lowest available quantities. Make a stock solution of each ME standard at an approximate concentration of 4 mg/mL (no need to be accurate) by weighing 4–8 mg inside a 2-mL screw top vial and dissolving in 1–2 mL of hexane. Cover the vial with a septum (glossy side facing the rim of the vial) and cap tightly. Use the standard to establish the retention time of the ME in your gas chromatographic system as described in Subheading 3.1. Store at −20°C (see Note 3). 7. Hexadecanal dimethyl acetal standard. Hexadecanal dimethyl acetal (16:0 DMA, Mr 286.5) is produced by methanolysis of plasmalogens (see Fig. 1d, f) carrying a 1-hexadecenyl group (see Note 4). Make a 4 mg/mL stock solution in hexane and store as described above (see Note 5). 8. Syringe, 10 μL, with pointed needle.

2.2. Analysis of Triacylglycerols and Total Phospholipids

1. Liquid nitrogen, mortar, and pestle for tissue pulverization. 2. Glass test tubes, small (3–5 mL). Use new tubes in each case and discard the used ones, as washing them with organic solvents to remove their lipids may be more expensive than buying them (let alone the effort). 3. Glass Pasteur pipettes and rubber bulb. 4. Glass screw top test tubes, small (5–7 mL), with PTFE-lined screw caps. 5. Glass funnel, small (fitting the opening of the screw top test tubes).

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Table 1 Methyl esters usually detected by gas chromatographic analysis of skeletal muscle lipids Name

Abbreviation of acyl group

Mr of methyl ester

Methyl laurate

12:0

214.4

Methyl myristate

14:0

242.4

Methyl myristoleate

14:1ω9

240.4

Methyl palmitate

16:0

270.5

16:1ω7

268.4

Methyl heptadecanoate

17:0

284.5

Methyl stearate

18:0

298.5

Methyl oleate

18:1ω9

296.5

Methyl cis-vaccenate

18:1ω7

296.5

Methyl linoleate

18:2ω6

294.5

Methyl γ-linolenate (all-cis-6,9,12)

18:3ω6

292.5

Methyl α-linolenate (all-cis-9,12,15)

18:3ω3

292.5

Methyl stearidonate

18:4ω3

290.5

Methyl arachidate

20:0

326.5

Methyl eicosenoate

20:1ω9

324.5

Methyl dihomo-γ-linolenate

20:3ω6

320.5

Methyl arachidonate

20:4ω6

318.5

Methyl 5,8,11,14,17eicosapentaenoate

20:5ω3

316.5

Methyl behenate

22:0

354.5

Methyl all-cis-7,10,13,16docosatetraenoate

22:4ω6

346.5

Methyl all-cis-4,7,10,13,16docosapentaenoate

22:5ω6

344.5

Methyl all-cis-7,10,13,16,19docosapentaenoate

22:5ω3

344.5

Methyl all-cis-4,7,10,13,16,19docosahexaenoate

22:6ω3

342.5

Methyl palmitoleate a

The methyl esters detected in a muscle depend on species and diet. Certain methyl esters may be only available by the name [fatty acid] methyl ester, for example, cis-vaccenic acid methyl ester, rather than methyl cis-vaccenate a Derived from methanolysis of internal standards

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6. Syringe, 10 μL, with blunt needle. 7. Scalpel with round blade. 8. Organic solvents: chloroform, methanol, ethanol (absolute), petroleum ether (boiling point range, about 40–60°C), diethyl ether, acetic acid. 9. Washing solvent: chloroform-methanol 2:1 (v/v). 10. Lipid extraction solvent: chloroform-methanol 2:1 (v/v) with 0.005% (w/v) 2,6-di-tert-butyl-4-methylphenol (aka butylated hydroxytoluene, or BHT) to prevent lipid peroxidation. 11. Triheptadecanoyl glycerol (triheptadecanoin) as triacylglycerol internal standard (abbreviated as 17:0 TG). 12. Diheptadecanoyl phosphatidyl choline (17:0 PC) as phospholipid internal standard (see Note 6). 13. Triacylglycerol and phospholipid internal standard solutions: Weigh out about 20 mg of each 17:0 TG and 17:0 PC into separate 4-mL screw top vials. Note weight to the fourth decimal point of the gram and place the vials on ice. Prepare two screw caps with septa in place. Then add 3 mL of cold chloroform to each vial and cap immediately and tightly (see Note 7). This will give two standard solutions of about 7 mg/mL, or 7 μg/μL, concentration. For greater accuracy, take into account the purity of the substances used (usually 99%). For the sake of subsequent calculations, let a and b be the concentrations of the two standard solutions, respectively. Store at −20°C. Stable for 1 year. 14. High-performance TLC plates of silica gel. Plates usually come in boxes of 25. Plate dimensions can be 10 × 10, 10 × 20, or 20 × 20 cm depending on the number of samples to be analyzed in each run. We recommend plates that are 20 cm on at least one dimension if you intend to run more than ten samples simultaneously. Another parameter to consider is the silica gel’s support: it may be glass, plastic, or aluminum. The main advantage of plastic and aluminum is that they can be cut to the desired dimensions with scissors. If you use a 20 × 20 cm plate with glass support, at least half of it will be wasted, since the developer needs only migrate by 10 cm (see Note 8). Handle TLC plates by their sides and supports. Do not touch the delicate silica gel surface. 15. Spotting guide: On a sheet of paper, draw a straight line parallel to one side at a distance of 1.5 cm. On that line, make 12 black dots with a marker pen that are 1.5 cm apart, starting 1.5 cm from one end. Number the dots (see Fig. 3). 16. TLC tank. Choose tank dimensions to accommodate the plates you have chosen. 17. Multiplate rack, optional.

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1.5 cm

1.5 cm

1

2

3

4

5

6

7

8

9

10

11

12

Fig. 3. Makeshift spotting guide for TLC.

18. Nitrogen gas, industrial grade. 19. Dichlorofluorescein spray reagent: 0.2% (w/v) dichlorofluorescein in ethanol. Make 100 mL at a time and store indefinitely at room temperature. 20. Spray bottle. 21. Spray box to prevent dichlorofluorescein from staining the lab’s surfaces. 22. Hair dryer, optional. 23. UV lamp. 24. Eye goggles against UV radiation. 25. Methanolic sodium methoxide, 0.5M. 26. Methanolic boron trifluoride, 10–15% (w/w) (1.3–2M). 27. Heating block reaching 100°C. A water bath reaching 50°C and a boiling water bath will do instead. 28. Microvolume inserts (if the gas chromatograph is equipped with autosampler). 2.3. Analysis of Triacylglycerols and Individual Phospholipids

You will need all materials described under Subheading 2.2. You will need a second TLC tank (see Subheading 2.2, item 16) for 2D TLC. Because 2D TLC requires 10 × 10 cm, we do not recommend TLC plates of other dimensions (10 × 20 or 20 × 20 cm) with glass support. Additionally, obtain the following: 1. Acetone. 2. Phospholipid standards: We have found seven phospholipids to be present at appreciable amounts (that is, at least 1% of total) in skeletal muscle: PC, lysophosphatidyl choline (LPC), phosphatidyl ethanolamine (PE), phosphatidyl serine (PS), phosphatidyl inositol (PI), CL, and sphingomyelin (SM). Ideally, they should all contain the heptadecanoyl group in order to be used as internal standards (just like 17:0 PC, already listed as Subheading 2.2, item 12). However, 17:0 LPC, PE, PS, PI, CL,

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and SM are either not commercially available or too expensive. Therefore, obtain LPC, PE, PS, PI, CL, and SM without regard to their acyl groups for use only in establishing the separation pattern of muscle phospholipids under 2D TLC (quantitation without internal standards will be described below). Make a 7 μg/μL solution of each in chloroform just like you did with 17:0 PC, although you do not need to be accurate with these. Store indefinitely at −20°C. 3. Methyl pentadecanoate (15:0 ME) to be used as external standard. Make a solution of approximately 2 mg/mL (no need to be accurate) by weighing 4–8 mg inside a 4-mL screw top vial and dissolving in 1–2 mL of hexane. Cap and store indefinitely at −20°C.

3. Methods 3.1. Gas Chromatography

In order to be able to go directly from the separation of lipid classes (by either 1D or 2D TLC) to the preparation and separation of MEs and DMAs by GC, you need to establish the operating parameters of the gas chromatograph (i.e., temperature program of the column, head pressure or flow rate of the carrier gas, and split ratio) in advance. Since GC is a demanding technique, we assume that the person who will carry out this analysis is familiar with the basic theory of GC and the operation of a gas chromatograph. Therefore, we will not provide instructions on, for example, how to install a column, how to set up gas flow rates, how to set up a temperature program, or how to inject a sample. Besides, these functions depend greatly on the particular instrument available. 1. You may begin with the operating parameters suggested by the manufacturer of the column you have obtained for the specific application (fatty acid ME analysis) or with the following parameters that we use with columns fitting the description under Materials (see Subheading 2.1, item 2): column temperature, 160–250°C at 5°C/min, then isothermic at 250°C for 10 min; flow rate of carrier gas, 1 mL/min; split ratio, 1:10. 2. Prepare a series of ME and DMA working solutions by diluting 3 μL of each ME and DMA stock solution with 200 μL of hexane in a 2-mL screw top vial (see Note 9). Cap and store at −20°C (see Note 3). 3. Inject 1 μL of each ME and DMA working solution separately with a 10-μL syringe with pointed needle either manually or through an autosampler. 4. Note the retention time of each ME and DMA. If necessary, adjust the operating parameters to achieve a distinct retention time for each component (see Note 10).

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5. Confirm that all MEs and 16:0 DMA are adequately separated by mixing 3 μL of each stock solution in a 2-mL screw top vial, adding hexane to 200 μL, and injecting 1 μL. Store the mixture at −20°C for possible future use. 3.2. Analysis of Triacylglycerols and Total Phospholipids 3.2.1. Lipid Extraction and Separation

1. Obtain and store muscle tissue specimens in a way that prevents lipid modification before analysis. Major threats are triacylglycerol and phospholipid hydrolysis and fatty acid oxidation. We recommend immersing the specimen in liquid nitrogen immediately after obtaining it, pulverizing with mortar and pestle in liquid nitrogen, and storing at −80°C (see Note 11). 2. Start the analysis by placing a TLC tank, with its lid on, in a place with constant room temperature and not exposed to air drafts (not in the fume hood). 3. Prepare the developer for TLC: If using a large tank (one that accommodates 20-cm wide plates), decant 86 mL of petroleum ether, 14 mL of diethyl ether, and 1 mL of acetic acid in a 250-mL conical glass flask under the fume hood operating at full speed. If using a small tank (one that accommodates 10-cm wide plates), mix 43 mL of petroleum ether, 7 mL of diethyl ether, and 0.5 mL of acetic acid in a 100-mL flask (see Note 12). 4. Mix well with a swirling motion and promptly pour off into the TLC tank after sliding the lid sideways just enough for the liquid to enter. Replace the lid immediately and turn the fume hood’s motor off. Grasp the tank with both hands and, while holding the lid in place with your index fingers, shake the tank from one side to the other for a few seconds to facilitate saturation of its atmosphere with developer vapors. We do not recommend lining the tank with filter paper. Let the tank stand no less than 1 h and no more than 2 h before chromatography to ensure reproducible separations (see Note 13). In the meantime, proceed to lipid extraction from the samples (see Note 14). 5. Prepare a mixture of the 17:0 TG and 17:0 PC standard solutions by mixing one volume of the former with four volumes of the latter in a 2-mL screw top vial kept on ice and capping immediately and tightly. You will need 5 μL of this mixture per sample, but, in any case, make at least 100 μL to protect the mixture’s composition against evaporation in the vial if the volume is too low. Keep on ice and discard at the end of the day. 6. Using a 10-μL syringe with pointed needle, pierce the septum of the vial and dispense 5 μL of the mixture (in effect, 1 μL of the 17:0 TG standard solution plus 4 μL of the 17:0 PC standard solution) at the bottom of as many small glass test tubes as the muscle samples you are going to analyze. Wait a minute for

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the solvent to evaporate before proceeding to the next step. Label the test tubes according to the muscle samples. 7. Take a muscle specimen out of the ultrafreezer, place the corresponding test tube in the analytical balance, tare, and promptly weigh out approximately 30 mg. Note weight to the third or, preferably, fourth decimal point of the gram in order to be able to express the lipid content per gram tissue with high accuracy (see Note 15). Return any remaining part of the specimen to the ultrafreezer. 8. Immediately add 570 μL of lipid extraction solvent [chloroformmethanol 2:1 (v/v) with 0.005% (w/v) BHT]. Vortex briefly and let stand for 5 min or longer with occasional vortexing (see Note 16). In the meantime, you may proceed with other muscle specimens. 9. Add 120 μL of distilled water and vortex vigorously for 1 min. 10. Briefly spin the test tube in a centrifuge to produce two clear phases with the muscle debris at the interphase. Lipids are contained in the lower phase. Handle the test tube gently so as not to disturb the separation of phases. 11. Attach a rubber bulb to a glass Pasteur pipette and squeeze the bulb almost completely. Immerse the tip of the pipette to the bottom of the test tube, taking care not to disturb the upper phase and interphase. Squeeze the bulb gently until you release one air bubble in order to push out any upper phase that has entered the tip of the pipette during immersion. Then gently release the pressure on the bulb to aspirate the lower phase. Take up as much of the lower phase as possible, but do not aspirate any of the muscle debris or upper phase. 12. Transfer the aspirate to another small glass test tube and evaporate the solvent under a stream of nitrogen (see Note 17). Discard the test tube containing the muscle debris and upper phase. This ends the extraction process. 13. If using a 20 × 20-cm silica gel plate with soft support, cut it in half with scissors carefully, taking care not to chip off too much of the silica gel layer. Use the cut sides of the two resulting plates as the far ends in the subsequent chromatography, that is, the ends toward which the developer migrates (see Note 18). If using any other kind of TLC plate, skip this step. If using a 20 × 20-cm plate with glass support, draw a line with ruler and pencil in the middle to mark the end of development. 14. You will need 10 cm along one side of the TLC plate(s) for the developer to migrate and (n + 1) × 1.5 cm along the other side, n being the number of samples to be spotted 1.5 cm apart and 1.5 cm from the sides. Accordingly, you may wish to (further) cut a plate for economy.

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15. Place the spotting guide on the bench. If using a plate with transparent support (that is, glass or plastic), place the plate on the spotting guide so that the sides of the two coincide. Now you can see the dots on the spotting guide under the plate. If using a plate with aluminum support, place the plate on the spotting guide so that the left sides of the two are aligned and the dots on the spotting guide barely appear along the bottom side of the plate. Then mark 1.5 cm from the bottom side on the left and right sides of the plate with a pencil. Thus, you will have the coordinates of where to spot. 16. Dissolve the dried lipid extract of the first sample in 30 μL of lipid extraction solvent. Make sure you retrieve all of the extract from the wall of the test tube by vortexing and rotating the test tube in your hands almost horizontally. 17. Draw 10 μL of the dissolved extract in a 10-μL syringe with blunt needle and bring the syringe over the first spotting position, the tip of the needle being a few millimeters above the plate. 18. Gently squeeze the plunger to create a medium-sized drop (of about 0.7 μL) and carefully lower the tip of the needle until the drop (not the needle) touches the plate and is absorbed by the silica gel. 19. Wait a few seconds until all of the solvent evaporates from the plate, then repeat the previous step until all of the syringe’s contents are spotted. Spot each drop on top of the previous one so that the spot formed does not exceed a few millimeters in diameter. You will find that it takes longer and longer for each drop to evaporate. You may speed up the evaporation by setting up a hair dryer next to the plate and directing the air at the spot (see Note 19). 20. By the end of spotting, a yellow-brownish spot will have formed on the plate. Then proceed to dissolving and spotting the next extract(s) by repeating steps 16–19 until you fill all available spotting positions on the plate. The first time you perform this analysis, spot 2 μL of each 17:0 TG and 17:0 PC standard solutions on separate positions in the middle of the plate in order to identify the triacylglycerol and phospholipid spots after chromatography. During subsequent analyses, you may spot only the 17:0 TG standard, since, as you will find out, phospholipids remain at the origin and are thus easily located. 21. Save the remaining extract in each test tube by covering the test tube with Parafilm and storing at 4°C, just in case you need to repeat the analysis. Stable for 1 month. 22. If you spot many samples, keep a record of which sample lies at which spot on the plate to avoid mix-up.

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23. If you have so many samples that you need additional plates, repeat steps 15–22. Unless you use a multiplate rack (see Note 20), you may develop up to two plates in a TLC tank. Mark each plate lightly with pencil by alphabet letters at the top to avoid mix-up. 24. Once spotting is over, bring the plate(s) next to the TLC tank. Hold the (first) plate by the middle of its far end with one hand and raise it so that it hovers horizontal above the TLC tank. Open the lid and lower the plate into the tank at a distance of 1–2 cm from one of its two long walls. When the plate touches the surface of the developer lower it a bit more until it lands on the bottom and let it rest gently against the wall, making sure it does not tip over (see Note 21). 25. If you have another plate to develop, repeat the previous step on the opposite wall of the tank immediately. Do not turn the tank around! Then promptly replace the lid. It is important not to let too much of the developer’s vapor escape the tank. You may wear a mask while the tank is open to avoid inhaling the fumes. However, do not turn the fume hood’s motor on! 26. The developer will rise (initially fast, then slower) on the plate, carrying the sample spots with it. When the developer reaches about one-half centimeter from the top of the plate(s) or from the pencil line you drew on the 20 × 20-cm plate(s) with glass support (in about 20 min), turn the fume hood’s motor on, open the lid of the tank, and pull out the plate(s). Place each plate, face up, under the fume hood and let it dry for about 15 min. In the meantime, dispose of the developer in the tank properly and let the tank dry under the fume hood. 27. Turn the fume hood’s motor off. Place the spray box inside the fume hood and place the plate nearly vertical (silica gel facing you) inside the spray box. Fill the spray bottle with the dichlorofluorescein spray reagent (see Note 22) and spray the plate evenly from a distance of about 25 cm, making sure it acquires a faint orange color without getting overly wet (see Note 23). 28. Turn the fume hood’s motor back on and let the plate dry completely (another 15 min). Bring the plate to the darkroom (or to a fairly dark place), put protective eye goggles on, and view the plate under a UV lamp. Bright yellow fluorescent spots will appear on a dark background, corresponding to the lipids present in each sample. Phospholipids will have remained at the origin, while triacylglycerols will have migrated halfway to the top. 29. Mark the contour of the phospholipid and triacylglycerol spots in each muscle sample with pencil, giving a slack of about 1 mm all around, and take the plate back to the lab.

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30. Prepare and label two screw top test tubes per muscle sample, one for triacylglycerols and another for phospholipids. 31. Fit a glass funnel on top of the first test tube and, using a scalpel with round blade, carefully scrape the phospholipid spot of the first muscle sample into the funnel. Shake the funnel lightly and then apply a gentle stream of nitrogen from all around the rim down to the neck to make sure that all silica gel flakes get to the bottom of the test tube. 32. Remove the funnel from the first test tube and clean it with a stronger stream of nitrogen to make sure no silica gel is carried over to the next test tube (see Note 24). Wipe the scalpel thoroughly with tissue paper for the same purpose. 33. Fit the funnel onto the next test tube and proceed with the phospholipid spot of the second muscle sample by repeating the previous two steps. Continue with the remaining samples. Then scrape off the triacylglycerol spots of all samples into their respective test tubes in the same way (see Note 25). Discard the TLC plate. 3.2.2. Preparation and Analysis of Methyl Esters and Dimethyl Acetals

1. To each screw top test tube containing triacylglycerols or phospholipids, add 0.5 mL of methanolic sodium methoxide and cap tightly. The liquid turns yellow, as it extracts the dichlorofluorescein from the silica gel. Vortex and heat at 50°C for 10 min (see Note 26). 2. Let all test tubes cool. To the ones containing phospholipids, add 0.5 mL of methanolic boron trifluoride, cap tightly, and heat at 100°C for 75 min (see Note 27). 3. Let the test tubes containing phospholipids cool. 4. Open all test tubes, taking care not to mix up their caps. To each test tube, add 1.5 mL of hexane, cap tightly, and vortex at full speed or shake vigorously for 1 min to extract the MEs and DMAs. 5. Let the test tubes stand for a few minutes and watch a sharp interphase form between the lower methanol phase (containing the silica gel and dichlorofluorescein) and the upper hexane phase containing the MEs and DMAs. Spin the test tubes briefly in a centrifuge if you are in a hurry. 6. Using a Pasteur pipette, remove as much of the upper phase as possible into a small glass test tube. Do not take any of the lower phase! Evaporate under a stream of nitrogen as in Subheading 3.2.1, step 12. Alternatively, this may be a good time to call it a day, especially if you are not in a hurry to start the ME and DMA analysis. In this case, you may just leave the test tubes overnight under the fume hood and let the hexane evaporate effortlessly.

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7. Contrary to plain test tubes, screw top test tubes and their caps are too expensive to be used only once. Therefore, rinse the tubes containing the lower methanol phase and silica gel thoroughly with tap water and then with chloroform-methanol 2:1. Let dry for future use. Treat the screw caps likewise. 8. Turn the gas chromatograph on. If you use manual injection, dissolve the dry residue in each test tube containing MEs from triacylglycerols in 50 μL of hexane just before injection. Mix thoroughly and inject 1 μL. 9. If you use an autosampler, dissolve the dry residue in each test tube containing MEs from triacylglycerols in 50 μL of hexane. Mix thoroughly and, using an automatic pipette, transfer the solution into a microvolume insert sitting inside a 2-mL screw top vial. Close the vial with septum and screw cap, place it in the autosampler and inject 1 μL (see Note 28). 10. Dissolve the dry residue in each test tube containing MEs and DMAs from phospholipids in 200 μL of hexane and inject 1 μL as above (see Note 29). 11. Use an appropriate software to acquire the gas chromatogram. Based on the retention times you have established in Subheading 3.1, assign the peaks in the chromatogram to MEs and DMAs and integrate the peaks so that you get a table containing an area value (in arbitrary units) for each identified ME and DMA. Depending on the capabilities of the data acquisition software, this can be done automatically, manually, or in part automatically and in part manually. 3.2.3. Calculations

Conversion of the peak integration data into muscle content values (that is, μmol lipid per gram muscle) can be done through either the data acquisition software of the gas chromatograph or a spreadsheet (such as Microsoft® Excel). Described below is the reasoning that has to be followed in any case. 1. Calculations are based on the premise that peak area is proportional to ME mass. Of pivotal importance, then, are the amount and area of 17:0 ME derived from methanolysis of the 17:0 TG and 17:0 PC internal standards that were added to the muscle samples in the beginning of the analysis (see Note 30). On the basis of proportionality, if M17 and A17 are the amount and area, respectively, of 17:0 ME and A is the area of an endogenous ME, then its amount, M, is given by the formula, M = M17 × A/A17. Let’s apply this calculation separately to the triacylglycerols and phospholipids of a muscle sample. 2. Begin with triacylglycerols. To calculate the amount of 17:0 ME derived from 17:0 TG that is present in the muscle triacylglycerol fraction, first multiply the 17:0 TG standard concentration,

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a (in μg/μL), by the volume added to each sample, that is, 1 μL (see Subheading 3.2.1, step 6), yielding a μg. Divide by the Mr of 17:0 TG (849.4) and get a/849.4 μmol. Since 1 mol of a triacylglycerol yields 3 mol of MEs, one gets 3a/849.4 μmol of 17:0 ME. Finally, by multiplying by the Mr of 17:0 ME (284.5), one gets 1.005a μg. This is M17. 3. Apply the formula, M = 1.005a × A/A17, to every ME identified in the chromatogram and get the amount of each ME in μg. Then divide by that ME’s Mr to express its amount in μmol, which is also μmol of the corresponding acyl group (or, more commonly, fatty acid) in muscle triacylglycerols. 4. You may divide the amount of each fatty acid by the amount of muscle tissue (either wet or dry) weighed in Subheading 3.2.1, step 7 to express the fatty acid content of muscle triacylglycerols in μmol/g or μmol/mg. 5. You may add the amounts of all fatty acids and divide the sum by 3 to get the triacylglycerol content of muscle in μmol/g or μmol/mg. 6. You may divide the amount of each fatty acid by the sum of fatty acids and then multiply by 100 to express the percentage molar distribution of fatty acids in muscle triacylglycerols (see Note 31). 7. You may use partial sums to calculate the amounts or percentages of fatty acid categories, such as saturated, unsaturated, ω6, etc. 8. Continue with phospholipids. To calculate the amount of 17:0 ME derived from 17:0 PC that is present in the muscle phospholipid fraction, first multiply the 17:0 PC standard concentration, b (in μg/μL), by the volume added to each sample, that is, 4 μL (see Subheading 3.2.1, step 6), yielding 4b μg. Divide by the Mr of 17:0 PC (762.2) and get 4b/762.2 μmol. Since 1 mol of PC yields 2 mol of MEs, one gets 8b/762.2 μmol of 17:0 ME. Finally, by multiplying by the Mr of 17:0 ME (284.5), one gets 2.986b μg. 9. Apply the formula, M = 2.986b × A/A17, to every ME identified in the chromatogram and get the amount of each ME in μg. Then divide by that ME’s Mr to express its amount in μmol, which is also μmol of the corresponding fatty acid in muscle phospholipids. 10. Apply the same formula to every DMA identified in the chromatogram and get the amount of each DMA in μg. Then divide by that DMA’s Mr to express its amount in μmol, which is also μmol of the corresponding fatty aldehyde in the muscle phospholipids. 11. You may divide the amount of each fatty acid (and aldehyde) by the amount of muscle tissue used in the analysis to express the fatty acid (and aldehyde) content of muscle phospholipids.

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12. You may add the amounts of all fatty acids and aldehydes, then divide the sum by 2 to get the approximate phospholipid content of muscle. “Approximate” refers to the fact that most, but not all, phospholipids contain two acyl groups in their structure (see Fig. 2). To get the exact phospholipid content of muscle, you need to know the amount of each individual phospholipid, which is determined in Subheading 3.3. 13. You may divide the amount of each fatty acid and aldehyde by the sum of fatty acids and aldehydes, then multiply by 100 to express the percentage molar distribution of fatty acids and aldehydes in muscle phospholipids (see Note 31). 14. You may use partial sums to calculate the amounts or percentages of fatty acid and aldehyde categories, such as saturated, unsaturated, ω6, etc.

3.3.1. Establishing the Separation Pattern of Phospholipids in 2D TLC

In order to be able to go directly from lipid extraction to phospholipid separation by 2D TLC, you need to establish the migration pattern of each phospholipid in advance. Although we provide such a pattern in Fig. 4, we strongly advise that you establish your own pattern. 1. Place two TLC tanks, with their lids on, in a place with constant room temperature and not exposed to air drafts (not in the fume hood). Number the tanks, 1 and 2.

CL

PE

1st dimension

3.3. Analysis of Triacylglycerols and Individual Phospholipids

PI

SM

PS

PC

LPC Origin

2nd dimension

Fig. 4. Separation pattern of muscle phospholipids by 2D TLC. First dimension, chloroformmethanol-acetic acid 10:5:1 (v/v/v); second dimension, chloroform-acetone-methanolacetic acid-water 10:4:2:2:1 (v/v/v/v/v) (9). CL cardiolipin; LPC lysophosphatidyl choline; PC phosphatidyl choline; PE phosphatidyl ethanolamine; PI phosphatidyl inositol; PS phosphatidyl serine; SM sphingomyelin.

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2. Prepare the developer for the first dimension of 2D TLC: If using a large tank (one that accommodates 20-cm wide plates), decant 60 mL of chloroform, 30 mL of methanol, and 6 mL of acetic acid in a 250-mL conical glass flask under the fume hood operating at full speed. If using a small tank (one that accommodates 10-cm wide plates), decant 30 mL of chloroform, 15 mL of methanol, and 3 mL of acetic acid in an 100-mL flask (see Note 12). 3. Mix well with a swirling motion and promptly pour off into tank #1 after sliding the lid sideways just enough for the liquid to enter. Replace the lid immediately and turn the fume hood’s motor off. Grasp the tank with both hands and, while holding the lid in place with your index fingers, shake the tank from one side to the other for a few seconds to facilitate saturation of its atmosphere with developer vapors. We do not recommend lining the tank with filter paper. Let the tank stand no less than 1 h and no more than 2 h before chromatography to ensure reproducible separations (see Note 13). 4. Prepare the developer for the second dimension of 2D TLC: If using a large tank, decant 50 mL of chloroform, 20 mL of acetone, 10 mL of methanol, 10 mL of acetic acid, and 5 mL of distilled water in a 250-mL flask under the fume hood operating at full speed. If using a small tank, mix 25 mL of chloroform, 10 mL of acetone, 5 mL of methanol, 5 mL of acetic acid, and 2.5 mL of distilled water in an 100-mL flask (see Note 32). 5. Mix and pour off into tank #2 just like you did with the developer for the first dimension. Let the tank stand no more than 2 h before chromatography (it will certainly stand 1 h). 6. You will need eight 10 × 10-cm TLC plates, that is, one for each of the seven phospholipids (PC, LPC, PE, PS, PI, CL, and SM) and one for the mixture of all seven. If your plates are not 10 × 10 cm, cut them carefully with scissors, taking care not to chip off too much of the silica gel layer. Use the cut sides of the resulting plates as the far ends in the subsequent chromatography, that is, the ends toward which the developer migrates (see Note 18). 7. Mix 120 μL of the 17:0 PC, 5 μL of the LPC, 40 μL of the PE, 5 μL of the PS, 10 μL of the PI, 15 μL of the CL, and 5 μL of the SM standard solutions (see Note 33) in a 2-mL screw top vial and cap promptly. 8. Place the spotting guide on the bench. If using a plate with transparent support (that is, glass or plastic), place the plate on the spotting guide so that the sides of the two coincide. Now you can see dot #1 on the spotting guide under the plate at its bottom left corner. If using a plate with aluminum support, place the plate on the spotting guide a bit to the right and up,

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so you can barely see the ends of the two perpendicular lines intersecting at dot #1. Thus, you will have the coordinates of where to spot. 9. Draw 10 μL of the phospholipid mixture in a 10-μL syringe with blunt needle and bring the syringe over the spotting position, the tip of the needle being a few millimeters above the plate. 10. Gently squeeze the plunger to create a medium-sized drop (of about 0.7 μL) and carefully lower the tip of the needle until the drop (not the needle) touches the plate and is absorbed by the silica gel. 11. Wait a few seconds until all of the solvent evaporates from the plate, then repeat the previous step until all of the syringe’s contents are spotted. Spot each drop on top of the previous one so that the spot formed does not exceed a few millimeters in diameter. You will find that it takes longer and longer for each drop to evaporate. You may speed up the evaporation by setting up a hair dryer next to the plate and directing the air at the spot (see Note 19). 12. Spot 5 μL of the 17:0 PC standard solution onto the next plate. If using small tanks, you may run two plates simultaneously, so stop here. If using large tanks you may run four plates simultaneously (two plates side by side on each long wall of the tank), so continue by spotting 5 μL of each LPC and PE standard solutions onto the next two plates. To avoid mix-up among plates, mark each one softly with pencil at its upper right corner. 13. Once spotting is over, bring the plates next to tank #1. Hold the first plate by the middle of its far end with one hand and raise it so that it hovers horizontal above the tank. Open the lid and lower the plate into the tank at a distance of 1–2 cm from one of its two long walls. When the plate touches the surface of the developer, lower it a bit more until it lands on the bottom and let it rest gently against the wall, making sure it does not tip over (see Note 21). 14. Place the second plate next to the first one (if you have a large tank) or opposite the first one (if you have a small tank). If you have a large tank, place the remaining plates on the opposite wall. Do not turn the tank around! Then promptly replace the lid. Work as fast as possible to minimize evaporation of the developer. You may wear a mask while the tank is open to avoid inhaling the fumes. However, do not turn the fume hood’s motor on! 15. The developer will rise (initially fast, then slower) on the plates, carrying the sample spots with it. When the developer reaches about one-half centimeter from the top of the plates

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(in about 35 min), turn the fume hood’s motor on, open the lid of the tank, and pull out the plates. Place the plates, face up, under the fume hood and let them dry for 20 min. In the meantime, dispose of the developer in the tank properly and let the tank dry under the fume hood. 16. Rotate the plates 90° counterclockwise and place them in tank #2 (just like you did with tank #1) so that now the origin lies at the bottom right corner. 17. When the developer in the second dimension reaches about one-half centimeter from the top of the plates (in another 35 min), turn the fume hood’s motor on, open the lid of the tank and pull out the plates. Let the plates dry for another 20 min, dispose of the developer, and let the tank dry. Then turn the fume hood’s motor off. 18. Repeat steps 1–17 with the remaining phospholipid standard solutions. 19. Place the spray box inside the fume hood and place the plates, one by one or two by two, nearly vertical (silica gel facing you) inside the spray box. Fill the spray bottle with the dichlorofluorescein spray reagent (see Note 22) and spray the plates evenly from a distance of about 25 cm, making sure they acquire a uniform faint orange color without getting overly wet (see Note 23). 20. Turn the fume hood’s motor back on and let the plates dry completely (about 15 min). Bring the plates to the darkroom (or to a fairly dark place), put protective eye goggles on, and view the plates under a UV lamp. 21. Seven bright yellow fluorescent spots of different shapes, corresponding to the seven phospholipids in mixture, should be visible on a dark background on the first plate. Place a transparency (or other transparent material) over the plate and outline each phospholipid spot with permanent pen. Also mark the origin. 22. One fluorescent spot should be visible on each of the remaining seven plates, each spot at a different position on the plate. Place the transparency on each of these plates so that the origins coincide and identify the seven phospholipids. In the end, you should produce something like Fig. 4 (see Note 34). 3.3.2. Lipid Extraction and Separation

This section resembles Subheading 3.2.1 in the way lipids are extracted from muscle and separated by 1D TLC. However, from that separation, you will only need the triacylglycerol spot, since the individual phospholipids will be obtained from 2D TLC. To accurately measure the minor phospholipids, you will need a higher amount of tissue than that needed for the analysis of total phospholipids.

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1. Follow Subheading 3.2.1, steps 1–4. 2. Prepare a mixture of the 17:0 TG and 17:0 PC standard solutions by mixing three volumes of the former with seven volumes of the latter in a 2-mL screw top vial kept on ice and capping immediately and tightly. You will need 10 μL of this mixture per sample, but, in any case, make at least 100 μL to protect the mixture’s composition against evaporation in the vial if the volume is too low. Keep on ice and discard at the end of the day. 3. Using a 10-μL syringe with pointed needle, pierce the septum of the vial and dispense 10 μL of the mixture (in effect, 3 μL of the 17:0 TG standard solution plus 7 μL of the 17:0 PC standard solution) at the bottom of as many small glass test tubes as the muscle samples you are going to analyze. Wait a minute for the solvent to evaporate before proceeding to the next step. Label the test tubes according to the muscle samples. 4. Take a muscle specimen out of the ultrafreezer, place the corresponding test tube in the analytical balance, tare, and promptly weigh out approximately 90 mg. Note weight to the third or, preferably, fourth decimal point of the gram (see Note 15). Return any remaining part of the specimen to the ultrafreezer. 5. Immediately add 1,710 μL of lipid extraction solvent [chloroform-methanol 2:1 (v/v) with 0.005% (w/v) BHT]. Vortex briefly and let stand for 5 min or longer with occasional vortexing (see Note 16). In the meantime, you may proceed with other muscle specimens. 6. Add 360 μL of distilled water and vortex vigorously for 1 min. 7. Follow Subheading 3.2.1, steps 10–16. 8. Draw 3 μL of the dissolved extract in a 10-μL syringe with blunt needle and bring the syringe over the first spotting position. 9. Gently squeeze the plunger to create a medium-sized drop (of about 0.7 μL) and carefully lower the tip of the needle until the drop touches the plate and is absorbed by the silica gel. 10. Wait a few seconds until all of the solvent evaporates from the plate, then repeat the previous step until all of the 3 μL of extract are spotted. Spot each drop on top of the previous one so that the spot formed does not exceed a few millimeters in diameter. 11. By the end of spotting, a yellow-brownish spot will have formed on the plate. Then proceed to dissolving and spotting the next extract(s) in the same way until you fill all available spotting positions on the plate. Also spot 2 μL of the 17:0 TG standard solution on a separate position in the middle of the plate in order to identify the triacylglycerol spot in each sample after chromatography.

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12. When you first try this method, we recommend that you perform the analysis of individual phospholipids by 2D TLC on the following day(s). Therefore, save the remaining extract in each test tube by covering the test tube with Parafilm and storing at 4°C. Stable for 1 month. If, later on, you feel confident to perform both procedures on the same day, just leave the extracts on the bench. 13. Follow Subheading 3.2.1, steps 22–28. 14. Mark the contour of the triacylglycerol spot in each muscle sample with pencil, giving a slack of about 1 mm all around, and take the plate back to the lab. 15. Prepare a screw top test tube for each muscle sample. 16. Apply Subheading 3.2.1, steps 31–33 to the triacylglycerol spots only. 17. Prepare for 2D TLC by following Subheading 3.3.1, steps 1–5. 18. You will need one 10 × 10-cm TLC plate per muscle sample. If your plates are not 10 × 10 cm, cut them carefully with scissors, taking care not to chip off too much of the silica gel layer. Use the cut sides as the far ends in the subsequent chromatography (see Note 18). 19. By now, the solvent of the lipid extract(s) used to isolate triacylglycerols by 1D TLC (step 8) has evaporated. So, redissolve the remaining dry extract of each muscle sample in 15 μL of lipid extraction solvent and spot 10 μL of the resulting solution on a plate by following Subheading 3.3.1, steps 8–11 (see Note 35). Spot up to two samples if using small TLC tanks and up to four samples if using large tanks. To avoid mixup among plates, mark each one softly with pencil at its upper right corner. 20. Perform 2D TLC by following Subheading 3.3.1, steps 13–17. 21. See the phospholipid spots on each plate by following Subheading 3.3.1, steps 19 and 20. Mark the contour of each spot with pencil, giving a slack of about 1 mm all around. Identify each phospholipid by comparing the pattern on the plate to that on the transparency created under Subheading 3.3.1. 22. Prepare and label seven screw top test tubes per muscle sample, one for each phospholipid. To the bottom of each test tube, decant accurately 5 μL of the 15:0 ME standard solution using a 10-μL syringe with pointed needle (see Note 36). 23. Apply Subheading 3.2.1, steps 31–33 to each phospholipid spot.

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1. To each screw top test tube containing SM, add 1 mL of methanolic boron trifluoride and cap tightly. The liquid turns yellow, as it extracts the dichlorofluorescein from the silica gel. Vortex and heat at 100°C for 75 min (see Note 37). 2. To each of the remaining screw top test tubes, that is, the ones containing triacylglycerols, PC, LPC, PE, PS, PI or CL, add 0.5 mL of methanolic sodium methoxide and cap tightly. Vortex and heat at 50°C for 10 min (see Note 38). 3. Let all test tubes cool. Then remove their caps, taking care not to mix them up. To each test tube, add 1.5 mL of hexane, cap tightly, and vortex at full speed or shake vigorously for 1 min to extract the MEs and DMAs. 4. Isolate and dry the MEs and DMAs by following Subheading 3.2.2, steps 5–7. 5. Turn the gas chromatograph on. If you use manual injection, dissolve the dry residue in each test tube in 50 μL of hexane just before injection. Mix thoroughly and, in the case of triacylglycerols and PC, inject 1 μL. In the case of PE, inject 3 μL. 6. In the case of all other phospholipids (LPC, PS, PI, CL, and SM), evaporate the solution under a stream of nitrogen. Gently direct the stream all around the wall of the test tube, forcing the solution to concentrate at the bottom. Then redissolve in 10 μL of hexane, mix, and inject 3 μL. 7. If you use an autosampler, dissolve the dry residue in each test tube in 50 μL of hexane. Mix thoroughly and, using an automatic pipette, transfer the solution into a microvolume insert sitting inside a 2-mL screw top vial. 8. In the case of triacylglycerols, PC and PE, close the vial with septum and screw cap, place it in the autosampler and inject 1 μL (in the case of triacylglycerols and PC) or 3 μL (in the case of PE). 9. In the case of all other phospholipids (LPC, PS, PI, CL, and SM), evaporate the solution in the microvolume insert under a stream of nitrogen, taking good care not to splatter the solution. Wash the corresponding test tube with another 50 μL of hexane, mix thoroughly, and transfer the solution into the microvolume insert. Reevaporate, dissolve in 10 μL of hexane, mix, and inject 3 μL (see Note 27). 10. Use an appropriate software to acquire the chromatogram. Based on the retention times you have established in Subheading 3.1, assign the peaks in the chromatogram to MEs and DMAs and integrate the peaks so that you get a table containing an area value (in arbitrary units) for each identified ME and DMA. Depending on the capabilities of the data acquisition software, this can be done automatically, manually, or in part automatically and in part manually.

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3.3.4. Calculations

Conversion of the peak integration data into muscle content values (that is, μmol lipid per gram muscle) can be done through either the data acquisition software of the gas chromatograph or a spreadsheet (such as Microsoft® Excel). Described below is the reasoning that has to be followed in any case. 1. Calculations are based on the premise that peak area is proportional to ME mass. Of pivotal importance, then, are the amount and area of 17:0 ME derived from methanolysis of the 17:0 TG and 17:0 PC internal standards that were added to the muscle samples in the beginning of the analysis (see Note 30). On the basis of proportionality, if M17 and A17 are the amount and area, respectively, of 17:0 ME and A is the area of an endogenous ME, then its amount, M, is given by the formula, M = M17 × A/A17. Let’s apply this calculation separately to the triacylglycerols and phospholipids of a muscle sample. 2. Begin with triacylglycerols. To calculate the amount of 17:0 ME derived from 17:0 TG that is present in the muscle triacylglycerol fraction, first multiply the 17:0 TG standard concentration, a (in μg/μL), by the volume added to each sample, that is, 3 μL (see Subheading 3.3.2, step 3), yielding 3a μg. Divide by the Mr of 17:0 TG (849.4) and get 3a/849.4 μmol. Since 1 mol of a triacylglycerol yields 3 mol of MEs, one gets 9a/849.4 μmol of the 17:0 ME. Finally, by multiplying by the Mr of 17:0 ME (284.5), one gets 3.014a μg. This is M17. 3. Apply the formula, M = 3.014a × A/A17, to every ME identified in the chromatogram and get the amount of each ME in μg. Then divide by that ME’s Mr to express its amount in μmol, which is also μmol of the corresponding acyl group (or, more commonly, fatty acid) in the muscle triacylglycerols. 4. You may divide the amount of each fatty acid by the amount of muscle tissue (either wet or dry) weighed in Subheading 3.3.2, step 4 to express the fatty acid content of muscle triacylglycerols in μmol/g or μmol/mg. 5. You may add the amounts of all fatty acids and divide the sum by 3 to get the triacylglycerol content of muscle in μmol/g or μmol/mg. 6. You may divide the amount of each fatty acid by the sum of fatty acids and then multiply by 100 to express the percentage molar distribution of fatty acids in muscle triacylglycerols (see Note 31). 7. You may use partial sums to calculate the amounts or percentages of fatty acid categories, such as saturated, unsaturated, ω6, etc. 8. Continue with PC. To calculate the amount of 17:0 ME derived from 17:0 PC that is present in the muscle PC fraction, first multiply the 17:0 PC standard concentration, b (in μg/μL),

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by the volume added to each sample, that is, 7 μL (see Subheading 3.3.2, step 3), yielding 7b μg. Divide by the Mr of 17:0 PC (762.2) and get 7b/762.2 μmol. Since 1 mol of PC yields 2 mol of MEs, one gets 14b/762.2 μmol of 17:0 ME. Finally, by multiplying by the Mr of 17:0 ME (284.5), one gets 5.226b μg. 9. Apply the formula, M = 5.226b × A/A17, to every ME identified in the chromatogram and get the amount of each ME in μg. Then divide by that ME’s Mr to express its amount in μmol, which is also μmol of the corresponding fatty acid in muscle PC. 10. Apply the same formula to every DMA identified in the chromatogram (see Note 39) and get the amount of each DMA in μ g. Then divide by that DMA’s M r to express its amount in μmol, which is also μmol of the corresponding fatty aldehyde in muscle PC. 11. You may divide the amount of each fatty acid (and aldehyde) by the amount of muscle tissue used in the analysis to express the fatty acid (and aldehyde) content of muscle PC. 12. You may add the amounts of all fatty acids and aldehydes, then divide the sum by 2 to get the PC content of muscle. 13. You may divide the amount of each fatty acid and aldehyde by the sum of fatty acids and aldehydes, then multiply by 100 to express the percentage molar distribution of fatty acids and aldehydes in muscle PC (see Note 31). 14. You may use partial sums to calculate the amounts or percentages of fatty acid and aldehyde categories, such as saturated, unsaturated, ω6, etc. 15. To calculate the amounts of the other phospholipids, for which no internal standards were included, we will use, in addition, the area of 15:0 ME (the external standard that was added in equal amounts to all scraped phospholipid spots after 2D TLC in Subheading 3.3.2, step 22). Let M15 be the amount of 15:0 ME added to each phospholipid, A15,PC be the area of 15:0 ME in the PC chromatogram, and A15,PE be the area of 15:0 ME in the PE chromatogram (see Note 40). Then, the amount, M, of an endogenous ME having an area, A, in the PE chromatogram will be M = M15 × A/A15,PE. However, from the PC chromatogram, M15 = M17 × A15,PC/A17. By substitution we get, M = M17 × A15,PC/A17 × A/A15,PE, or M = M17 × A/A17 × A15,PC/A15,PE. This formula is like the one given in step 1, except that it contains an additional term that links the PE chromatogram (from which A is taken) to the PC chromatogram (from which A17 is taken), thus normalizing for the different areas of the external standard in the two chromatograms.

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16. Apply the formula, M = 5.226b × A/A17 × A15,PC/A15,PE, to every ME identified in the PE chromatogram and get the amount of each ME in μg. Then divide by that ME’s Mr to express its amount in μmol, which is also μmol of the corresponding fatty acid in muscle PE. 17. Apply steps 10–14 to PE. 18. Apply the same calculations to all of the remaining phospholipids with two exceptions. First, you do not have to bother with DMAs in these. Second, when coming to step 12, you will have to divide the sum of fatty acids in CL by 4 in order to get the CL content of muscle (see Fig. 2i); do nothing with the sum of fatty acids in LPC and SM, as each contains one acyl group.

4. Notes 1. Other lipid classes, such as (nonesterified) fatty acids and diacylglycerols, may be analyzed by the same methodology, but the amounts of tissue required will be higher than the ones dictated herein because of the low abundance of these lipid classes. 2. Fatty acids and their acyl groups are often abbreviated as [number of carbon atoms]:[number of double bonds], occasionally followed by an indication of the position of double bonds. 3. You may store the solutions indefinitely. However, certain MEs in solution, especially the polyunsaturated ones, deteriorate with time, resulting in their peaks decreasing and, eventually, vanishing from the gas chromatograms. If you notice this, discard the solution and make a fresh one. 4. In general, an 1-alkenyl group attached by ether linkage to glycerol in a plasmalogen is hydrolyzed to 1-alkenol, which is isomerized to the more stable aldehyde. A dimethyl acetal is produced by methylation of that aldehyde. 5. Based on the presence, in the gas chromatograms of the MEs derived from PC and, in particular, PE, of certain peaks that differ in retention times from the 18:0, 18:1ω9 and 18:1ω7 MEs by as much as 16:0 DMA differs in retention time from 16:0 ME, we have tentatively identified these peaks as 18:0, 18:1ω9 and 18:1ω7 DMAs (Mr 314.5, 312.5, and 312.5, respectively). Look for such peaks in your chromatograms. Unfortunately, we have not been able to find commercially available standards for these compounds. 6. Triacylglycerols and phospholipids with acyl groups bearing other odd numbers of carbon atoms, such as 15, 19, or 21, may be used as internal standards instead.

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7. Keep a small bottle of chloroform in the refrigerator or coldroom to minimize evaporation during preparation of the internal standard solutions. Rinse the pipette tip once or twice with the cold chloroform before dispensing it into the vials to make sure you deliver the right volume. 8. Keep the TLC plates in their box in a dry place, as silica gel is highly hygroscopic. Plates exposed to high humidity or kept in an opened box for over a year may need to be activated by placing in an oven at 110°C for 15–30 min prior to use. 9. Because organic solvents are volatile, it is preferable to remove an aliquot of a lipid standard solution by piercing the septum of its vial with a syringe with pointed needle, rather than by opening the vial and using an automatic pipette. However, always remember to promptly replace a pierced septum with a new one before storing a vial, or the solvent will gradually evaporate through the hole in the septum, even at −20°C. 10. Inject the larger and more unsaturated MEs first, as these are more difficult to separate than the smaller and saturated ones. Thus, you may save time by making any necessary adjustments earlier than later. 11. Alternatively, you may postpone the pulverization until the day of analysis. A pulverized tissue ensures higher sample homogeneity and more efficient lipid extraction. 12. The idea is to obtain a solvent depth of about 5 mm in the tank. 13. Prepare the developer fresh and use it only once. If, for any reason, you do not use it on the day of preparation discard it, as you cannot trust its composition the next day. 14. Depending on the number of samples, experience, dexterity, and number of available hands, lipid extraction and spotting on the TLC plate(s) may take from one-half to more than 2 h. In the latter case, start the day with lipid extraction (step 5) and prepare the developer later so that it does not have to wait for over 2 h. 15. If using lyophilized tissue, weigh out approximately one-fourth the recommended amount of wet tissue and note weight to the fourth decimal point of the gram. 16. If the muscle is not pulverized, make sure that its pieces are less than 2 mm across to achieve complete extraction of their lipids. If this is not the case, break the pieces apart within the extraction solvent with a spatula. If this is not feasible, homogenize the sample. 17. We create a fine stream of nitrogen by attaching a glass Pasteur pipette to the tubing supplying the nitrogen. We then bring the tip of the pipette about 1 cm from the surface of the liquid

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and adjust the gas flow so as to agitate the surface but not splatter the liquid. Since the evaporation cools the liquid and this slows down the process, we keep the test tube in a beaker with tap water at room temperature or up to 37°C. Alternatively, you may wish to hold the test tube in your palm during the evaporation process. 18. Draw a faint line with ruler and pencil on the silica gel to be able to cut the plate straight. Make sure the ruler is clean and does not touch the silica gel extensively. 19. The hair dryer should be turned off when delivering each drop, or the air current will blow the drop away. Also take care not to blow the plate and spotting guide away. 20. A multiplate rack can hold up to six plates and lets you develop them simultaneously in one TLC tank. All you have to do is load the plates on the rack and then carefully lower the rack into the tank. However, take care not to let the side that carries the silica gel touch the margin of the rack holding the plate in place, since the developer rises there through capillary action more rapidly than on the rest of the plate, resulting in abnormal developer front and aberrant lipid separation. 21. It is good for the plate to touch the surface of the developer horizontally so that the developer forms a horizontal front as it rises and all samples migrate uniformly. However, if the plate enters the developer sideways, do not panic and, under no circumstance, raise the plate in order to attempt a better “landing.” Just let the plate sit on the bottom. All you will get is a slightly crooked developer front and a slightly uneven migration pattern. Nevertheless, most probably, you will still be able to discern the lipid spots of interest. 22. Other means of locating lipid spots on a TLC plate, such as spraying with sulfuric acid solution or exposing to iodine vapors, are unsuitable for the subsequent ME and DMA analysis. Sulfuric acid chars all lipids, while iodine reacts with double bonds, thereby altering the unsaturated fatty acids. 23. The spray reagent may remain in the spray bottle if used daily. However, if the spray bottle is left unused for several days, the spray reagent may dry in the nozzle; then dichlorofluorescein will block the nozzle. If this happens, return the spray reagent to its container and unblock the nozzle by rinsing and spraying with ethanol. To prevent blocking from happening, remember to return the spray reagent to its container after use, rinse the spray bottle with ethanol, and spray some of it to clean the nozzle. 24. Do not wash the funnel between samples, as silica gel flakes will stick to it. However, do wash the funnel with chloroformmethanol 2:1 at the end of the day.

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25. If using a plate with soft support, you may cut it with scissors during the scraping sequence in any way you find convenient in order to bring distant spots close to the funnel and prevent silica gel flakes from falling outside the funnel. 26. This base-catalyzed methanolysis is according to Kramer et al. (8) and results in the full conversion of triacylglycerols and glycerophospholipids (see Fig. 2b–i) into MEs and DMAs. However, SM requires acid-catalyzed methanolysis which is performed in the next step on the phospholipid fraction only. 27. Heating a methanol-based solution at 100°C may cause its rapid evaporation unless it is tightly closed. Therefore, at about 5 min of heating tighten the caps (wear heat-resistant gloves!), as the caps tend to get loose. Check the level of methanol inside the test tubes from time to time. If you notice a leak, take the test tube out of the heating apparatus and let it cool briefly. Unscrew the cap and inspect both the cap’s lining and the test tube’s rim. If the cap’s lining is damaged replace the cap. If the test tube’s rim is chipped, transfer the contents into another test tube with a glass Pasteur pipette. In any case, add methanol to about 1 mL and continue the incubation. Make a note of any problematic test tubes, as, despite all efforts, they may produce aberrant data; this will necessitate repetition of the analysis. 28. Microvolume inserts require meticulous cleaning before being reused. We recommend rinsing five times with chloroformmethanol 2:1. 29. The fraction of the ME and DMA solution to be injected into the gas chromatograph depends on the split ratio chosen and on the triacylglycerol and phospholipid content of the muscle sample. Thus, you may change the volume of hexane in which you dissolve the ME and DMA residue, the volume injected into the gas chromatograph or the split ratio to achieve the best possible balance between getting signals of the minor components well above the noise and not overloading the column with the major components. 30. It is good for the area of 17:0 ME to be neither too small nor too big relative to the areas of the endogenous MEs in a chromatogram. Therefore, once you get your first chromatograms, try to adjust the amounts of the 17:0 TG and PC internal standards added to subsequent similar samples so that the area of 17:0 ME is about half the area of the most abundant ME (which is usually 16:0). 31. If you only intend to determine the percentage molar distribution of fatty acids (and aldehydes) in muscle triacylglycerols and phospholipids, you may omit the internal standards from the analysis. All you need to do then is divide the area of each ME

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(and DMA) in the chromatogram by that compound’s Mr and use the quotients to calculate percentage distribution. 32. This 2D TLC system is according to Kester et al. (9). 33. These proportions simulate the usual phospholipid distribution in skeletal muscles and are suggested instead of equal amounts of all phospholipids in order to produce a 2D TLC pattern that is as close to the natural one as possible. 34. Slight differences in the positions of some phospholipids between the plate containing all phospholipids and the plates containing individual phospholipids should be expected, since the migration of one compound in a chromatographic system may be affected by interactions with other compounds in the mixture and since the chromatographic conditions are rarely identical between two runs. 35. If you are unable to recover 10 μL of extract, dissolve the dry residue in 20, rather than 15, μL of lipid extraction solvent. 36. Methyl pentadecanoate will be used as external standard to enable calculation of the amounts of LPC, PE, PS, PI, CL, and SM in conjunction with the 17:0 PC internal standard. 37. Boron trifluoride affords acid-catalyzed methanolysis, which is necessary to break the amide linkage of a fatty acid to sphingosine in SM. The conditions for this reaction have been optimized in our laboratory. 38. This base-catalyzed methanolysis is according to Kramer et al. (8) and results in the full conversion of triacylglycerols and glycerophospholipids (see Fig. 2b–i) into MEs and DMAs. 39. As implied by Note 5, phosphatidal choline and phosphatidal ethanolamine comigrate with PC and PE, respectively, during 2D TLC. 40. Although the same amount of 15:0 ME is present with each phospholipid, its area will differ from chromatogram to chromatogram because a different fraction of each preparation was injected into the chromatograph. References 1. Abbott SK, Else PL, Hulbert AJ (2010) Membrane fatty acid composition of rat skeletal muscle is most responsive to the balance of dietary n-3 and n-6 PUFA. Br J Nutr 103:522–529 2. Nikolaidis MG, Mougios V (2004) Effects of exercise on the fatty acid composition of blood and tissue lipids. Sports Med 34:1051–1076 3. Bruce Å (1974) Skeletal muscle lipids. II. Changes in phospholipid composition in man from fetal to middle age. J Lipid Res 15:103–108 4. Bruce Å (1974) Skeletal muscle lipids. III. Changes in fatty acid composition of individual

phosphoglycerides in man from fetal to middle age. J Lipid Res 15:109–113 5. Hulbert AJ, Turner N, Storlien LH et al (2005) Dietary fats and membrane function: implications for metabolism and disease. Biol Rev Camb Philos Soc 80:155–169 6. Muoio DM (2010) Intramuscular triacylglycerol and insulin resistance: guilty as charged or wrongly accused? Biochim Biophys Acta 1801:281–288 7. Folch J, Lees M, Sloane-Stanley GH (1957) A simple method for the isolation and purification

19 of total lipids from animal tissues. J Biol Chem 226:497–509 8. Kramer JKG, Fellner V, Dugan MER et al (1997) Evaluating acid and base catalysts in the methylation of milk and rumen fatty acids with

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special emphasis on conjugated dienes and total trans fatty acids. Lipids 32:1219–1228 9. Kester M, Schliselfeld LH, Bárány M (1984) Minor phospholipids in human muscle. Mol Physiol 5:71–84

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Chapter 20 Proteomic Analysis of Dystrophic Muscle Caroline Lewis, Philip Doran, and Kay Ohlendieck Abstract Mass spectrometry-based proteomics had a major impact on the global characterization of skeletal muscles and has decisively enhanced the field of neuromuscular pathology. Proteomic profiling of x-linked muscular dystrophy has identified a large number of new signature molecules involved in fiber degeneration. Here, we describe the difference in-gel electrophoretic analysis of the dystrophic diaphragm muscle from the MDX mouse model of Duchenne muscular dystrophy. This chapter summarizes the various experimental steps involved in muscle proteomics, such as sample preparation, fluorescence labeling, isoelectric focusing, second-dimension slab gel electrophoresis, image analysis, in-gel digestion and electrospray ionization mass spectrometry. Key words: Difference in-gel electrophoresis, Duchenne muscular dystrophy, Mass spectrometry, MDX, Proteomics, Two-dimensional gel electrophoresis

1. Introduction Over the last decade, skeletal muscle proteomics has clearly established itself as a distinct discipline within the field of basic and applied myology (1–3). The mass spectrometry-based cataloging of the muscle proteome and the global comparative analysis of developing, differentiating, degenerating, and aging contractile tissues has decisively advanced our understanding of skeletal muscle function in health and disease (4). The most common neuromuscular disorder and most frequent gender-specific genetic disease of childhood is represented by Duchenne muscular dystrophy. This highly progressive muscle-wasting disease is due to primary genetic abnormalities in the gene that encodes the large membrane cytoskeletal protein dystrophin (5). In healthy skeletal muscle fibers, the molecular linkage between the extracellular matrix and

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the membrane cytoskeleton is provided by dystrophin isoform Dp427 and its associated glycoprotein complex, consisting of sarcoglycans, dystroglycans, syntrophins, dystrobrevins, sarcospan, and various receptors (6–8). Dystrophin is an early marker of muscle development (9–11) and deficiency in its Dp427 isoform causes severe disruption of embryonic myogenesis. The lack of full-length dystrophin results in the disturbance of the developmental skeletal muscle patterning process (12). In both Duchenne patients and Dp427-deficient mice, the expression of dystrophin-associated glycoproteins is greatly reduced in the postnatal musculature (13, 14). The disintegration of the dystrophin-glycoprotein complex is believed to weaken the sarcolemmal membrane and render it more susceptible to microrupturing. Faulty membrane repair mechanisms involving Ca2+-leak channels (15) are probably at the core of downstream alterations that trigger pathophysiological levels of cytosolic Ca2+-levels (16). The proteomic profiling of an established animal model of X-linked muscular dystrophy, the MDX mouse, has revealed numerous novel biomarkers of dystrophinopathy (17–20). Signature molecules associated with the cellular stress response, energy metabolism, and the contractile apparatus had not previously been identified by conventional biochemical and cell biological surveys to be altered in dystrophic muscle tissues (21). This emphasizes the analytical power of mass spectrometry-based proteomic investigations. Proteomics suggests itself as an unbiased analytical tool to investigate complex pathological mechanisms. High-resolution two-dimensional gel electrophoresis with immobilized pH gradients is a frequently used protein separation method in modern proteomics (22–24) and is often combined with fluorescent labeling technology (25). Fluorescence difference in-gel electrophoresis (DIGE) was developed by Minden and coworkers (26) and represents one of the most effective biochemical methods to directly compare protein expression levels between distinct proteomes (27–29). Here, we describe in detail the application of the DIGE technique for comparing normal vs. dystrophic skeletal muscle preparations.

2. Materials 2.1. Equipment

All equipment is from Amersham/GE Healthcare unless otherwise stated. 1. IPG DryStrip reswelling tray. 2. IPGphor IEF unit. 3. Manifold. 4. Sample loading cups.

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5. Gel casting box. 6. Cassette racks. 7. Ettan DALTtwelve multiple vertical slab gel electrophoresis apparatus. 8. Glass plates suitable for DIGE analysis. 9. Typhoon Trio variable mode imager. 10. ImageScanner UMax. 11. Ettan spot picker workstation. 12. Vortex Genie-2 (Scientific Industries). 13. Stuart SSL4 shaker (Lennox Laboratory Supplies Ltd.). 14. Heto speedvac concentrator. 15. Eppendorf Model 5417R centrifuge. 16. Agilent 6340 Ion Trap LC mass spectrometer using electrospray ionization (Agilent Technologies). 17. Nanoflow Agilent 1200 series system, equipped with a Zorbax 300SB C18 Mm; 4 mm 40 nL precolumn was used for the separation of peptides (Agilent Technologies). 2.2. Reagent Solutions

All reagents were from Amersham Biosciences/GE Healthcare unless otherwise stated. 1. CyDye DIGE fluor minimal dye Cy2. 2. CyDye DIGE fluor minimal dye Cy3. 3. CyDye DIGE fluor minimal dye Cy5. 4. Immobilized pH gradient (IPG) strips. 5. IPG buffer. 6. Iodoacetamide. 7. Destreak agent. 8. Laemmli-type buffer system (Biorad). 9. Protein molecular mass markers (Biorad). 10. Ultrapure Protogel acrylamide stock solution (National Diagnostics). 11. Protease inhibitors (Roche). 12. Sequencing grade-modified trypsin (Promega). 13. All other chemicals used were of analytical grade (Sigma). 14. All solutions should be prepared with ultrapure water.

2.2.1. Preparation of Crude Skeletal Muscle Extracts

1. Lysis buffer: 9.5 M urea, 4% CHAPS, 1% pH 3–10 ampholytes, 100 mM dithiothreitol. In order to prevent excess proteolytic degradation of sensitive muscle proteins, lysis buffer was supplemented with commercially available protease inhibitors

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(see Note 1). This solution can be dispensed into 1 mL aliquots and stored at −20°C. 2. DIGE lysis buffer (pH 8.5): 9.5 M urea, 4% CHAPS, 30 mM Tris–HCl, pH 8.5 (see Note 2). This solution can be dispensed into 1 mL aliquots and stored at −20°C. 3. Reducing lysis buffer (for addition to sample after dye labeling): 9.5 M urea, 4% CHAPS, 2% IPG buffer pH 3–10, 130 mM dithiothreitol. The buffer should be supplemented with commercially available protease inhibitors (see Note 1). This solution can be dispensed into 1 mL aliquot and stored at −20°C. 2.2.2. Rehydration of IPG Strips

1. Rehydration buffer: To 1 mL lysis buffer, add 12 ML Destreak and 0.002% Bromophenol Blue (see Note 3).

2.2.3. Second-Dimension Gel Electrophoresis

1. Equilibrium buffer: 8 M urea, 30% glycerol, 2% SDS, 50 mM Tris–HCl pH 8.8, 0.002% Bromophenol Blue (see Note 4). Solution can be aliquot and stored at −20°C. 2. DTT equilibrium buffer: Add 100 mg dithiothreitol per 10 mL of equilibrium buffer. Make this solution freshly prior to use in gel electrophoresis. 3. IA equilibrium buffer: Add 125 mg iodoacetamide per 10 mL of equilibrium buffer. Make this solution freshly prior to use in gel electrophoresis. 4. 10× SDS buffer: 25 mM Tris, 192 mM glycine, 0.1% SDS. Store this solution at room temperature. 5. Sealing solution: 1% agarose in 1× SDS buffer, and Bromophenol Blue. Heat solution until agarose has properly dissolved. Store the solution at room temperature (see Note 5).

2.2.4. Protein Visualization Using Silver Staining

1. Fixative solution: 30% ethanol, 10% acetic acid. 2. Rinse solution: 20% ethanol. 3. Sensitizing solution: 0.8 mM sodium thiosulfate. Make this solution freshly prior to use in silver staining. 4. Staining solution: 12 mM silver nitrate. Make this solution freshly prior to use in silver staining. 5. Developing solution: 30 g/L sodium carbonate, 250 ML/L formaldehyde, 125 ML/L of 10% sodium thiosulphate. Make this solution freshly prior to use in silver staining. 6. Stop solution: 40 g/L Trizma base, 20 mL/L acetic acid.

2.2.5. Reduction and Alkylation of Silver-Stained Proteins

1. DTT solution: 10 mM DTT in 100 mM ammonium bicarbonate. Make this solution freshly prior to use in silver staining and store at room temperature. 2. IA solution: 55 mM iodoacetamide in 100 mM ammonium bicarbonate. Make this solution freshly prior to use in silver staining and store at room temperature.

20 2.2.6. In-Gel Digestion of Protein Spots for Mass Spectrometric Identification

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1. Trypsin buffer: 20 Mg of sequencing grade-modified trypsin in 1.5 mL buffer. Stock solution of trypsination buffer with a total volume of 10 mL consists of 1 mL of 100 mM ammonium bicarbonate, 1 mL of acetonitrile, and 8 mL of deionized water (see Note 6). 2. Extraction buffer: 1:2 (v/v) formic acid/acetonitrile. Make this solution freshly and use it immediately for peptide extraction.

3. Methods A frequent problem with proteomic studies, that use a combination of two-dimensional gel electrophoresis and mass spectrometry for the identification of distinct protein species, is the contamination of samples with keratin protein. To keep potential impurities to a minimum, researchers are advised to wear protective gloves during the preparation and handling of protein samples. In addition, electrophoretic and analytical solutions should be prepared and stored in designated facilities with at least semiclean analytical status. Electrophoretic separation steps are ideally performed under designated fume hoods or in special rooms that lack excess air passage. Mass spectrometric analyses should be carried out in a special proteomics suite that is air-conditioned and kept free from potential contaminants. 3.1. Preparation of Crude Muscle Extracts

1. Weigh muscle tissue from normal and dystrophic mice (see Note 7). Since the mdx diaphragm exhibits severe dystrophic changes, we have focused our proteomic studies on this subtype of skeletal muscle (19–21). 2. Place muscle tissue in liquid nitrogen and grind to a powder with mortar and pestle. 3. Add muscle powder to lysis buffer at a ratio of 1:10 (see Note 8). 4. Briefly vortex the solution. The suspension is then incubated on a rocker for 1 h at room temperature, with gentle vortexing every 10 min for 30 s. 5. Centrifuge at 20,000 × g for 20 min at 4°C. Save the proteincontaining middle layer, discarding the pellet and uppermost fatty layer. 6. Carry out a reliable protein quantification of muscle extracts to be analyzed (see Note 9). 7. Dispense protein extracts into aliquots of 50 ML and store at −80°C.

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3.2. Labeling of Muscle Proteins with Fluorescent CyDyes

1. Commercially available CyDye stock solutions are resuspended in anhydrous dimethylformamide (DMF) to give a final concentration of 1 mM dye (see Note 10). Stock solutions can be stored in the dark at −80°C. 2. Prior to labeling, briefly vortex the vials containing individual CyDyes and centrifuge them at 12,000 × g for 10 s. 3. Dilute dyes 1:4 (v/v) with DMF to make a working solution of 200 pmol/ML. Working solutions can be stored in the dark at −80°C. 4. Check whether the sample pH-value is at pH 8.5 prior to the labeling reaction (see Note 11). 5. Add 1 ML of dye per 25 Mg muscle protein. For the differential analysis of the normal vs. the dystrophic diaphragm proteome, label 50 Mg of normal diaphragm extract using Cy3 dye, 50 Mg of dystrophic mdx diaphragm extract using Cy5 dye, and 50 Mg of pooled internal standards using the Cy2 dye. 6. Briefly vortex the samples, centrifuge them at 12,000 × g for 10 s and then incubate the suspension on ice in the dark for 30 min. 7. Stop the labeling reaction by addition of 1 ML of 10 mM lysine per 25 Mg of protein. Briefly vortex the samples, centrifuge them at 12,000 × g for 10 s, and then incubate them on ice in the dark for 10 min. Samples can be used immediately for electrophoretic separation or stored in the dark at −80°C.

3.3. Rehydration of First-Dimension Gel Strips

1. Dispense 450 ML of rehydration buffer into the reservoir slots of the Ettan IPGphor DryStrip reswelling tray. 2. Remove plastic backing from 24 cm-long IPG strips of pH 3–10 by peeling from the (−) end and push the (+) end towards the top of strip holder. 3. Place strips gel side down into rehydration buffer and rehydrate for at least 12 h (see Note 12).

3.4. Isoelectric Focusing

1. Transfer strips to manifold gel side up on the Ettan Multiphor II (GE Healthcare). Lift by the (−) end and place the (+) end of strips towards the (+) end marked on the IPGphor. 2. Cover strips with cover fluid by addition of 108 mL of drystrip cover fluid over entire manifold. 3. Place wicks, wet with deionised water, onto ends of strips. 4. Place sample loading cups onto strips. 5. For analytical gels: Following DIGE labeling, add an equal volume of reducing lysis buffer. Pool the samples together to give 150 Mg of total combined labeled extract per strip as per experimental design. Subsequently pipette protein into sample loading

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cups and carry out the same isoelectric focusing procedure as described below. 6. For preparative gels: Pipette 200 Mg of the protein extract into cups. Position the electrodes. Isoelectric focusing is carried out at 20°C as follows: 4 h step at 80 V, 2 h step at 100 V, 1.5 h step at 500 V, 1.5 h step at 1,000 V, 1 h step at 2,000 V, 1 h step at 4,000 V, 2 h step at 6,000 V, and a 2.5-h step at 8,000 V. After completion of the first-dimensional gel separation, IEF strips can be stored at −80°C. 3.5. SecondDimensional Electrophoresis

1. Incubate strips in 10 mL DTT equilibrium buffer while rocking for 15 min. 2. Pour off solution and incubate while rocking in 10 mL IA equilibrium buffer for 15 min. 3. Pour off solution, wash strips briefly in 1× SDS running buffer before placing strips (+) end to left, gel side facing out, onto a 12.5% resolving slab gel (see Note 13). 4. Strips should be pressed against gel when adding warmed overlay sealing solution, so to eliminate air bubbles. 5. Place gels in the Ettan DALTtwelve tank and carry out electrophoresis at 0.2 W/gel for 1 h, followed by 0.4 W/gel for 1 h and then 1.5 W/gel overnight until dye front runs off. 6. Following electrophoresis, carefully remove gels from plates and mark one corner of gel to track orientation. Strips should remain with gels as each contains a unique number to allow tracking of samples. DIGE gels should be stored in darkness to protect the fluorescence signal of individually labeled muscle proteins. 7. Preparative pick gels should ideally be run at the same time as analytical DIGE gels as to eliminate any potential technical discrepancies arising from the second-dimension separation step.

3.6. Protein Visualization Using Silver Staining

1. For this mass spectrometry-compatible method, approximately 200 mL of solution is needed per gel and staining step. 2. Place gels in clean glassware and add fixative solution for a minimum of 30 min. Gently agitate gels during the fixation step (see Note 14). If necessary, gels can be left overnight in fixing solution. 3. Rinse gels twice for 10 min with rinse solution, then rinse twice with fresh deionized water for 10 min with gentle agitation. 4. Wash gel with sensitizing solution for 1 min (see Note 15). 5. Rinse gel with deionized water for 10 min with gentle agitation. 6. Remove water and add staining solution for 20 min to 2 h (see Note 16).

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7. Rinse with deionized water for 10 s and agitate in developing solution until a 2D-spot pattern is properly recognized. 8. When suitably developed, gels can be placed into stop solution. Prior to densitometric scanning, gels should be rinsed in deionized water (see Note 17). 3.7. Image Analysis of Protein Spot Patterns

1. Scan CyDye-labeled gels using a suitable variable mode imager, such as the Amersham Biosciences/GE Healthcare Typhoon Trio apparatus. 2. For image acquisition, scan Cy2, Cy3, and Cy5-labeled muscle proteins at wavelengths of 488, 532, and 633 nm, respectively. Photomultiplier tube PMT-values should be optimized so that the volume of the most abundant protein spot is between 80,000 and 99,000 when scanned at a resolution of 100 Mm (see Note 18). 3. The normal and dystrophic mdx sample images are then evaluated using 2D gel analysis software, such as Progenesis SameSpots analysis software (NonLinear Dynamics, Newcastle upon Tyne, UK; software version 3.2.3) and are normalized against their corresponding pooled image. 4. Two-dimensional gels are aligned to the reference image. See Fig. 1 for a representative DIGE master gel. 5. Following detection of spots, gels are placed into groups (normal vs. mdx images) and analyzed to determine significant changes in the abundance of distinct 2D-spots. 6. Paired Student’s t-test values are calculated for each protein spot across all gels. An ANOVA score of 0.5 is required for spots

Fig. 1. DIGE master gel for the comparative analysis of normal vs. dystrophic mdx diaphragm extracts. Shown is a Cy2-labeled master gel of the total soluble protein complement from normal diaphragm vs. dystrophic diaphragm tissue. The pH-values of the first-dimension gel system and molecular mass standards (in kDa) of the second dimension are indicated on the top and on the left of the panels, respectively.

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to be included in the subsequent detailed evaluation of changes in protein expression patterns. Then principal component analysis (PCA) was verified with changes displaying power of <0.8 being removed from the analysis. All remaining changed protein spots that exhibit a fold-change of 1.5 or greater and meet the significance criteria should be visually checked on the aligned gels to ensure feasibility. 7. Silver-stained preparative gels are scanned using a suitable instrument, such as the ImageScanner UMax from Amersham Biosciences/GE Healthcare. Images are added to the Progenesis software programme as a pick gel and aligned to analytical DIGE-based gels. 3.8. Excision of Protein Spots from TwoDimensional Gels

1. If an automated spot-picking workstation, such as Ettan spot picker from Amersham Biosciences/GE Healthcare, is available, then a spot-picking map of interest can be exported from the Progenesis software programme to the automated device. Proteins can be excised into 96-well plastic plates. 2. Alternatively, spots of interest can be picked manually from two-dimensional gels. Protein spots can be conveniently removed and transferred with the help of sterile pipette tips with their ends cut off. Place the tip over the individual protein spot, press firmly down through the gel, take the plug up into the tip and aspirate it into a fresh plastic tube.

3.9. Destaining of Silver-Stained Preparative Protein Spots

1. Prepare stock solutions of 30 mM potassium ferricyanide and 100 mM sodium thiosulfate. 2. Immediately prior to use, mix stock solutions 1:1 (v/v) and add 30 ML per gel plug. 3. Incubate with gentle shaking at room temperature until color disappears from gel plugs. 4. Wash plugs 3 times with deionized water for 10 min each, followed by the removal of all liquid.

3.10. Reduce and Alkylate Preparative Silver-Stained Spots

1. Add 500 ML of neat acetonitrile and incubate gel plugs for 10 min while shaking. Briefly spin down the suspension and remove liquid. 2. Add 30 ML of DTT solution and incubate gel plugs for 30 min at 56°C while shaking. 3. Chill tubes to room temperature and add 500 ML of acetonitrile and incubate gel plugs for 10 min at room temperature. Remove all liquid. 4. Add 30 ML of IA solution and incubate gel plugs for 20 min at room temperature. 5. Shrink gel plugs with acetonitrile and remove all liquid (see Note 19).

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3.11. In-Gel Digestion of Protein for Mass Spectrometric Identification

1. Add 50 ML of trypsin buffer and incubate gel plugs for 30 min at 4°C. 2. Add more buffer to fully cover gel plugs and incubate proteins for a further 90 min at 4°C (see Note 20). 3. Incubate overnight at 37°C (see Note 21). 4. Add 100 ML of extraction buffer and incubate proteins for 15 min at 37°C with shaking. 5. Using gel-loading tips, transfer all supernatant fractions into individual 0.2 mL plastic tubes. 6. Dry down peptides in a standard speedvac concentrator (see Note 22).

3.12. Electrospray Ionization-Mass Spectrometric Analysis

1. Reconstitute peptides in 12 ML of 0.1% formic acid. 2. Samples should then be briefly vortexed, sonicated for 5 min, and briefly centrifuged. 3. To remove any gel particles, centrifuge samples for 20 min in cellulose spin filter tubes at 14,000 × g. 4. Using fresh tips for each sample, pipette samples to individually labeled LC-MS vials. 5. To identify muscle-associated proteins of interest, analyze peptide mixtures on an ion trap LC mass spectrometer by injecting 5 ML of sample. 6. Although conditions have to be usually optimized with specific mass spectrometers, using a 10-min gradient of 5–100% acetonitrile/0.1% formic acid and a post run of 1 min through a Zorbax 300SB C18 Mm column gives reliable results with muscle proteins. 7. Separation of peptides can, for example, be achieved with a nanoflow Agilent 1200 series system, equipped with a Zorbax 300SB C18 5 Mm, 4 mm 40 nL precolumn, and an Zorbax 300SB C18 5 Mm, 43 mm × 75 Mm analytical reversed phase column using HPLC-Chip technology. 8. Mobile phases should be (A): 0.1% formic acid, and (B): 90% acetonitrile and 0.1% formic acid. Load samples into the enrichment at a capillary flow rate set to 4 ML/min with a mix of A and B at a ratio 19:1. 9. Elute tryptic peptides with a linear gradient of 5–70% solvent B over 6 min, 70–100% for 1 min and 100% for 1 min with a constant nano pump flow of 0.60 mL/min. A 1-min posttime of Solvent A should be used to remove sample carryover. 10. Set the capillary voltage to 2,000 V. The flow rate and the temperature of the drying gas should be 4 L/min and 300°C, respectively.

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11. In order to identify distinct protein species, utilize intern-based database search engines, such as Mascot MS/MS Ion search from Matrix Science (London, UK; NCBI database, release 20100212).

4. Notes 1. A suitable protease inhibitor cocktail for the preparation of crude skeletal muscle extracts would be the usage of one tablet of “Complete Mini” (Roche) per 10 mL of buffer. 2. The pH-value of the DIGE lysis buffer should be verified as being pH 8.5 and adjusted, if necessary, using 30 mM Tris–HCl. 3. In order to prevent the potential interference with fluorescent signals, omit Bromophenol Blue dye in DIGE gels. 4. The equilibrium buffer is a stock solution. Dithiothreitol or iodoacetamide must be added prior to use. 5. Sealing solution should be melted prior to use. 6. Make this solution freshly and use it immediately for trypsination of protein extracts. 7. Approximately 100 mg wet weight of diaphragm tissue can be obtained from adult mice for comparative proteomic studies. 8. For DIGE analysis, place samples into DIGE lysis buffer (which contains no reducing agents) and add equal volumes of 2× lysis buffer to protein extracts before placing them onto IEF strips. 9. Suitable protein quantification assays are for example the commercially available 2-D Quant Kit (Amersham Biosciences/GE Healthcare). This kit is designed for the accurate determination of protein concentration in samples to be analyzed by high-resolution two-dimensional gel electrophoresis. 10. It is recommended that a fresh batch of DMF is used for the generation of a new stock solution of fluorescent dyes. 11. Fresh samples should be made up in DIGE lysis buffer and may need no adjustment with respect to their pH-values. 12. When laying first-dimension strips down, ensure no bubbles remain trapped under the IEF strip, as rehydration can then not occur evenly. 13. SDS running buffer helps to slide the strip smoothly onto gels and so reduces air bubbles forming between strip and gel. 14. Glassware must be used with the silver staining technique, as silver binds to plastic dishes. Ideally, glassware is cleaned with 70% ethanol, then 10% acetic acid, and then washed with deionized water.

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15. Do not touch gels during sensitizing or staining steps in order to minimize the potential for fingerprint marks. 16. Following staining, dispose toxic silver nitrate waste correctly. 17. Washing gels with water removes acetic acid and this greatly reduces the formation of air bubbles between the gel and flatbed scanner. 18. Optimized PMT-values guarantee that no 2D spot will be saturated on the gel, therefore allowing accurate and quantifiable analysis. 19. Samples are now ready for in-gel digestion. Alternatively, they can be stored at −20°C. 20. Incubating at 4°C allows the slow and efficient diffusion of trypsin into gel plugs. 21. Following digestion, peptide mixtures can be stored at −20°C. 22. If necessary, dried-down extracts can be stored at −20°C. References 1. Isfort RJ (2002) Proteomic analysis of striated muscle. J Chromatogr B 771:155–165 2. Doran P, Gannon J, O’Connell K, Ohlendieck K. (2007) Proteomic profiling of aged and pathological skeletal muscle (review). Int J Mol Med 19:547–564 3. Doran P, O’Connell K, Gannon J, Ohlendieck K. (2007) Proteomic profiling of animal models mimicking skeletal muscle disorders. Proteomics Clin Appl 1:1169–1184 4. Ohlendieck K (2010) Proteomics of skeletal muscle differentiation, neuromuscular disorders and aging. Expert Rev Proteomics 7:283–296 5. Monaco AP, Neve RL, Colletti-Feener C, Bertelson CJ, Kurnit DM, Kunkel LM (1986) Isolation of candidate cDNAs for portions of the Duchenne muscular dystrophy gene. Nature 323:646–650 6. Campbell KP (1995) Three muscular dystrophies: loss of cytoskeleton-extracellular matrix linkage. Cell 80:675–679 7. Ohlendieck K (1996) Towards an understanding of the dystrophin-glycoprotein complex: linkage between the extracellular matrix and the subsarcolemmal membrane cytoskeleton. Eur J Cell Biol 69:1–10 8. Ervasti JM, Sonnemann KJ (2008) Biology of the striated muscle dystrophin-glycoprotein complex. Int Rev Cytol 265:191–225 9. Clerk A, Sewry CA, Dubowitz V, Strong, PN (1992) Characterisation of dystrophin in fetuses at risk for Duchenne muscular dystrophy. J Neurol Sci 111:82–91

10. Wessels A, Ginjaar IB, Moorman AF, van Ommen GJ (1991) Different localization of dystrophin in developing and adult human skeletal muscle. Muscle Nerve 14:1–7 11. Clerk A, Strong PN, Sewry CA (1992) Characterisation of dystrophin during development of human skeletal muscle. Development 114:395–402 12. Merrick D, Stadler LK, Larner D, Smith J (2009) Muscular dystrophy begins early in embryonic development deriving from stem cell loss and disrupted skeletal muscle formation. Dis Model Mech 2:374–388 13. Ohlendieck K, Campbell KP (1991) Dystrophinassociated proteins are greatly reduced in skeletal muscle from mdx mice. J Cell Biol 115:1685–1694 14. Ohlendieck K, Matsumura K, Ionasescu VV, Towbin JA, Bosch P, Weinstein SL, Sernett SW, Campbell KP (1993) Duchenne muscular dystrophy: deficiency of dystrophin-associated proteins in the sarcolemma. Neurology 43:795–800 15. Krueger J, Kunert-Keil C, Bisping F, Brinkmeier H (2008) Transient receptor potential cation channels in normal and dystrophic mdx muscle. Neuromuscul Disord 18:501–513 16. Mallouk N, Jacquemond V, Allard B (2000) Elevated subsarcolemmal Ca2+ in mdx mouse skeletal muscle fibres detected with Ca2+activated K+ channels. Proc Natl Acad Sci USA 97:4950–4955 17. Ge Y, Molloy MP, Chamberlain JS, Andrews PC (2003) Proteomic analysis of mdx skeletal

20 muscle: Great reduction of adenylate kinase 1 expression and enzymatic activity. Proteomics 3:1895–1903 18. Doran P, Dowling P, Lohan J, McDonnell K, Poetsch S, Ohlendieck K (2004) Subproteomics analysis of Ca2+-binding proteins demonstrates decreased calsequestrin expression in dystrophic mouse skeletal muscle. Eur J Biochem 271:3943–3952 19. Doran P, Dowling P, Donoghue P, Buffini M, Ohlendieck K (2006) Reduced expression of regucalcin in young and aged mdx diaphragm indicates abnormal cytosolic calcium handling in dystrophin-deficient muscle. Biochim Biophys Acta 1764:773–785 20. Doran P, Martin G, Dowling P, Jockusch H, Ohlendieck K (2006) Proteome analysis of the dystrophin-deficient MDX diaphragm reveals a drastic increase in the heat shock protein cvHSP. Proteomics 6:4610–4621 21. Lewis C, Carberry S, Ohlendieck, K (2009) Proteomic profiling of x-linked muscular dystrophy. J Muscle Res Cell Motil 30:267–279. 22. Carrette O, Burkhard PR, Sanchez JC, Hochstrasser DF (2006) State-of-the-art

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two-dimensional gel electrophoresis: a key tool of proteomics research. Nast Protoc 1:812–823 23. Weiss W, Gorg A (2009) High-resolution twodimensional electrophoresis. Methods Mol Biol 564:13–32 24. Friedman DB, Hoving S, Westermeier R (2009) Isoelectric focusing and two-dimensional gel electrophoresis. Methods Enzymol 463: 515–540 25. Viswanathan S, Unlu M, Minden JS (2006) Two-dimensional difference gel electrophoresis. Nat Protoc 1:1351–1358 26. Minden JS, Dowd SR, Meyer HE, Stuehler K (2009) Difference gel electrophoresis. Electrophoresis 30:S156–S161 27. Marouga R, David S, Hawkins E (2005) The development of the DIGE system: 2D fluorescence difference gel analysis technology. Anal Bioanal Chem 382:669–678 28. Timms JF, Cramer R (2008) Difference gel electrophoresis. Proteomics 8:4886–4897 29. Sapra R (2009) The use of difference in-gel electrophoresis for quantitation of protein expression. Methods Mol Biol 492:93–112

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Part VI Experimental Approaches in Calcium Imaging of Skeletal Muscle

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Chapter 21 Detection of Calcium Release via Ryanodine Receptors Jerry P. Eu and Gerhard Meissner Abstract The ryanodine receptor ion channels (RyRs) release Ca2+ from the endo/sarcoplasmic reticulum in a variety of nonvertebrate and vertebrate species including flies, crustaceans, birds, fish, and amphibians. They are most abundant in skeletal and cardiac muscle, where in response to an action potential, the release of Ca2+ ions from the sarcoplasmic reticulum through the RyRs into the cytoplasm leads to muscle contraction (i.e., excitation-contraction coupling). Here, we describe how to determine their cellular location using isoform-specific antibodies, their protein levels using an in vitro (3H)ryanodine-binding assay, and their cellular release of Ca2+ using RyR-specific channel agonists and inhibitors. Key words: Ryanodine receptors, Ca2+ release channels, Immunofluorescent/immunohistochemical localization, (3H)ryanodine binding, Ryanodine receptor agonists and inhibitors

1. Introduction The release of Ca2+ ions from intracellular stores regulates a wide variety of biological functions including muscle contraction, secretion, apoptosis, and gene expression. The release of Ca2+ is predominantly mediated by two related Ca2+ release channels, the ryanodine receptors (RyRs) (1–3) and inositol 1,4,5-trisphosphate receptors (IP3Rs) (4). Both are localized in the endoplasmic reticulum and in striated muscle in a specialized subcompartment, the sarcoplasmic reticulum. Additional intracellular organelles that store and release Ca2+ include mitochondria and acidic endosomalrelated structures. The RyRs are 2,200 kDa multiprotein complexes composed of four 560-kDa RyR subunits and four small FK506-binding protein (FKBP) subunits (1–3, 5). They are regulated by multiple endogenous effectors that include Ca2+, Mg2+, ATP, protein kinases,

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calmodulin, and reactive oxygen and nitrogen molecules. Exogenous effectors found to affect RyR function include ryanoids, toxins, xanthines, anthraquinones, phenol derivatives, dantrolene, local anesthetics, and polycationic and sulfhyryl reacting reagents. The highly specific interaction of RyRs with the plant alkaloid ryanodine has given them their name to distinguish them from other intracellular Ca2+ release mechanisms (6). Ryanodine binds with high affinity to RyRs and modifies their single channel conductance in a highly characteristic way by inducing a longlasting subconductance state at low concentrations (<1 MM) and closed channels at elevated concentrations (>10 MM) (7). The RyR family of Ca2+ release channels is composed of at least three isoforms. In mammals, RyR1 is the dominant isoform in skeletal muscle. RyR2 is found in high levels in cardiac muscle. RyR3 was initially identified in brain, but is expressed in many tissues including smooth muscle and brain. In amphibian and avian skeletal muscle, RyR1 and RyR3 are both expressed. In skeletal muscle, an action potential initiates L-type Ca2+ channel (Cav1.1) protein conformational changes that open the closely apposed RyR1s through a direct physical interaction (8). In cardiac muscle, an action potential results in the influx of extracellular Ca2+ through Cav1.2 channels (9). Both mechanisms lead to the release of Ca2+ from the SR and subsequent muscle contraction. The release of Ca2+ via RyR2 following an influx of Ca2+ in the heart is referred to as Ca2+-induced Ca2+ release (CICR). Sequestration of released Ca2+ by the SR Ca2+-transporting ATPase (SERCA) and extrusion by the Na+-Ca2+ exchanger restore the myofibrillar Ca2+ concentration from 10−6–10−5 to ~10−7 M, causing muscle to relax.

2. Materials 2.1. Immuno fluorescent /Immuno histochemical Localization of RyR Isoforms

1. Isoform-specific polyclonal antibodies recognizing mammalian RyRs (Alomone Labs and Millipore). 2. Nonisoform-specific mouse monoclonal anti-RyR antibody (C34 clone) and anti-RyR2 antibody (C3-33 clone) (Thermo Scientific). These antibodies recognize RyRs from various vertebrate species. 3. Species-specific fluorochrome-conjugated goat antimouse IgG and goat antirabbit IgG secondary antibodies (Jackson ImmunoResearch Laboratories). 4. Chamber glass slides (Nalge Nunc International). 5. VectaStain Elite kits and Mouse-on-mouse kits (Vector Labs). 6. Tissue-Tek Optimum Cutting Temperature (OTC) Compound and Cryomold (Sakura Finetek). 7. 4c,6-diamidino-2-phenylindole (DAPI).

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8. Washing solution: Phosphate-buffered saline (PBS): 140 mM NaCl, 2 mM KCl, 1.5 mM KH 2PO4, 15 mM Na 2HPO4, pH 7.4. 9. Fixing solution: 2% paraformaldehyde in PBS. 10. Quenching solution: 0.75% glycine in PBS. 11. Permealizing solution: 0.01% SDS and 0.05% Triton-X in PBS. 12. Blocking solution: 10% goat serum in PBS. 13. Antibody solution: 1% goat serum in PBS. 14. ProLong Antifade solution (Invitrogen). 2.2. Determination of RyR Protein Levels Using (3H)RyanodineBinding Assay

1. (3H)Ryanodine (Perkin Elmer Life Sciences). 2. Ryanodine (Calbiochem-EMD4Biosciences). 3. Protease inhibitor cocktail. 4. Polyethyleneimine (50% solution in water). 5. Tissue homogenizer. 6. Glass/glass homogenizers. 7. Glass microfiber filters (25 mm diameter, Whatman GFB). 8. 0.5 mL microcentrifuge tubes (with cap). 9. Scintillation vials and scintillation liquid. 10. Bicinchoninic acid (BCA, Pierce). 11. Filtration apparatus (e.g., single or ten-place filtration manifold with stainless steel chimney weights to hold prewetted 25 mm filters in place; Hoefer). 12. House vacuum or vacuum pump. 13. 4 mL homogenization solution (0.1 M NaCl, 0.3 M sucrose, 100 MM EGTA, 1 mM oxidized glutathione (GSSG), 20 mM imidazole, pH 7) for 100 mg tissue (cellular pellet) (see Note 1). Adjust to pH 7 with 0.1 M HCl or 0.1 M NaOH. Place on ice and add protease inhibitor cocktail (1:100 dilution) to homogenization solution before use. 14. (3H)Ryanodine-binding solutions for determination of Bmax value (a) Stock solution (0.6 M KCl, 20 mM imidazole, pH 7, 2 mM GSSG): Combine 0.42 mL of 2 M KCl, 1.7 mg (GSSG), 28 ML of 1 M imidazole, pH 7. Add deionized water to make 1.4 mL solution and check pH. If necessary, readjust to pH 7 with 0.1 M HCl or 0.1 M KOH. Add protease inhibitor cocktail (1:50 dilution). (b) Stock solution containing 60 nM (3H)ryanodine: Place 42 ML of 1 MM (3H)ryanodine in ethanol in 10 mL glass tube and evaporate ethanol by placing a Pasteur pipette connected to a nitrogen tank several centimeters above the

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sample. Prior to evaporating ethanol, adjust N2 flow by placing Pasteur pipette above a 1 mL ethanol control solution in glass tube. N2 should indent ethanol solution without causing splashing. After removal of ethanol from (3H) ryanodine sample, add 0.7 mL stock solution. To fully dissolve (3H)ryanodine, keep solution at room temperature for 20–30 min with occasional slight shaking. (c) Solution for determination of total (3H)ryanodine binding: 0.3 mL of stock solution containing 60 nM (3H)ryanodine, 3 ML of 10 mM EGTA,12 ML of 10 mM CaCl2. (d) Solution for determination of nonspecific (3H)ryanodine binding: 0.3 mL of stock solution containing 60 nM (3H) Ryanodine, 15 ML of 100 mM EGTA, 18 ML of 100 MM ryanodine in H2O. (e) Wash solution (0.1 M KCl, 1 mM imidazole, pH 7): Add 5 mL of 2 M KCl, 0.1 mL of 1 M imidazole, pH 7–95 mL deionized H2O. (f) Homogenates (see Subheading 3.3). 2.3. Detection of Ca2+ Transients Mediated by RyRs (see Note 2)

1. An inverted microscope equipped with a 40× oil objective suitable for UV light source. 2. A xenon lamp coupled to a monochromator (e.g., DeltaRam X from Photon Technology International) or a filter wheel that allows rapid excitation filter changes (Prior Scientific) exciting Fluo-4 at 465 nm and Fura-2 alternatively at 340 and 380 nm (see Notes 3 and 4). 3. Standard fluorescence emission filter sets (10). 4. A charge-coupled device camera to capture emitted fluorescence in the outline cytoplasmic areas. 5. Imaging software (e.g., EasyRatioPro (PTI) or SimplePCI) (Hammamatsu Photonics). 6. Perfusion setup (e.g., cFlow, Cell MicroControls). 7. Fluo-4 acetoxymethyl ester (Fluo-4-AM) and Fura-2AM. 8. Glass-bottom culture dishes. 9. Ryanodine. 10. RyR-activating agents caffeine and 4-chloro-m-cresol (4-CmC, for activation of RyR1 or RyR2 (11)). 11. PBS: 140 mM NaCl; 2 mM KCl; 1.5 mM KH2PO4; 15 mM Na2HPO4, pH 7.4. 12. Krebs-Ringer-Henseleit (KRH) buffer: 125 mM NaCl, 5 mM KCl, 1.2 mM KH2PO4, 6 mM glucose, 1.2 mM MgCl2, 2 mM CaCl2, and 25 mM HEPES, pH 7.4. 13. M199 media (Gibco).

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3. Methods 3.1. Localization of RyRs in Isolated Cells (12)

1. Wash the cultured cells or freshly isolated cells in individual wells of chamber glass slides (see Note 5) with PBS twice, fix with 2% paraformaldehyde in PBS for 15 min, and then wash with PBS two more times. 2. Quench the remaining paraformaldehyde with the quenching solution for 30 min. Afterward, wash the cells once with PBS, treat them with permeabilizing solution for 15 min, and wash them once more. 3. Apply the blocking solution for 30 min. Then wash off the blocking solution. 4. Probe the cells with individual anti-RyR antibody in 1% goat serum/PBS for more than 1 h. (see Note 6). The suggested dilutions for these antibodies are provided by vendors. 5. After washing off the primary antibody, incubate the cells with the corresponding fluorochrome-labeled secondary antibody (7.5 ng/ML) in 1% goat serum/PBS for 1 h. 6. Wash the cells once more. The cells can also be counterstained with DAPI which stains the nuclei. Mount them on ProLong Antifade solution and cover the slides with cover glasses. 7. 12 h later, seal the edge of the cover glass by applying a nail polish. Store the slide in −20°C freezer until viewing.

3.2. Immunofluorescent/Immuno histochemical Detection of RyRs in Tissues (see Note 7)

1. Freshly isolated tissues should be immersed in OCT Compound in Cryomold and rapidly frozen in liquid nitrogen. Cut the frozen tissues into 5 Mm sections onto glass slides and store at −80°C until the day of study. 2. Dry the tissue sections in room temperature over 10 min. Fix them with 2% paraformaldehyde in PBS solution for 10 min, then wash two more times with PBS. 3. Quench the remaining paraformaldehyde with quenching solution and then apply the permealizing solution for 30 min (see Note 8). 4. Block the tissue sections with the blocking solution for 30 min and then wash them with PBS once. 5. For immunofluorescent staining, incubate the tissue sections with the primary antibody in 1% goat serum for more than 1 h (see Note 6). Wash off the primary antibody, then incubate the tissue sections with the corresponding fluorochromelabeled secondary antibody (7.5 ng/ML) in 1% goat serum/ PBS for 1 h. 6. The tissue sections can also be counterstained with DAPI which stain the nuclei. Mount them on ProLong Antifade solution and preserve the slides until viewing.

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7. For enhanced detection of RyRs in tissues expressing low levels of RyRs, immunohistochemical staining may be preferred (13). Incubate the tissue sections with individual primary antibody for more than 1 h, and then develop the staining using the VectaStain Elite kit according to the manufacturer’s instructions. Each slide can then be lightly counterstained with hematoxylin-eosin. 3.3. Preparations of Homogenates

1. Add tissue or cellular pellets to 30–40 volumes of homogenization solution, place homogenizer probe in solution, and homogenize at preset speed of 13,000 rpm for twice for 5 s or until tissue is fully broken up. 2. Take three 5 ML aliquots for protein determination using BCA method, and store aliquots of homogenate at −80°C. Store one 150 ML aliquot for (3H)ryanodine-binding assay.

3.4. ( 3H)RyanodineBinding Assay (see Notes 9–12)

1. Add to each of three 0.5 mL microcentrifuge tubes, 70 ML of solutions for determination of total and nonspecific (3H)ryanodine binding (6 vials per homogenate). Add 0.5 mL of (3H)ryanodine binding stock solution to 0.15 mL thawed homogenate and homogenize using glass/glass homogenizer. Add 70 ML of diluted homogenates to microcentrifuge tube, cap tubes, shake gently, and keep for 4 h in the dark at room temperature (~24°C). 2. Add 25 ML of Stock solution containing 60 nM (3H)ryanodine into 2 vials each and 5 mL scintillation liquid (2 vials total) for determination of radiospecificity (disintegrations per min (dpm)/nmol). 3. Add 1 g polyethyleneimine (50% solution in water) to 25 mL deionized water. Add stir bar and stir until a homogenous solution is obtained. Add six Whatman GFB filters to solution, and soak filters for 1 h or more at room temperature. 4. Place filter preincubated with 2% polyethyleneimine in water on filter support screen, place on top of filter chimney weight, and apply vacuum. 5. Add 120 ML sample to 0.72 mL ice-cold deionized H2O in glass tube (kept on ice), place diluted sample on center of filter, and wash filter 3 times with 5 mL of ice-cold 100 mM KCl, 1 mM imidazole, pH 7.0 solution. 6. Place filter into scintillation vial. After all samples have been filtered, add 5 mL scintillation liquid. Place scintillation vials on a shaker overnight, and count total and nonspecifically bound (3H)ryanodine, using a scintillation counter.

3.5. Detection of Ca2+ Transients Mediated by RyRs (see Notes 2 and 13)

1. Culture cells in glass-bottom culture dishes. For studies of freshly isolated cells, allow 1 h for the cells to adhere after plating the cells.

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Fig. 1. Demonstration of functional recombinant RyR1 in HEK cells. HEK293 cells expressing RyR1 were loaded with Fluo-4AM and subsequently challenged with 5 mM caffeine. Ca2+ transients were measured in two cells with DeltaRam X from Photon Technology International, using EasyRatioPro (PTI) (A. Chakraborty, unpublished studies).

Certain cell types may need precoating of the glass-bottom dishes with an extra cellular matrix material (see Note 5). 2. Wash off the culture media with PBS. 3. Load the cells with 5 MM Fluo-4-AM or 10 MM Fura-4-AM in M199 media for 1 h. Then wash the cells with KRH buffer to remove excess Ca2+ indicator. 4. Functional RyRs can be detected by applying 5–40 mM of caffeine (final concentration) (14) or 0.5 mM 4-CmC (for RyR1 and RyR2) (11). See Fig. 1 (see Note 14). 5. For determination of RyRs’ role in Ca2+ responses provoked by ligands, one may compare the ligand-induced Ca2+ responses of cells with or without pretreatment of 100–200 MM ryanodine (15).

4. Notes 1. Prepare all solutions using deionized water and analytical grade reagents. Store all stock solutions at 4°C. 2. RyRs may spontaneously open resulting in an elevated Ca2+ concentration over short distances (16). Formation of localized Ca2+ release events (Ca2+ sparks) can be detected after loading with Fluo-4AM using laser scanning confocal microscope (17). 3. Fluo-4 has higher sensitivity for Ca2+ transients than the traditional ratiometric dyes such as Fura-2 and Indo-1. Ratiometric Ca2+ dyes, however, are considered more accurate because the end results are less affected by technical issues such as dye load-

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ing. For a more complete discussion of various Ca2+ dyes and the setups required, see references (10) and (18). 4. In cells using Fura-2, intracellular Ca2+ concentrations can be calculated using the equation (Ca)i = Kd (F380max/F380mn (R−Rmin) (Rmax−R)). A dissociation constant (Kd) of 224 nM is used for the binding of Ca2+ to Fura-2 at 37°C (19, 20). Rmax and Rmin are determined by the consecutive addition of 10 MM ionomycin and 50 mM EGTA. 5. Allow freshly isolated cells to adhere to the glass slides under normal cell culture conditions for about an hour. If necessary, coat glass slides with extracellular matrices (e.g., laminin). 6. It is recommended that each experiment also contains at least one negative control in which the primary antibody is omitted or the primary antibody is pretreated with a blocking peptide for that antibody (13). In addition, for detection of RyRs in tissues, small pieces of muscles from the cardiac ventricle and diaphragm can be used as positive controls for RyR2 and RyR1/RyR3, respectively (21). 7. Formalin-fixed tissues, even after applying antigen-retrieval method, may not be ideal for immunofluorescent/immunohistochemical detection of RyRs as formalin fixation and storage can significantly decrease the antigenicity of RyR proteins. 8. Before probing the mouse tissue sections with a mouse antiRyR antibody, the tissue sections should be treated using the Mouse-on-mouse kit according to manufacturers’ instructions to reduce background staining from endogenous immunoglobulins in the tissues. 9. Calculation of Bmax value [3 H]Ry (pmol/ml) dpm / 0.025ml r 40

dpm nonspecific ) / filter

pmol bound [3 H]Ry / mg protein  r

(dpm total

mgprotein / filter

10. The tetrameric RyRs bind one ryanodine molecule with nanomolar affinity. Accordingly, a Bmax value of 1 pmol/mg protein corresponds to the presence of 1 pmol RyR per mg cellular protein. 11. Typical values for adult mouse skeletal muscle and cardiac muscle are 0.05–0.2 pmol/mg cellular protein. These values are well within the detection limit of the assay (0.01 pmol/mg protein) to allow quantification of RyR protein levels. 12. Sodium dodecyl sulfate gel electrophoresis (SDS PAGE) and immunoblot analysis are an alternative method to quantify RyR protein levels. It is recommended that homogenate and

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membrane fractions are solubilized in the Laemmli separation system (22) using a 3–12% linear gradient polyacrylamide gel and 3% stacking gel to allow a sufficient migration of the high molecular weight RyR polypeptide into the separating gel. Samples are denatured for 10 min at 65°C in 0.1 M Tris–HCl, pH 6.8, containing 2% SDS, 5% B-mercaptoethanol, and 10% glycerol. For immunoblots, the separated proteins from SDS PAGE are electrophoretically transferred to PVDF membranes for 1 h at 400 mA and then for 20 h at 1,500 mA, using transfer buffer which contains 0.1% SDS. To ascertain complete transfer, gels are stained with 0.1% Coomassie Brilliant Blue R-250 in 50% methanol, 10% acetic acid and destained with 10% methanol, 15% acetic acid. 13. Following the demonstration of RyR expression in certain cell types, the general strategy to determine whether RyRs play a role in a physiological response in those cells is to first demonstrate that RyRs are functional using RyR-activating agents caffeine and/or 4-CmC. Subsequently, the physiological Ca2+ response should be suppressed or eliminated by pretreating these cells with a high dose of ryanodine (100–200 MM) for more than 5 min. Alternatively, one can compare the physiological Ca2+ responses of cells deficient in RyRs and their “wildtype” counterparts. 14. Since the activation of RyRs by caffeine and 4-CmC is reversible, it is possible to remove these reagents and rechallenge the RyR-expressing cells with other agents (11).

Acknowledgment Support by NIH grants to JE (HL081825) and GM (AR018687 and HL073051) is gratefully acknowledged. References 1. Franzini-Armstrong C, and Protasi F (1997) Ryanodine receptors of striated muscles: a complex channel capable of multiple interactions. Physiol Rev 77:699–729 2. Hamilton SL, Serysheva II (2009) Ryanodine receptor structure: progress and challenges. J Biol Chem 284:4047–4051 3. Meissner G (2002) Regulation of mammalian ryanodine receptors. Frontiers in Bioscience 7:d2072–2080 4. Patterson RL, Boehning D, Snyder SH (2004) Inositol 1,4,5-trisphosphate receptors as signal integrators. Annu Rev Biochem 73:437–465

5. Fill M, and Copello JA (2002) Ryanodine receptor calcium release channels. Physiol Rev 82:893–922 6. Sutko JL, Airey JA, Welch W, Ruest L (1997) The pharmacology of ryanodine and related compounds. Pharmacol Rev 49:53–98 7. Xu L, Tripathy A, Pasek DA, Meissner G (1998) Potential for pharmacology of ryanodine receptor/calcium release channel. Ann New York Acad Sci 853:130–148 8. Rios E, Pizarro G (1991) Voltage sensor of excitation-contraction coupling in skeletal muscle. Physiol Rev 71:849–908

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9. Fabiato A (1983) Calcium-induced release of calcium from the cardiac sarcoplasmic reticulum. Am J Physiol 245:C1–C14 10. Poeni M (2000) Flourescent calcium indicators based on BAPTA. in Calcium Signaling (Putney JW, Ed.), CRC Press, 1–46 11. Fessenden JD, Feng W, Pessah IN, Allen PD. (2006) Amino acid residues Gln4020 and Lys4021 of the ryanodine receptor type 1 are required for activation by 4-chloro-m-cresol. J Biol Chem 281:21022–21031 12. Du W, Stiber JA, Rosenberg PB, Meissner G, Eu JP (2005) Ryanodine receptors in muscarinic receptor-mediated bronchoconstriction. J Biol Chem 280:26287–26294 13. Du W, McMahon TJ, Zhang ZS, Stiber JA, Meissner G, Eu JP. (2006) Excitationcontraction coupling in airway smooth muscle. J Biol Chem 281:30143–30151 14. Wang Y, Xu L, Duan H, Pasek DA, Eu JP, Meissner G (2006) Knocking down type 2 but not type 1 calsequestrin reduces calcium sequestration and release in C2C12 skeletal muscle myotubes. J Biol Chem 281:15572–15581 15. Dai JM, Kuo KH, Leo JM, Pare PD, van Breemen C, Lee CH (2007) Acetylcholineinduced asynchronous calcium waves in intact

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human bronchial muscle bundle. Am J Respir Cell Mol Biol 36:600–608 Cheng H, Lederer WJ (2008) Calcium sparks. Physiol Rev 88:1491–1545 Pouvreau S, Royer L, Yi J, Brum G, Meissner G, Rios E, Zhou J (2007) Ca2+ sparks operated by membrane depolarization require isoform 3 ryanodine receptor channels in skeletal muscle. Proc Natl Acad Sci USA 104:5235–5240 Patel S, Robb-Gaspers LD, Thomase AP (2000) Measuring Single Cells and subcellular Ca2+ signaling. In Calcium Signaling (Putney JW, Ed.), CRC Press, pp. 343–364. Grynkiewicz G, Poenie M, Tsien RW (1985) A new generation of Ca2+ indicators with greatly improved fluorescence properties. J Biol Chem 260:3440–3445 Quinn SJ, Williams GH, Tillotson DL (1988) Calcium oscillations in single adrenal glomerulosa cells stimulated by angiotensin II. Proc Natl Acad Sci USA 85:5754–5758 Flucher BE, Conti A, Takeshima H, Sorrentino V (1999) Type 3 and type 1 ryanodine receptors are localized in triads of the same mammalian skeletal muscle fibers. J Cell Biol 146:621–630 Laemmli UK (1970) Cleavage of structural proteins during the assembly of the head of bacteriophage T4. Nature 227:680–685

Chapter 22 Measurement of Calcium Release Due to Inositol Trisphosphate Receptors in Skeletal Muscle Mariana Casas, Francisco Altamirano, and Enrique Jaimovich Abstract Calcium transients elicited by IP3 receptors upon electrical stimulation of skeletal muscle cells (slow calcium signals) are often hard to visualize due to their relatively small amplitude compared to the large transient originated from ryanodine receptors associated to excitation-contraction coupling. The study of slow calcium transients, however, is relevant due to their function in regulation of muscle gene expression and in the process of excitation-transcription coupling. Discussed here are the procedures used to record slow calcium signals from both cultured mouse myotubes and from cultured adult skeletal muscle fibers. Key words: Myotubes, Cultured muscle fibers, Calcium dyes, Electrical stimulation, Gene expression, Slow calcium signals

1. Introduction The notion of inositol 1,4,5 trisphosphate (IP3) receptors (IP3Rs) releasing calcium upon stimulation of skeletal muscle was originated from Julio Vergara’s laboratory (1) who suggested that muscle fibers were capable of producing IP3 in response to electrical stimulation (ES) and proposed a role for IP3 in the process of muscle excitation-contraction coupling. Several laboratories followed that lead (reviewed in (2)) and reached the conclusion that calcium release by IP3 was not fast enough to account for the needs of contracting sarcomeres. The evidence gathered nevertheless pointed to the fact that all the molecular machinery needed to synthesize and degrade IP3 was present in the muscle cells and actually calcium was released by IP3 in permeabilized fibers (3) and this release depended on membrane potential! A first hint of a new calcium signal in cultured muscle cells was published in 1994 (4) when

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calcium transients elicited by potassium depolarization of myotubes were shown to have two components, the slower one highly sensitive to nifedipine. The location of type 1 receptor in cultured myotubes was studied (5, 6), together with detailed description of IP3dependent slow calcium transients (see also (7–9)) in rat and mouse myotubes with an important component of the signal occurring at the myonuclei (10). The IP3-dependent slow calcium signal was associated to signaling pathways leading to transcription factor activation. MAP kinases and CREB phosphoryation as well as increase in mRNA for early genes c-fos, c-jun, and egr-1 was confirmed (6, 11, 12) and a role for IP3 regulating the activity of transcription factors in skeletal muscle cells was proposed. This role was reinforced when studying expression of specific genes as interleukin 6 (IL6, (13)) and more than a 100-genes studied using microarrays (and confirming some of them using RT-PCR) (14). More recently, a mechanism involving Cav1.1 voltage sensor (12), ATP release from the muscle cell through pannexin 1 channels, P2Y purinergic receptors (15), G protein, PI3K, and PLC (8) has been proposed (see Note 2). The recent work in adult muscle fibers (16) shows the presence of IP3-dependent calcium transients in adult muscle fibers, sharing many characteristics of the signals described in cultured myotubes (see Fig. 2, Note 3). The methods described below show how to detect slow, IP3dependent calcium signals in both cultured muscle cells and in cultured adult skeletal muscle fibers.

2. Materials 2.1. Cultures of Myoblasts from Neonatal Skeletal Muscle

1. 5–8 neonatal mice (1–3 days old). 2. Sterile surgical material. 3. Phosphate-buffered saline (PBS), 0.22 Mm filtered. 4. Collagenase type II (Worthington Biochemical) solution at 1 mg/mL in PBS, 0.22 Mm filtered. 5. Proliferation medium: F-10 Nutrient mixture medium (Gibco) supplemented with 5 ng/mL human FGF-basic (Preprotech), 20% bovine growth serum (BGS) or 20% fetal bovine serum (FBS, Hyclone), and 1× Penicillin-Streptomycin-Glutamine solution (Gibco). 6. Differentiation medium: DMEM-low glucose (Gibco) supplemented with 4% horse serum (HS, Gibco) and 1× PenicillinStreptomycin-Glutamine solution (Gibco). 7. Collagen type I rat tail, high concentration (BD Bioscience) solution at 0.2 mg/mL. 8. Nytex filter.

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9. Petri dishes, 100 mm. 10. Trypsin-EDTA solution, 1× (Hyclone). 2.2. Isolation of Adult Muscle Fibers from Mice

1. Phosphate-buffered saline (PBS), sterile. 2. Dissection chamber filled with Silgar (60 cm Petri dishes are good enough). 3. Steel pins. 4. Pasteur pipettes cut to obtain pipettes at different diameters (from 0.5 to 3 mm), fire polished. Choose pipettes made of thick glass to avoid closing them during the fire polish. 5. Dissection instruments: iridissection scissors, tweezers. 6. Mice from 3 to 20 weeks old. 7. Collagenase type II (Worthington Biochemical). 8. Matrigel (prepared in our lab from published protocols (17)). 9. Horse serum (HS, Gibco). 10. Dulbecco’s modified Eagle medium (DMEM, Gibco) with 1% penicillin/streptomycin. 11. Dulbecco’s modified Eagle medium with 1% penicillin/streptomycin supplemented with 10% HS.

2.3. Calcium Signal Measurement

1. Krebs physiological solution: 140 mM NaCl, 5 mM KCl, 1 mM CaCl2, 1 mM MgCl2, 5.6 mM glucose, 10 mM HEPES-Tris pH 7.4. Krebs solution without Ca2+ has essentially the same composition, but contains 2 mM MgCl2 and 0.5 mM EGTA. 2. Calcium indicator Fluo3-AM: 2–5 MM in Krebs solution. Fluo3-AM solution is made by dissolving 50 Mg of lyophilized compound in 20 ML of 20% pluronic acid in dimethyl sulfoxide (DMSO) and then adding Krebs solution to obtain the desired Fluo3 concentration. 3. Electrical stimulator (e.g., Grass S48). 4. Confocal or epifluorescence microscope with image acquisition capability. 5. 25 MM nifedipine. 6. 5 MM Xestospongin C or 5 MM Xestospongin B.

3. Methods 3.1. Cultures of Myoblasts from Neonatal Skeletal Muscle

Primary myoblasts can be easily isolated from neonatal mice and grown in proliferation medium. Myoblasts can be differentiated into myotubes with a reduction of serum and can be used for experimental calcium determination on day 3–5 of differentiation (18).

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1. Sacrifice the neonatal mice by rapid decapitation with surgical scissors and then submerge the body in 70% ethanol for a few seconds in order to kill the skin bacteria and fungus. 2. Remove the hind limbs with scissors and remove the skin with surgical tweezers. Then with small tweezers remove the muscle tissue and discard the bones. Cut the muscle into small pieces and enzymatically dissociate by adding collagenase solution (10 mL) for 15 min at 37°C. Then, repetitively aspirate the solution with a sterile syringe (without needle) in order to mechanically dissociate the tissue, and leave the suspension at 37°C for an additional 15 min to complete the tissue disintegration. Add 10 mL of proliferation medium, filter the solution with Nytex paper (in order to remove the undigested material), and centrifuge for 10 min to pellet the cells. Carefully resuspend the pellet in 10 mL of proliferation medium. 3. In order to remove fibroblasts, preplate the cell suspension in 100 mm Petri dishes at 37°C, 5% CO2. After 1 h incubation, remove the cells in suspension (myoblasts) and preplate on PBS washed, collagen-treated 100 mm Petri dishes for another 30 min to remove fibroblasts. Plate the cell suspension in collagen-treated Petri dishes and incubate at 37°C, 5% CO2. Change the proliferation medium daily until you get a population highly enriched in myoblasts. 4. Trypsinize the cells and repeat the preplating on plastic Petri dishes if fibroblasts remain in the cell culture (1 h, 37°C, 5% CO2). Plate the cell suspension in order to increase the myoblast population for further experiments. 5. For long-term storage, myoblasts can be frozen in liquid nitrogen. Trypsinize a 100-mm dish at 70–80% of confluence, pellet the cells, and resuspend in 1 mL of proliferation media supplemented with 10% culture grade DMSO. Freeze the cells using an isopropanol chamber at -80°C and then in liquid nitrogen. 3.2. Myotube Differentiation

1. For calcium determinations by fluorescence microscopy, plate the cells on matrigel-coated fluorescence suitable plastic covers (in glass covers the myotubes are easily detached upon spontaneous contraction) with 30% confluence. 2. Change the proliferation medium daily until the culture has a confluence of 70–80%. 3. In order to differentiate the myoblasts, change the media to differentiation medium and incubate the cells at 37°C, 5% CO2. Change the medium daily and you will obtain differentiated myotubes at day 2–4.

3.3. Isolation of Adult Muscle Fibers from Mice

1. Sacrifice the animals by cervical dislocation. 2. Remove skin from hind limb by cutting it from the proximal end of the limb (above the knee) until the ankle. Place the

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limbs without skin in PBS and fix it with steel pins to a dissection chamber filled with Silgar; add PBS to cover the piece. Perform the following steps under a low amplification microscope and under a laminar flow hood. 3. Expose the flexor digitorum brevis (FDB) muscle by delicately cutting the skin in the hind paw. Muscle dissection must be done avoiding cutting muscle fibers. 4. Place the muscles in a solution of DMEM with 1% penicillin/ streptomycin (without serum) and with 450–500 U/mL of collagenase at 37°C for 90 min. Three to four milliliter of solution is enough for 2 FDB muscles. No shaking is needed. 5. Prepare plates or coverslips covered with Matrigel 30 min before seeding fibers. 10–15 ML of Matrigel is enough for a 35 mm Petri dish. 6. Before collagenase digestion is finished, prepare two 35 mm Petri dishes by coating them with HS or a solution of 5% BSA. Remove the excess and place 2 mL of 10% HS in DMEM. This procedure prevents fibers from sticking to the Petri dish during the dissociation step. 7. After collagensase digestion, place muscles in a Petri dish prepared as above and dissociate fibers by passing the muscle through the fire-polished Pasteur pipettes, successively from large to small diameters. 8. Seed the isolated fibers on Matrigel-coated cover slips. Place the desired quantity of fibers in a small volume of medium, wait 5–10 min for fibers to sediment, and adhere to the Matrigel and then delicately add DMEM supplemented with 10% HS. Fibers can be used immediately, but are fragile because of collagenase digestion. We usually do experiments 20–30 h after seeding. This protocol is based on a protocol described in (19). 3.4. Calcium Signal Measurement

The choice of a Ca2+ probe must take into account the magnitude of the signal to be recorded. For a slow calcium signal, probes with high affinity, like Fluo3 (Kd = 390 nM) and Fluo4 (Kd = 350 nM), should be used. They have the sensitivity needed to detect small amounts of Ca2+ and display up to 100-fold Ca2+-dependent fluorescence enhancement (dynamic range), allowing visualization of low amplitude Ca2+ variations. Figure 1 shows calcium signals obtained in mouse myotubes (see Notes 1 and 2 for more details). Figure 2 shows a representative calcium signal evoked by electrical stimuli and the respective kinetic analysis (see Note 3). Nevertheless, they are not adequate for ratiometric measurements or Ca2+ concentration quantization. 1. Wash the covers with myotubes (2–4 days after change to differentiation medium) with 37°C prewarmed Krebs buffer

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Fig. 1. Calcium transient evoked by electrical stimuli in differentiated C57Bl/6 myotubes. Cells were preloaded with 5 MM Fluo-3AM and stimulated at 250 pulses of 0.5 ms of duration at 20 Hz frequency in Krebs buffer at RT. Image acquisition was obtained with an inverted fluorescence microscope equipped with a CCD cooled camera every 0.7 s for 3 min. Representative Fluo-3AM fluorescence images are shown at the indicated times and the quantification of the fluorescence for fast and slow calcium signals is shown at the bottom. (a) Myotube showing mainly EC-coupling-related Ca2+ signal (representative of 8% of cells at this stage). (b) Myotube showing fast calcium transient and a prominent slow calcium signal (representative of 68% of cells at this stage). (c) More differentiated myotube (note the larger amount of nuclei and branching) showing a fused signal with a fast and a slow component (representative of 12% of cells at this stage). Bar indicates the electrical stimuli duration.

solution. Do not carry out this step for adult muscle fibers for risk of detachment. Simply remove the excess medium from coverslips and load it with the Ca2+ probe. 2. Load the cells with a Krebs solution containing 2–5 MM of Fluo3-AM. Incubate the cells at 37°C for 25 min. In the case of adult muscle fibers, incubate 30 min at room temperature (20–22°C). 3. Wash the cells with Krebs solution and place the coverslips in a microscope chamber. 4. Apply ES with a couple of platinum electrodes connected through an isolation unit to a stimulator. Different duration and frequencies for trains of 0.3 ms square pulses can be used in adult fibers (0.5 ms for myotubes). During stimulation experiments, maintain the myotubes or fibers in Krebs solution at 21–23°C. Experiments with no extracellular calcium can be performed in the same Krebs solution without calcium and supplemented with 0.5 mM EGTA. 5. With the chamber of myotubes or fibers in a confocal or epifluorescence microscope, set the excitation source at

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Fig. 2. Slow Ca2+ signal induced by electrical stimuli in adult skeletal muscle fibers. Isolated muscle fibers from mouse FDB, loaded with Fluo3-AM, were stimulated with a train of 270 pulses (0.3 ms each) at 20 Hz. Images were obtained by confocal microscopy. (a) Images of fiber fluorescence before, during, and at different times after application of electrical stimulus (end of stimulation indicated by time = 0 s). (b) The graph shows the variations in relative fluorescence values with time in a fiber stimulated as in (a). We observed a first fast Ca2+ signal related to contraction when a tetanic train was applied (bar in the graph) followed by a slower signal, observed as a delayed return to basal fluorescence levels after tetanic stimulus. Experiments were done in the presence of standard Krebs solution containing 1 mM calcium. Experimental points could be fitted by a double exponential function (equation and continuous line in (b)) as shown for a typical record in the graph (b). (c) Single twitch can be fitted by a single exponential decay (equation and continuous line in (c)) as shown for a representative record. (d) When muscle fibers are incubated for 30 min with 20 MM Xestospongin B, a specific inhibitor of IP3R (16), slow calcium signal is completely abolished (filled squares, mean of n = 4). Empty squares correspond to mean of four signals obtained from control fibers.

488 nm with the minor laser or lamp emission in order to avoid photobleaching. Collect the emitted light at 526 nm. 6. Start acquiring images. Take several images before stimulation to estimate the basal calcium and then stimulate the cells with the electrical field. Acquire for 2 min after ES in case of adult muscle fibers and for 5 min for myotubes. Collect the fluorescence images every 1.0–2.0 s and analyze frame by frame. This

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acquisition speed is short enough to resolve the slow signal. For rapid image acquisitions, needed to resolve a fast Ca2+ signal (related to EC coupling) or a slow signal after a short tetanus, data must be collected by line scan of fibers every 1–2 ms. These data can be analyzed with the software program WinWCP (J Dempster, Strathclyde University). 7. Frame-by-frame fluorescence image analysis can be made using the public domain Image J software (NIH, Bethesda). The average cell fluorescence, F, is calculated by setting a region of interest (ROI) for the image series and normalized to its initial or preintervention value F0 as (F−F0)/F0. 8. To block the dihydropyridine receptor (DHPR), add 25 MM nifedipine to the physiological medium for 30 min. To specifically block IP3 receptors, apply either 5 MM Xestospongin C or 5 MM Xestospongin B (see Fig. 2) for 20 or 30 min, respectively, to fiber preparations. Both toxins have been reported to be effective (20), although some variability between batches makes it advisable to test the alternative toxin in case of negative results. We have not studied in detail IP3-dependent, slow calcium signals during muscle development and differentiation. It is worth noting though that calcium signals tend to evolve in myotubes from a distinct late calcium transient, to a fused signal comprising the fast and slow calcium transients (Fig. 1). The latter is also the case for adult skeletal muscle fibers (Fig. 2). These observations suggest that the shape of the slow calcium transient may be related to structural characteristics of the skeletal muscle cell, which may include the degree of development of the transverse tubular system.

4. Notes 1. As mentioned before, we studied the calcium kinetics in neonatal myotubes after several stimuli. We found that ES induces RyR and IP3R calcium release to the cytosol in a biphasic manner (8). In order to discriminate these signals, we used Fluo-3AM single-wavelength calcium indicator that has a 390-nM Kd for calcium. In summary, we incubated the cells with the probe for several minutes and then studied the calcium transients evoked by electrical stimuli in live cells by fluorescence microscopy. Calcium increase during the fast signal (RyR-dependent) occurs rapidly, has a sustained plateau during tetanic stimuli, and a fast decay at the end of the stimuli. Slow calcium (IP3Rdependent) signals are more variable in both the onset and the amplitude of the signal. We illustrate these events in Fig. 1 and show representative calcium measurements obtained in

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differentiated myotubes from C57 mice. A small proportion (8%) of cells has only the fast calcium component with no evidence of the slow calcium signal (Fig. 1a). The slow calcium signal appears as a second increase after excitation-contraction coupling or fast calcium signal in the majority of cells at this stage (Fig. 1b, 68% of the cells tested). In more differentiated myotubes, judged by size, abundance of nuclei, and branching, slow Ca2+ signal is observed as a slow decay of the fast calcium transient similar to that seen in adult myofibers (Fig. 1c, 14% of the cells). 2. Primary differentiated myotubes are a suitable model to study muscle physiology, gene expression, and calcium transient evoked by several stimuli or basal calcium modifications (8, 13, 15, 21, 22). We demonstrated that tetanic ES in myotubes produces a biphasic increase in intracellular Ca2+. The first increase or fast Ca2+ transient is related to excitation-contraction coupling. The second increase or slow Ca2+ transient is generated by DHPR/GBG protein/PI3K/PLC activation and IP3 receptor (IP3R) Ca2+ release and the participation of ATP release and ATP signaling by purinergic receptors (8). 3. In adult muscle fibers, a slow Ca2+ signal is visualized as a delayed return to basal fluorescent levels after the end of the tetanus (16). This posttetanic signal can be fitted to a double exponential decay. This kind of analysis allows for some quantification of the signal, characterization, and comparison in different experimental situations. In this respect, image acquisition speed has a direct influence on time constants decay calculation, imposing a limit for this time constant determination. In order to have an actual value for time constant, line scan records (500–1,000 Hz acquisition speed) are recommended. The slow Ca2+ signal doesn’t have an important Ca2+ entry component, as no differences are observed in signal when experiments are done in Krebs without Ca2+ and 0.5 mM EGTA. The slow Ca2+ signal is dependent on number of pulses and frequency of the electrical stimulus. At a fixed frequency, the slow Ca2+ signal increases with an increase in the number of pulses applied, becoming difficult to detect when the number of pulses is less than 20. Slow signal has a maximum amplitude at 10–20 Hz, being smaller at lower and higher frequencies (16).

Acknowledgments This work was supported by FONDECYT grant N° 1080120 and FONDAP grant N° 15010006; CONICYT 24100066 Doctoral Support (F.A.).

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22. Eltit, J. M., Yang, T., Li, H., Molinski, T. F., Pessah, I. N., Allen, P. D., and Lopez, J. R. (2010) RyR1-mediated Ca2+ leak and Ca2+ entry determine resting intracellular Ca2+ in skeletal myotubes, J Biol Chem 285, 13781– 13787.

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Chapter 23 Detection of Calcium Sparks in Intact and Permeabilized Skeletal Muscle Fibers Noah Weisleder, Jingsong Zhou, and Jianjie Ma Abstract Ca2+ sparks are the elementary units of Ca2+ signaling in striated muscle fibers that appear as highly localized Ca2+ release events through ryanodine receptor (RyR) Ca2+ release channels in the sarcoplasmic reticulum (SR). While these events are commonly observed in resting cardiac myocytes, they are rarely seen in resting skeletal muscle fibers. Since Ca2+ spark analysis can provide extensive data on the Ca2+ handling characteritsics of normal and diseased striated muscle, there has been interest in developing methods for observing Ca2+ sparks in skeletal muscle. Previously, we discovered that stress generated by osmotic pressure changes induces a robust Ca2+ spark response confined in close spatial proximity to the sarcolemmal membrane in wild-type intact mammalian muscles. Our studies showed these peripheral Ca2+ sparks (PCS) were altered in dystrophic or aged skeletal muscles. Other methods to induce Ca2+ sparks include permeabilization of the sarcolemmal membrane with detergents, such as saponin. In this chapter, we will discuss the methods for isolation of muscle fibers, the techniques for inducing Ca2+ sparks in these isolated fibers, and provide guidance on the analysis of data from these experiments. Key words: Calcium, Spark, Burst, Skeletal muscle, Permeabilization, Saponin, Osmotic shock, Osmotic stress, Myocyte, Muscle fiber, Ryanodine receptor

1. Introduction The elementary units of Ca2+ release from sarcoplasmic reticulum (SR) in striated muscle fibers are discreet events known as Ca2+ sparks. Ca2+ sparks were first discovered in cardiac muscle as localized quantal Ca2+ release events originating from arrays of ryanodine receptor (RyR) Ca2+ release channels inserted into the SR membrane. In cardiac muscle, they represent the elemental units of Ca2+-induced Ca2+ release (CICR) necessary for cardiac contractility (1–3). The discovery of Ca2+ sparks revolutionized our understanding of the physiology and pathophysiology of Ca2+ signaling in cardiac

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and smooth muscles (4–7). Ever since the discovery of Ca2+ sparks in cardiac muscle, investigators have had difficulty in detecting Ca2+ sparks in adult mammalian skeletal muscle where voltage-induced Ca2+ release (VICR) is the dominant mode of Ca2+ release from the SR (8). Initial studies detecting Ca2+ sparks in skeletal muscle were performed with amphibian muscle (9, 10) or neonatal mammalian skeletal muscle (11) where they were attributed to the activity of the type 3 RyR (12), a RyR isoform present in mammalian skeletal muscle principally during development (13). While rare observations of Ca2+ sparks were made in resting intact mammalian muscle fibers (11, 14), until recently, significant numbers of events were only observed in skeletal fibers whose sarcolemmal integrity was disrupted by various physical or skinning methods (15, 16). Here we present one such method where saponin detergent is used to permeabilize the membrane of muscle fibers isolated from the extensor digitorum longus (EDL) muscle of rats. Because Ca2+ spark signaling rarely appears in intact mammalian muscle fibers, it was questioned if Ca2+ sparks appear in mammalian skeletal muscle at all, and if so, what sort of physiological role they would play in skeletal muscle function. In 2003, we began a series of experiments that would reveal mammalian skeletal muscle display Ca2+ sparks at the periphery of the muscle fiber, directly under the sarcolemma. During these studies, we discovered that transient hypoosmotic stress (and nonphysiological levels of hyperosmotic stress) induced peripheral Ca2+ sparks (PCS) adjacent to the sarcolemmal membrane in intact muscle fibers (17–19). Our studies suggested a physiological role for PCS in skeletal muscle by linking enhanced spark activity to exercise (17, 20). Further studies showed that this Ca2+ spark response was altered in different states, including muscular dystrophy (17) and aging muscle (21, 22). Additionally, we found osmotic stressinduced Ca2+ signaling is altered in a mouse model of amyotrophic lateral sclerosis (23), another disease state with compromised muscle function. Several other groups have gone on to examine the phenomena of PCS in mammalian skeletal muscle using variations on this osmotic stress approach or simply through examination of such fibers without stimulation. Multiple investigators have used this technique to confirm and expand our findings of elevated PCS events in dystrophic muscle from mdx mice (24–26) and to link this altered response in dystrophic muscle to increased levels of oxidative stress (25). Additional studies show that PCS are modified by the redox state of the cell, pointing to the physiological relevance of PCS (27). Further evidence of physiological regulation of PCS comes from a series of studies that show dihydropyridine receptor (DHPR) modulates the characteristics of PCS (28). Other investigators show that removal of inhibition by DHPR is

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essential for induction of PCS (29). These findings correspond nicely with other studies illustrating an essential role for DHPR in the repression of spontaneous Ca2+ sparks in differentiating skeletal myotubes (30) and in general regulation of RyR function in many cell types (31). Here we present our protocols to measure Ca2+ sparks in permeabilized EDL muscle fibers and intact flexor digitorum brevis (FDB) muscle fibers treated with hypoosmotic stress. These specific methods detail how to isolate muscle fibers from each of these muscle types (Subheadings 3.1 and 3.2) and how to induce Ca2+ sparks in these fibers (Subheadings 3.3 and 3.4). Additionally, we provide a brief tutorial on recording and analyzing Ca2+ data while providing additional resources for further details on this extensive topic (Subheading 3.5).

2. Materials Water reaching “ultrapure” requirements (18 M7 per cm, TOC < 10 ppb) was used for all solutions prepared for these studies. 2.1. FDB Muscle Fiber Isolation

1. To prepare the dissection chamber, add 8 mL liquid Sylgard (DOW CORNING, Sylgard® 184 Silicone Elastomer Kit) into an 100-mm cell culture dish and wait 48 h to let the Sylgard become solid. Store at room temperature. 2. Isotonic Tyrode Solution: 140 mM NaCl, 5 mM KCl, 10 mM HEPES (free acid), 5.5 mM D-glucose, 2.5 mM CaCl2, 2 mM MgCl2. Adjust pH to 7.2 with NaOH. Filter sterilize. Osmolality of this solution should be measured at 290 ± 5 mOsm. Store at 4°C for up to 2 months. Solution should be warmed to room temperature before use. 3. Minimal Ca2+ Tyrode Solution: 140 mM NaCl, 5 mM KCl, 10 mM HEPES (free acid), 5.5 D-glucose, 2 mM MgCl2. Adjust pH to 7.2 with NaOH. Filter sterilize. Osmolatity of this solution should be measured at 280 ± 5 mOsm. Store at 4°C for up to 2 months. Solution should be warmed to room temperature before use. 4. Digestion Solution I: Minimal Ca2+ Tyrode Solution supplemented with 2 mg/mL collagenase type I (#4196 from Worthington). Solution should be prepared in advance and 0.75 mL aliquots should be stored in 1.5 mL snap-cap tubes at −20°C for up to a month. Solution should be warmed to 37°C before use. 5. Dissecting Tools: Heavy Mayo dissecting scissors, curved iris dissection scissors, Dumont 46 blunt forceps, Noyes spring scissors, dissecting pins.

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2.2. EDL Muscle Fiber Isolation

1. Modified Krebs Solution: 136 mM NaCl, 5 mM KCl, 10 mM HEPES, 10 mM glucose, 2.6 mM CaCl2, 1 mM MgCl2. Adjust pH to 7.0 with NaOH. Osmolatity of this solution should be measured at 310 ± 5 mOsm. Filter sterilize. Store at 4°C for up to 2 months. Solution should be warmed to room temperature before use. 2. Relaxing Solution: 150 mM K-glutamate, 10 mM HEPES, 2 mM MgCl2, 1 mM EGTA. Adjust pH to 7.0 with NaOH. Filter sterilize. Store at 4°C for up to 2 months. Solution should be warmed to room temperature before use. 3. Digestion Solution II (in mM): 136 mM NaCl, 5 mM KCl, 10 mM HEPES, 10 mM glucose, 10% fetal bovine serum (FBS), 2 mg/mL collagenase type I (#4196 from Worthington). Adjust pH to 7.0 with NaOH. Solution should be made fresh daily. 4. Dissection tools: Moria spring scissors, Moria ultra fine-tipped forceps, and dissecting pins.

2.3. Ca2+ Spark Imaging of Intact FDB Fibers

1. 35 mm Delta TPG dishes. 2. Fluorescent Ca2+ imaging dyes (fluo-4AM, cell permeant, or fluo-3AM, cell permeant) are prepared as 1 mM stocks in DMSO and individual tubes are prepared with 10 ML of stock per tube. Individual tubes are stored desiccated in the dark at −20°C for up to 3 months. 3. Hypotonic Tyrode Solution: 70 mM NaCl, 5 mM KCl, 10 mM HEPES (free acid), 2.5 mM CaCl2, 2 mM MgCl2. Adjust pH to 7.2 with NaOH. Filter sterilize. Osmolatity of this solution should be measured at 170 ± 2 mOsm. Store at 4°C for up to 2 months. Solution should be warmed to room temperature before use. 4. Conventional laser scanning confocal microscope with t40× objective lens with a minimum of a 1.2-NA. The confocal microscope must be configured for an excitation at wavelength of 488 nm and an emission range at 510–580 nm for recording the fluorescence signal from fluo-3 or fluo-4. 5. Perfusion system with at least two independent perfusion channels capable of perfusing >1 mL/min of solution through a single perfusion tip of >0.2 mm in diameter. 6. Three-axis micromanipulator capable of accommodating the perfusion system.

2.4. Ca2+ Spark Imaging of Permeabilized EDL Fibers

1. Custom made 100 ML Lucite chamber with a glass bottom (16, 32, 33). Commercially available glass bottom dishes can also be used. Some possibilities include Delta TPG dishes, MatTek glass bottom dishes, or confocal imaging chambers.

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2. Fluorescent Ca2+ imaging dyes (fluo-4, pentapotassium salt, or fluo-3, pentapotassium salt) are prepared as 10 mM stocks in millipure water and individual tubes are prepared with 10 ML of stock per tube. Individual tubes are stored desiccated in the dark at −20°C for up to 3 months. 3. Saponin Solution: 150 mM K-Glutamate, 10 mM HEPES, 2 mM MgCl2, 1 mM EGTA. Adjust pH to 7.0 with NaOH. Add 0.002% saponin, 4% dextran, and 50 MM fluo-4 (or fluo3) pentapotassium salt (see Note 1). Store at 4°C for up to 2 weeks. Solution should be warmed to room temperature before use. 4. Internal Solution: 97 mM K2SO4, 10 mM trizma maleate, 10 mM Na2PC (phosphocreatine), 5 mM Na2ATP, 5 mM glucose, 1 mM EGTA, 0.32 mM CaCl2 (200 nM free Ca2+), 10.7 mM MgCl2 (2 mM free Mg2+), 8% dextran, pH 7.0. Make 10 mL stock solution and individual tubes are prepared with 1 mL of stock per tube. Store at −20°C for up to 1 year. Before the experiment, add 100 MM fluo-4 (or fluo-3) into 1 mL internal stock solution (see Notes 2 and 3). 5. Conventional laser scanning confocal microscope with t40× objective lens with a minimum of a 1.2-NA. The confocal microscope must be configured for an excitation at wavelength of 488 nm and an emission range at 510–580 nm for recording the fluorescence signal from fluo-3 or fluo-4.

3. Methods All manipulations of the experimental preparations were conducted at room temperature unless otherwise noted in the procedure. Adherence to the recommended temperatures for digestion of anatomical muscles with collagenase and for storage of muscle fibers is important for the success of the protocol as a whole. 3.1. FDB Muscle Fiber Isolation

1. Euthanize the mouse by CO2 inhalation followed by cervical dislocation. Remove the foot from the carcass using a pair of heavy dissecting scissors to cut through the leg slightly above the ankle joint. 2. Fill a Sylgard dissection chamber with a sufficient quantity of Minimal Ca2+ Tyrode Solution to fully submerge the mouse foot. The foot is then pinned, bottom side facing up, to the dissection chamber using dissection pins. Remove the FDB from the foot using a dissection scope if necessary (see Note 4). 3. Use forceps to transfer the FDB muscle into a thawed aliquot of Digestion Solution I warmed to 37°C. Always handle muscles by the tendons to avoid additional damage.

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4. Incubate the tube containing the FDB upright at 37°C with orbital shaking at 160 rpm for 60–90 min (see Note 5). 5. Wash digested FDB muscle by using blunt forceps to transfer the muscle through three dishes containing at least 1 mL of Minimal Ca2+ Tyrode Solution. 6. Transfer digested FDB muscle into a 1.5 mL snap-cap tube containing 700 ML of Isotonic Tyrode Solution using blunt forceps. 7. Cut the tip of a standard 20 ML plastic micropipette tip with a razor blade so that the diameter of the tip is just large enough to allow the muscle to be drawn through the tip. 8. Slowly draw the tendon end of the digested FDB muscle through the pipette tip 5–6 times until it passes through with no resistance. It should be possible to observe portions of the muscle coming loose from the muscle bundle. Allow these released pieces to settle at the bottom of the tube and do not disturb with subsequent pipetting (until step 12 if necessary). This process is known to some as “tituration.” 9. Cut another plastic micropipette tip to a new diameter that matches the current width of the FDB muscle. 10. Repeat steps 8 and 9 until the muscle is completely broken into smaller pieces. Let sit for 5 min at room temperature (see Note 6). 11. Flick the bottom of the tube with a finger to suspend the pieces of the muscle in the Tyrode Solution. Draw out 35 ML with a cut 200 ML plastic micropipette tip and add to the center of a 35-mm Delta TPG dish containing 1 mL of Isotonic Tyrode Solution. The tip should be placed against the glass bottom and the muscle fibers added slowly so that they do not diffuse throughout the dish. 12. Examine the muscle fibers using a dissection microscope. This will allow determination if the muscle fibers have undergone sufficient tituration and if these can be used for experimentation (see Note 7). Additionally, the number of useful fibers can be determined for preparation of additional dishes (see Note 8). 13. Additional aliquots of the FDB muscle fibers can be added until there are at least three useful fibers on the dish. The presence of three fibers on the dishes makes it likely that at least one will survive loading with Ca2+ indicator dye. 14. FDB fibers can be added to more dishes using the volume added to the initial dish as a guide to plate at least three useful fibers. Usually three dishes are prepared at a time for Ca2+ imaging. The tube of muscle fibers can be stored at 4°C for plating fibers into additional dishes later in the day.

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1. Euthanize the rat (Sprague–Dawley) with CO2 inhalation. Separate the EDL muscle at the tendon and dissect it out from the leg (see Note 9). Pin down the muscle to the Sylgard dissection chamber filled with Digestion Solution II at room temperature. The muscle should be stretched to the length that it was in the body of the animal. 2. Place the Sylgard dissection chamber in an orbital shaker (10 rpm) at 37°C for 60 min. The EDL muscle will appear only partially digested with ragged edges but no loss of portions of the muscle into the Digestion Solution. 3. Wash out the Digestion Solution from the chamber by changing to Krebs Solution 3 times. 4. Let the digested muscle rest in Krebs Solution plus 10% FBS for at least 2 h at 4°C. 5. We used a custom-built glass bottom chamber for these experiments. If a custom chamber is not available, commercial glass bottom dishes can also be used. To prepare the glass bottom for imaging, apply ~1 ML of grease (DOW CORNING high vacuum grease) at both ends of the glass bottom ~7 mm apart. Place a small piece of double-side tape (1.5 × 3 mm) on the top of each spot of grease and then add a similar amount of the grease on the top of the tape. Then fill the glass bottom chamber/dish with Relaxing Solution. 6. Replace the Krebs Solution in the Sylgard dissection chamber with Relaxing Solution. Use Moria spring scissors to make a small cut at one end of the EDL muscle. From the location of the small cut, use Moria ultra fine-tipped forceps to separate a small muscle bundle (containing 3–5 muscle fibers) about 1 cm in length from the whole EDL muscle (do not cut it off from the EDL). Then, use two Moria ultra fine-tipped forceps to peel one muscle fiber from this small bundle. The digestion should be sufficient to loosen the connective tissue around the muscle bundles. So it should not be difficult to separate a single muscle fiber from the bundle. Otherwise, it is necessary to optimize the digestion conditions by increasing the digestion time. Finally, cut out a single fiber segment (about 1 cm). Use a glass Pasteur pipette to transfer the fiber segment to the imaging chamber filled with the Relaxing Solution (see Note 10). 7. Position either end of the EDL fiber on one of the pieces of tape covered with grease. Then fix the fiber in place against the glass bottom by placing another small piece of double-side tape (1.5 × 3 mm) at each end of the fiber.

3.3. Ca2+ Spark Imaging of Intact FDB Fibers

1. Transfer 500 ML of Isotonic Tyrode Solution from dishes with plated FDB fibers to a tube of fluo-4 in DMSO. Mix 3 times by pipetting and add back to the dish drop-wise.

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2. Incubate for 60 min at room temperature in the dark. 3. Load two channels of the perfusion system, one with Isotonic Tyrode Solution and other with Hypotonic Tyrode Solution. 4. Wash the fibers by gently removing 500 ML of Isotonic Tyrode Solution from the dish and replacing with fresh Isotonic Tyrode Solution. Repeat this process a total of 4 times. 5. The dish can now be transferred to the confocal microscope. Select a target fiber for experimentation (see Note 8). The tip of the perfusion system should be positioned directly adjacent to the target muscle fiber using the micromanipulator controls. 6. Begin the flow of Isotonic Tyrode Solution from the perfusion system onto the muscle fiber to test if the FDB fiber remains in place. 7. Start acquisition of fluo-4 fluorescent signal using the confocal microscope. Individual manufacturers will have significantly different software to control the microscope. Thus, this should be done according to manufacturer’s directions. General guidelines on the collection of data can be found in Subheading 3.5. 8. After collecting at least 60 s of baseline recordings, switch the perfusion solution to Hypotonic Tyrode Solution for 60 s. The muscle fiber should swell in volume in the presence of the Hypotonic Tyrode Solution. 9. After 60 s of exposure to Hypotonic Tyrode Solution, switch the perfusion solution back to the Isotonic Tyrode Solution. As the muscle fiber shrinks back to the original volume, there will be a robust Ca2+ spark response directly under the sarcolemma of the muscle fiber (Fig. 1a). Other osmotic stress methods can be used to induce Ca2+ sparks as well (see Note 11). 10. Record and analyze Ca2+ spark data as described in Subheading 3.5. 3.4. Ca2+ Spark Imaging of Permeabilized EDL Fibers

1. Mount the imaging chamber on the stage of a confocal microscope. 2. Replace the Relaxing Solution with the Saponin Solution, and monitor the appearance of the fluo-4 fluorescence inside the fiber in the confocal microscope system. Immediately after the appearance of fluo-4 inside the fiber (usually about 2–3 min after the addition of the Saponin Solution), wash the muscle fiber 3 times with the Relaxing Solution plus 4% dextran. 3. Replace the Relaxing Solution with the Internal Solution (see Note 12). 4. Record Ca2+ sparks using a confocal scanning microscope. Recording conditions and analysis of these recordings are discussed in Subheading 3.5.

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Fig. 1. (a) A xy field scan image of an intact FDB muscle fibers loaded with Fluo-4-AM. When perfused with isotonic Tyrode solution (left ), the fiber only displays minimal Ca2+ sparks. Perfusion of hypotonic Tyrode solution causes the fiber to swell (center) and a return to perfusion of isotonic Tyrode solution induces robust Ca2+ sparks in the periphery of the muscle fiber (right ). (b) A xt line scan image along the dotted line in (a) shows the varying kinetics that occur in individual Ca2+ sparks following osmotic stress.

3.5. Analysis of Ca2+ Sparks Imaging from Isolated Muscle Fibers

Ca2+ sparks can be recorded under xy confocal scan mode to evaluate the distribution and frequency of Ca2+ sparks (15, 30). Figure 2a shows a representative two-dimensional (xy) field scan image obtained from a permeabilized EDL fiber with a conventional confocal scanning microscope. Note that Ca2+ sparks inside the muscle fiber. For more precise analysis of the kinetics and morphology, individual Ca2+ sparks images can be collected using a confocal line scan (xt) mode. Figure 2b shows a representative xt line scan image obtained by repeatedly scanning the permeabilized EDL fiber (shown in Fig. 2a) along the dashed line for 512 times at a time interval of 1.25 ms. Traces a, b, and c are quantitative indication of the changes of fluorescence intensity (F/F0) along the area of Ca2+ sparks. By collecting a large number of such traces, the morphology of individual Ca2+ sparks can be evaluated, i.e., the amplitude, rise time, duration, time to the peak, full width of half magnitude

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Fig. 2. (a) A xy field scan image of a permeabilized EDL muscle fiber loaded with the internal solution containing 100 MM fluo-4, 200 nM free Ca2+, and 2 mM free Mg2+. (b) A xt line scan image of the same muscle fiber obtained through repeatedly scanning the fiber along the dashed line. Traces a, b, and c are the quantitative measurement of Ca2+ sparks by evaluating the changes of fluo-4 fluorescence intensity (F/F0) along the area occupied by Ca2+ sparks.

(FWHM), and full duration of half magnitude (FDHM) of the recorded sparks (16, 32, 33). Cheng et al. (1999) developed an automated algorithm to detect and measure sparks in line scan images without human intervention (34). The algorithm was coded in the image-processing language IDL (Research System, Boulder, CO). This method was further refined by González et al. (2000) for analyzing sparks recorded from amphibian skeletal muscle (35). Further refinement led to an optimized computer routine to characterize Ca2+ sparks in permeabilized rat EDL muscle fibers (16). In the case of Ca2+ sparks induced in FDB fibers by hypoosmotic stress, the unique kinetics of Ca2+ release (19, 20) (Fig. 1b) are better assessed using another semi-automated IDL routine, sparkfit, which provides flexibility to select certain events not recognized by automatic detection programs (17).

4. Notes 1. Before beginning this step, prepare 1% saponin stock solution (1 mL) and Relaxing Solution plus 4% dextran (referred to as relaxing + dextran, 20 mL) and store them at 4°C. Before an experiment, add 2 ML 1% saponin and 50 MM fluo-4 to 1 mL of this relaxing + dextran stock. Because it can be difficult to

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suspend saponin evenly inside a solution, vigorously vortex the saponin stock solution before use and also vigorously vortex the final saponin solution before adding it to the chamber. 2. Ca2+ sparks in permeabilized mammalian skeletal muscle fibers were first recorded by Kirsch et al. (2001) using an internal solution containing glutamate as the major anion (15). We found that muscle fibers immersed in such internal solution with glutamate produced events transiently at a very low frequency, and some fibers did not show sparks at all. Substitution of glutamate with SO42− as the major anion caused an increase in spark frequency for any given fibers (16). 3. The concentration of free Ca2+ and Mg2+ in an internal solution is critical for inducing Ca2+ sparks in permeabilized muscle fibers. We found that 200–300 nM free Ca2+ and 2 mM free Mg2+ produced maximal frequency of Ca2+ sparks in mammalian muscle fibers (32). The concentrations of free Ca2+ and Mg2+ were calculated using public domain program WinMaxC 2.10 (http:// www.stanford.edu/~cpatton/maxc.html). Kd values of all Ca2+ and Mg2+ buffers (SO42−, ATP, PC, EGTA) were from Martel and Smith (36) and extrapolated within WinMaxC. 4. While the dissection of the mouse foot can be done with the naked eye, some find it easier to use a dissection scope during the removal of the FDB. Effective dissection can best be done by removing the skin with a pair of curved dissection scissors. The FDB is the most superficial muscle of the foot and appears in the center on the foot once the skin is removed. It is attached to the medial tubercle of the calcaneum by a thick tendon that then runs through the mass of the muscle and branches into four thinner tendon strands that eventually insert into the middle phalanx of the four lateral toes. Once the thicker portion of the tendon near the ankle joint has been identified, it can be grasped with a pair of blunt forceps just above the union between the tendon and the mass of the FDB muscle. The tendon can then be cut near the ankle using curved dissection scissors while maintaining grip on the tendon using the blunt forceps. Once the tendon is cut, the forceps can be used to pull at the muscle and begin to peel it off of the layer of tendon on which is rests. The FDB is held in place by connective tissue fascia that will provide resistance as you begin to pull the muscle free from the foot. This connective tissue can be removed using a pair of spring scissors to make short, sweeping cuts along the edges of the FDB with the points of the scissors always running out and away from the FBD proper. Continue this approach until you reach the point when the FDB connects with the tendon layer that runs beneath the FDB. The FDB can then be removed from the animal using a single cross-cut of the spring scissors as far down as possible towards the point where the FDB joins this lower tendon.

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5. Determination of the appropriate time of digestion for each lot of collagenase used is a critical step for the success of FDB fiber isolation. In general, 60 min is sufficient. The best indication that the FDB has been properly digested is that the edges of the muscle itself begin to appear ragged. If the muscle begins to fall apart in the tube of collagenase or upon transfer into Tyrode Solution before titration, then the digestion has gone far too long. 6. The FDB fibers used for experiments are fragments of longer muscle fibers that have been broken apart by tituration of the fiber. The ends of these fibers must reseal to allow the fiber to remain intact. This membrane resealing takes a few minutes. So allow the fibers to sit before plating in Isotonic Tyrode Solution. 7. If there has not been sufficient tituration of the muscle, many of the muscle fibers will appear in clumps when examined under the microscope (Fig. 3a). To remedy this situation, a 10-ML plastic micropipette tip can be used to conduct a series of 4–5 additional gentle tituration strokes that pull through the FDB fragments in the bottom of the snap-cap tube. If this does not reduce the number of muscle fibers clumps, then it is likely the FDB was not digested with collagenase for a sufficient amount of time. 8. It is important to determine how many useful fibers are on a dish before beginning to load Ca2+ indicator dye into the fibers. To be useful for the measurement of Ca2+ sparks, a fiber must be firmly attached to the bottom of the dish and display intact morphology. Intact morphology can be assessed by the following characteristics: (1) a straight, rod-like appearance (2) a length between 70 and 100 Mm, (3) a width between 8 and 18 Mm, (4) a clear, uniform striation pattern, and (5) a smooth sarcolemmal membrane with no patches of “wrinkled” membrane. The size of the fiber may be altered in aged mice, or in transgenic and disease models such as dystrophic animals. Some curvature of the fiber may be acceptable as long as the membrane is not wrinkled at the site where the fiber bends. See Fig. 3, for example, of isolated FDB fibers with proper morphology and common types of defects. 9. The origin tendon of the EDL is found at the lateral condyle of the tibia, while the insertion tendon passes under the extensor retinaculum ligament. Generally, it is easier to identify the insertion tendon, grasp the tendon with a pair of blunt forceps, and then cut the tendon while holding the forceps to keep tension of the EDL. Then you can run a separate pair of forceps under the length of the muscle to remove it from the weak layer of fascia surrounding the muscle. The origin tendon can be removed from the body using a pair of spring scissors.

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Fig. 3. (a) Clumped muscle fibers indicate that the FDB muscle is not sufficiently digested or requires additional disruption with a pipette. (b) Hypercontracted muscle fibers result from excessive pipetting of muscle fibers. (c) Damaged FDB fibers (arrow ) are generally wider than health fibers (center ) and lack the smooth membrane surface of healthy fibers. (d) Curled ends of FDB fibers (arrow ) should exclude such fibers from use in Ca2+ spark experiments. (e) Wrinkled sarcolemmal membranes (arrow ) also should exclude FDB fibers from use in experiments. (f) Wide ends of FDB fibers (arrow ) indicate a fiber that has not properly resealed at the end and should be excluded from experimentation. (g) Excessively long or bent FDB fibers so are not optimal for use in Ca2+ spark experiments. (h) Slight bending or twisting of an FDB fiber usually does not exclude a fiber from use in experiments. (i) An example of an ideal FDB fiber for experimentation. The long, rod-like structure, smooth membrane, and average size are all characteristics that indicate a fiber useful for experimentation.

10. After dissecting out a single fiber from the EDL muscle, the Relaxing Solution in the dissection chamber should be immediately replaced with Krebs Solution plus 10% FBS, and the muscle should be returned to a refrigerator (4°C) immediately. The digested EDL muscle kept in Krebs Solution plus 10% FBS at 4°C can be used repeatedly in 24 h.

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11. Other investigators have used a number of different solutions of varying composition to induce Ca2+ sparks. Frequently, these are hyperosmotic solutions that can induce Ca2+ sparks immediately following perfusion of the solution and shrinking of the fiber. We have previously used a high Ca2+ solution to induce hyperosmotic conditions and trigger Ca2+ sparks (17). Others have perfused solutions using sucrose to elevate osmolality and observed similar effects (29). Hyperosmotic approaches using buffers with an osmolality >420 mOsm are effective and valid, but in some cases can result in more damage to the fiber than the hypoosmotic approach detailed here. Some of the solutions used, particularly the high Ca2+ solution, also go beyond the physiological range that may be experienced by cells in vivo. These effects can elevate the intracellular Ca2+ level of the muscle fiber and complicate imaging and analysis of Ca2+ spark signaling. 12. The concentration of free Ca2+ and Mg2+ in an internal solution is critical for inducing Ca2+ sparks in permeabilized muscle fibers. We found that 200–300 nM free Ca2+ and 2 mM free Mg2+ usually gave the maximal frequency of Ca2+ sparks in mammalian muscle fibers (32).

Acknowledgment This work was supported by NIH grants to Drs. Weisleder (AR54793), Zhou (AR57404) and Ma (AG28614, HL69000 and AG28856), and MDA funding to Dr. Zhou (MDA4351). The authors thank Ms. Andoria Tjondrokoesoemo for helpful comments during the preparation of the final manuscript. References 1. Cheng H, Lederer WJ, Cannell MB (1993) Calcium sparks: elementary events underlying excitation-contraction coupling in heart muscle. Science 262:740–744 2. Wang SQ, Stern MD, Rios E, Cheng H (2004) The quantal nature of Ca2+ sparks and in situ operation of the ryanodine receptor array in cardiac cells. Proc Natl Acad Sci USA 101: 3979–3984 3. Wier WG, ter Keurs HE, Marban E, Gao WD, Balke CW (1997) Ca2+ ‘sparks’ and waves in intact ventricular muscle resolved by confocal imaging. Circ Res 81:462–469 4. Kamishima T, Quayle JM (2003) Ca2+ -induced Ca2+ release in cardiac and smooth muscle cells. Biochem Soc Trans 31:943–946

5. Nelson MT, Cheng H, Rubart M, Santana LF, Bonev AD, Knot HJ, Lederer WJ (1995) Relaxation of arterial smooth muscle by calcium sparks. Science 270:633–637 6. Zhuge R, Fogarty KE, Baker SP, McCarron JG, Tuft RA, Lifshitz LM, Walsh JV, Jr (2004) Ca(2+) spark sites in smooth muscle cells are numerous and differ in number of ryanodine receptors, large-conductance K(+) channels, and coupling ratio between them. Am J Physiol Cell Physiol 287:C1577–1588 7. Zhuge R, Fogarty KE, Tuft RA, Lifshitz LM, Sayar K, Walsh JV, Jr (2000) Dynamics of signaling between Ca(2+) sparks and Ca(2+)activated K(+) channels studied with a novel image-based method for direct intracellular

23 measurement of ryanodine receptor Ca(2+) current. J Gen Physiol 116:845–864 8. Rossi AE, Dirksen RT (2006) Sarcoplasmic reticulum: the dynamic calcium governor of muscle. Muscle Nerve 33:715–731 9. Klein MG, Cheng H, Santana LF, Jiang YH, Lederer WJ, Schneider MF (1996) Two mechanisms of quantized calcium release in skeletal muscle. Nature 379:455–458 10. Tsugorka A, Rios E, Blatter LA (1995) Imaging elementary events of calcium release in skeletal muscle cells. Science 269:1723–1726 11. Shirokova N, Garcia J, Rios E (1998) Local calcium release in mammalian skeletal muscle. J Physiol 512:377–384 12. Ward CW, Schneider MF, Castillo D, Protasi F, Wang Y, Chen SR, Allen PD (2000) Expression of ryanodine receptor RyR3 produces Ca2+ sparks in dyspedic myotubes. J Physiol 525:91–103 13. Sutko JL, Airey JA, Murakami K, Takeda M, Beck C, Deerinck T, Ellisman MH (1991) Foot protein isoforms are expressed at different times during embryonic chick skeletal muscle development. J Cell Biol 113:793–803 14. Conklin MW, Barone V, Sorrentino V, Coronado R (1999) Contribution of ryanodine receptor type 3 to Ca(2+) sparks in embryonic mouse skeletal muscle. Biophys J 77:1394–1403 15. Kirsch WG, Uttenweiler D, Fink RH (2001) Spark- and ember-like elementary Ca2+ release events in skinned fibres of adult mammalian skeletal muscle. J Physiol 537:379–389 16. Zhou J, Brum G, Gonzalez A, Launikonis BS, Stern MD, Rios E (2003) Ca2+ sparks and embers of mammalian muscle. Properties of the sources. J Gen Physiol 122:95–114 17. Wang X, Weisleder N, Collet C, Zhou J, Chu Y, Hirata Y, Zhao X, Pan Z, Brotto M, Cheng H, Ma J (2005) Uncontrolled calcium sparks act as a dystrophic signal for mammalian skeletal muscle. Nat Cell Biol 7:525–530 18. Ward CW, Lederer WJ (2005) Ghost sparks. Nat Cell Biol 7:457–459 19. Weisleder N, Ferrante C, Hirata Y, Collet C, Chu Y, Cheng H, Takeshima H, Ma J (2007) Systemic ablation of RyR3 alters Ca2+ spark signaling in adult skeletal muscle. Cell Calcium 42:548–555 20. Weisleder N, Ma J (2006) Ca2+ sparks as a plastic signal for skeletal muscle health, aging, and dystrophy. Acta Pharmacol Sin 27:791–798 21. Weisleder N, Brotto M, Komazaki S, Pan Z, Zhao X, Nosek T, Parness J, Takeshima H, Ma J (2006) Muscle aging is associated with compromised Ca2+ spark signaling and segregated

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intracellular Ca2+ release. J Cell Biol 174: 639–645 22. Weisleder N, Ma J (2008) Altered Ca2+ sparks in aging skeletal and cardiac muscle. Ageing Res Rev 7:177–188 23. Zhou J, Yi J, Fu R, Liu E, Siddique T, Rios E, Deng HX (2010) Hyperactive intracellular calcium signaling associated with localized mitochondrial defects in skeletal muscle of an animal model of amyotrophic lateral sclerosis. J Biol Chem 285:705–712 24. Lovering RM, Michaelson L, Ward CW (2009) Malformed mdx myofibers have normal cytoskeletal architecture yet altered EC coupling and stress-induced Ca2+ signaling. Am J Physiol Cell Physiol 297:C571–580 25. Shkryl VM, Martins AS, Ullrich ND, Nowycky MC, Niggli E, Shirokova N (2009) Reciprocal amplification of ROS and Ca(2+) signals in stressed mdx dystrophic skeletal muscle fibers. Pflugers Arch 458:915–928 26. Teichmann MD, Wegner FV, Fink RH, Chamberlain JS, Launikonis BS, Martinac B, Friedrich O (2008) Inhibitory control over Ca(2+) sparks via mechanosensitive channels is disrupted in dystrophin deficient muscle but restored by mini-dystrophin expression. PLoS One 3:e3644 27. Martins AS, Shkryl VM, Nowyck, MC, Shirokova N (2008) Reactive oxygen species contribute to Ca2+ signals produced by osmotic stress in mouse skeletal muscle fibres. J Physiol 586:197–210 28. Apostol S, Ursu D, Lehmann-Horn F, Melzer W (2009) Local calcium signals induced by hyperosmotic stress in mammalian skeletal muscle cells. J Muscle Res Cell Motil 30:97–109 29. Pickering JD, White E, Duke AM, Steele DS (2009) DHPR activation underlies SR Ca2+ release induced by osmotic stress in isolated rat skeletal muscle fibers. J Gen Physiol 133:511–524 30. Zhou J, Yi J, Royer L, Launikonis BS, Gonzalez A, Garcia J, Rios E (2006) A probable role of dihydropyridine receptors in repression of Ca2+ sparks demonstrated in cultured mammalian muscle. Am J Physiol Cell Physiol 290:C539–553 31. Dirksen RT (2002) Bi-directional coupling between dihydropyridine receptors and ryanodine receptors. Front Biosci 7:d659–670 32. Zhou J, Launikonis BS, Rios E, Brum G (2004) Regulation of Ca2+ sparks by Ca2+ and Mg2+ in mammalian and amphibian muscle. An RyR isoform-specific role in excitation-contraction coupling? J Gen Physiol 124:409–428 33. Zhou J, Brum G, Gonzalez A, Launikonis BS, Stern MD, Rios E (2005) Concerted vs. sequential. Two activation patterns of vast

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arrays of intracellular Ca2+ channels in muscle. J Gen Physiol 126:301–309 34. Cheng H, Song LS, Shirokova N, Gonzalez A, Lakatta EG, Rios E, Stern MD (1999) Amplitude distribution of calcium sparks in confocal images: theory and studies with an automatic detection method. Biophys J 76:606–617

35. Gonzalez A, Kirsch WG, Shirokova N, Pizarro G, Stern MD, Rios E (2000) The spark and its ember: separately gated local components of Ca(2+) release in skeletal muscle. J Gen Physiol 115:139–158 36. Martell AE, Smith RM (1974) Critical Stability Constants. Plenum Press, New York.

Chapter 24 Analysis of Calcium Transients in Cardiac Myocytes and Assessment of the Sarcoplasmic Reticulum Ca2+-ATPase Contribution Anand Mohan Prasad and Giuseppe Inesi Abstract Ca2+ signaling plays an essential role in several functions of cardiac myocytes. Transient rises and reductions of cytosolic Ca2+, permitted by the sarcoplasmic reticulum Ca2+ ATPase (SERCA2) and other proteins, control each cycle of contraction and relaxation. Here we provide a practical method for isolation of neonatal rat cardiac myocytes and measurement of Ca2+ transients in cultured cardiac myocytes, yielding information on kinetic resolution of the transients, variations of cytosolic Ca2+ concentrations, and adequacy of intracellular Ca2+ stores. We also provide examples of experimental perturbations that can be used to assess the contribution of SERCA2 to Ca2+ signaling. Key words: Cardiac myocytes, Cytosolic Ca2+ transients, Sarcoplasmic reticulum Ca2+ ATPase, Excitation-contraction coupling

1. Introduction Controlled oscillations of cytosolic Ca2+ constitute signals for numerous cellular functions. Ca2+ signaling serves as a common mechanism to couple membrane excitation to intracellular functions in most biological tissues (1, 2). This mechanism is based on: (a) a high gradient between extracellular (mM) and cytosolic (<0.1 MM) Ca2+ concentrations, permitting fluxes of signaling Ca2+ through selective channels from extracellular fluids and intracellular stores into the cytosol, and (b) the presence of intracellular proteins, such as calmodulin and troponin, that activate specific functions upon Ca2+ binding.

Joseph X. DiMario (ed.), Myogenesis: Methods and Protocols, Methods in Molecular Biology, vol. 798, DOI 10.1007/978-1-61779-343-1_24, © Springer Science+Business Media, LLC 2012

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Fig. 1. Diagram of Ca2+ signaling in excitation-contraction coupling of heart muscle.

In cardiac muscle, variations of cytosolic Ca2+ are involved in several signaling functions including activation of transcription and contraction (3). It is shown in Fig. 1 that each contractile cycle of cardiac myocytes begins with electrical depolarization of the plasma membrane, which allows influx of a small quantity of extracellular Ca2+ through a voltage-dependent channel. This Ca2+ triggers release of a much larger quantity of internal Ca2+ from the sarcoplasmic reticulum (SR) stores (“Ca2+ induced Ca2+ release”) through the ryanodine receptor channel. The subsequent rise of cytosolic Ca2+ allows Ca2+ binding to troponin and contractile activation of actomyosin. Relaxation is then produced following membrane repolarization and removal of cytosolic Ca2+ by the ATP-dependent Ca2+ pump associated with SR membranes. The SR internal stores are thereby refilled with Ca2+ to be released for the next cycle. Cytosolic Ca2+ is also removed (to a lesser extent) by the Na+/Ca2+ exchanger (NCX) and the plasma membrane Ca2+ ATPase (PMCA). The cardiac SR Ca2+ ATPase was first isolated (4) with a microsomal fraction of cardiac muscle, in association with membrane vesicles exhibiting Ca2+ transport coupled to utilization of ATP, and producing relaxation of contractile models by reducing the Ca2+ concentration in the medium to values lower than 0.1 MM. The cardiac SR ATPase derives from one of three known SERCA genes, yielding the prevalent SERCA2a isoform (5), but also the

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Fig. 2. Cardiac myocytes isolated for neonatal rats (left ) or adult rabbits (right ). The preparation from neonatal rats and adult rabbits was obtained as described by Prasad and Inesi and Dani et al. (8, 9), respectively. Note the homogeneously healthy population in large microscopic fields, which we find least troublesome to obtain from neonatal rats or adult rabbits. Left panel figure adapted from Prasad and Inesi, (8) Am J Physiol Cell Physiol with permission from Am Physiol Soc. Right panel figure from a preparation obtained as described (9). Line bars correspond to 50 (a) and 100 (b) μm.

SERCA2b (6) and SERCA2c (7) isoforms. SERCA2 turned out to be most important in excitation/contraction coupling of cardiac myocytes, as SERCA2 fills intracellular stores with Ca2+ to be released for contractile activation, and in turn sequesters cytosolic Ca2+ to allow relaxation. Cytosolic Ca2+ transients can be studied conveniently by microscopy of cultured cardiac myocytes loaded with fluorescent indicators, using a high-speed fluorescence imaging system. Fluorescence emission from single cells is measured following double wavelength excitation, whereby dye calibration yields estimates of cytosolic Ca2+ concentration. Although these measurements can be obtained from selected single myocyte, it is very important to obtain preparations yielding homogeneous populations of healthy cells in culture (Fig. 2), since collection of samples for parallel molecular or biochemical assays (see below) requires a large number of cells that must be comparable to those used for Ca2+ transients. In our experience, least troublesome preparations can be obtained from neonatal rats (Fig. 2 left panel) or adult rabbits (Fig. 2 right panel). The cultured myocytes can then be subjected to electrical field stimulation or exposed to pharmacological agents to elicit Ca2+ transients and gain information on the kinetics and extent of cytosolic Ca2+ rise and decay.

2. Materials 2.1. Primary Cell Cultures (Neonatal Rat Cardiac Myocytes)

1. Gelatin (Sigma G-2500; 300 bloom): Make a 1% gelatin solution by dissolving 5 g gelatin powder in 500 mL of ddH2O. Dissolve in microwave and then autoclave. Aliquot into sterile 50 mL tubes. Store in a refrigerator.

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2. Mouse laminin: Use 20 Mg/mL of PBS (phosphate buffer saline) buffer to coat the culture dishes or glass coverslips. 3. DMEM (with high glucose, L-glutamine, 110 mg/L sodium pyruvate). 4. M199 (with Hanks’ salts, L-glutamine, 25 mM HEPES buffer). 5. Horse Serum (HS) (heat-inactivated, mycoplasma-tested, EIAtested, virus-tested). 6. Fetal Bovine Serum (FBS) (heat-inactivated, performancetested, mycoplasma-tested, virus-tested). 7. Penicillin Streptomycin (Pen-Strep). Add 5 mL of Pen-Strep to each 500 mL bottle of medium. 8. Fungizone. Add 5 mL of Fungizone to each 500 mL bottle of medium. 9. ITS solution: Defined medium containing insulin, transferrin, and selenium substitute for serum. Usually available as 100× solutions. 10. 5-bromo-2-deoxyuridine (BrdU). 11. Plating Medium: DMEM/M199, 5% FBS, 10% HS. Make 500 mL of plating medium by combining 340 mL of DMEM (containing Pen-Strep and fungizone), 85 mL of M199 (containing Pen-Strep and fungizone), 25 mL of FBS, 50 mL of HS, 100 MM 5-bromo-2-deoxyuridine (BrDU), and filter sterilize with a 0.22 Mm filter. Store in a refrigerator. 12. Maintenance Medium: (80%DMEM/20% M 199). Combine 400 mL of DMEM (containing Pen-Strep and fungizone), 100 mL of M199 (containing Pen-Strep and fungizone), 5 mL of ITS, 100 MM BrDU. Filter through a 0.22 Mm filter and store in a refrigerator. 13. Bovine serum albumin (BSA), 0.1%. 14. Vitamin C, 0.1 mM. 15. Vitamin B12, 2 Mg/mL. 16. Ads Buffer: 116 mM NaCl, 20 mM HEPES, 1 mM NaH2PO4 (monobase), 5.5 mM Glucose, 5.4 mM KCl, 0.8 mM MgSO4·7H2O, 0.001 g Phenol red, pH 7.35 ± 0.005. 17. Prepare 10× Ads buffer without phenol red and adjust the pH to 7.35 ± 0.005 with 1 N NaOH. Filter sterilize the buffer and store in a refrigerator. While making 1× Ads buffer, add phenol red and then again adjust the pH to 7.35 ± 0.005 with 1 N NaOH. Filter sterilize and store in the refrigerator. 18. Collagenase: (Type 2) Worthington # CLS 2. (see Note 1). 19. Pancreatin. 20. Enzyme medium (Digestion mix). Add 25 mg collagenase (0.357 mg/mL) and 20 mg pancreatin (0.286 mg/mL) to 70 mL

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of 1× Ads buffer and warm up at 37°C waterbath. Make sure that the enzymes are dissolved completely. Filter sterilize the digestion mix. 21. Sterile nylon net filters (Millipore, 25 mm, 100 U pore size; Millipore Swinnex filter holders, 25 mm). 22. Mat Tek culture dishes (35 mm). 23. Glass coverslips. 2.2. Calcium Transients

1. Ringer’s Potassium Bicarbonate Buffer Solution: 10.0 mM HEPES, 135.0 mM NaCl, 4.0 mM KCl, 1.0 mM MgSO4, 1.0 mM Na2HPO4, 1.8 mM CaCl2, 0.5 mM EGTA, 25.0 mM NaHCO3, 10.0 mM glucose, pH 7.35 with 1 N NaOH or 1 N HCl. 2. To prepare Ringer’s buffer, make 10× buffer solution (pH 7.35) without adding CaCl2, EGTA, NaHCO3, and glucose. Separately prepare 10× CaCl2, 10× EGTA, 10× NaHCO3, and 10× glucose. These solutions can be kept at 4°C for weeks. On the day of experiment, prepare 1× Ringer’s buffer by adding NaHCO3, glucose, CaCl2 (for calcium buffer), or EGTA (for calcium free buffer). Warm up the buffer to room temperature. Readjust the pH to 7.35 with 1 N NaOH or 1 N HCl. 3. 10 mM caffeine. 4. 1 MM thapsigargin (TG). 5. 135.0 mM N-methyl-D-glucamine. 6. Fura-2AM, cell permeant: Prepare 1 mM solution in DMSO and keep frozen for storage. 7. Fluo-4AM, cell permeant: Prepare 1 mM solution in DMSO and keep frozen for storage. 8. Fluo-3AM, cell permeant: Prepare 50 Mg/100 ML of DMSO and dilute 10 ML to 1 mL. 9. Anhydrous DMSO. 10. Pluronic-F127, a nonionic, surfactant polyol that facilitates the solubilization of water-insoluble dyes: To prepare, weigh 57 mg Pluronic-127 and add 228 ML sterile anhydrous DMSO (under hood). Incubate at 37°C for 1 h to ensure dissolution. 11. On day of use, prepare 1 mL of dye solution containing 11.5 ML of Pluronic F-127 (final conc, 0.2%) and 1 ML of Fura-2AM (final conc, 1.0 MM), or 4.4 ML of 1 mM Fluo-3AM (final conc, 4.4 MM) or 1 ML of 1 mM Fluo-4AM (final conc, 1.0 MM). Keep solution in the dark. Add 12.5 ML of dye mix (e.g., Fura-2AM + Pluronic F 127) to 987.5 ML of 1× Ringer’s buffer solution to make 1 mL of dye solution. 12. Ion Wizard high-speed fluorescence imaging system with a MYO100 Myocam (Ion Optix Corporation; Milton MA).

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Fluorescence System Interface that allows communication between IonWizard and most of the peripheral devices, including PMT, MyoPacer, HyperSwitch, MyoPacer field stimulator for electrical field stimulation, a circulating bath for stage temperature control (Cell MicroControls mTCII Temperature Controller & Heater), and IonWizard-Core and Analysis software for recording.

3. Methods 3.1. Isolation of Neonatal Rat Cardiac Myocytes (Primary Cell Cultures (8))

1. Precoat plates with 1% gelatin for overnight at 37°C prior to beginning the isolation of heart cells. 2. Bathe neonatal rats in 70% ethanol, open the chest, and remove hearts. Transfer hearts to 15 mL cold 1× Ads buffer. Wash hearts with additional 15 mL 1× Ads. Transfer the hearts to a dish of 1× Ads buffer. After all the hearts are collected, trim the hearts of excess atria, fat, and connective tissue (see Note 2). Cut hearts into small pieces. 3. Aspirate off the Ads buffer and add 10 mL of enzyme medium. Transfer hearts into a sterile 15 mL tube and incubate on a rotator for 30 min at 37°C at low speed. Discard the supernatant from the first digestion (see Note 3). 4. Add 10 mL of fresh enzyme medium to digestion tube and incubate at 37°C for 15 min. 5. Gently triturate hearts using a 10-mL pipet and incubate 5 min. 6. Collect supernatant into a 15-mL tube with 1 mL HS. 7. Spin down the cells at 500 g for 5 min. Discard supernatant and resuspend pellet in 2 mL HS. Loosen cap and incubate in a CO2 incubator until all digestions are completed. 8. To digestion tube, add fresh enzyme medium and repeat steps 4–7 until all the tissue is digested completely (about six total digestions for a total of about 2 h). 9. Filter all the digestions through sterile Nylon Net Filters into a 50-mL tube (see Note 4). Centrifuge at 500 g for 5 min. Discard supernatant and resuspend in 20 mL serum-containing plating medium. Plate onto an uncoated 150 mm culture plate and incubate in CO2 incubator for 1–2 h (see Note 5). 10. Determine the cell number. Plate the cells onto gelatin-coated dishes or laminin-coated glass coverslips or Mat Tek dishes. Incubate cells in plating medium at 37°C with 5% CO2. 11. Cell seeding: 0.5–1×106 cells/P-35 dish/2 mL volume, 2.5–3.5 × 106 cells/P-60 dish/5 mL volume, 5.0–7.0 × 106 cells/P-100 dish/10 mL volume, and 350,000 cells per glass cover slips or Mat Tek dish.

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12. Twenty four hours after plating, change the medium to maintenance medium with 0.1% BSA, 0.1 mM vitamin C, and 2 Mg/mL vitamin B12 (but no FBS) (“serum-free medium”). Maintain the myocytes at 37°C under 5% CO2. 3.2. Calcium Transients Measurements

Cardiac myocytes from neonatal rat (Fig. 2; left panel) or adult rabbit (9) hearts (Fig. 2; right panel) are grown on culture dishes with laminin-coated glass coverslips or Mat Tek culture dishes. In preparation for the calcium transients measurements, the myocytes are seeded at 350,000 cells per laminin-coated glass coverslips or Mat Tek culture dishes. When ready for the measurements: 1. Remove medium. 2. Wash 2 times with room temperature Ringer’s Potassium Bicarbonate Solution. 3. Add 200 ML of calcium dye solution (e.g., Fura-2, AM). 4. Incubate for 10 min at room temperature in the dark (covered by box). 5. Wash 2 times with dye-free Ringer’s Potassium Bicarbonate Solution. 6. Following wash with dye-free Ringer’s Potassium Bicarbonate Solution, place the coverslips with Fura-2 loaded cells or Mat Tek dishes containing loaded cells in a special chamber mounted on an Olympus 1 × 70 inverted microscope, and connected to a circulating bath with Ringer’s Potassium Bicarbonate Solution maintained at 30 ± 2°C temperature. 7. Perform measurements using the Ion Wizard high-speed fluorescence imaging system. Subject the cells to field stimulation of 10 V and 20-ms duration at 1 Hz frequency. 8. Measure the fluorescence emission from single cells using 380 or 340 nm excitation. 9. Perform dye calibration and processing as described previously in detail (10). 10. Calculate cytoplasmic-free Ca2+ from background-corrected fluorescence ratios (R = F340/F380) using the equation [Ca2+] = Kd [(R–Rmin)/(Rmax–R)) × Q (11). Estimate Rmax (F340/F380) at the end of each experiment in the presence of 1 mM Ca2+ and Rmin (F340/F380) in the presence of 1 mM EGTA with no Ca2+. Q is the ratio Rmin/Rmax at 380 nm. Kd = 225 nM. 11. To measure total releasable Ca2+ from SR in single cells loaded with Fura-2 (1 MM), stimulate the cells at 1.0 Hz for at least 1 min before exchanging the external Ringer’s Potassium Bicarbonate Solution with medium containing 10 mM caffeine and 1 MM TG. For these measurements, replace Na+ by N-methyl-D-glucamine (see Note 6).

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12. Measure fluorescence emission continuously using 380 or 340 nm excitation. Obtain computer-assisted analysis of the transients and that would include kinetic parameters for their rise and decay, peak height, and duration (see Note 7). 3.3. Contribution of SERCA2 to Ca2+ Transients in Cardiac Myocytes

The prominent role sustained by SERCA2 in excitation/contraction coupling is reflected both in physiological and pathological features of cardiac muscle. In fact, abnormalities of intracellular Ca2+ handling play a crucial role in the pathogenesis of heart failure (12). In this regard, studies of Ca2+ transients in cardiac myocytes may be very helpful to establish and clarify the involvement of SR calcium ATPase in alterations of cardiac signaling and excitation-contraction coupling. Specific inhibitors and various agonists can be used to assess the contribution of SERCA2 to Ca2+ transients. In addition, the availability of cDNA clones has rendered possible gene transfer into cultured cells, and studies of the functional consequences of SERCA overexpression or downregulation in situ. Severe alterations of Ca2+ signaling and contractile function have been demonstrated following specific inhibition of SERCA2 transport activity with TG (8, 13, 14), reduction of expression by a SERCA2 gene null mutation (15), and SERCA2 gene silencing with short interference RNA (16). The effects of SERCA inhibition with TG, and SERCA downregulation by adrenergic hypertrophy, are clearly shown in Fig. 3a. On the other hand, increased levels of Ca2+ transport ATPase in transgenic mouse heart (17–19), or heterologous SERCA expression in myocytes (10, 20), yield clear enhancement of Ca2+ signaling heights and kinetics in cardiac myocytes (Fig. 3b).

Fig. 3. Cytosolic Ca2+ transients in myocytes subjected to electrical field stimulation. As indicated in the figure (a), the myocytes were exposed for 3 days to 20 MM phenylephrine to produce adrenergic hypertrophy, or to 10 nM thapsigargin (TG) for specific SERCA inhibitions (8). Note the increase in resting cytosolic Ca2+, the reduction in Ca2+ release, and the slower Ca2+ signal decay induced by phenylephrine and TG. The effect of exogenous SERCA1 expression (10) on Ca2+ signaling kinetics is shown in (b). Figure (a) adapted from Prasad and Inesi (8), Am J Physiol Cell Physiol with permission from Am Physiol Soc. Figure (b) adapted from Cavagna et al. (10), Journal of Physiology, with permission from John Wiley and Sons.

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Fig. 4. SERCA protein levels and Ca2+ transport activity in total myocyte homogenates. The myocytes were exposed for 3 days to 20 MM phenylephrine or to 10 nM TG as described for Fig. 3. Note that reduction of transport activity is reduced as a consequence of SERCA downregulation (i.e., reduction of SERCA protein level) in the phenylephrine-treated myocytes, but to direct SERCA inhibition (i.e., no SERCA protein level reduction) in the myocytes treated with TG (8). Figure is adapted from Prasad and Inesi (8), Am J Physiol Cell Physiol with permission from American Physiological Society.

It is most important to consider that the involvement of SERCA in Ca2+ signaling alterations can be best evaluated by means of parallel measurements of SERCA2 protein levels by Western blotting as well as ATP-dependent Ca2+ transport (Fig. 4), which can be easily performed with homogenates of the cultured myocytes used for measurements of Ca2+ transients, and subjected to the same experimental manipulations.

4. Notes 1. For collagenase, pretest the lot before using. Not all lots work. 2. After collecting all the hearts, trim the hearts of excess atria, fat, and connective tissue. This will greatly reduce the number of fibroblasts. 3. First digestion of heart cells with enzyme medium should be discarded. The first digestion allows the removal of broken cells and blood cell. 4. Before filtering all the digestions through sterile Nylon Net Filters into a 50-mL tube, remember to prewet the filtration unit.

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5. After resuspension of cells in 20 mL serum-containing plating medium, plate into an uncoated 150 mm culture plate and incubate in CO2 incubator for 1–2 h. This is a preplating step which may help reduce the number of cardiac fibroblasts. Before plating, count the cells, remove all the gelatin from the culture dishes, and then plate the cells. 6. While measuring total releasable Ca2+ from SR in single cells loaded with Fura-2 (1 MM), replace Na+ by N-methyl-Dglucamine to prevent Ca2+ exchange with sodium calcium exchanger. 7. Using Ion Wizard high-speed fluorescence imaging system with its dedicated software, it is also possible to measure sarcomere shortening, thereby assessing patterns of contractile activation (21).

Acknowledgment This and related work were supported by National Institutes of Health Grant NHBLI RO301-69830. References 1. Carafoli E (2002) Calcium signaling: a tale for all seasons. Proc Natl Acad Sci 99:1115–1122 2. Clapham DE (2007) Calcium signaling. Cell 131:1047–1058 3. Bers DM (2008) Calcium cycling and signaling in cardiac myocytes. Ann Rev Physiol 70:23-49 4. Inesi G, Ebashi S, Watanabe S (1964) Preparation of vesicular relaxing factor from bovine heart tissue. Am J Physiol 207: 1339–1344 5. Zarain-Herzberg A, MacLennan DH, Periasamy M (1990) Characterization of rabbit cardiac sarco(endo)plasmic reticulum Ca2+-ATPase gene. J Biol Chem 265:4670–4677 6. Lytton J, Westlin M, Burk SE, Shull GE, MacLennan DH (1992) Functional comparisons between isoforms of the sarcoplasmic or endoplasmic reticulum family of calcium pumps. J Biol Chem 267:14483–14489 7. Dally S, Bredoux R, Corvazier E, Anderson JP, Clausen JD,Dode L, Fanchaouy M, Gelebart P, Monceau V, Del Monte F, Gwathmey JK, Hajjar R, Chaabane C, Bobe R, Raies A, Enouf J (2006) Ca2+-ATPases in non-failing and failing heart: evidence for a novel cardiac sarco/endoplasmic reticulum Ca2+-ATPase 2 isoform (SERCA2). Biochem J 395:249–258

8. Prasad AM, Inesi G (2009) Effects of thapsigargin and phenylephrine on calcineurin and protein kinase C signaling functions in cardiac myocytes. Am J Physiol Cell Physiol 296:C992–C1002 9. Dani AM, Cittadini A, Inesi G (1979) Calcium transport and contractile activity in dissociated mammalian heart cells. Am J Physiol 237:C147–C155 10. Cavagna M, O’Donnell JM, Sumbilla C, Inesi G, Klein MG (2000) Exogenous Ca2+-ATPase isoform effects on Ca2+ transients of embryonic chicken and neonatal rat cardiac myocytes. J Physiol 528:53–63 11. Grynkiewicz G, Poenie M, Tsien RY (1985) A new generation of Ca2+ indicators with greatly improved fluorescence properties. J Biol Chem 260:3440–3450 12. Flesch M, Schwinger RH, Schnabel P, Schiffer F, van Gelder I, Bavendiek U, Südkamp M, Kuhn-Regnier F, Böhm M (1996) Sarcoplasmic reticulum Ca2+ATPase and phospholamban mRNA and protein levels in end-stage heart failure due to ischemic or dilated cardiomyopathy. J Mol Med 74:321–332 13. Kirby MS, Sagara Y, Gaa S, Inesi G, Lederer WJ, Rogers TB (1992) Thapsigargin inhibits

24 contraction and Ca2+ transient in cardiac cells by specific inhibition of the sarcoplasmic reticulum Ca2+ pump. J Biol Chem 267:12545–12551 14. Prasad AM, Ma H, Sumbilla C, Lee DI, Klein MG, Inesi G (2007) Phenylephrine hypertrophy, Ca2+-ATPase (SERCA2), and Ca2+ – signaling in neonatal rat cardiac myocytes. Am J Physiol Cell Physiol 292:C2269–C2275 15. Periasamy M, Reed TD, Liu LH, Ji Y, Loukianov E, Paul RJ, Nieman ML, Riddle T, Duffy JJ, Doetschman T, Lorenz JN, Shull GE (1999) Impaired cardiac performance in heterozygous mice with a null mutation in the sarco(endo) plasmic reticulum Ca2+-ATPase isoform 2 (SERCA2) gene. J Biol Chem 274:2556–2562 16. Seth M, Sumbilla C, Mullen SP, Lewis D, Klein MG, Hussain A, Soboloff J, Gill DL, Inesi G (2004) Sarco(endo)plasmic reticulum Ca2+ATPase (SERCA) gene silencing and remodeling of the Ca2+ signaling mechanism in cardiac myocytes. Proc Natl Acad Sci USA 101:16683–16688 17. He H, Giordano FJ, Hilal-Dandan R, Choi DJ, Rockman HA, McDonough PM, Bluhm WF, Meyer M, Sayen MR, Swanson E, Dillman WH (1997) Overexpression of the rat sarcoplasmic reticulum Ca2+ATPase gene in the heart of

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transgenic mice accelerates calcium transients and cardiac relaxation. J Clin Invest 100: 380–389 18. Baker DL, Hashimoto K, Grupp IL, Ji Y, Reed T, Loukianov E, Grupp G, Bhagwhat A, Hoit B, Walsh R, Marban E, Periasamy M (1998) Targetedoverexpression of the sarcoplasmic reticulum Ca2+ATPase increases cardiac contractility in transgenic mouse hearts. Circ Res 83: 1205–1214 19. Loukianov E, Ji Y, Grupp IL, Kirkpatrick DL, Baker DL, Loukianova T, Grupp G, Lytton J, Walsh RA, Periasamy M (1998) Enhanced myocardial contractility and increased Ca2+ transport function in transgenic hearts expressing the fast-twitch skeletal muscle sarcoplasmic reticulum Ca2+-ATPase Circ Res 83:889–897 20. Hajjar RJ, Kang JX, Gwathmey JK, Rosenzweig A (1997) Physiological effects of adenoviral gene transfer of sarcoplasmic reticulum calcium ATPase in isolated rat myocytes. Circulation 95:423–429 21. Lundblad A, Gonzalez-Serratos H, Inesi G, Swanson J, Paolini P (1986) Patterns of sarcomere activation, temperature dependence, and effect of ryanodine in chemically skinned cardiac fibers. J Gen Physiol 87:885–905

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Part VII Analysis of Gene Promoter Transcriptional Activity

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Chapter 25 Analysis of Muscle Gene Transcription in Cultured Skeletal Muscle Cells Charis L. Himeda, Phillip W.L. Tai, and Stephen D. Hauschka Abstract The mechanisms by which muscle gene expression is initiated and maintained are not fully understood. Muscle genes are regulated by combinatorial interactions between numerous transcription factors bound to enhancers and promoters, and their associated protein complexes. Among the most important are the MyoD and MEF2 transcription factor families, but dozens of other factors play important regulatory roles, and many additional transcription factors are certain to be involved. Expression of muscle-specific genes varies among different anatomical muscles and in fast- vs. slow-twitch fiber types, suggesting different mechanisms of regulation in response to diverse physiological cues. Thus, identifying novel transcriptional regulators and interactions is key to understanding how different cells establish the muscle phenotype; it is also critical for developing methods to combat diseases such as muscular dystrophy. Using Muscle creatine kinase as a model, we outline the key steps involved in identifying muscle gene control elements, their binding factors, and mechanisms of transcriptional activation and repression. The basic principles described here can also be applied to the transcriptional analysis of other cell-type specific genes. Key words: Skeletal muscle, Transcriptional regulation, Control elements, Transcription factors, Muscle creatine kinase, Chromatin immunoprecipitation, Quantitative proteomics

1. Introduction The commitment of cells to the myogenic lineage and their subsequent differentiation into mature myocytes are determined by a complex network of signaling molecules and their downstream effectors. Extracellular factors such as insulin, insulin-like growth factors I and II (IGF-I and -II), sonic hedgehog (Shh), transforming growth factor-β1 (TGF-β1), basic fibroblast growth factor (bFGF), and Wnts (1–5) trigger intracellular signaling cascades that culminate in the activation of metabolic enzymes and structural proteins necessary for myogenesis and the repression of myogenic

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inhibitors. Committed myoblasts are marked by expression of Myf5 and Pax3/7, transcription factors that are required for expression of MyoD, the founding member of the Myogenic Regulatory Factor (MRF) family of bHLH transcription factors (6). MyoD, in concert with the MADS-box transcription factor MEF2 and a host of other regulators (e.g., Six4/5, SRF, MAZ, KLF3), activates a cascade of muscle differentiation genes to give rise to mature muscle fibers (7–10). While the critical myogenic roles of MRFs and MEF2 have been well established, the exact mechanisms by which muscle gene expression is initiated and maintained are still poorly understood. Gene expression is determined by combinatorial interactions between transcription factors bound to enhancers and promoters, and their associated protein complexes. Expression of muscle-specific genes varies among different anatomical muscles and in fast- vs. slow-twitch fiber types, suggesting different mechanisms of regulation in response to different physiological cues. Thus, identifying novel transcriptional regulators and interactions is key to understanding how different cells establish the muscle phenotype; it is also critical for developing methods to combat diseases such as muscular dystrophy. Many muscle-specific genes are controlled by multiple enhancers which direct expression in different cell lineages and in response to different physiological signals. For example, expression of MyoD is controlled by two separate enhancers. The proximal enhancer directs expression in differentiated skeletal muscle during embryogenesis and maintains expression in adult skeletal muscle (11). In contrast, the distal enhancer is required for correct expression in the hypaxial myotome and limb buds, but is not known to play roles in adult skeletal muscle (12). Myf5 is regulated by four separate enhancers which direct expression in discrete skeletal muscle precursor cell populations in the embryo (13–16). Structural muscle genes such as Muscle creatine kinase (MCK) also have multiple enhancers that control expression in fast vs. slow fiber types (17). With respect to experimental strategies for studying muscle gene regulation, it is important to emphasize that both cell culture and transgenic studies were required to delineate and test the functions of the multiple enhancers in the Myf5 and MCK genes (13–16). Enhancers and promoters contain binding sites for diverse transcription factors, and our understanding of these sequence motifs is constantly evolving. Although MyoD and Myogenin recognize simple E-boxes (CANNTG), they have preferences for certain internal and flanking bp that occur frequently in muscle genes (18, 19). In contrast to the relatively similar sequences of E-boxes, transcription factors such as TEF1 have been shown to recognize motifs that are surprisingly divergent (20, 21), and our work has expanded the spectrum of known binding motifs for both Six4 and MAZ (8, 9). Additionally, we and others have found that multiple

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factors are capable of regulating transcription via the same or similar binding motifs. It is likely that this redundancy, as well as competition for binding sites, allows for both robustness and fine-tuning of gene expression levels in response to different upstream signals. Although often difficult and labor-intensive, identifying control elements and their cognate transcription factors is only the first step toward understanding mechanisms of transcriptional regulation. Transcriptional control is a complex, dynamic process in which multiple interactions between protein complexes – some of them separated by distances of up to 1 Mb (22) – are vital to establishing correct gene expression patterns. Transcription factor interactions play a key role in stabilizing factor binding to DNA, increasing or decreasing accessibility of binding sites, and helping to recruit members of the basal Pol II machinery to the promoter (23). Transcription factors regulating muscle-specific genes exhibit a range of expression profiles, from skeletal muscle-specific (the MyoD family) to ubiquitous (Sp1, MAZ, AP2). With a number of widely expressed regulators, muscle cells employ several mechanisms to ensure that these factors are active in the right time and place. MyoD, for example, heterodimerizes with E proteins to activate its target genes in differentiating muscle; however, it is also expressed in proliferating myoblasts prior to the onset of differentiation. Several regulatory mechanisms exist in myoblasts to keep MyoD from activating differentiation genes prematurely: (1) negative regulators sequester either MyoD or its E protein partners, (2) HDAC1 associates with MyoD, and (3) hyperphosphorylation of MyoD by cell cycle effectors targets it for rapid degradation (24–27). Likewise, Type II HDACs associate with MEF2 in replicating myoblasts and are shuttled out of the nucleus at the onset of differentiation (28). By contrast, Serum Response Factor (SRF) activates distinct sets of target genes in both proliferating and differentiating muscle. In replicating myoblasts, phosphorylation of SRF by PKCα facilitates its association with Ets factors, allowing it to activate immediate early genes; when myoblasts withdraw from the cell cycle, phosphorylation of SRF by PKCα declines, allowing it to activate muscle genes and help drive the program of muscle differentiation (29). Interactions of transcription factors with DNA are stabilized through both homotypic and heterotypic interactions. For example, binding of SRF to the skeletal α-actin promoter is stabilized through the interactions of SRF homodimers binding to multiple CArG boxes (30). Likewise, MyoD binds with higher affinity to paired E-boxes or E-boxes adjacent to another transcription factor binding site (Pbx/Meis, MEF2, Sp1) (10). Many transcription factors have been shown to synergize with each other in transactivating muscle genes, and in some cases, synergy requires only a single control element. For example, SRF has been shown to recruit Nkx2.5 to CArG sites (31), and we have

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recently shown that KLF3 recruits SRF to KLF-binding motifs (7). Mechanisms of synergy include recruitment of coactivators such as p300/CBP and PCAF, which function as histone acetyltransferases (HATs), directly modifying chromatin structure to increase accessibility of binding sites. Through interactions with Pbx/Meis, MyoD may serve as such a pioneer factor, recruiting chromatin remodelers to initiate target gene expression (32). DNA-bound factors can also recruit corepressors, such as CtBP2 and histone deacetylases (HDACs). In proliferating myoblasts, for example, DNA-bound MEF2 recruits Class II HDACs to repress the expression of muscle differentiation genes (33). In addition to cofactor recruitment, many transcription factors (MyoD, SRF, Sp1) are also capable of making direct contacts with the basal Pol II machinery (34–36). We have used MCK as a paradigm for understanding musclespecific gene transcription since it is specifically and abundantly expressed in striated muscle and is one of the major hallmarks of muscle differentiation. MCK is also expressed at different levels in different anatomical skeletal muscles and in skeletal vs. cardiac muscle (37, 38). This allows for the identification of control elements and binding factors important for expression in different muscle types, as well as those specific to each myogenic lineage. While results in cultured cells need to be confirmed in vivo, skeletal muscle cell lines such as MM14 and C2C12 continue to serve as useful model systems for identifying regulatory regions and their composite control elements. Additionally, nonmuscle cell lines such as 10T1/2 and COS cells are convenient systems in which to test novel synergistic interactions without the confounding presence of multiple myogenic proteins or high levels of the factors being tested. The following protocol outlines the basic steps involved in analyzing mechanisms of muscle gene transcription, using MCK as a model. In two expanded sections, we focus on the analysis of protein-DNA interactions by chromatin immunoprecipitation (ChIP), and selective enrichment of transcription factors for identification by quantitative proteomics.

2. Materials 2.1. Identifying Regulatory Regions and Control Elements in Muscle Genes

1. Standard cloning equipment and reagents. 2. Muscle cell culture equipment and reagents. 3. Equipment and reagents for reporter gene expression assays (e.g., enzyme assays, quantitative RT-PCR, and immunofluorescent microscopy).

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2.2. Identifying Transcription Factors Binding Muscle Control Elements

Standard molecular biology equipment and reagents are required. Access to a mass spectrometry facility is required for proteomic identification of transcription factor candidates.

2.3. Identifying Transcription Factor Interactions

Standard molecular biology equipment and reagents are required.

3. Methods 3.1. Identifying Regulatory Regions and Control Elements in Muscle Genes

1. Define candidate regulatory regions. Multi-species sequence alignments (e.g., using ClustalW) (39) can provide strong clues to the locations of regulatory regions. However, since exact or close matches to transcription factor binding motifs can occur solely due to random sequence probabilities (e.g., the core MyoD E-box sequence CANNTG occurs approximately once every 256 bp), we typically focus on conserved 50–400 bp regions containing clusters of at least three highly conserved sequence motifs (see Note 1). 2. Test candidate regulatory regions for activity. Depending on the size of the candidate sequence, it should be inserted into a plasmid or BAC upstream of a convenient reporter cDNA (e.g., chloramphenicol acetyltransferase (CAT), placental alkaline phosphatase (PAP), or luciferase). Putative regulatory regions are then examined by deletion analysis using convenient restriction sites. Muscle cell lines (see Note 2) can be transiently transfected using a standard Ca phosphate method (40) or lipofection-based methods (e.g., Invitrogen), and reporter activity of the deletion construct compared to that of the wild-type (see Note 3). After delineating the boundaries of a regulatory region, it is useful to test activity following ligation to either the native gene promoter or a heterologous promoter such as thymidine kinase (41) (see Note 4). 3. Define candidate control elements. Highly conserved sequence motifs within a defined regulatory region are likely to correspond to functional elements (see Note 5), and candidate sequences can be searched against transcription factor binding site databases such as TRANSFAC (42). Database searches often return long lists of possible binding factors, and a literature search of specific binding sites and expression profiles can help determine the most likely candidates. If a conserved sequence does not match any motif in the database, it might represent a binding site for a novel transcription factor, or (more likely) a divergent motif for a known factor (see Note 6).

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4. Test candidate control elements for activity. Putative cis elements are usually tested by mutating or deleting in the context of their native enhancer/promoter as described above (see Note 7). 3.2. Identifying Transcription Factors Binding Muscle Control Elements

1. Test control elements for transcription factor binding. Electrophoretic mobility shift assays (EMSA), also known as gel-shift assays, are a quick and economical way to verify that functional elements are recognized by specific nuclear factors. Gel-shift assays should be performed using labeled probes that contain the target control element and appropriate competitors (see Note 8). Nuclear extracts from myoblasts, differentiated myocytes, and nonmuscle cells can be used to determine developmental and cell-type specificity of binding factors (see Note 9). 2. Determine identity of candidate transcription factor(s). If the target control element matches the binding motif for one or several transcription factors for which specific antibodies exist, gel supershift or ChIP assays (Subheading 3.4) can be performed to verify the candidate(s). 3. Determine identity of unknown transcription factor(s). In some cases, a target control element will not match any known binding motif or will match binding motifs for many transcription factors. cDNA cloning and yeast 1-hybrid screening (43, 44) have traditionally been used to identify unknown transcription factors binding isolated control elements. A more recent strategy is selective enrichment of candidates followed by either mass spectrometry or quantitative proteomics (7–9, 45, 46) (Subheading 3.5). While these screening methods are often difficult and labor-intensive, they nonetheless have the potential to provide a relatively unbiased identification of novel DNA-binding factors. 4. Verify the in vivo relevance of candidate transcription factors (see Note 10).

3.3. Identifying Transcription Factor Interactions

1. Determine cofactor interactions. Once a transcription factor has been established as a muscle gene regulator, the proteins it associates with can provide important clues to its mechanism of action. Cofactor candidates can be identified using yeast 2-hybrid (43) or proteomic methods (47), and candidate interactions can be validated via coimmunoprecipitation or GST pull-down studies. Determine regions/sequences required for interaction by testing truncated/mutated forms of each factor in any of the above assays. To test whether two factors can associate on a single binding site, gel supershift assays can also be performed (see Note 11).

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2. Determine synergistic/repressive interactions. Interacting factors can be tested for their ability to cooperate in activating or repressing expression of a reporter construct in cell culture (see Note 12). Determine regions/sequences required for synergy/ repression by testing truncated/mutated forms of each factor in the reporter assay. Chimeric proteins (containing the DNAbinding domain of one factor and the activation/repression domain of another) can also be used to test the ability of a cofactor to directly activate/repress transcription factor activity (48). 3.4. Focus on Chromatin Immunoprecipitation

ChIP analysis is increasingly becoming a requirement in gene transcription studies. While techniques such as EMSA and plasmid reporter assays rely on the analysis of exogenous, nonchromatinized DNA, ChIP allows the analysis of the native chromatin architecture of endogenous genes. This includes dynamic modifications to histones and DNA, as well as occupancy by transcription factors and accessory proteins. Briefly, chromatin from fixed nuclei contains DNA cross-linked to protein complexes, which can be sheared into fragments (~200–1,000 bp) and precipitated using specific antibodies and Protein A-coupled beads. Immunoprecipitated chromatin fragments can then be assayed for the presence of enriched target genomic sequences by PCR using flanking primers. With the proper controls, an investigator can demonstrate the presence of a specific factor or modification at any genomic locus in myoblast or myocyte cultures, or cultures subjected to different physiological manipulations. Choosing the cell source: Because ChIP requires a large amount of nuclear material from a uniform cell source, many ChIP studies are performed using established cell lines (e.g., MM14 or C2C12 skeletal muscle cell lines, or fibroblasts induced to become muscle) (7, 9, 49, 50). However, one must keep in mind that these cell lines display incompletely overlapping expression profiles of muscle genes, a phenomenon that is reflected by different factor occupancy at enhancers and promoters. For example, in an inducible skeletal muscle model (a fibroblast line infected with retrovirus expressing MyoD), the MCK enhancer is occupied by MyoD during early differentiation and by Myogenin during late differentiation (49). Although this result was replicated in mouse embryos, ChIP studies in MM14 and C2C12 cultures show that both MyoD and Myogenin are present at the MCK enhancer well after terminal differentiation (>60 h post-switch) (17). In a recent study analyzing genome-wide binding of MyoD, the choice of cell source greatly affected the number of bound loci (MyoD occupied >50% more genomic regions in primary myotubes than in C2C12 cells or in fibroblasts induced to become muscle) (51). These discrepancies serve to emphasize that a “best” cell culture system for ChIP analysis

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will always be a point of conjecture, since multiple skeletal muscle culture models, including those mentioned above, have proven to be useful paradigms for understanding myogenesis. For ChIP assays of differentiated MM14 myocytes, we recommend using the Fast ChIP protocol as described (52) and as modified for skeletal myocytes (7, 17). Finding the right controls: Reproducibility of ChIP analysis is dependent on several factors that are difficult to control. Among these are: (1) extent of chromatin fixation, (2) quality of chromatin fragmentation, and (3) quality of the specific lot of antibody/antisera. Unfortunately, many antibodies that work well for Western analysis or immunostaining do not work well for ChIP. In practice, ChIP data can vary significantly between experiments, while still maintaining consistent trends of enrichment in test vs. control conditions. In cases where interexperimental variability is high, ChIP data are sometimes presented as single experiments that are representative of many that were performed with reproducible trends. While ChIP data are often presented as semiquantitative PCR analysis of gel bands, quantitative PCR offers increased sensitivity and accuracy. Many studies also present ChIP data as enrichment of the target region by a specific antibody compared to input DNA (a diluted, untreated sample). However, this method does not take into account the specificity of the antibody or the quality of wash steps following immunoprecipitation. Since most antibodies have some degree of nonspecific binding, enrichment by nonspecific IgG or preimmune sera serves as a better control. Because ChIP typically detects binding to genomic regions located within ~500 bp 5¢ or 3¢ of the target binding site(s) (due to the production of partially overlapping genomic fragments during sonication), it is virtually impossible to determine which motifs are bound by a particular target factor when the motifs are closely spaced (such as the two E-boxes in the MCK enhancer, which are separated by only 22 bp). Furthermore, since contamination by factor binding to similar sites on larger fragments is a concern, it is important to test enrichment with primers to regions outside of the target region. For example, ChIP for MyoD or Myogenin, which recognize E-boxes (CANNTG), should be controlled for with primers for flanking regions that contain either nonfunctional E-boxes or no E-box motifs (see Fig. 1). Global analysis of factor binding: ChIP analysis uses a top-down (factor-to-chromatin) approach to promoter analysis, whereas techniques such as EMSA and yeast one-hybrid screening use a bottom-up approach (DNA sequence-to-factor). Recently developed ChIP-based techniques are allowing investigators to ask increasingly global questions about transcription factor targets. In ChIP-on-chip, immunoprecipitated chromatin fragments are hybridized to promoter or genomic tiling arrays, allowing researchers to assess the occupancy of >1,000 genomic regions by a specific

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Fig. 1. ChIP for MyoD and Myogenin at the MCK locus. MCK contains three major regulatory regions: a 5¢-enhancer (Enh), proximal promoter (PP), and intronic region (MR1) containing a short intronic enhancer (SIE). There are 35 E-boxes (arrowheads) within the region shown, including five with proven transcriptional activity (two within the Enh, one within the PP, and two within the SIE). To demonstrate enrichment of MyoD and Myogenin at the SIE (17), primers were designed to amplify this region as well as a positive control region (Enh), and two negative control regions (Exon1/Intron1 boundary (Ex1/In1) and Exon2 (Ex2)). Primers to a distant genomic region containing no E-boxes were used as an additional negative control (17).

factor (53). Using this approach, MEF2 has been demonstrated to regulate a network of Drosophila genes that help specify muscle identity during development (54). The genome-wide application of ChIP has expanded even further with ChIP-seq (55), a more sensitive technique which does not depend on a tiling array of known regions. In ChIP-seq, enriched chromatin fragments are ligated to short oligonucleotide tags, allowing them to be sequenced. While ChIP-on-chip studies revealed the binding of MyoD at ~100–200 different gene promoters (56, 57), a ChIP-seq study demonstrated enrichment of MyoD at >60,000 nonrepetitive regions in the genome (~41–74% of all genes) (58). ChIP-based methods can also be used to observe in vivo enhancer-promoter looping, a mechanism which was previously predicted, but not formally shown to occur until recently (59). Techniques such as chromosome conformation capture (3C), 3C on chip (4C), and 3C carbon copy (5C) have revealed interactions between distant intra- and interchromosomal regions, and detailed protocols for these techniques have been described (60–64). Such data lend evidence to the theory of nuclear neighborhoods (65), in which genes that are transcribed at the same time in a cell may share a tethered transcriptional machine through common enhancers. What does ChIP data really mean? As the global utility of ChIP continues to expand, interpretation of the data remains a significant challenge. While it is generally assumed that transcription factor binding to chromatin indicates a functional interaction, this may not always prove to be the case. For example, a recent ChIPseq analysis demonstrated constitutive, genome-wide binding of MyoD in both myocytes and myoblasts (51). The functional consequences of this are unclear, since regions where MyoD binds in myoblasts correlate with a marker of open chromatin, yet MyoD does not mediate expression of genes within these regions (51). The ability of ChIP to detect indirect interactions of proteins with DNA as well as interactions of proteins from distant genomic

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locations poses a further complication. Using ChIP-on-chip, it was recently determined that ~33% of validated SRF gene targets contain no known SRF binding motif (CArG box) (66). There are several possible explanations for this surprising result: (1) SRF binds a noncanonical motif; (2) SRF is recruited indirectly through another factor binding to DNA (7); or (3) proteins associated with the target locus interact with SRF bound to distant CArG boxes. Similarly, in a recent ChIP-seq study, MEF2 was found to be enriched at approximately −400 to +50 in the mouse MCK promoter, a region that lacks any obvious MEF2 binding motif (17). In this case, it is likely that MEF2 is recruited by MyoD to the MCK promoter E-box (67), although interactions between MyoD associated with the promoter E-box and MEF2 bound to a distant A/T-rich site cannot be ruled out. Thus, while ChIP has proven to be a powerful tool for shedding light on protein-DNA interactions, a full understanding of gene regulation will require additional tools and knowledge gained from multiple sources. 3.5. Focus on Selective Enrichment of Transcription Factors for Identification by Quantitative Proteomics

In contrast to EMSA and ChIP analysis, quantitative proteomics has the potential to provide a relatively unbiased identification of candidate DNA-binding factors. When Isotope-Coded Affinity Tags are used, the differential incorporation of stable isotopes in two samples allows the relative abundance of proteins in the two samples to be determined. This strategy has been used to identify a number of transcriptional regulators, including several in skeletal muscle (7–9). While the advantages and caveats of this approach have been well described, less has been written regarding the steps preceding isotopic labeling. In this section, we highlight issues regarding the selective enrichment of transcription factors for quantitative proteomic identification. Enriching candidate transcription factors: One of the advantages of quantitative proteomics is that target factors do not have to be purified to homogeneity in order to be identified. Nonetheless, because the resolution capacity of mass spectrometers is limited (i.e., a target peptide can be obscured by contaminating peptides of similar mass/charge), at least partial enrichment of the target factor is recommended. The simplest and most successful enrichments have been achieved using specific DNA affinity chromatography preceded (in some cases) by partial purification (45, 68–74). Binding, washing, and elution of the target factor must be optimized for multiple conditions, including time, temperature, salt concentration, and amount of competitor DNA. Once target factor recovery is estimated (e.g., by calculating densitometry of gel-shift bands), the required amount of source material can be determined (see below). We have enriched target transcription factors using biotinylated oligonucleotides coupled to streptavidin-linked magnetic beads (7–9). Due to the large number of proteins co-purifying with the target factor(s), a selective enrichment strategy (see Fig. 2) was

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Fig. 2. Selective enrichment of transcription factors for quantitative proteomic identification. Experimental strategy used for selective enrichment of factors. Refer to Subheading 3.5 for details.

used to exclude common contaminating proteins via a normalization step. In this strategy, equal numbers of beads were coupled to oligonucleotides containing either a wild-type or a mutant target site. These were incubated with equal volumes of the same nuclear extract, washed, and the bound proteins eluted at the optimal salt concentration. This strategy resulted in two samples, one enriched for the target factor, and both containing equal amounts of nonspecific co-purifying proteins. The selectively enriched factor(s) were then identified by quantitative proteomics. An important attribute of quantitative proteomic studies is that multiple candidates are disclosed if several different factors can associate with the same control element or with sequences overlapping or immediately flanking the element. For example, in studies aimed at identifying factors bound to the MCK MPEX sequence, peptides from nine different transcription factors were identified, suggesting that multiple factors compete for occupancy of overlapping sites (9). Enriching candidate cofactors: If an unknown transcriptional cofactor or other interacting protein is the target, a slightly different strategy can be employed. In this case, an antibody specific to the known transcription factor bait can be cross-linked to protein Sepharose beads, and the beads incubated with nuclear extracts, washed, and bound proteins eluted. Although this strategy has not been reported for the identification of muscle cofactors, it has been

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successfully used in conjunction with mass spectrometry to identify novel factors associating with yeast TFIID (47). If no antibody to the transcription factor bait exists, an antibody to a protein tag can be cross-linked to the beads, and nuclear extracts from cells overexpressing a tagged form of the transcription factor can be used. However, in this case, it is important to verify that incorporation of the tag does not interfere with any known protein-protein interactions. In either case, the selectively enriched fraction would then be compared to the control fraction via quantitative proteomics. Source material: what type and how much? Proteomic identification of transcriptional regulators requires large amounts of source material, considering that: (1) identification by mass spectrometry requires at least 1 pmol of the target factor, and (2) transcription factors represent only a small fraction of total cellular proteins and their purification entails significant loss. Since intact nuclei are difficult to isolate directly from adult or even newborn muscle, cultured cells are a more amenable starting material. We have successfully identified factors binding control elements in the MCK promoter using nuclear extracts from ~1 × 109 MM14 myocytes grown on gelatin-coated plates (7, 9). However, growing large quantities of muscle cells is a time-consuming and laborious process. To avoid this in an earlier study, we were able to use a nonmuscle cell line (HeLa cells) that contained a factor with identical gel-shift mobility and sequence-specific binding properties as the target factor in muscle cells (8). In this case, it was essential to verify the relevance of the candidate factor in muscle through functional studies (see Note 10). For any cell type, once enrichment conditions have been optimized, the number of required cells can be determined based on recovery of the target factor in pilot studies. For example, in our successful identification of a transcription factor from HeLa cells (8), we estimated that there were ~2 × 108 molecules of the target factor per μg of nuclear extract. This was calculated based on the ratio of bound:free probe in gel-shift assays, assuming one molecule of factor per oligonucleotide. Estimating that each cell contains ~2.8 × 10−5 μg of nuclear protein and estimating ~40% recovery of the target factor (based on gel-shift pilot studies), we needed at least 2.5 × 108 cells to recover 1 pmol. Because equal amounts of nuclear extracts needed to be incubated with wild-type and mutant oligonucleotide-coupled beads, 5 × 108 cells were minimally required to recover 1 pmol of the target factor. Allowing for 50% loss during the quantitative proteomic steps raised the minimum amount of cells to 1 × 109. To increase our chances of success, we decided to use 5× this amount, or 5 × 109 HeLa cells. Evaluation of transcription factor candidates: Although quantitative proteomics is a powerful tool for transcription factor identification, subsequent functional studies are required to substantiate the involvement of specific candidates in muscle gene regulation (see Note 10).

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4. Notes 1. Alignments of this type using sequences of mammalian MCK genes disclose highly conserved noncoding sequences of ~200 bp (upstream enhancer), ~350 bp (proximal promoter), and ~100 bp (intronic enhancer) (38, 40). 2. Skeletal muscle cell lines such as MM14, C2C12, and L6 are well-established model systems for studying muscle gene regulation (75–77). None of these myoblast lines is identical to primary myoblasts, and they are typically aneuploid; thus, the relative abundance of transcription factors differs from that in diploid myoblasts. However, studies of primary myoblast cultures are complicated by the presence of nonmuscle cells. Although these cells should not express reporter genes driven by muscle promoters, they will express reporters driven by ubiquitous promoters such as CMV if these are used for normalization. This problem can be circumvented via the procedure in Note 3. 3. To control for variation in both transfection efficiency and extent of differentiation, it is important to transfect muscle cells with a normalization construct in addition to the test construct (78). In our experiments, we have used test constructs expressing the CAT reporter gene and normalization constructs expressing the human PAP reporter gene under the control of the wild-type MCK enhancer-promoter. We now use the more sensitive Dual-Luciferase Reporter Assay System (Promega), which utilizes Firefly luciferase as the test reporter, and Renilla luciferase as the normalization reporter. For each test construct, we typically perform at least three independent transfections of three plates each. 4. Interestingly, the MCK upstream enhancer functions well with either the 80-bp MCK basal promoter or with a heterologous promoter (41), while the MCK short intronic enhancer is only active in the context of the full MCK proximal promoter (17). 5. The sequences and relative positions of the seven known control elements in the MCK enhancer are highly conserved among mammalian species, whereas sequences between these elements are poorly conserved (79, 80). 6. We and others have found that several transcription factors regulating muscle genes recognize sequences that diverge substantially from the established binding motif. For example, the Trex site in the MCK enhancer is bound by Six4, a homeodomain protein of the Six/sine oculis family, in skeletal muscle, and Six5 in cardiac muscle (8). Six proteins recognize MEF3 motifs in the regulatory regions of their target genes; however,

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because the MCK Trex site deviates from the previously established MEF3 sequence in 2 out of 7 bp, this relationship was not identifiable by in silico screening against the TRANSFAC database. In more recent studies, we showed that MAZ and KLF3, two zinc-finger transcription factors that regulate the MCK gene and other muscle genes, also recognize a divergent spectrum of sequences (8, 9), and TEF-1 has been shown to bind both MCAT elements and A/T-rich motifs (20, 21). 7. A candidate control element is usually tested through two transverse mutations (pyrimidine to purine and vice-versa) or one mutation and one deletion. It is important to verify that the alteration does not create a new binding motif by searching the changed sequence against the TRANSFAC database. 8. Probes should be end-labeled and contain the target control element with at least 5 bp flanking each side (usually ~20– 40 bp long). Shorter probes are better for verifying specific factor binding to the relevant control element, as the presence of multiple binding sites can complicate data interpretation. Unlabeled competitors (used at 50- to 100-fold molar excess) should be the same length as the probe, and mutant competitors should contain the same altered bp that disrupted activity in a reporter assay (see Note 3). Additional probes containing a spectrum of altered sequences can be used to examine factor binding specificity (7, 9). Note that multiple specific binding complexes can correspond to different proteins, isoforms, cofactors, or degradation products. Many nonspecific complexes are also typically detected. 9. Nuclear extracts are prepared as described (81). Extracts from skeletal myocytes are typically made from 20 to 40 100-mm dishes containing ~5 × 106 cells (myonuclei) each. Anticipated yields are ~130 μg nuclear protein per 107 myonuclei. Nuclear extracts are adjusted to ~1–2 μg protein per μl. Because 1–2 μg extracts are used per EMSA lane, and because 10–15 lanes are used in typical experiments, nuclear extracts are frozen at −80°C in working aliquots of ~20–50 μL. Nuclear extract aliquots should be freeze-thawed no more than 3× and are usable for at least a year. 10. If antibodies for candidate transcription factors exist, they can be tested in gel supershift studies (EMSA using a specific antibody to either “supershift” or abolish the factor-probe complex) to confirm in vitro binding of the candidate to the target control element, and in ChIP studies to demonstrate in vivo binding to the target regulatory region. In such studies, nonimmune antisera should always be included as a negative control. Additional evidence for a candidate factor’s involvement can be obtained by

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overexpression or repression via transgenic, viral-mediated transduction, or cell culture transfection strategies, followed by assessment of effects on the candidate’s presumed function. However, if the factor is part of a family with a high degree of functional overlap among members, depletion may result in compensation by another family member (82). Furthermore, if steady-state levels of the factor are above those needed for maximal expression of a reporter gene, overexpression may not result in increased gene expression. This problem can potentially be circumvented by conducting these experiments in cells that do not express the muscle factor of interest (7). 11. Evidence of ternary complex formation (transcription factor and cofactor associated with a single control element) can be difficult to detect in gel-shift assays, which may not mimic in vivo conditions closely enough for any but the most stable interactions to be detected. 12. Synergy between muscle factors is often difficult to determine in muscle cells, which may contain high amounts of the factors being tested. Thus, other cell types that are devoid of many muscle transcription factors (fibroblasts, COS cells, SL2 cells) are often used as a convenient system for testing interactions. For example, we used COS-7 cells to demonstrate a novel synergistic interaction between KLF3 and SRF (7).

Acknowledgments Research described in this article was supported by grants from the National Institutes of Health RO1-AR18860, 1P01-NS046788, and RO1-HL64387, and by a grant from the Muscular Dystrophy Association. References 1. Pirskanen, A., Kiefer, J. C., and Hauschka, S. D. (2000). IGFs, insulin, Shh, bFGF, and TGFbeta1 interact synergistically to promote somite myogenesis in vitro. Dev Biol 224, 189–203. 2. Stern, H. M., Lin-Jones, J., and Hauschka, S. D. (1997). Synergistic interactions between bFGF and a TGF-beta family member may mediate myogenic signals from the neural tube. Development 124, 3511–23. 3. Stern, H. M., and Hauschka, S. D. (1995). Neural tube and notochord promote in vitro myogenesis in single somite explants. Dev Biol 167, 87–103.

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Chapter 26 Analysis of Fiber-Type Differences in Reporter Gene Expression of b-Gal Transgenic Muscle Phillip W. L. Tai, Catherine L. Smith, John C. Angello, and Stephen D. Hauschka Abstract β-galactosidase (β-gal) is among the most frequently used markers for studying a wide variety of biological mechanisms, e.g., gene expression, cell migration, stem cell conversion to different cell types, and gene silencing. Many of these studies require the histochemical detection of relative β-gal levels in tissue crosssections mounted onto glass slides and visualized by microscopy. This is particularly useful for the analysis of promoter activity in skeletal muscle tissue since the β-gal levels can vary dramatically between different anatomical muscles and myofiber types. The differences in promoter activity can be due to a myofiber’s developmental history, innervation, response to normal or experimental physiological signals, and its disease state. It is thus important to identify the individual fiber types within muscle cross-sections and to correlate these with transgene expression signals. Here, we provide a detailed description of how to process and analyze muscle tissues to determine the fiber-type composition and β-gal transgene expression within cryosections. Key words: β-galactosidase, Cryosectioning, Muscle fibers, Freezing artifacts, Fiber type, Myosin heavy chain, Antibody, Fast-twitch, Slow-twitch

1. Introduction The bacterial reporter gene β-galactosidase (β-gal) has been used for many years as a convenient marker for gene expression studies in vitro and in vivo (1–4), as a means of identifying marked cells following transplantation (5), for tracing cell lineages in conjunction with Cre recombinase strategies (6), and as a reporter for promoter activity (2). In the latter case, the transcriptional activity of musclespecific promoters in adult transgenic mice can be quantitatively or qualitatively gauged by the sarcoplasmic accumulation of β-gal. Researchers can thus compare the relative X-gal staining intensities Joseph X. DiMario (ed.), Myogenesis: Methods and Protocols, Methods in Molecular Biology, vol. 798, DOI 10.1007/978-1-61779-343-1_26, © Springer Science+Business Media, LLC 2012

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of experimental muscle tissues to analyze the activity of transgenic regulatory sequences during development, in the adult, and/or under specific experimental or disease conditions. However, many muscle gene promoters are not uniformly active in all anatomical muscle or in all muscle fibers. For example, the muscle creatine kinase (MCK) gene, which is active in all striated muscle types, is expressed at varying amounts in different anatomical muscles (7–9). Fast-twitch skeletal muscles (muscle groups that depend on anaerobic (glycolytic) respiration) have several-fold higher levels of MCK than slow-twitch muscles, (muscles that depend on aerobic (oxidative) respiration) and transcribe about six times more MCK mRNA than cardiac muscle. Adult mammalian skeletal muscle is comprised of a heterogeneous bundling of muscle fibers. Limb and epaxial skeletal muscles in small mammals such as mice contain four distinct fiber types: a single slow-twitch type (Type-I, oxidative), and three distinct fast-twitch types (Type-IIa, oxidative-glycolytic, Type-IId (sometimes referred to as Type-IIx), glycolytic, and Type-IIb, glycoytic) (10, 11). Type-IIa fibers are sometimes also classified as intermediatetwitch fibers. The physiological and metabolic differences between these different fiber types have been well characterized (12). It is reasonable to assume that the expression of most musclespecific genes is directly or indirectly linked to a fiber’s metabolic state. Therefore, in vivo promoter analysis should correlate transcriptional activity with the fiber types in which the promoter is active. The most robust method of classifying muscle fiber types is by immunostaining for myosin heavy chain (MYHC) isotypes (13, 14). In mammals, there are four predominant muscle MYHC isoforms (MYHC1, MYHC2A, MYHC2D/X MYHC2B), which are expressed in Type-I, Type-IIa, Type IId/x, and Type-IIb fibers, respectively. The antibodies BA-D5, SC-71, and BF-F3 (which recognize the myosin isotypes MYHC1, MYHC2A, and MYHC2B, respectively) have been used in many studies and in various mammalian models (13, 15–17). Therefore, by taking successive serial sections through muscles and analyzing them for β-gal activity and MYHC isotypes, researchers can determine whether the transcriptional activities of enhancer and/or promoter sequences correlate with fiber type identity. Here, we provide step-by-step instructions for: generating serial cryosections of mouse muscle tissue (see Subheading 2.1.1), generating and collecting BA-D5, SC-71, and BF-F3 monoclonal antibodies from hybridoma cultures ( see Subheading 3.2 ) using these in conjunction with fluorochrome-labeled secondary antibodies to identify muscle fiber types (see Subheading 3.3), and X-gal staining to assay β-gal transgene expression (see Subheading 3.4). Figure 1 illustrates the ability to distinguish the four fiber types in a mouse gastrocnemius cryosection. In this example, three

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Fig. 1. Adult mouse gastrocnemeus stained using mAbs BA-D5, SC-71, and BF-F3, which mark Type-I (blue ), Type-IIa (red ), and Type-IIb (green) fibers, respectively. Fibers that are negative for immunostaining (gray ) are considered to be Type-IId/x fibers.

channels in the visible spectrum (blue, red, and green) are used to visualize the three MYHC isoforms (MYHC1, MYHC2A, and MYHC2B, respectively). Fibers that lack or have very weak fluorescent signals are considered to be Type-IId/x fibers or MYHC2D/X-positive (see Fig. 1). Figure 2 illustrates the ability to compare and contrast the fiber type composition within a tibialis anterior (TA) cross-section (see Fig. 2a) to the intensity of β-gal transgene expression in the same muscle sample (see Fig. 2b). In addition to this method of tissue preparation, we have also included a subsection (see Subheading 2.1.2), which describes a procedure for tissue pretreatment that can decrease the extent of freezing-induced artifacts. Figure 3 illustrates the impact of freezing-induced artifacts on the overall quality of muscle cryosections. Freezing artifacts can distort muscle fiber borders and cause nonuniform X-gal staining. Analyzing muscle samples of transgenic lines that express β-gal at relatively low levels, such as the ROSA26 strain (2, 18), would be particularly problematic (see Fig. 3a). The pretreatment includes a prolonged fixation step, followed by an extended cryopreservation step in a 30% sucrose solution. Unfortunately, this treatment sacrifices the ability to obtain MYHC immunostained images and may reduce X-gal staining intensity (see Fig. 3b). This step benefits studies that do not require immunostaining, but

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Fig. 2. Serial TA muscle sections of a β-gal expressing transgenic mouse (6.5MCKΔMR1-β-gal) (23) treated by (a) immunostaining to identify the fiber type composition of the section (Type-I (blue), Type-IIa (red ), Type-IId/x (gray ), and Type-IIb (green)), and (b) treated by X-gal staining to visualize the distribution of β-gal expression. This transgenic tissue contains a higher X-gal staining intensity in the regions of the TA that contain more Type-IIb fibers.

Fig. 3. Comparison of X-gal stained 10 μm cryosections of ROSA26 quadriceps muscles under different preparative conditions. (a) Samples fixed in 4% PFA overnight followed by cryoprotection in 30% sucrose overnight exhibit distinct muscle fiber borders and uniform X-gal staining. (b) Samples that were not fixed or cryoprotected in sucrose exhibit numerous freezing artifacts throughout individual fibers and ill-defined fiber borders. Scale bar = 100 μm, insets are at 2× higher magnification.

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which do require fiber-to-fiber resolution and the visualization of lower-levels of β-gal such as might be needed in vector delivery studies (4, 19) that utilize β-gal as the reporter gene. The need for high-quality fiber staining across the entire muscle cross-section is even more critical for in vivo studies involving β-gal mRNA knockdown (20, 21), where the experimental manipulation itself is subject to penetrance variability. The long fixation times that tissue samples are subjected to may compromise the quality of other histological methods. For example, immunofluorescence is not compatible with the pretreatment step described in Subheading 2.1.2 because many antibody epitopes are destroyed by even short fixation periods, and fixatives can cause background fluorescence in muscle tissues. It is thus recommended to users who wish to apply the pretreatment step that they be aware that the long fixation time necessary for optimal X-gal staining may compromise the ability to use the same treated tissues in conjunction with other histological assays.

2. Materials 2.1. Cryofreezing and Cryosectioning Muscle Tissues 2.1.1. Cryofreezing Without Fixation

1. β-gal expressing mouse. The ROSA26 (Gt[ROSA]26Sor) gene trap line, which expresses β-gal ubiquitously in all tissues, can be used as a control (The Jackson Laboratory). 2. Gum Tragacanth (Sigma-Aldrich). 3. OCT (Sakura-Finetek). 4. 2-Methylbutane (Isopentane). 5. Liquid N2. 6. Mortar and pestle. 7. 10 × 10 × 5 mm plastic tissue Cryomold cassettes (SakuraFinetek). 8. 100 × 50 mm Pyrex dish. 9. 150 × 150 × 50 mm Styrofoam container (exact dimensions are not critical). 10. Glass slides. 11. Subzero thermometer (Digi-Sense Type-J Thermocouple) (Oakton Instruments). 12. Cryostat (CM1805 UV) (Leica).

2.1.2. Cryofreezing with Fixation and Cryopreservation (Pretreatment Step)

This section requires all of the materials described Subheading 2.1.1 in addition to the following materials:

in

1. Phosphate-buffered saline (PBS). 2. Paraformaldehyde (PFA). A 4% PFA in PBS solution (w/v) can be made for long-term storage at −20°C.

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3. Sucrose. A stock of 30% sucrose in sterile dH2O can be stored at room temperature. 4. Nutator. Any rocking platform that can perform gentle mixing will suffice. 2.2. Generating Monoclonal Antibodies from Cultured Hybridomas

1. Hybridoma lines BA-D5, SC-71, and BF-F3. (American Type Culture Collection). 2. Dulbecco’s modified Eagle medium (DMEM/High Glucose) (Hyclone). 3. Fetal bovine serum (FBS) (Gemini Bio-Products). 4. Penicillin–streptomycin (Sigma-Aldrich). 5. 150 mm TC-treated culture dish. 6. 0.22 μm Stericup filter unit (Millipore Corp). 7. HiTrap Protein G HP columns (GE Healthcare Bio-Sciences). 8. HiTrap IgM Purification HP columns (GE Healthcare BioSciences). 9. Slide-A-Lyzer Dialysis Cassettes (Pierce).

2.3. Fiber-Type Staining

1. Bovine serum albumin (BSA). 2. PBS (Sigma-Aldrich). 3. Tween-20 (Sigma-Aldrich). 4. Alexa Flour 350 goat anti-mouse IgG2b (Invitrogen). 5. Alexa Fluor 594 goat anti-mouse IgG1 (Invitrogen). 6. Alexa Fluor 488 goat anti-mouse IgM (Invitrogen). 7. Prolong Antifade Kit (Invitrogen). 8. PAP pen (Sigma-Aldrich). 9. Coplin jars. Alternatively, the smaller Five-Slide mailers (Fisher Scientific) can be used to save on reagent. 10. Humidity chamber. 11. Glass cover slips.

2.4. b-gal Staining

1. PBS. 2. PFA. 3. Monobasic sodium phosphate, NaH2PO4•H2O. 4. Dibasic sodium phosphate, Na2HPO4. 5. Magnesium Chloride, MgCl2. 6. Sodium deoxycholate. 7. NP-40 (Igepal-CA630). 8. Dimethylformamide (DMF).

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9. X-gal. A 25 mg/mL stock in DMF can be prepared and stored at −20°C. 10. Potassium ferricyanide, K3[Fe(CN)6]. A 0.5 M stock in sterile dH2O can be stored at room temperature for no longer than a month. It is recommended that this stock be made fresh whenever possible. 11. Potassium ferrocyanide, K4[Fe(CN)6]•3H2O. A 0.5 M stock in sterile dH2O can be stored at room temperature for no longer than a month. It is recommended that this stock be made fresh whenever possible. 12. 37% formaldehyde. 13. Gelvatol (Air Products). 14. Monopotassium phosphate, KH2PO4. 15. Sodium chloride, NaCl. 16. Sterile dH2O. 17. Five-Slide mailers (Fisher Scientific). 18. Glass cover slips.

3. Methods 3.1. Collecting, Cryofreezing, and Cryosectioning Muscle Tissues 3.1.1. Cryofreezing (Without Fixation)

This section describes a protocol that is optimal for muscle fiber immunostaining. It lacks the pretreatment steps (see Subheading 2.1.2) for optimal X-gal staining because these are incompatible with the immunostaining method described in Subheading 3.3. 1. Prepare the OCT: 10% gum tragacanth mixture. Use a mortar and pestle to first mix the 10% gum tragacanth solution (w/v). The consistency should be similar to toothpaste. Mix 2 parts OCT with 1 part 10% gum tragacanth (see Note 1). The final mixture should thus be 3.33% gum tragacanth and 66.67% OCT. Store the OCT: 10% gum tragacanth mixture at 4°C. 2. Label 10 × 10 × 5 mm plastic tissue Cryomold cassettes (see Note 2). Each cassette can hold several muscle tissues. For example, the 10 × 10 × 5 mm cassettes can accommodate two adjacent TA muscles (see Note 3). 3. Fill cassettes with chilled OCT: 10% gum tragacanth. Mix the compound thoroughly just before use, as gum tragacanth tends to separate from the OCT after a day of storage at 4°C. 4. Place the cassettes on ice. A metal block placed on ice offers a cool and even surface. Prechilling the cassette on ice in this manner will prevent the OCT: 10% gum tragacanth (2:1) solution

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from becoming too liquescent, and will allow tissues to be positioned with greater ease. Additionally, the colder temperatures will slow the degradation of β-gal enzymatic activity. 5. Dissect tissues of interest and blot on filter paper to remove blood and excess moisture. Avoid rinsing tissues in PBS (see Notes 4–6). Transfer tissues into the chilled cassettes. Allow tissues to set in the cassettes for ~30 min. Gum tragacanth is a natural desiccant and will draw out additional liquid from the tissue to minimize freezing artifacts. 6. Prepare liquid nitrogen-chilled isopentane bath. Chill isopentane to about −150°C. A simple bath setup consists of a 100 × 50 mm Pyrex dish filled with ~150 mL of isopentane, set in a larger Styrofoam container (150 × 150 × 50 mm) that is deep enough to fill with liquid N2 to a level equal to that of the isopentane, yet shallow enough so that evaporating liquid N2 does not visually obscure the cassettes in the bath. The isopentane should partially freeze so that the bottom and the rim of the isopentane bath are frozen, and a pool of liquid isopentane large enough to float a cassette on remains (see Note 7). A subzero thermometer can be used to measure the actual temperature of the bath. 7. Ensure that tissues are oriented correctly in the OCT: 10% gum tragacanth compound. The tissues should be arranged so that fibers run perpendicular to the cassette surface for crosssections or set parallel to the cassette surface for longitudinal sections (see Note 8). 8. Float cassettes in the prechilled isopentane and allow the tissues to completely freeze (2–3 min) (see Note 9). 9. Place frozen cassettes immediately into a −80°C freezer for storage, or in the prechilled cryostat chamber for immediate sectioning. Some protocols recommend burying cassettes in dry ice pellets before transferring to −80°C; however, this could result in small fluctuations in temperature and may result in freezing artifacts. Therefore, a direct transfer from chilled isopentane to a −80°C freezer is ideal. Tissues can remain at −80°C indefinitely. When transferring cassette samples from −80°C to the cryostat, allow at least 30 min for the sample to acclimate to the cutting temperature. Cutting at too low a temperature will result in chipping the sample. After warming to −25°C, do not restore at −80°C. Doing so can cause additional formation of ice crystals, and thus produce more freezing artifacts. 10. Mount tissue blocks on cryostat chuck with OCT. Follow manufacturer’s recommended procedures for operating the cryostat. 11. Cut tissues at a desired thickness and temperature (see Note 10).

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12. Mount tissue slices onto room temperature glass slides (see Note 11). Static attraction should pull the section onto the glass slide when they are in close proximity with each other. Proteins in the sample should allow the muscle sections to stay firmly adhered to the slide, thus there is no immediate requirement to fix at this step. Air dry at room temperature and place in −80°C for long-term storage, where they can remain indefinitely. 3.1.2. Cryofreezing with Fixation and Cryopreservation (Pretreatment)

This section provides tissue cryopreservation pretreatment steps that will produce superior sections for X-gal staining, described in Subheading 3.3, but is incompatible with the immunohistochemistry method of Subheading 3.2. 1. Fill labeled multiwell plate or Eppendorf tubes with ~1 mL of PBS and chill on ice. 2. Dissect muscles of interest and place them immediately into the PBS. Store no longer than 30 min. 3. Transfer tissues into 1.5 mL Eppendorf tubes containing 1 mL of 4% PFA in PBS. Rock on a Nutator (or a rocking platform) at 4°C for 3 h to overnight. 4. Transfer tissues to 12 mL conical tubes containing 10 mL PBS. 5. Wash tissues three additional times with 10 mL of PBS at 4°C and rock for the following times: 2, 10, and 30 min. 6. Blot excess buffer from tissues and place them in a 1.5 mL Eppendorf tube containing 1 mL of 30% sucrose. Rock overnight 4°C (see Note 12). 7. Proceed to steps 1–12 of Subheading 3.1.1.

3.2. Preparation of Monoclonal Antibodies

This section provides detailed instructions for generating the BA-D5, SC-71, and BF-F3 antibodies from cultured hybridomas. 1. Inoculate the hybridoma culture lines BA-D5, SC-71, and BF-F3 separately in DMEM/high glucose supplemented with 10% FBS and penicillin–streptomycin, and grow to near confluence in 150-mm tissue culture dishes. 2. Rinse the cultures 1× with serum-free medium (SFM) and then begin mAb production in 25 mL of SFM. Incubate for 2–3 days, depending on cell viability. 3. Collect and centrifuge the medium at 130 × g for 5 min. 4. Filter-sterilize medium through a 0.22 μm Stericup filter unit. 5. Concentrate monoclonal antibodies (mAb) using HiTrap columns according to manufacturer’s instructions. BA-D5 and SC-71 are IgGs, while BF-F3 is an IgM. Therefore, use HiTrap Protein G HP columns to concentrate mAbs BA-D5 and SC-71, and HiTrap IgM Purification HP columns to concentrate the mAb BF-F3.

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6. Collect eluent in 1 mL fractions. 7. Pool the first and second highest concentrated fractions containing the eluent as determined by Bradford assay. 8. Dialyze the pools using the Slide-A-Lyzer dialysis cassettes against two changes of 1.5 L PBS for 24 h at 4°C. The protein concentration as determined by Bradford assay should be between 100–200 mg/mL. 9. Subdivide into aliquots whose volumes are appropriate for amounts needed for typical sample numbers to be immunostained. Store at −20°C. Avoid multiple freeze-thaw cycles when using antibodies since this tends to lower the antibody potency. 3.3. Fiber-Type Staining

The following section provides instructions for treating the mounted cryosections from Subheading 3.1 for visualization by fluorescence microscopy. Four-channel fluorescence microscopy allows visual identification of the Type-I, Type-IIa, Type-IId, and Type-IIb fiber types on a single section (Type-IId fibers remain nonimmunostained after exposure to the three monoclonal antibodies). A MYHC2D/X-specific monoclonal is available (22) and can be applied to this protocol as well. 1. Construct a humidity chamber. A humidity chamber is simply any lidded storage box that is of reasonable size that can accommodate the number of slides you wish to treat. The chamber is lined at the bottom with paper towels that are moistened with water. Staining slides can be placed on top of a rack so that the slides themselves are not in direct contact with any moisture. 2. Prepare fresh Blocking Buffer (1% BSA, 0.05% Tween-20 in PBS). For short-term storage, keep at 4°C. 3. Prepare the primary antibody mixture. Calculate the volume of mAbs required for immunostaining and dilute the BA-D5, SC-71, and BF-F3 mAbs in Blocking Buffer (see Note 13). Approximately, 30–60 μL of antibody mixture is required to cover most sections from mouse muscles (see below). Prepare mAbs mixture on the day of use. When working with antibodies, always keep them chilled on ice until they are ready to be placed on the sample. 4. Remove slides from −80°C storage and air dry. 5. Circle section with a PAP pen leaving enough room around edges of section to aspirate off liquid. This will reduce the amount of antibody usage by keeping liquid pooled in a small droplet. 6. Immerse sections in blocking buffer for 20–30 min at room temperature in Coplin jars. 7. Carefully aspirate excess liquid off slides, without touching the sections.

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8. Add diluted mAbs to sections and incubate slides in the humidity chamber for 1 h at room temperature. 9. Secondary antibody mixture. Calculate the volume of secondary antibody required for the number of sections being stained with each mAb type (30–60 μL per section). Dilute Alexa Flour 350 goat anti-mouse IgG2b, Alexa Fluor 594 goat antimouse IgG1, and Alexa Fluor 488 goat anti-mouse IgM 1:100 in Blocking Buffer. Keep mixture on ice. 10. Rinse slides 4× in PBS in Coplin jars at room temperature. Slide mailers can be used to reduce the volumes of rinse and wash solutions. 11. Wash slides 3× for 5 min in Blocking Buffer in Coplin jars at room temperature. 12. Carefully aspirate excess liquid off sections. 13. Add 50 μL of secondary antibody to sections and incubate slides in the humidity chamber for 30 min at room temperature. 14. Rinse slides 4× in PBS in Coplin jars at room temperature. 15. Wash slides 3× for 5 min in Blocking Buffer in Coplin jars at room temperature. 16. Wash slides 2× for 5 min in PBS in Coplin jars at room temperature. 17. Rinse slides 2× in dH2O. 18. Remove excess liquid from slides by aspiration. 19. Mount coverslips using ProLong Antifade Kit reagent according to manufacturer’s instructions. Let slides harden overnight on a flat surface, and avoid exposure to light. 20. Visualize by fluorescence microscopy. The fluorescent signal will remain strong for ~1 week. 3.4. b-gal Staining

This section describes the X-gal staining method for mounted sections obtained in Subheading 3.1. To facilitate correlations between X-gal and MYHC-stained fibers, this procedure should be performed on sections that are as contiguous as possible to the sections selected for fiber-type identification (Subheading 3.2). 1. Prepare Gelvatol mounting medium. First, prepare the Gelvatol-buffered saline solution (25% (w/v) Gelvatol, 10 mM KH2PO4/Na2HPO4 (pH 7.2), 140 mM NaCl in dH2O) and stir at 37°C for several hours. Add glycerol in an amount equal to one-half the total volume of the Gelvatol-buffered saline solution and stir overnight at room temperature. Centrifuge the Gelvatol solution to remove undissolved particles. Pipette the supernatant into smaller 1 mL aliquots. Check pH of Gelvatol solution. It should be between pH 6 and 7. Store Gelvatol solution at 4°C for up to 1 year. Do not leave Gelvatol uncapped for longer than necessary to avoid evaporation.

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2. Prepare Wash Buffer (100 mM monobasic sodium phosphate/ dibasic sodium phosphate, 2 mM MgCl2, 0.01% sodium deoxycholate, 0.02% NP-40 in sterile dH2O, buffer at pH 7.3). Store at room temperature for up to 6 months. 3. Prepare X-gal staining solution (5 mM potassium ferricyanide, 5 mM potassium ferrocyanide, 1 mg/mL X-gal in wash buffer). Always prepare this fresh and keep on ice until used. 4. Remove slides from −80°C storage and air dry. 5. Fix slides in 4% PFA in PBS for 15 min at 4°C. 6. Wash 3× for 5 min each with Wash Buffer at room temperature. 7. Stain by immersing slides into freshly prepared X-gal staining solution and incubate for 2–48 h at 37°C in the dark. Check periodically for the intensity of blue staining (see Note 14). 8. Postfix in 10% buffered formalin (1 part 37% formaldehyde and 9 parts PBS) for 5 min at 4°C. 9. Wash in PBS 3× for 5 min at room temperature. 10. Rinse in H2O and remove excess liquid on the sections by blotting/touching edge of slide on paper towel. 11. Mount glass cover slip with Gelvatol medium. 12. Visualize by brightfield microscopy. The X-gal stain will not fade over time.

4. Notes 1. The gum tragacanth is difficult to mix uniformly in the OCT (it usually takes several days) and mixing usually causes the accumulation of bubbles. Heating at 60°C for several min followed by a high-speed centrifugation can be used to expel bubbles. However, the simplest process is to mix the solution several times daily, and allow the bubbles to naturally rise to the top. 2. Use a “super-permanent” marker to label cassettes since exposure to the isopentane may remove the ink of standard lab markers. 3. Avoid crowding the tissue block. Tissues should be placed at least 1 mm away from the edges of the cassette. Tissues do not section well if they are placed immediately adjacent to the block edges. 4. IACUC rules governing the euthanization of animals by CO2, lethal intraperitoneal injection of anesthesia (e.g. lethal dosage of ketamine and xylazine mixture), or by cervical dislocation may vary between approved protocols. These procedures do not appear to affect staining outcomes if muscle samples are handled rapidly.

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5. Exposing tissues to additional moisture without proper cryoprotection pretreatment (see Subheading 3.1.2) exacerbates freezing artifacts. 6. There is no preferred technique for dissecting different anatomical muscles other than to avoid tearing the tissue, since this negatively affects overall histology. For small muscles such as the soleus, it is recommended that the entire muscle be dissected (from origin to insertion), since the extra tissue length allows room for “practice cuts” in cryosectioning steps before satisfactory sections are mounted onto slides. Visible connective tissue and fat should be removed from the surrounding muscle sample, since inconsistencies in the sample can lead to snagging of the sample during cryosectioning, as the microtome is sensitive to differences in the cutting material. For example, the optimal cutting temperature for fatty tissue ranges from −25 to −35°C, while the optimal cutting temperature for muscle ranges from −15 to −25°C. 7. If the entire isopentane bath freezes, it can be rapidly thawed by touching it with the handle of a large wrench, without having to remove the isopentane bath from the liquid N2. Avoid injury by using the proper protection for handling liquid N2. 8. Since the cutting surface begins at what is the bottom of the cassette, tissue(s) may be placed as close to the bottom as possible. This will limit the number of empty/practice cuts necessary to reach the tissue during cryosectioning. 9. Float 1–2 cassettes at a time to avoid sinking adjacent cassettes. If cassettes sink to the bottom of the bath, they may freeze to the frozen isopentane. If this happens, forced removal of the cassette may damage the cassette, since the plastic is brittle at these extremely low temperatures. Removal of the isopentane bath dish from the liquid nitrogen may be necessary to thaw the frozen isopentane enough to facilitate the removal of a sunken cassette. If a cassette does not float, but sinks immediately to the bottom and makes contact with the frozen isopentane, there is a good chance that the sample may crack. 10. It is recommended that a first-time user cut 10 μm sections at −20°C. These parameters can then be modified depending on the tissue and personal preference. Since tissues are embedded in a OCT: 10% gum tragacanth compound, the blocks are more susceptible to chipping or flaking compared to blocks mounted in OCT alone. Because of this, it is recommended that sections be cut using the antiroll bar that is included with the CM1805 UV cryostat models, rather than assisting the section off the chunk and blade using a paintbrush. Cutting at extremely low temperatures causes chafing of the block, while cutting at warmer temperatures causes gumming of the section. Unfortunately, the cryosectioning process is more of an “art

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form” than a technical method. Acquiring this skill is best done through proper training and unlimited practice. 11. Ideally, sections should be cut and mounted onto several slides. Sections treated for immunostaining (see Subheading 3.3) and β-gal treatment (see Subheading 3.4) are performed on separate slides; therefore, sections to be compared should be as contiguous as possible, since morphological differences throughout the length of the muscle tissue may complicate the ability to identify the same fibers between serial sections. The more consecutive the sections, the easier it is to relate sections treated for X-gal staining to sections treated for fibertype immunostaining. 12. These volumes are appropriate for mouse muscles that range from 10 mg (soleus muscle, wet weight) to 300 mg (quadriceps muscle, wet weight). 13. It is recommended that the overall strength or working dilution of the mAbs be tested on sample muscle sections before applying to experimental slides. When mAbs are prepared according to Subheading 3.2, typical dilutions range from 1:10 to 1:1,000. 14. For tissues that express low amounts of the β-gal transgene, such as the Rosa26 strain, staining can take as long as 48 h at 37°C in the dark. Keep slides rocking in the stain solution to attain even staining.

Acknowledgments Miki Haraguchi and Paul Gregorevic are thanked for their initial technical assistance and very helpful advice; Joel R. Chamberlain is thanked for providing a ROSA26 mouse; and Robert E. Welikson, Charis L. Himeda, and Joel R. Chamberlain are thanked for their critical comments on earlier versions of the manuscript. This research was supported by grants from the NIH RO1-AR18860 and 1P01-NS046788 to SDH and by an NIH Developmental Biology Training Grant 5732-HD07183 to PWLT. References 1. Rosenthal, N. (1987). Identification of regulatory elements of cloned genes with functional assays. Methods Enzymol 152, 704–20. 2. Friedrich, G., and Soriano, P. (1991). Promoter traps in embryonic stem cells: a genetic screen to identify and mutate developmental genes in mice. Genes Dev 5, 1513–23.

3. Hauser, M. A., Robinson, A., HartiganO’Connor, D., Williams-Gregory, D. A., Buskin, J. N., Apone, S., Kirk, C. J., Hardy, S., Hauschka, S. D., and Chamberlain, J. S. (2000). Analysis of muscle creatine kinase regulatory elements in recombinant adenoviral vectors. Mol Ther 2, 16–25.

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4. Gregorevic, P., and Chamberlain, J. S. (2005). Functional enhancement of skeletal muscle by gene transfer. Phys Med Rehabil Clin N Am 16, 875–87, vii-viii. 5. Fan, Q., Yee, C. L., Ohyama, M., Tock, C., Zhang, G., Darling, T. N., and Vogel, J. C. (2006). Bone marrow-derived keratinocytes are not detected in normal skin and only rarely detected in wounded skin in two different murine models. Exp Hematol 34, 672–9. 6. Badea, T. C., Hua, Z. L., Smallwood, P. M., Williams, J., Rotolo, T., Ye, X., and Nathans, J. (2009). New mouse lines for the analysis of neuronal morphology using CreER(T)/loxPdirected sparse labeling. PLoS One 4, e7859. 7. Yamashita, K., and Yoshioka, T. (1991). Profiles of creatine kinase isoenzyme compositions in single muscle fibres of different types. J Muscle Res Cell Motil 12, 37–44. 8. Johnson, J. E., Wold, B. J., and Hauschka, S. D. (1989). Muscle creatine kinase sequence elements regulating skeletal and cardiac muscle expression in transgenic mice. Mol Cell Biol 9, 3393–9. 9. LaFramboise, W. A., Guthrie, R. D., Scalise, D., Elborne, V., Bombach, K. L., Armanious, C. S., and Magovern, J. A. (2003). Effect of muscle origin and phenotype on satellite cell muscle-specific gene expression. J Mol Cell Cardiol 35, 1307–18. 10. Scott, W., Stevens, J., and Binder-Macleod, S. A. (2001). Human skeletal muscle fiber type classifications. Phys Ther 81, 1810–6. 11. Larsson, L., Edstrom, L., Lindegren, B., Gorza, L., and Schiaffino, S. (1991). MHC composition and enzyme-histochemical and physiological properties of a novel fast-twitch motor unit type. Am J Physiol 261, C93–101. 12. Zierath, J. R., and Hawley, J. A. (2004). Skeletal muscle fiber type: influence on contractile and metabolic properties. PLoS Biol 2, e348. 13. Gregorevic, P., Meznarich, N. A., Blankinship, M. J., Crawford, R. W., and Chamberlain, J. S. (2008). Fluorophore-labeled myosin-specific antibodies simplify muscle-fiber phenotyping. Muscle Nerve 37, 104–6. 14. Schiaffino, S., Gorza, L., Sartore, S., Saggin, L., Ausoni, S., Vianello, M., Gundersen, K., and Lomo, T. (1989). Three myosin heavy chain

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isoforms in type 2 skeletal muscle fibres. J Muscle Res Cell Motil 10, 197–205. 15. Gorza, L. (1990). Identification of a novel type 2 fiber population in mammalian skeletal muscle by combined use of histochemical myosin ATPase and anti-myosin monoclonal antibodies. J Histochem Cytochem 38, 257–65. 16. Sokoloff, A. J., Yang, B., Li, H., and Burkholder, T. J. (2007). Immunohistochemical characterization of slow and fast myosin heavy chain composition of muscle fibres in the styloglossus muscle of the human and macaque (Macaca rhesus). Arch Oral Biol 52, 533–43. 17. Town, S. C., Putman, C. T., Turchinsky, N. J., Dixon, W. T., and Foxcroft, G. R. (2004). Number of conceptuses in utero affects porcine fetal muscle development. Reproduction 128, 443–54. 18. Zambrowicz, B. P., Imamoto, A., Fiering, S., Herzenberg, L. A., Kerr, W. G., and Soriano, P. (1997). Disruption of overlapping transcripts in the ROSA beta geo 26 gene trap strain leads to widespread expression of beta-galactosidase in mouse embryos and hematopoietic cells. Proc Natl Acad Sci USA 94, 3789–94. 19. Hauser, M. A., Amalfitano, A., Kumar-Singh, R., Hauschka, S. D., and Chamberlain, J. S. (1997). Improved adenoviral vectors for gene therapy of Duchenne muscular dystrophy. Neuromuscul Disord 7, 277–83. 20. Yu, J., and McMahon, A. P. (2006). Reproducible and inducible knockdown of gene expression in mice. Genesis 44, 252–61. 21. Kanzler, B., Haas-Assenbaum, A., Haas, I., Morawiec, L., Huber, E., and Boehm, T. (2003). Morpholino oligonucleotide-triggered knockdown reveals a role for maternal E-cadherin during early mouse development. Mech Dev 120, 1423–32. 22. Lucas, C. A., Kang, L. H., and Hoh, J. F. (2000). Monospecific antibodies against the three mammalian fast limb myosin heavy chains. Biochem Biophys Res Commun 272, 303–8. 23. Tai, P. W., Fisher-Aylor, K. I., Himeda, C. L., Smith, C. L., Mackenzie, A. P., Helterline, D. L., Angello, J. C., Welikson, R. E., Wold, B. J., and Hauschka, S. D. (2011). Differentiation and fiber type-specific activity of a muscle creatine kinase intronic enhancer. Skelet Muscle 1, 25.

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Chapter 27 Determination of Gene Promoter Activity in Skeletal Muscles In Vivo Sarah M. Senf and Andrew R. Judge Abstract The use of nonviral (plasmid DNA) gene delivery into skeletal muscle has increased significantly in recent years. The procedure is used to overexpress wild-type proteins, express mutant proteins, or knock down endogenous proteins. These manipulations can identify the role of a specific protein in muscle cell biology and physiology. The same procedure of plasmid DNA gene delivery can be used to introduce a gene promoter reporter construct. Such constructs contain a defined sequence of a gene promoter that regulates the expression of a “reporter.” This reporter is easily measured and reflects the in vivo transcriptional activity of the gene promoter sequence under study. The gene promoter can be mutated at known transcription factor-binding sites, truncated to identify specific regions of the gene promoter that are required for transcription, or introduced into skeletal muscle with an expression plasmid for a protein believed to regulate the gene’s transcription. Therefore, the use of such gene promoter reporters allows for an in-depth physiological assessment of the gene’s transcriptional regulation. Key words: Gene promoter reporter, Gene transcriptional activity, Gene regulatory region, Skeletal muscle, Plasmid injection, Electrotransfer

1. Introduction The study of gene expression and gene regulation is important to our understanding of the multifaceted regulatory networks that control biological and physiological processes. Gene expression analyses, using either northern blots or reverse transcriptasequantitative polymerase chain reaction (RT-qPCR), measure mRNA levels and provide important information regarding gene transcription

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from their native chromosomal environment. While information can be obtained regarding gene regulation through mRNA expression analysis, the use of gene promoter reporter systems has greatly contributed to the understanding of gene regulation. Gene promoter reporters consist of a gene promoter sequence cloned into a reporter vector upstream of a reporter gene, thereby regulating the reporter’s expression. Following introduction of the promoter reporter construct into cells, measurement of either the reporter’s expression or activity reflects the activity of the promoter sequence. Common reporters include luminescent markers, such as luciferase, or fluorescent markers, such as green fluorescent protein (GFP). Such constructs are widely used in cultured muscle cells and provide important mechanistic data regarding gene regulation. However, their use in whole muscle, in vivo, provides information regarding gene promoter activity in a physiologically relevant environment (1–4). The ability to use gene promoter reporter plasmids in vivo depends on the transfection of whole muscle, which can be achieved through direct injection and electroporation (electrotransfer). This procedure of plasmid injection and electroporation into skeletal muscle in vivo was recently described in detail ((5) or see Wu and Kandarian in Chap. 13). A potential downside of studying gene promoter regulation using a promoter reporter plasmid is that plasmid DNA remains extrachromosomal. Therefore, any chromosomal regulatory information is lost (6). Moreover, regulatory regions cloned into reporter vectors are typically fragments of the 5c flanking region, which discounts any regulatory regions (1) upstream of the fragment cloned (2), in the 3c flanking region, and (3) that are intronic, which may also regulate gene transcription. Therefore, comparisons between gene promoter reporter activity and endogenous mRNA expression can be informative in this regard since changes in endogenous mRNA may be an outcome of genomic DNA regulation at any of these regulatory regions. However, it is important to recognize that mRNA levels may also be influenced by factors which regulate mRNA stability. Therefore, disparities between gene promoter reporter activity and mRNA levels may, in some cases, actually highlight the involvement of posttranscriptional regulatory mechanisms in the regulation of gene expression. One major advantage of using gene promoter reporters in the study of transcriptional regulation is the ability to study specific DNA sequences thought to be responsible for gene transcription. After a DNA sequence believed to be involved in the regulation of a specific gene has been identified, a simple way of identifying regulatory regions is to create a series of truncated versions of the promoter region and compare reporter activity in response to an appropriate stimulus. This can significantly shorten the unknown regulatory region that is relevant to the stimulus of interest.

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Scanning of this unknown region for transcription factor consensus sequences, which can be accomplished by using various software programs, can further assist in narrowing the unknown regulatory region to several potential base-pair sequences. Site-directed mutagenesis of these specific sequences can subsequently identify whether these sites are required for gene transcription. The following methods describe the use and measurement of gene promoterdependent luciferase reporters in skeletal muscle, in vivo.

2. Materials 2.1. Gene Promoter Reporter Plasmid Preparation

1. A reporter plasmid with the promoter (or regulatory) region of a gene of interest cloned upstream, and driving expression of luciferase, or a detectable marker (reporter gene). 2. Chemically competent bacteria (we typically use either DN5A or top ten strains of Escherichia coli). 3. SOC medium: 2% tryptone, 0.5% yeast extract, 10 mM NaCl, 2.5 mM KCl, 10 mM MgCl2, 10 mM MgSO4, 20 mM glucose. 4. Luria broth (LB) media (per 1 L): 10 g tryptone, 5 g yeast extract, 10 g NaCl. 5. LB-agar (per 1 L): 10 g tryptone, 5 g yeast extract, 10 g NaCl, 15 g bacto-agar. 6. Antibiotic (commonly ampicillin or kanamycin): Reconstitute ampicillin to 50 mg/mL and kanamycin to 10 mg/mL, both in ddH2O. 7. A commercially available, endotoxin-free plasmid DNA kit (we use Qiagen Plasmid Mega or Maxi kits, which provide DNA yields of up to 500 Mg or 2.5 mg, respectively). 8. Phosphate-buffered saline (PBS; per 1 L): 8 g NaCl, 0.2 g KCl, 1.44 g Na2HPO4, 0.24 g KH2PO4. 9. Agarose. 10. Ethidium bromide. 11. Tris/Borate/EDTA (TBE) buffer (5× stock, per 1 L; working solution is 1×): 54 g Tris base, 27.5 g boric acid, 20 mL of 0.5 M EDTA. 12. Nucleic acid sample loading buffer (we use Bio-Rad Nucleic Acid Sample Loading Buffer, 5×: 50 mM Tris–HCl, pH 8.0, 25% glycerol, 5 mM EDTA, 0.2% bromophenol blue, 0.2% xylene cyanole FF). Xylene cyanole runs ~3,000 bp (±200 nucleotides) and bromophenol blue runs at 300 bp (±100 nucleotides).

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2.2. Gene Reporter Plasmid Injection and Electroporation

1. 28-guage 3/10 cc insulin syringe. 2. Surgical tools (scalpel and blade, hemostats, forceps). 3. Electroporation generator (we use the BTX-Harvard Apparatus ECM 830 Square Wave Electroporator). 4. Calipers with electrodes for electroporation (for rat soleus muscles we use BTX tweezertrodes with a 7 mm diameter that have a stainless steel circular electrode at the tip). 5. Sutures: 5–0 sutures (fascia), 4–0 sutures (skin).

2.3. Luciferase Activity

1. Tissue homogenizer and homogenizing tubes. 2. Passive lysis buffer (PLB; Promega). 3. Luciferase Assay System (Promega). 4. Luminometer (we use the GloMax 20/20 single tube Luminometer (Promega)).

3. Methods 3.1. Gene Promoter Reporter Plasmid Preparation

1. Thaw TOP10 E. coli cells on ice. Add 25 ML of cells to a prechilled polypropylene 14 mL round bottom culture tube and add 1–2 ML of reporter plasmid DNA. Mix by flicking the tube gently. Do not vortex. Incubate on ice for 20 min. 2. Heat-shock the mixture by submerging the tube in a 42°C water bath for 50 s. Place back on ice. 3. Add 200 ML of prewarmed (37°C) SOC media to the mixture tube. Shake at 250 rpm for 1 h at 37°C. 4. Streak two different mixture amounts (factor of 10) on a selective agar plate containing the appropriate antibiotic (see Note 1) and incubate overnight at 37°C. 5. The next morning prepare a starter culture. Pick an isolated colony (we use a 1 mL pipette tip) from the plate that was incubated overnight and place into a 14 mL round bottom falcon tube containing 3 mL of LB media and the appropriate antibiotic. Shake at 250–300 rpm for ~8 h at 37°C. 6. Prepare a larger culture using a 1/500 dilution of starter culture into a larger volume of LB media containing the appropriate antibiotic, using a flask that is 5× the liquid volume. Shake at 250–300 rpm for 12–16 h at 37°C. 7. Harvest bacterial cultures and isolate reporter plasmid DNA using an endotoxin-free plasmid preparation kit (we use either Qiagen EndoFree Plasmid Mega or Maxi Kits depending on the amount of plasmid DNA needed). Follow manufacturer’s instructions.

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8. Following precipitation of plasmid DNA, redissolve the plasmid DNA pellet in an appropriate volume of 1× PBS (physiological pH of 7.4) to resuspend the plasmid DNA (see Note 2). Do not vortex or pipette the pellet up and down to promote resuspension, as this may cause shearing of the DNA. To promote resuspension of the plasmid DNA, place the suspension at 4°C overnight. The next morning, briefly centrifuge the plasmid DNA suspension at 1,000 × g for 1 min at 4°C and then read the concentration (see Note 3). 3.1.1. Reading Plasmid Concentrations

1. To determine plasmid DNA concentration, measure the absorbance of plasmid DNA through UV spectrophotometry at 260 nm. Since protein, which should be minimized for in vivo injections, absorbs light maximally at 280 nm, the A260/A280 ratio should also be calculated. A260/A280 ratio ~1.8 is recommended. 2. As stated in the Qiagen Endofree Plasmid Purification Handbook, and from our own experience, A260 (absorbance) readings of plasmid DNA are the most accurate between 0.1 and 1.0. Therefore, to determine the plasmid concentration, it is necessary to dilute a small volume (e.g., 3 ML) of plasmid DNA in 1× PBS (e.g., 300 ML) into a separate tube for absorbance readings. Mix the diluted plasmid thoroughly by inverting the tube several times. Transfer diluted plasmid into a cuvette and place in spectrophotometer. Enter the dilution volumes (e.g., 3 ML sample + 300 ML diluent) and read absorbance. If the A260 reading is significantly greater than 1.0, repeat the procedure using a higher volume of diluent. Record concentration (Mg/ML) and A260/A280 ratio. Repeat this step at least twice more to confirm plasmid concentration. 3. Remove a small volume (0.5 ML) of plasmid DNA and linearize using a restriction enzyme that will cut the plasmid at a single site. Run this linearized plasmid DNA and a small volume of your uncut plasmid DNA on an ethidium bromide stained 1% agarose gel (0.5 mg agarose/50 mL 1× TBE). This will help confirm that the correct plasmid was amplified. Supercoiled, uncut plasmid DNA is necessary for transfection and may migrate slightly faster than the linearized plasmid, which will migrate to the expected size of the plasmid.

3.1.2. Determining Plasmid Concentrations for Injections

1. For each rat soleus muscle of ~100 mg in mass, inject 50 Mg of promoter reporter (see Note 4) plasmid DNA diluted in 50 ML 1× PBS (see Note 5). Therefore, the plasmid concentration is 50 Mg/50 ML, or 1.0 Mg/ML. Dilute the plasmid DNA accordingly in 1× PBS and be sure to reread the diluted plasmid at least three times to ensure that the concentration is correct. 2. If injecting two different plasmids (a promoter reporter plasmid plus an expression plasmid), see Note 6.

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3.2. Gene Reporter Plasmid Injection and Electroporation

1. Before beginning the procedure, remove the plasmid preparation to be injected from the 4°C refrigerator and allow it to come to room temperature so that a cold solution is not injected into the muscle. 2. Under adequate anesthesia, shave the hind limbs and prepare the lateral side of the hind limbs for surgery using three alternating preparations of Betadine and 70% ethanol (for rinsing), to create a sterile surgical site. 3. To isolate the rat soleus muscle (see Note 7), make a small incision (~0.5 in. long) on the lateral side of the lower leg using a new blade, penetrating the skin first, then the underlying fascia. Enter the incision site with sterilized hemostats and isolate the soleus muscle on both its anterior surface and its posterior surface by blunt dissection (see Fig. 1). 4. Inject 50 Mg of endotoxin-free gene promoter reporter plasmid in a total volume of 50 ML of sterile 1× PBS using a 28-guage 3/10 cc insulin syringe. To do this, insert the syringe near the distal myotendinous junction and push it along the longitudinal axis of the muscle, toward the proximal myotendinous junction. Inject the plasmid DNA evenly throughout the longitudinal axis of the muscle during syringe withdrawal. 1 min following injection, deliver electric pulses, using an electric pulse generator (Electro Square porator ECM 830, BTX), by placing two paddle-like electrodes on each side of the muscle. Deliver five electric pulses at 75 V/cm, duration of 20 ms, and interpulse interval of 200 ms (see Note 8). 5. Suture the fascia incision using 5–0 sutures and suture the skin with 4–0 sutures. The entire procedure (i.e., plasmid injection into both solei) takes approximately 10 min. 6. Take some of the remaining plasmid from the tube used to inject the muscles and separate in an agarose gel (1% for 90 min) to verify that the plasmid has not become degraded, as this could negatively affect the transfection efficiency. This postinjection verification provides a final confirmation on the plasmid’s integrity.

3.3. Luciferase Activity

1. Seven to 10 days following plasmid injection, remove the soleus muscle (see Note 9), immediately freeze in liquid nitrogen, and store at −80°C until processing. 2. Thoroughly homogenize the whole soleus muscle in 1 mL of passive lysis buffer (Promega; see Note 10). 3. Centrifuge the muscle homogenate at 5,000 × g for 20 min at 4°C to pellet the cell debris, and transfer the resulting supernatant to a clean tube. 4. For manual luminometers, add 100 ML of Luciferase Assay Reagent into 1.5 mL eppendorf tubes, one tube per sample.

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Fig. 1. Representative images to illustrate the injection of rat soleus muscle with a gene promoter reporter plasmid. Tape the foot down to anchor the limb in place, such that the lateral side of the lower limb is visible and accessible (a). Make an incision through the skin (b) and then through the fascia of the intramuscular septum at the insertion of the biceps femoris (c). Enter the incision site with hemostats and blunt dissect the soleus on both its anterior (d) and posterior (e) surface. Evenly inject the gene promoter reporter plasmid along the longitudinal axis of the muscle (f).

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Please refer to the Promega handbook if using other types of luminometers. 5. Program the luminometer to perform a 2-s measurement delay followed by a 10-s measurement read for luciferase activity, as indicated in the Promega Technical Bulletin for the Luciferase Assay System. 6. Add 10 ML of muscle homogenate to a luminometer tube containing the luciferase assay reagent (see Note 11). Mix by pipetting several times. 7. Place the tube in the luminometer, measure luminescence, and record the reading (see Fig. 1). 8. Alternative means to determine gene promoter-dependent luciferase activity or expression include in vivo bioluminescent imaging and immunohistochemical analysis (see Note 12).

4. Notes 1. Both agar plates and LB media containing antibiotics should include antibiotics at the recommended working concentrations: 100 Mg/mL for ampicillin, and 50 Mg/mL for kanamycin. 2. Plasmid DNA resuspended in PBS is not protected from nucleases. It is therefore extremely important to plan the plasmid preparation such that the plasmid is resuspended in sterile PBS no more than 2–3 days before injections and stored at 4°C. Alternatively, if the plasmid DNA needs to be prepared at an earlier time point, the plasmid DNA pellet can be resuspended in endotoxin-free Tris–EDTA (TE) buffer and stored at −20°C to protect against nucleases. The DNA can then be precipitated out of TE and resuspended in sterile PBS 2–3 days prior to injection. TE should not be used for plasmid DNA injections, in vivo, since it compromises the transfection efficiency (7). Importantly, gene reporter activity should only be compared between muscles from animals injected concurrently with promoter reporter plasmid made from the same plasmid batch (i.e., made from the same plasmid prep). 3. Maximum capacity of QIAGEN-tips for Maxi preps: ~500 Mg DNA. Maximum capacity of QIAGEN-tips for Mega Preps: ~2,500 Mg DNA. Therefore, following resuspension of DNA pellets in the recommended volume of 1× PBS, plasmid DNA concentrations will be ~3 Mg/ML following Maxi Prep and ~3–4 Mg/ML following Mega Prep (if maximum amount of DNA is recovered). Refer to QIAGEN handbook for guidelines to increase plasmid DNA amplification and recovery.

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4. The microgram amount of a gene promoter reporter injected into skeletal muscle significantly affects luciferase activity (8). Similarly, the amount of CMV-driven luciferase expression is dependent upon the dose injected (9). We inject 50 Mg based on data from these referenced studies. 5. There is evidence that luciferase activity, driven by the SV 40 promoter, is increased (7), but also that luciferase expression, driven by a CMV promoter, is unaltered (9), with the volume of vehicle injected (for a constant plasmid DNA amount). We inject 50 ML into a rat soleus of ~100 mg since we believe this is the approximate capacity of the muscle (i.e., the muscle swells but does not “leak,” whereas volumes greater than 50 ML tend to “leak” from the muscle). 6. When injecting two plasmids (usually a reporter plasmid plus an expression plasmid or the respective empty vector) to ensure that the total volume injected remains at 50 ML, prepare the promoter reporter plasmid stock at double the concentration, or 2.0 Mg/ML. Prior to analyses of expression plasmids on gene promoter reporters, in vivo, dose-response analyses are necessary to determine the amount of expression plasmid needed to obtain optimal levels of protein overexpression. Levels of protein overexpression should be within physiologically attainable ranges to prevent toxic or off-target effects. We typically find that anywhere from 5–40 Mg of expression plasmid is sufficient to optimally increase our protein of interest. However, when the size of the plasmid and size of the expressed protein are large, greater amounts of plasmid DNA may be necessary to optimally express the protein of interest. If injecting only an expression plasmid into a soleus, and the desired amount is 10 Mg in 50 ML 1× PBS, the plasmid would be concentrated to 0.2 Mg/ML. Therefore, if coinjecting an expression plasmid plus a reporter plasmid, prepare the expression plasmid (and empty vector) at double the concentration (0.4 Mg/ML). Add equal amounts of promoter reporter plasmid stock (2.0 Mg/ML in this case – i.e., double the concentration that would be injected if only injecting the promoter reporter) and expression plasmid (or empty vector) stock (0.4 Mg/ML) to obtain a final injection mixture containing 1.0 Mg/ML of promoter reporter and 0.2 Mg/ML of expression or control plasmid. Keeping injected plasmid DNA concentrations and volume amount identical between each soleus muscle is critical to obtain consistent reporter-driven luciferase measurements that are comparable across muscles. Furthermore, although cotransfection of renilla luciferase constructs in cell culture is routinely used to normalize for transfection efficiency and differences in cell numbers between wells, the use of these constructs in skeletal muscle, in vivo,

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has clearly been shown to correlate poorly with the activity of firefly luciferase promoter constructs (8). Therefore, in our in vivo experiments we do not normalize our firefly luciferase gene promoter reporter data to renilla luciferase. 7. Aside from the soleus muscle, the tibialis anterior (TA) muscle is easily accessible and can be injected with gene promoter reporter plasmids, similar to the soleus. TA isolation and plasmid DNA injection have previously been detailed (5). 8. It is essential to ensure that the electroporation parameters selected do not cause muscle damage. Therefore, cross-sections should be taken from various parts along the longitudinal axis of a muscle that has been injected and electroporated, and these sections stained with hematoxylin and eosin to check for muscle damage and inflammatory cell infiltration. 9. Luciferase expression has been detected as soon as 2 min following injection of a CMV promoter-driven luciferase plasmid. Luciferase expression peaked at 7–14 days postinjection and remained significantly increased 120 days postinjection (9). Therefore, although we remove muscles 7–10 days following injection of a gene promoter reporter plasmid in our work, a much broader time frame could be studied. 10. The whole muscle is used as the whole muscle was injected with the promoter reporter plasmid. The same homogenizing volume is used for all muscles as each soleus muscle was injected with an equal amount of promoter reporter plasmid in a standardized volume of PBS, and the soleus muscles of rats of the same age and weight are approximately the same muscle mass and contain approximately equal number of muscle fibers at the time of injection. We homogenize in PLB, since this buffer contains an antifoam agent to prevent excessive bubbling of the sample, which otherwise occurs during the muscle homogenization process. The presence of bubbles in the sample may contribute to inconsistencies in the detection of light output and, therefore, luciferase measurements. 11. Luciferase assay reagent should be at room temperature for luciferase measurements. The linear dynamic range of the luminometer to be used for measuring luciferase activity should be determined prior to determining sample luciferase activity. The dynamic range extends from the minimal detectable concentration to the concentration in which the detector no longer responds to an increase. The linear dynamic range is important for quantitative luciferase measurements and refers to the concentration range over which the detector responds in a linear fashion to increased substrate.

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Fig. 2. Representative gene (atrogin-1) data collected in muscle extracts from weight bearing and cast immobilized rats. The increase in atrogin-1 mRNA expression (a) during cast immobilization is closely mimicked by an increase in luciferase activity of an atrogin-1 promoter reporter plasmid during the same conditions, as measured using a luciferase assay system (b). Immunostaining for luciferase protein in soleus muscle cross-sections, although not quantitative, also shows increased expression following cast immobilization (c, d). Muscle cross-sections were fixed in 4% paraformaldehyde, incubated with anti-luciferase (1:1,000; Sigma) followed by Alexa Fluor 488 (Invitrogen) fluorescent dye-conjugated secondary antibody, and visualized with fluorescence microscopy.

12. The distinct advantage of in vivo bioluminescent imaging is the capability to collect repeated measures within the same animal. However, this imaging requires specialized equipment, such as the IVIS system from Xenogen. Animals imaged using this system are anesthetized, placed on a platform and injected with D-luciferin. After a set period of time following this injection, the luminescence emitted from the muscles of animals is integrated for a set period of time and acquired. The pseudocolored images captured represent the luminescent signal, are overlayed on a photographic image of the animal, and are analyzed with appropriate software. More detailed methods of this procedure and representative images can be found in several publications (10–12). Immunohistochemical analysis of luciferase expression (see Fig. 2) is mostly used for confirmation of luciferase activity measurements (13), but also provides the potential advantage that serial sections can be taken from the muscle for fiber typing or alternative histological analysis that can then be associated with, or compared to, luciferase expression.

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Acknowledgment This work was supported by National Institute of Arthritis and Musculoskeletal and Skin Diseases Grant AR056418. References 1. Schertzer JD, and Lynch, GS (2008) Plasmidbased gene transfer in mouse skeletal muscle by electroporation. Methods Mol Biol 433:115–125. 2. Kitsis RN, and Leinwand, LA (1992) Discordance between gene regulation in vitro and in vivo. Gene Expr 2:313–318. 3. Hiraoka K, Koike, H, Yamamoto, S et al (2003) Enhanced therapeutic angiogenesis by cotransfection of prostacyclin synthase gene or optimization of intramuscular injection of naked plasmid DNA. Circulation 108:2689–2696. 4. Mitchell-Felton H, and Kandarian, SC (1999) Normalization of muscle plasmid uptake by Southern blot: application to SERCA1 promoter analysis. Am J Physiol 277:C1269–1276. 5. Manthorpe M, Cornefert-Jensen, F, Hartikka, J et al (1993) Gene therapy by intramuscular injection of plasmid DNA: studies on firefly luciferase gene expression in mice. Hum Gene Ther 4:419–431. 6. Kang JH, and Chung, JK (2008) Moleculargenetic imaging based on reporter gene expression. J Nucl Med 49 Suppl 2:164S–179S. 7. Akimoto T, Sorg, BS, and Yan, Z (2004) Realtime imaging of peroxisome proliferator-activated receptor-gamma coactivator-1alpha promoter activity in skeletal muscles of living mice. Am J Physiol Cell Physiol 287:C790–796.

8. Bao S, Liu, MJ, Lee, B et al (2010) Zinc modulates the innate immune response in vivo to polymicrobial sepsis through regulation of NF-kappaB. Am J Physiol Lung Cell Mol Physiol 298:L744–754. 9. Acharyya S, Villalta, SA, Bakkar, N et al (2007) Interplay of IKK/NF-kappaB signaling in macrophages and myofibers promotes muscle degeneration in Duchenne muscular dystrophy. J Clin Invest 117:889–901. 10. Dodd SL, Hain, B, Senf, SM et al (2009) Hsp27 inhibits IKKbeta-induced NF-kappaB activity and skeletal muscle atrophy. FASEB J 23:3415–3423. 11. Senf SM, Dodd, SL, and Judge, AR (2009) FOXO signaling is required for disuse muscle atrophy and is directly regulated by Hsp70. Am J Physiol Cell Physiol 298:C38–45. 12. Senf SM, Dodd, SL, McClung, JM et al (2008) Hsp70 overexpression inhibits NF-kappaB and Foxo3a transcriptional activities and prevents skeletal muscle atrophy. FASEB J 22: 3836–3845. 13. Mitchell-Felton H, Hunter, RB, Stevenson, EJ et al (2000) Identification of weight-bearingresponsive elements in the skeletal muscle sarco(endo)plasmic reticulum Ca2+ ATPase (SERCA1) gene. J Biol Chem 275:23005– 23011.

Part VIII RNA-Mediated Gene Regulation

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Chapter 28 Determination of MiRNA Targets in Skeletal Muscle Cells Zhan-Peng Huang, Ramón Espinoza-Lewis, and Da-Zhi Wang Abstract MicroRNAs (miRNAs) are a class of small ~22 nucleotide noncoding RNAs which regulate gene expression at the posttranscriptional level by either destabilizing and consequently degrading their targeted mRNAs or by repressing their translation. Emerging evidence has demonstrated that miRNAs are essential for normal mammalian development, homeostasis, and many other functions. In addition, deleterious changes in miRNA expression were associated with human diseases. Several muscle-specific miRNAs, including miR-1, miR-133, miR-206, and miR-208, have been shown to be important for normal myoblast differentiation, proliferation, and muscle remodeling in response to stress. They have also been implicated in various cardiac and skeletal muscular diseases. miRNA-based gene therapies hold great potential for the treatment of cardiac and skeletal muscle diseases. Herein, we describe methods commonly applied to study the biological role of miRNAs, as well as techniques utilized to manipulate miRNA expression and to investigate their target regulation. Key words: MicroRNA, miRNA, Muscle, Gene expression, Posttranscriptional regulation, Muscle disease

1. Introduction Formation, development, and physiology of skeletal muscle are of the utmost importance for the normal locomotion of an organism. Abnormal development, damage, or deterioration of skeletal muscle might result in muscle atrophy, paralysis, or even death. Skeletal muscle disorders are a group of diseases caused by different mechanisms, including defects in structural proteins, disorganization of the sarcomeres, and/or perturbed regulation of growth/maturation signaling pathways (1). These diseases can be classified as: 1) Neuromuscular, such as multiple sclerosis

Joseph X. DiMario (ed.), Myogenesis: Methods and Protocols, Methods in Molecular Biology, vol. 798, DOI 10.1007/978-1-61779-343-1_28, © Springer Science+Business Media, LLC 2012

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or 2) musculoskeletal, such as Duchenne Muscular Dystrophy, myotubular myopathy, and others. Elucidation of mechanisms that regulate muscle determination, differentiation, and proliferation is an important prerequisite to developing therapeutic strategies to correct or to circumvent skeletal muscle defects that accompany neuromuscular disease. Recently, a large class of small ~22 nucleotide (nt) noncoding RNAs have been discovered and are collectively referred to as microRNAs (miRNAs). To date, thousands of miRNA genes have been identified in multiple organisms from plants and nematodes, to fish and mammals (2). Similar to protein-encoding genes, the expression of a miRNA begins with the transcription of the miRNA gene by RNA polymerase II. Though many miRNAs are under the control of their own promoter, some miRNA genes are found in clusters sharing a single promoter (3); others are encoded within an intron and are coexpressed along with the host gene (4). After transcription within the nucleus, the large primary miRNA transcript is processed by the microprocessor complex (Drosha/DGCR8) into a hairpin intermediate commonly referred to as a pre-miRNA. However, a small subgroup of miRNAs found within short introns is known to bypass this step (5). Pre-miRNAs are then exported from the nucleus to the cytoplasm by the nuclear transporter exportin-5 (6); in the cytoplasm, they are further processed into miRNA duplexes by the cytosolic RNase III enzyme Dicer (7). Finally, the functional strand of the miRNA duplex is loaded into the RNA-induced silencing complex (RISC) to facilitate targeted mRNA degradation and/or translational repression (8). Evolutionarily conserved miRNAs have been identified in multiple eukaryotes from the worm Caenorhabditis elegans, to the fruit fly Drosophila melanogaster, to the mouse Mus musculus, and to the human Homo sapiens. The C. elegans genome contains a single mammalian miR-1 ortholog (9), whereas in higher eukaryotes there are multiple genes encoding miR-1 (identical coding sequences of miR-1-1 and miR-1-2). miR-206, expressed specifically in skeletal muscles, is related to miR-1 and differs from miR-1 by only four nucleotides (10). Several mammalian miRNAs, including the miR1/206 and miR-133 families, and miR-208a/b, are specifically expressed in cardiac and skeletal muscle (11–13). miR-1 is known to regulate skeletal muscle differentiation and proliferation in C2C12 myoblasts (12) as well as the neuromuscular junction in C. elegans (14). In addition, miR-1 expression is dependent upon the activity of the transcription factors Serum Response Factor (SRF) and MyoD, as evidenced in both D. melanogaster (15) and M. musculus (11). Together, these results strongly suggest that both the regulation and the function of miR-1 are conserved throughout eukaryotic evolution, and miR-1 plays an important role in several

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processes, such as cell proliferation, differentiation, migration, and apoptosis during both normal development and disease progression (16, 17). Intriguingly, even though both miR-1 and miR-133 genes are clustered together and cotranscribed as a single primary transcript, they represent two distinct miRNAs, each with its own biological function (12). miR-1 overexpression has been shown to promote skeletal muscle myoblast differentiation in cultured C2C12 myoblasts (12). miR-1 also significantly impairs normal cardiac development (11, 12), inducing arrhythmias through negative regulation of Kcnj2 and Gja1 (18). Conversely, miR-133 induces cell proliferation and represses myogenic gene expression (12). miR-208 is specifically expressed in the myocardium and is required for stress-dependent cardiac growth and remodeling (13, 19). miR-206 is uniquely expressed in skeletal muscle cells (20); although its functions are not fully understood, they include a potential role in muscular hypertrophy, maintaining the ratio between DMHC and EMHC through regulating the activity of the retinoic acid receptor alpha (RXRD), a potential role in satellite cell specification through the regulation of Pax3, and a potential role in the switching of the fiber types by downregulating Utrophin, which could compensate for the loss of dystrophin in Duchenne muscular dystrophy syndrome (20). The expression of many miRNAs is altered under pathological conditions. Subsets of miRNAs are found to be both positively and negatively regulated in clinical human samples and animal models of cardiac and skeletal myopathies (21–24). In vivo overexpression of miR-195 in cardiomyocytes is sufficient to cause dilated cardiomyopathy and heart failure in the mouse (22). In addition, dystrophin-deficient mice were found to have significantly decreased expression of miR-133a and miR-206 (25). Together, these results indicate that proper expression of miRNAs is necessary for both normal development and function of skeletal and cardiac muscles. Strategies commonly used to investigate the biological function of a particular miRNA include both gain-of-function and loss-offunction approaches. Gain-of-function studies are usually performed in vitro, where cells can be transiently transfected with an expression construct encoding the pre-miRNA. Alternatively, synthetic miRNA duplexes and virus-based miRNA expression systems may also be employed. In vitro loss-of-function studies can be accomplished with either 2c-O-methyl miRNA antisense oligonucleotides or locked nucleic acid (LNA)-miRNA antisense oligonucleotides, which will block the function of an endogenous miRNA. The in vivo determination of a miRNA’s function is best examined utilizing conventional transgenic and gene knockout strategies. Recently, a lentivirus targeting strategy that overexpresses short RNA fragments

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containing multiple miRNA target sequences has been shown to phenocopy a genetic miRNA knockout mouse (26). In addition, intravenous delivery of cholesterol-modified miRNA antisense oligonucleotides (antagomiRs) can inhibit miRNA function in vivo (27). These molecular approaches are invaluable in elucidating the biological function of miRNAs and may potentially lend themselves to future gene-based therapies. A key to the understanding of the molecular mechanism of miRNA function is to identify miRNA targets. miRNAs are known to repress their targets primarily by targeting the 3c UTRs of their target transcripts. However, identification of such targets has proved to be challenging in animals, primarily due to imperfect sequence match between miRNAs and their regulatory targets. Information technologies and bioinformatics databases are very useful tools to identify putative miRNA targets. In particular, computational algorithms for miRNA targets are publicly available through the world wide web (i.e., Pictar (28), miRanda (29), TargetScan (30), etc.). These algorithms allow investigators to search for possible miRNA target sites in the 3c UTR of a candidate mRNA, or to predict possible regulatory targets of a specific miRNA. To date, it has been recognized that a single miRNA could target multiple mRNA transcripts (31, 32). On the other hand, the 3c UTR of a gene might have multiple target sites for different miRNAs. Thus, miRNAs offer themselves as one additional layer in the posttranscriptional regulation of gene expression. Microarray technology is most commonly utilized for the basic purpose of comparing mRNA expression levels between two or more samples (i.e., dystrophic muscle vs. normal muscle) (21). The results are obtained in terms of expression folds either for upregulated or down-regulated genes. These up- or down-regulated genes are of most interest since they are the ones showing a dynamic expression. The results from microarray analyses are available through databases (i.e., NCBI database) from which one can extract the specifics for a gene. A further step in this technology is that of miRNA microarrays; these provide results for the up- or down-regulation of miRNAs in the compared samples. It is conceivable that a combination of conventional mRNA microarray and a miRNA microarray on the same sets of samples could be a powerful approach and will allow us to determine the correlation of miRNAs and their regulatory targets. In this chapter, we will first describe how to document the expression of miRNAs by northern blot and qPCR analyses. We will then describe the method to define the regulatory targets of a miRNA using luciferase reporter assays. Finally, we will detail how to manipulate the expression level of muscle miRNAs in the C2C12 myoblast cell line and how to determine their biological function in muscle proliferation and differentiation.

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2. Materials 2.1. Detecting the Expression of Muscle miRNAs by Northern Blotting and Quantitative RT-PCR Analyses 2.1.1. Northern Blot

1. Hoefer SE 400 vertical slab gel electrophoresis unit. 2. Hoefer TE77 Semidry transfer unit. 3. UV stratalinker 1800 (Stratagene). 4. Trizol Reagent (Invitrogen). 5. 40% Acrylamide: AccuGel 29:1 (National Diagnostics). 6. 10× TBE buffer: 0.9 M Tris base, 0.9 M boric acid, 0.02 M EDTA (pH 8.0), autoclave for 20 min. 7. Urea (molecular biology grade). 8. 10% (w/v) Ammonium persulfate solution (APS). Aliquot and store at −20ºC. 9. N,N,N,Nc-Tetramethyl-ethylenediamine (TEMED). 10. Formamide. 11. Bromophenol blue solution: 10% (w/v) bromophenol blue. 12. Filter paper, sheet, grade 3, 460 × 570 MM (Whatman). 13. Zeta-Probe GT genomic tested blotting membranes (Bio-Rad). 14. T4 polynucleotide kinase (PNK). 15. Mini Quick Spin Oligo Columns (Roche). 16. Adenosine 5c-triphosphate [J-32P], 3,000 Ci/mmol. 17. Anti-miRNA probe: the synthetic antisense oligonucleotide of the target miRNA. 18. Diethylpyrocarbonate (DEPC)-treated water: 1 mL DEPC in 1 L double-distilled H2O. Stir at room temperature for 1 h and autoclave. 19. 1 M phosphate buffer: 71 g of anhydrous Na2HPO4, 4 mL of 85% H3PO4. Add DEPC-treated water to 1 L. 20. Hybridization buffer: 0.5 M phosphate buffer, 1 mM EDTA at pH 8.0, 7% (w/v) of sodium dodecyl sulfate (SDS), 1% (w/v) of bovine serum albumin (BSA), in DEPC-treated water. 21. 20 × SSC: 3 M sodium chloride and 300 mM tri-sodium citrate dihydrate, pH 7.0. 22. Wash buffer: 1× SSC supplemented with 0.1% SDS. 23. Stripping buffer: 0.1× SSC supplemented with 0.1% SDS. 24. Storage phosphor screen (Amersham).

2.1.2. Quantitative RT-PCR

1. TaqMan MicroRNA Reverse Transcription Kit (Applied Biosystems). 2. TaqMan Universal PCR Master Mix, No AmpErase UNG (Applied Biosystems). 3. TaqMan MicroRNA Assays Kit (Applied Biosystems).

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2.2. Studying the Regulation of Muscle miRNAs on Their Targets by Luciferase Reporter Assays

1. Mouse genomic DNA. 2. Primers for reporter construction: HDAC4-UTR-F, 5c-ATCGGAGCTCCAGCACTGGTGATAGACTTGG-3c; HDAC4-UTR-R, 5c-GTCTTATTGAACTTATTCTTAAGC TCGAGATCG-3c; HDAC4-Mut-F, 5c-GTTTCTTTCCT CAGATTTAAAATTCTTCACTGGTCACAGCCACG-3c; HDAC4-Mut-R, 5c-GTGACCAGTGAAGAATTTTAAATCT GAGGAAAGAAACACAACC-3c. 3. PfuTurbo DNA polymerase. 4. pGL3cM vector (modified by Chen JF and Wang DZ, the backbone is the pGL3-Control vector, Promega). 5. SacI restriction endonuclease. 6. XhoI restriction endonuclease. 7. T4 DNA ligase. 8. pRL-TK Vector for Renilla luciferase reporter (Promega). 9. NucleoBond plasmid Maxi kit (Macherey-Nagel). 10. HEK293T cells (ATCC). 11. CELLSTAR 12- and 24-well tissue culture plate. 12. Growth medium for HEK293T cells: Combine 1 L of 1× Dulbecco’s modified Eagle medium (DMEM, high glucose with L-glutamine), 110 mL of fetal bovine serum (FBS), and 11 mL of 100× penicillin G–streptomycin (10,000 units penicillin; 10,000 Pg streptomycin). 13. 1× Trypsin–EDTA: 0.25% Trypsin, 1 mM EDTA/4Na. 14. Lipofectamine LTX and Plus Reagent (Invitrogen). 15. Opti-MEM I Reduced Serum Medium (Gibco). 16. miR-1 miRIDIAN miRNA mimic (Dharmacon). 17. 10× Phosphate-buffered saline (PBS) solution: 80.6 mM sodium phosphate, 19.4 mM potassium phosphate, 27 mM KCl and 1.37 M NaCl at pH 7.4. 18. Dual-luciferase reporter assay system (Promega).

2.3. Overexpression and Knockdown of Muscle miRNAs in Cell Lines

1. Primers for miR-22 overexpression vector construction: miR22-F 5c-TAGCAGGTACCTTATTCAAGAACCCCTCA TTAG-3c, miR22-R 5c-GTATCTCTAGATTTCCCTCCCA TAAAGCCAT-3c. 2. pcDNA3.1(+) vector (Invitrogen). 3. anti-miR-22 probe: antisense oligonucleotide to miR-22. 4. C2C12 cells (ATCC). 5. KpnI restriction endonuclease. 6. XbaI restriction endonuclease. 7. 2c-O-methyl miR-133 antisense oligonucleotide (Dharmacon).

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8. Growth medium for C2C12 cells: DMEM medium with 10% FBS and 1% penicillin G–streptomycin. 9. Differentiation medium for C2C12 cells: DMEM medium with 2% horse serum and 1% penicillin G–streptomycin. 10. Complete Protease Inhibitor Cocktail Tablets (Roche). 11. Cell lysis buffer: 40 mM Tris–HCl (pH7.5), 150 mM NaCl, 1% (v/v) Triton ×-100, Complete Protease Inhibitor Cocktail Tablet (1 tablet/50 mL). 12. Mini-PROTEAN 3 Electrophoresis Cell (Bio-Rad). 13. 1× SDS–PAGE running buffer: 25 mM Tris, 200 mM glycine; 0.1% (w/v) SDS. 14. 5× protein loading buffer: 10% (w/v) SDS, 10 mM beta-mercapto-ethanol, 20% (v/v) glycerol, 0.2 M Tris–HCl (pH6.8), 0.05% (w/v) bromophenol blue. 15. PVDF membrane. 16. Transfer buffer: 25 mM Tris, 200 mM glycine, 20% (v/v) methanol. 17. Odyssey blocking buffer (LI-COR Biosciences). 18. Anti-SRF antibody from rabbit (Santa Cruz Biotechnology). 19. IRDye goat-anti-rabbit secondary antibody (LI-COR Biosciences).

3. Methods 3.1. Detecting the Expression of Muscle miRNAs by Northern Blot and Quantitative RT-PCR Analyses

1. Prepare total RNA from tissue or cultured cells with Trizol Reagent according to manufacturer’s protocol (see Note 1). 2. Prepare 15% denaturing gel for electrophoresis separation of miRNAs. Carefully wash, dry, and assemble the Hoefer SE 400 vertical slab gel electrophoresis unit. Prepare denaturing gel containing 18.75 mL of 40% acrylamide, 2.5 mL of 10× TBE buffer, 12.5 mL of DEPC-treated water, and 20 g of urea. Mixture may need to be gently heated in 37°C water bath in order for urea to completely dissolve. To polymerize, add 400 PL of 10% APS; 40 PL of TEMED, mix well, and quickly pour. Allow the gel to polymerize for 1 h. 3. Prerun denaturing gel for 30 min at 200 V. Use 0.5× TBE for running buffer. 4. Prepare RNA samples for electrophoresis. Mix the RNA sample (40 Pg) 1:1 (v/v) with formamide, and incubate at 65°C for 10 min. Chill RNA on ice for 3 min and add 2 PL of bromophenol blue solution. Mix well.

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5. Load the sample(s) into the well(s) and run the gel at 250 V (see Note 2). Use 0.5× TBE for running buffer. Voltage can be stopped when the loading dye reaches the bottom of the plate. 6. Transfer the RNA from the gel to a membrane with Hoefer TE77 Semidry transfer unit. Soak the membrane and six pieces of filter paper in 0.5× TBE. Set up the transfer in the order from top (−) to bottom (+) as: three pieces of filter paper, gel, membrane, three pieces of filter paper (see Note 3). Transfer with constant current (0.8 mA/cm2 of gel area) for 1 h. 7. After transfer, wash the membrane with 0.5× TBE and perform UV crosslink using the auto crosslink option. 8. Prepare isotope-labeled probe for hybridization. Mix 5 PL of adenosine 5c-triphosphate [J-32P], 5 PL of 1 PM anti-miRNA probe, 2 PL of 10× PNK buffer, 1 PL of T4 polynucleotide kinase, and 7 PL of double-distilled water and incubate at 37°C for 1 h. 9. Purify the [J-32P]-labeled probe using a mini Quick Spin Oligo Column according to manufacturer’s protocol (see Note 4). 10. Prehybridize the membrane for 1 h at 37°C with 5–10 mL of hybridization buffer. 11. Add the labeled anti-miRNA probe into the hybridization buffer and incubate overnight at 37°C. 12. Remove the hybridization buffer and wash the membrane 3 times with wash buffer (10 min per wash). 13. Expose the membrane to the storage phosphor screen for 4–24 h. The length of exposure depends upon strength of signal and will vary with different miRNA probes (see Note 5). 14. Scan the screen with Typhoon phosphor-imager (see Figs. 1a, 3b).

Fig. 1. Determination of miR-1 expression during C2C12 myoblast cell differentiation by (a) northern blotting analyses and (b) qPCR assays. Total RNAs isolated from C2C12 myoblasts which are switched into differentiation conditions at indicated time points (day-0 (D0) to day-7 (D7)) were used for northern blot and qPCR according to the protocols described in this chapter. U6 snRNA serves as a loading control.

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15. If you want to probe the membrane with a different miRNA probe or if you want to store the membrane for long term, add the membrane to heated stripping buffer (>95°C). Incubate for approximately 10 min while rocking. 16. After stripping, rinse membrane with fresh stripping buffer, and allow drying. The membrane may be reprobed immediately following the steps outlined above. The membrane can also be stored at −20°C for future use. 17. To measure the expression of muscle miRNAs by quantitative RT-PCR, dilute total RNA sample to 2 ng/PL and perform reverse transcription. Mix the following reagents, including 0.15 PL 100 mM dNTPs, 1 PL MultiScribe reverse transcriptase, 1.5 PL 10× reverse transcription buffer, 0.2 PL RNase inhibitor, 4.15 PL nuclease-free water, 5 PL diluted RNA sample, and 3 PL RT primer (see Note 6), into a 0.2 mL polypropylene PCR tube on ice. Briefly centrifuge the mixture and incubate the tube on ice for 5 min. Perform reverse transcription in thermal cycler with the following program: 16°C for 30 min Ȣ 42°C for 30 min Ȣ 85°C for 5 min Ȣ hold at 4°C. The cDNA can be used immediately or store in −20°C for further use. 18. Perform quantitative PCR by mixing the following reagents: 1 PL 20× TaqMan MicroRNA Assay (a mixture contains both PCR primer pair and TaqMan probe for the specific miRNA), 1.5 PL product from RT reaction, 10 PL TaqMan 2× universal PCR master mix (no AmpErase UNG), and 7.5 PL nucleasefree water, into a 0.2 mL quantitative PCR tube on ice. Briefly centrifuge the mixture and perform quantitative PCR in realtime PCR system with the default program for TaqMan quantitative PCR (see Fig. 1b). 3.2. Studying the Regulation of Muscle miRNAs on Their Targets by Luciferase Reporter Assays

Here we show an example using the luciferase reporter vectors which contains either the wild-type or the mutant HDAC4 3c UTR. 1. Generate the ~400 bp HDAC4 gene 3c UTR DNA fragment containing the seed sequence for miR-1 by PCR reaction using mouse genomic DNA as the template and the HDAC4-UTR-F and HDAC4-UTR-R primers. The SacI and XhoI sites are introduced at the 5c and 3c-ends, respectively, by the PCR primers. The UTR PCR products are cloned into the SacI/XhoI sites of the pGL3cM vector (see Note 7). The resulting LucWT-UTR reporter contains the wild-type 3c UTR of the HDAC4 gene (see Fig. 2a, b). 2. Generate the Luc-Mut-UTR reporter by using the plasmid generated in step 1 as a template and introducing mutations with HDAC4-Mut-F and HDAC4-Mut-R primers (see Fig. 2b).

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Fig. 2. Luciferase reporter assays of miR-1 and HDAC4-3c UTR. (a) Graphic representation of the modified luciferase vector containing the 3c UTR of the HDAC4 gene. (b) Sequence comparison among HDAC4 wild-type 3c UTR, HDAC4 mutant 3c UTR sequence, and miR-1. (c) miR-1 significantly represses the luciferase reporter activity of HDAC4 wild-type 3c UTR, but not that of mutant 3c UTR.

3. Prepare high-quality Luc-WT-UTR reporter, Luc-Mut-UTR reporter, and pRL-TK plasmid for reporter assays with NucleoBond Plasmid Maxi Kit. These plasmids will be used for HEK293T cell transfection. 4. At 1 day before transfection, plate HEK293T cells in a 24-well plate at 5 × 104 cells per well in 500 PL of growth medium (see Note 8). This will yield 50–80% confluence at the day of transfection. 5. To generate the transfection complex for one well, add 25 ng reporter (either Luc-WT-UTR or Luc-Mut-UTR), 25 ng pRLTK plasmid, and 0.5 PL of 10 PM miRIDIAN miRNA mimic to 100 PL of Opti-MEM I Reduced Serum Medium and mix gently. To this mixture, add 0.5 PL of PLUS Reagent, mix gently, and incubate for 5–10 min at room temperature. Finally, add 1.25 PL of Lipofectamine LTX Reagent, mix gently, and incubate for 30 min at room temperature.

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6. Add the transfection complex (~100 PL) to the well. Mix gently by rocking the plate back and forth. 7. At 24 h after transfection, remove cell culture medium and wash cells twice with 1× PBS. 8. Lyse cells with 100 PL of 1× passive lysis buffer by gentle shaking at room temperature for 15 min. 9. Mix 20 PL of cell lysate with 50 PL of firefly luciferase substrate and measure the firefly luciferase activity with a scintillation counter. 10. Add 50 PL Stop&Glo reagent into the mixture from step 9 and measure the Renilla luciferase activity with a scintillation counter. 11. Normalize firefly luciferase activity with Renilla luciferase activity and plot results (see Fig. 2c). 3.3. Overexpression and Knockdown of Muscle miRNAs in Cell Lines

In this section, we describe steps to overexpress miR-22 in HEK293T cells and to knockdown miR-133 in C2C12 myoblasts. 1. For the overexpression study, use PCR to generate a ~350 bp DNA fragment containing the intact hairpin for the miR-22 precursor plus the flanking sequences on both ends (see Note 9). Use mouse genomic DNA as the template. The KpnI and XbaI sites are introduced at the 5c and 3c-ends, respectively, by the PCR primers. Clone the PCR product into the KpnI/XbaI sites in the pcDNA3.1(+) vector (see Note 10). The resulting construct is termed the miR-22 overexpression vector (see Fig. 3a). 2. Prepare high-quality plasmid for transfection with NucleoBond Plasmid Maxi Kit. 3. Transfect the miR-22 overexpression vector into HEK293T cells following the steps outlined in Subheading 3.2. Use 6-well plates for the transfection and adjust the amount of transfection reagents accordingly. 4. Extract total RNA from cells 48 h after transfection using the Trizol reagent according to manufacturer’s protocol. 5. Evaluate the overexpression of miR-22 by northern blot according to the protocol described in Subheading 3.1 (see Fig. 3b). Similarly, the miR-22 overexpression vector can also be evaluated in other cells such as C2C12 myoblasts. 6. For miR-133 knockdown study, plate C2C12 myoblasts in a 6-well plate at 2 × 105 cells per well in 2 mL of growth medium 1 day before transfection. This will yield 50–80% confluence at the day of transfection. 7. Transfect C2C12 myoblasts with 200 nM 2c-O-methyl miR133 antisense oligonucleotides (see Note 11). Adjust the amount of transfection reagents accordingly.

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Fig. 3. Overexpression and knockdown of miRNAs in C2C12 myoblast cells. (a) The strategy for the construction of miR-22 overexpression vector. P miRNA hairpin precursor; F flanking sequences. (b) Verification of miR-22 overexpression in HEK293T cells by northern blot analyses. Lane 1, cells transfected with the miR-22 overexpression vector; Lane 2, cells transfected with a control vector. U6 snRNA serves as a loading control. (c) Western blot showing SRF protein expression repressed by miR-133. Lane 3, C2C12 cells transfected with control 2c-O-methyl oligonucleotide; Lane 4, C2C12 cells transfected with 2c-O-methyl miR-133 antisense oligonucleotide. E-tubulin serves as a loading control.

8. Change growth medium 4–6 h after transfection and continue to culture the cells for an additional 24 h. 9. 24 h after transfection, replace growth medium with differentiation medium and culture the cells for an additional 12 h. 10. Confirm miR-133 knockdown by northern blotting analysis according to the protocol described in Subheading 3.1. 11. Prepare cell lysate with cell lysis buffer (100 PL per well) and examine the up-regulation of SRF, a target regulated by miR133 (12), by western blot analysis. 12. Prepare SDS–PAGE gel for electrophoresis separation of proteins. Carefully wash, dry, and assemble the Bio-Rad MiniPROTEAN 3 Electrophoresis Cell. Prepare 9% running gel containing 2.4 mL of 30% acrylamide, 2 mL of 1.5 M Tris– HCl (pH8.8), 3.5 mL of double-distilled water, and 80 PL of 10% SDS. To polymerize, add 40 PL of 10% APS, 5.5 PL of

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TEMED, mix well, and quickly pour. Allow the gel polymerize for 30 min. Then, prepare stacking gel containing 0.53 mL of 30% acrylamide, 0.5 mL of 1 M Tris–HCl (pH 6.8), 2.97 mL of double-distilled water, and 40 PL of 10% SDS. To polymerize, add 20 PL of 10% APS, 4 PL of TEMED, mix well, and quickly pour on the top of the polymerized running gel. Allow the gel polymerize for 30 min. 13. Prepare protein samples for SDS–PAGE electrophoresis. Mix the cell lysis sample (50 Pg) with 5× protein loading buffer, and incubate at 95°C for 5 min. 14. Load the sample(s) into the well(s) and run the gel at 100 V. Use 1× running buffer. Voltage can be stopped when the loading dye reaches the bottom of the plate. 15. Transfer the protein from the gel to the PVDF membrane with 1× transfer buffer. Wet the PVDF membrane with methanol. Soak the PVDF membrane, gel, and six pieces of filter papers in 1× transfer buffer for 10 min. Set up the transfer in the order from cathode (−) to anode (+) as: three pieces of filter paper, gel, membrane, three pieces of filter paper (see Note 12). Transfer at either 100 V for 3 h or 30 V overnight at 4°C. 16. After transfer, incubate the membrane with Odyssey blocking buffer for 1 h at room temperature. 17. Dilute SRF first antibody with 1:500 dilution into Odyssey blocking buffer and incubate the membrane with diluted first antibody overnight. 18. Wash the membrane with 1× PBS 3 times at room temperature (15 min per wash). 19. Dilute IRDye goat-anti-rabbit secondary antibody with 1:7,500 dilution into Odyssey blocking buffer and incubate the membrane with diluted second antibody for 1 h at room temperature. 20. Wash the membrane with 1× PBS 3 times at room temperature (15 min per wash). 21. Scan the membrane with the Odyssey infrared imaging system (see Fig. 3c).

4. Notes 1. RNase(s) rapidly degrade RNA and are abundant in the environment. When extracting total RNA from samples, RNasefree tubes, DEPC-treated water, and solutions made with DEPC-treated water are highly recommended. RNA samples can be preserved in pellet for more than 1 year if stored in 100% ethanol at −80°C.

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2. Prior to loading the RNA sample into the denaturing gel, wash the well by flushing with 0.5 × TBE running buffer. Excess urea in the well will prevent the RNA sample from sinking to the bottom of the well. 3. Exclude air bubbles when assembling the “sandwich” for RNA transfer. 4. It is important to have enough protection when conducting the isotope-related experiments. Always wear personal protective equipment when handling radioisotopes. 5. Besides the phosphor-imager system, northern blot can also be imaged with X-ray autography. In general, the membrane needs to be exposed to film for 1 day to 1 week. 6. Up to five different miRNA RT primers can be added in one reverse transcription reaction. In this case, the total volume of all the RT primers should be 3 PL for one reaction. It is noticed that some combinations of miRNA RT primers may not work well. Test the combination before performing experiments. 7. To generate the pGL3cM vector, the multiple cloning site (MCS) is removed from pGL3-control vector by KpnI/BglII digestion and filled in by Klenow. The 53 bp oligonucleotide containing the MCS is then introduced into the XbaI site. 8. At least 12 wells are needed for one experiment to examine four combinations of transfection reagents including Luc-WTUTR reporter and miR-1 miRIDIAN mimic, Luc-WT-UTR reporter and control miRIDIAN mimic, Luc-Mut-UTR reporter and miR-1 miRIDIAN mimic, and Luc-Mut-UTR reporter and control mimic. Each combination of transfection reagents is performed in triplicate. 9. Different cloning strategies can be applied to generate a miRNA overexpression vector. In this protocol, our strategy is to clone the fragment containing the whole hairpin (miRNA precursor) and a 100–150 bp flanking sequence on both the 5c and 3c ends of the miRNA sequence. Alternatively, the fulllength noncoding transcript can be cloned into the expression vector. However, this is only applicable for miRNAs generated from a nonprotein-coding gene. 10. Besides pcDNA3.1(+), other expression vectors can be used to construct a miRNA overexpression plasmid. Virus-based expression vectors have already been reported for miRNA overexpression (33). 11. In this protocol, a 2c-O-methyl miRNA antisense oligonucleotide is used to knockdown the endogenous miRNA. Alternatively, LNA antisense oligonucleotides can be used to obtain similar effects. 12. Exclude air bubbles when assembling the “sandwich” for protein transfer.

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Acknowledgments We thank members of the Wang laboratory for discussion and support. Research in the Wang lab was supported by the March of Dimes Birth Defect Foundation, National Institutes of Health and Muscular Dystrophy Association. ZP Huang is a postdoctoral fellow and DZ Wang is an Established Investigator of the American Heart Association. References 1. Wagner, K.R. (2002) Genetic diseases of muscle. Neurol Clin. 20, 645–678. 2. Bartel, D.P. (2004) MicroRNAs: genomics, biogenesis, mechanism, and function. Cell. 116, 281–297. 3. He, L., Thomson, J.M., Hemann, M.T., Hernando-Monge, E., Mu, D., Goodson, S., Powers, S., Cordon-Cardo, C., Lowe, S.W., Hannon, G.J., and Hammond, S.M. (2005) A microRNA polycistron as a potential human oncogene. Nature. 435, 828–833. 4. Rodriguez, A., Griffiths-Jones, S., Ashurst, J.L., and Bradley, A. (2004) Identification of mammalian microRNA host genes and transcription units. Genome Res. 14, 1902–1910. 5. Ruby, J.G., Jan, C.H., and Bartel, D.P. (2007) Intronic microRNA precursors that bypass Drosha processing. Nature. 448, 83–86. 6. Yi, R., Qin, Y., Macara, I.G., and Cullen, B.R. (2003) Exportin-5 mediates the nuclear export of pre-microRNAs and short hairpin RNAs. Genes Dev. 17, 3011–3016. 7. Hutvágner, G., McLachlan, J., Pasquinelli, A.E., Bálint, E., Tuschl, T., and Zamore, P.D. (2001) A cellular function for the RNAinterference enzyme Dicer in the maturation of the let-7 small temporal RNA. Science. 293, 834–838. 8. Schwarz, D.S., Hutvágner. G., Du. T., Xu. Z., Aronin. N., and Zamore, P.D. (2003) Asymmetry in the assembly of the RNAi enzyme complex. Cell. 115, 199–208. 9. Lee, R.C., and Ambros, V. (2001) An extensive class of small RNAs in Caenorhabditis elegans. Science. 294, 862–864. 10. Williams, A.H., Liu, N., van Rooij, E., and Olson, E.N. (2009) MicroRNA control of muscle development and disease. Curr Opin Cell Biol. 21, 461–469. 11. Zhao, Y., Samal, E., and Srivastava, D. (2005) Serum response factor regulates a muscle-specific microRNA that targets Hand2 during cardiogenesis. Nature. 436, 214–220.

12. Chen, J.F., Mandel, E.M., Thomson, J.M., Wu, Q., Callis, T.E., Hammond, S.M., Conlon, F.L., and Wang, D.Z. (2006) The role of microRNA-1 and microRNA-133 in skeletal muscle proliferation and differentiation. Nat Genet. 38, 228–233. 13. van Rooij, E., Sutherland, L.B., Qi, X., Richardson, J.A., Hill, J., and Olson, E.N. (2007) Control of stress-dependent cardiac growth and gene expression by a microRNA. Science. 316, 575–579. 14. Simon, D.J., Madison, J.M., Conery, A.L., Thompson-Peer, K.L., Soskis, M., Ruvkun, G.B., Kaplan, J.M., and Kim, J.K. (2008) The microRNA miR-1 regulates a MEF-2dependent retrograde signal at neuromuscular junctions. Cell. 133, 903–915. 15. Kwon, C., Han, Z., Olson, E.N., and Srivastava, D. (2005) MicroRNA1 influences cardiac differentiation in Drosophila and regulates Notch signaling. Proc Natl Acad Sci USA. 102, 18986–18991. 16. Callis, T.E., and Wang, D.Z. (2008) Taking microRNAs to heart. Trends Mol Med. 14, 254–260. 17. Chen, J.F., Callis, T.E., and Wang, D.Z. (2009) microRNAs and muscle disorders. J Cell Sci. 122, 13–20. 18. Yang,B., Lin, H., Xiao, J., Lu, Y., Luo, X., Li, B., Zhang, Y., Xu, C., Bai, Y., Wang, H., Chen, G., and Wang, Z. (2007) The muscle-specific microRNA miR-1 regulates cardiac arrhythmogenic potential by targeting GJA1 and KCNJ2. Nat Med. 13, 486–91. 19. Callis, T.E., Pandya, K., Seok, H.Y., Tang, R.H., Tatsuguchi, M., Huang, Z.P., Chen, J.F., Deng, Z., Gunn, B., Shumate, J., Willis, M.S., Selzman, C.H., Wang, D.Z. (2009) MicroRNA-208a is a regulator of cardiac hypertrophy and conduction in mice. J Clin Invest. 119, 2772–86. 20. McCarthy, J.J. (2008) MicroRNA-206: the skeletal muscle-specific myomiR. Biochim Biophys Acta. 1779, 682–691.

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21. Eisenberg, I., Eran, A., Nishino, I., Moggio, M., Lamperti, C., Amato, A.A., Lidov, H.G,, Kang, P.B., North, K.N., Mitrani-Rosenbaum, S., Flanigan, K.M., Neely, L.A., Whitney, D., Beggs, A.H., Kohane, I.S., and Kunkel, L.M. (2007) Distinctive patterns of microRNA expression in primary muscular disorders. Proc Natl Acad Sci. USA. 104, 17016–17021. 22. van Rooij, E., Sutherland, L.B., Liu, N., Williams, A.H., McAnally, J., Gerard, R.D., Richardson, J.A., and Olson, E.N. (2006) A signature pattern of stress-responsive microRNAs that can evoke cardiac hypertrophy and heart failure. Proc Natl Acad Sci. USA. 103, 18255–18260. 23. van Rooij, E., Sutherland, L.B., Thatcher, J.E., DiMaio, J.M., Naseem, R.H., Marshall, W.S., Hill, J.A., and Olson, E.N. (2008) Dysregulation of microRNAs after myocardial infarction reveals a role of miR-29 in cardiac fibrosis. Proc Natl Acad Sci. USA. 105, 13027–13032. 24. Tatsuguchi, M., Seok, H.Y., Callis, T.E., Thomson, J.M., Chen, J.F., Newman, M., Rojas, M., Hammond, S.M., and Wang, D.Z. (2007) Expression of microRNAs is dynamically regulated during cardiomyocyte hypertrophy. J Mol Cell Cardiol. 42, 1137–1141. 25. McCarthy, J.J., Esser, K.A., and Andrade, F.H. (2007) MicroRNA-206 is overexpressed in the diaphragm but not the hindlimb muscle of mdx mouse. Am J Physiol Cell Physiol. 293, C451–457. 26. Gentner, B., Schira, G., Giustacchini, A., Amendola, M., Brown, B.D., Ponzoni, M., and

Naldini, L. (2009) Stable knockdown of microRNA in vivo by lentiviral vectors. Nat Methods. 6, 63–66. 27. Krützfeldt, J., Rajewsky, N., Braich, R., Rajeev, K.G., Tuschl, T., Manoharan, M., and Stoffel, M. (2005) Silencing of microRNAs in vivo with ‘antagomirs’. Nature. 438, 685–689. 28. Krek, A., Grun, D., Poy, M.N., Wolf, R., Rosenberg, L., Epstein, E.J., MacMenamin, P., da Piedade, I., Gunsalus, K.C., Stoffel, M. & Rajewsky, N. (2005). Combinatorial microRNA target predictions. Nat Genet. 37, 495–500. 29. John, B., Enright, A.J., Aravin, A., Tuschl, T., Sander, C. & Marks, D.S. (2004). Human MicroRNA targets. PLoS Biol. 2, e363. 30. Lewis, B.P., Burge, C.B. & Bartel, D.P. (2005). Conserved seed pairing, often flanked by adenosines, indicates that thousands of human genes are microRNA targets. Cell. 120, 15–20. 31. Selbach, M., Schwanhäusser, B., Thierfelder, N., Fang, Z., Khanin, R., Rajewsky, N. (2008) Widespread changes in protein synthesis induced by microRNAs. Nature. 455, 58–63. 32. Baek, D., Villén, J., Shin, C., Camargo, F.D., Gygi, S.P., Bartel, D.P. (2008) The impact of microRNAs on protein output. Nature. 455, 64–71. 33. Stegmeier, F., Hu, G., Rickles, R.J., Hannon, G.J., Elledge, S.J. (2005) A lentiviral microRNA-based system for single-copy polymerase II-regulated RNA interference in mammalian cells. Proc Natl Acad Sci. USA. 102, 13212–13217.

Chapter 29 shRNA-Mediated Gene Knockdown in Skeletal Muscle Muriel Golzio, Jean-Michel Escoffre, and Justin Teissié Abstract RNA interference appears as a promising tool for therapeutic gene silencing to block protein expression. A long-lived silencing is obtained through the in situ expression of shRNA. A safe approach is to use a physical method such as in vivo electropulsation with plate electrodes. This is presently validated in muscles by the in vivo coelectrotransfer of plasmids specifically coding for expression and silencing of a fluorescent protein. No long-lived tissue damage is observed by the proper choice of the electric pulsing parameters and the amount of injected plasmids. Using a noninvasive fluorescence imaging assay, electrodelivery in mouse muscles is observed to induce complete silencing over more than 2 months in a specific way. The proper choices of the plasmids (sequence, promoter, and relative amounts) appear as key parameters in the successful long-term silencing. Key words: Plasmids, Gene electrotransfer, GFP, Muscle, Silencing, shRNA

1. Introduction RNA interference can be achieved by using chemically synthesized small interfering RNA (siRNA) or short hairpin RNA (shRNA) expressing plasmid. Previous works showed that the delivery of chemically synthesized siRNA resulted in strong and sequencespecific inhibition of gene expression in vitro (1, 2) and in vivo (3, 4). However, chemically synthesized siRNAs had several drawbacks beside their expensive cost such as a transient gene expression silencing due to their short lived stability in vivo (5, 6). To overcome these limitations, the delivery of shRNA expression cassettes appeared as a more suitable approach. The development of these expression cassettes required safe and efficient in vivo-targeted delivery methods. Viral vectors have been reported as highly efficient methods for shRNA delivery to several tissues (7, 8). But safety concerns remain

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present concerning their clinical use. Direct delivery of plasmids is another approach. DNA plasmids are composed entirely of covalently closed circles of double-stranded DNA with no associated proteins. Commercially available efficient, highly transfectable, and simple-to-use plasmids have been designed such as psiRNA™ and pCpG-siRNA™. These plasmids were designed by inserting a DNA fragment of approximately 50 mer. After transcription from the human H1 or 7SK RNA polymerase III promoter, this insert generated short RNAs with a hairpin structure (shRNAs) (9, 10). shRNAs were more stable than chemically synthesized siRNAs and their expression within the cells allowed long-lasting silencing of target gene expression (11). DNA plasmids are more suitable for large volume production and quality control than viral vectors. Moreover, the safety advantages of DNA plasmid are lack of integration and low immunogenicity. Plasmid DNA can be a highly attractive molecule for gene therapy, but it must be associated with safe, efficient, and targeted delivery as gene delivery with plasmid vectors is highly inefficient if DNA plasmids are not associated with chemical or physical methods (12, 13). During the 90’s, in vivo electrotransfer appeared as a promising tool for exogenous drugs and nucleic acids delivery. Moreover, this nonviral method offered the advantages as reduction of toxicity, safety, and friendly use (14). In vivo, electrotransfer allowed efficient delivery of DNA plasmids and other large molecules like proteins (15) and antisense oligonucleotides (16). Indeed, a wide range of tissues were targeted including skin (17), liver (18), lung (19), skeletal (20, 21) and cardiac muscle (22), kidney (23), joints (24), brain (25), and retina (26). But skeletal muscle attracted a lot of attention, muscle being considered as a first-choice cellular factory. Expression of episomal plasmids could be long-lived in muscle tissue (20, 21). Delivery was targeted to the volume of tissue localized between the electrodes, where the electric field is applied (27, 28). In living animals, the quantitative follow-up of reporter gene expression is very important to monitor the therapeutic gene expression in targeted tissues and to assess the effectiveness of delivery methods. In vivo optical imaging is a noninvasive method which can detect and follow the reporter gene activity on the same animal as a function of time (29, 30). Indeed, working on the same animal brings a reduction of the number of experimental animals and increases the accuracy of statistical analysis. Exogenous gene expression of fluorescent proteins such as enhanced green fluorescent protein (eGFP) can be detected directly in living animals by means of a fluorescence macroscope coupled to a cooled charged-coupled device camera (CCD camera). In this study, we established the proof of concept of the effectiveness of electrotransfer for the targeted delivery of DNA plasmid coding for shRNA in adult mice using tibialis cranialis muscle as a

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model system. The resulting gene silencing was monitored in living animals by time-lapse fluorescence imaging. Silencing of gene expression was observed over a 2-month period.

2. Materials 2.1. Plasmids

Plasmids were produced in the Cayla facility (Invivogen). Identity was confirmed by agarose gel. Contamination with RNA was not observed and the majority of plasmids was present as covalently closed circles. 1. pCLEF14-EGFP plasmid contains eGFP cDNA under the control of the elongation factor 1A (EF 1A) promoter. pUC18 plasmid contains the cDNA of subunit A of LacZ under the control of the pLac promoter. Prepared plasmids from Escherichia coli with a maxiprep endotoxin-free cartridge (Qiagen). 2. psiRNA25-EGFP or psiRNA25-SCR: psiRNA25 is an RNA polymerase III-based plasmid which contains the human 7SK RNA Pol III promoter. Design the DNA fragments coding for scramble shRNA (SCR) or shRNA against eGFP (EGFP) mRNA by a siRNA design algorithm, named siRNA Wizard (http://www.sirnawizard.com). DNA sequence of eGFP shRNA is (relative position to ATG), GCAAGCTGACCCTGAAGT TCACCACCTGAACTTCAGGGTCAGCTTGC and DNA sequence of SCR is GCATATGTGCGTACCTAG CATTCAAGAGATGCTAGGTACGCACATATGC. 3. pCpG76-EGFP or pCpG76-SCR: pCpG76 is a plasmid that combines a CpG-free plasmid backbone with shRNA expression cassette of psiRNA25 plasmids. In the expression cassette, mCMV enhancer sequence is added on upstream of the 7SK promoter. This plasmid is designed for long-lasting expression of shRNA in vivo as the plasmid does not induce inflammatory responses (31) and gene silencing by methylation in vertebrate hosts (32). Use them as a control for specificity of the shRNA construct.

2.2. In Vivo Experiments

All animal studies were conducted in accordance with the principles and procedures outlined by the European convention for the protection of vertebrate animals used for experimentation. 1. Female Balb/c mice (Charles River) 9–10 weeks old at the beginning of the experiments, weighing 20–25 g, maintained at constant room temperature with 12-h light cycle in a conventional animal colony. Before the experiments, subject mice to an adaptation period of at least 10 days. 2. Veet Cream (Reckitt Benckiser).

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3. Hamilton syringe with a 26G needle. 4. Isoflurane. 5. Anesthetic machine with air compressor and isoflurane vaporizer (Xenogene). 6. Electropulsator PS 10 CNRS or Betatech electropulsator. Monitor all parameters on line with an oscilloscope. An electronic switch cutting the pulse as soon as its intensity is 5 Amp obtains safety against current surge. 7. Plate parallel stainless steel electrodes (length 1 cm, width 0.6 mm). 8. Conducting paste (Eko-gel). 2.3. Noninvasive Visualization of Gene Expression and/or Gene Silencing

1. A fluorescence macroscope (Leica). The fluorescence excitation was obtained with an EL6000 lamp, GFP, or the G filters. 2. Cooled CCD Camera Coolsnap HQ (Roper Scientific). 3. The MetaVue software (Universal) drives the CCD camera from a Dell computer under Windows XP and allows quantitative analysis of the GFP fluorescence level.

3. Methods In this study, electrotransfer of plasmid encoding the GFP reporter gene was used to show the proof of concept of the efficiency of in vivo electro-administration of specific shRNA after intramuscular injection in adult mice. We compared treatment groups using psiRNA25-EGFP or pCpG76-EGFP, electric field alone, and nonrelevant psiRNA25-SCR or pCpG76-SCR. It is important to use the same volumes for injection to obtain reliable results. The first step was to determine the kinetics of GFP gene expression and then to determine whether the injection of different ratios of GFP plasmid to shRNA plasmid affected GFP expression. The expression of the fluorescent reporter gene was determined by in vivo fluorescence macroscopy to quantify the fluorescence on the digitized images. 3.1. In Vivo Electrotransfer

1. Two days before the treatment, shave one of the legs with the cream (see Note 1). 2. Anesthetize mice by isoflurane inhalation (see Note 2). 3. In total, mix 5 Mg of pCLEF14-EGFP (1 Mg/ML) in PBS (see Note 3) with 10 Mg (Ratio target/shRNA, 1/2), 25 Mg (Ratio target/shRNA, 1/5), 50 Mg (ratio target/shRNA, 1/10) of plasmid expressing shRNA or pUC18 (control conditions). Adjust the DNA mix to a 25 ML final volume with PBS. Slowly

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Fig. 1. Experimental procedure. The animal is under anesthesia during all the experimental procedures. (a) The plasmid solution (25 ML) is injected in the tibialis muscle. The mouse leg is previously shaved. (b) The electrodes are set around the leg and four electric pulses of 120 V lasting 20 ms at 1 Hz are applied. (c) The electrode set is turned to a perpendicular position around the leg. Four electric pulses of 120 V lasting 20 ms at 1 Hz are again delivered.

(about 15 s) inject solutions (see Note 4) with a Hamilton syringe through a 26G needle into the tibialis muscle in mice (see Fig. 1a). 4. Obtain a good electric contact between the skin and the electrodes by use of a conducting paste (see Note 5). 5. Around 30 s following injection, fit the plate parallel electrodes around the leg (see Note 6). The fixed 6 mm gap distance between the electrodes allows a good contact with the skin surface. Electrode position can be easily changed by a rotation of the electrode set around the muscle. A 90° rotation brings a direction of the field in a perpendicular direction, so-called crossed directions (see Note 7). Apply electric pulses of 120 V in two sets of four rectangular wave pulses (see Note 8) of crossed directions, lasting 20 ms at 1 Hz (see Fig. 1b, c). 3.2. Fluorescence Data Acquisition and Analysis

The electrically mediated GFP gene transfer in the mouse muscle is detected directly on the anesthetized animal by digitized macroscopy. Fluorescent muscle fibers are observed through the skin. This procedure allows monitoring of reporter gene expression on the same animal by time-lapse fluorescence imaging. The GFP fluorescence from the muscle is quantitatively evaluated at different days and thereafter with weekly intervals until the GFP fluorescence is no longer detectable. 1. Anesthetize the mouse. 2. Hold the leg and place it under the fluorescence macroscope. Observe the whole muscle as a 12 bits 1.3 M pixels image with a cooled CCD Camera. Drive the camera from a Dell computer with MetaVue software. Take a normal light picture (see Note 9). 3. Obtain the fluorescence excitation with an EL6000 lamp. Set the exposure time at 1 s with no binning. Acquire two different

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Fig. 2. Whole-muscle fluorescence imaging. (a) The electrotransfected muscle is observed by fluorescence macroscopy with the GFP filter. GFP and autofluorescence emissions are detected. (b) The same muscle is observed with the G filter. Only the autofluorescence emission is detected. (c) On the resulting image (a, b), only the GFP emission is detected and quantified into the region of interest (ROI in white).

pictures by selecting the GFP (see Fig. 2a) and the G filters (see Fig. 2b). 4. From the transmission picture, locate and gate the tibialis cranialis muscle to give the region of interest (ROI). Subtract the picture of muscle with G filter (autofluorescence signal) from the image of muscle with GFP filter. This operation suppresses most of the autofluorescence. On the resulting image (see Fig. 2c), the tibialis cranialis muscle is located and gated to give the ROI. The mean fluorescence in the gated area (whole muscle) is quantitatively determined. 5. Treat 4–6 legs of different animals for each condition. Treat only one leg per animal to avoid cross-reaction between the successive treatments. Statistically evaluate differences between mean fluorescence levels measured in the experiments by using an unpaired Student t-test using the Prism software (version 4.02).

4. Conclusion Mice muscles were electropulsed with a mixture of pCLEF14EGFP plasmid and pUC18, psiRNA25-EGFP, psiRNA25-SCR, pCpG76-EGFP, or pCpG76-SCR at various ratios (1:2, 1:5, and 1:10) with a constant amount of pCLEF14-EGFP plasmid (5 Mg). GFP fluorescence is quantified by time-lapse fluorescence imaging on the same animal. Fluorescence is present only in the tibialis cranialis muscle that is electropulsed. It remains detectable during a long period (more than 70 days). When anesthetized mice are electropulsed after injection of pCLEF14-EGFP plasmid mixed with control plasmid (psiRNA25-SCR, pCpG76-SCR, pUC18),

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Fig. 3. Visualization of the cotransfection GFP/shRNA (plasmid ratio 1:2). Whole muscle imaging after cotransfer with pUC18 (a), psiRNA25-SCR (b), and psiRNA25-EGFP (c) plasmids on day 4 and after cotransfer with psiRNA25-SCR (d) and psiRNA25-EGFP (e) plasmids on day 30. White bar represents 0.4 cm.

GFP expression is detected 24 h after electrotransfer and remains present until day 72 (see Figs. 3a, b, d and .5a, b, d). Varying the quantity of shRNA expressing plasmid is achieved to evaluate the efficiency of these different plasmids. The cotransfer of pCLEF14EGFP plasmid and pCpG76-EGFP or psiRNA25-EGFP plasmid in a ratio 1:2 induces a partial silencing of GFP expression during the first 23 days (see Figs. 3c and 4). However, after day 23, GFP expression is not detected until day 72 (see Figs. 3e and 4). Codelivery of shRNA against GFP completely silences GFP expression when a ratio larger than 1/5 is used (see Fig. 5c, d, f). When the pCLEF14-EGFP plasmid is cointroduced with pCpG76-SCR and whatever the ratio between these two plasmids, the GFP expression is higher than after the cointroduction with pUC18 or psiRNA25-SCR plasmid during 70 days (see Figs. 4 and 5f).

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Fig. 4. Quantification of the fluorescence intensity (plasmid ratio 1:2). Mean fluorescence changes after the different treatments pUC18 (filled diamond ), psiRNA25-SCR (square) and psiRNA25-EGFP (filled square), pCpG76-SCR (filled triangle), and pCpG76-EGFP (triangle) plasmids (±SD) are plotted as a function of time. N = 4.

Fig. 5. Cotransfection (plasmid ratio GFP/shRNA of 1/5). Whole-muscle imaging on day 6 after cotransfer with pUC18 (a), psiRNA25-SCR (b) and psiRNA25-EGFP (c), pCpG76-SCR (d), and pCpG76-EGFP (e) plasmids. White bar represents 0.4 cm. Mean fluorescence changes after the different treatments pUC18 (filled diamond ), psiRNA25-SCR (square) and psiRNA25-EGFP (filled square), pCpG76-SCR (filled triangle), and pCpG76-EGFP (triangle) plasmids (±SD) are plotted as a function of time (f). N = 6.

A specific enhancing effect in expression (fluorescence) is associated with the pCpG construct after cointroduction with the EGFPcoding plasmid (33). The described intramuscular injection of shRNA is safe for the animal. A muscle contraction is observed when the electric field pulse is delivered. No local burns, edema, or loss of limb functions are observed. Intramuscular injection of GFP plasmid DNA induces

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GFP expression in the injected muscles of Balb/C mice after pulse delivery for more than 70 days, peaking at day 20. Codelivery of shRNA against GFP completely silences GFP expression when a ratio larger than 1:5 is used and partially silences the GFP expression with a ratio 1:2 in a statistically significant way.

5. Notes 1. The cream should be used 2 days before the fluorescence imaging because some components fluoresce under blue excitation. This cream should be used carefully, as it may cause some irritations in the skin of the leg. Rinse the cream with lots of water. 2. Isoflurane inhalation is safe; mice recover very fast after the electrical treatment. It can be used every day for in vivo imaging, with no pain for the observed animal. 3. PBS was used to avoid an osmotic shock due to the injection of the plasmid solution. 4. A too rapid injection could give false-positive muscle fibers. This injection needle must be parallel to the fibers (34). 5. Conductive paste is very important to ensure a good electrical contact with the skin. One should pay attention that the paste is not continuous between the two electrodes as the field will pass through the paste and not through the muscle. 6. Carefully clean the surface of the electrodes at the end of the experiments to avoid rusting due to the electrochemical reactions associated with the pulses. 7. Changing the orientation of the cells in the electric field led to a higher effect in enhancement of transfection in vivo in intact animals. The vectorial property of the electric field and the electrophoretic migration of DNA during pulses lead to a specific transport of DNA on only one pole of the permeabilized cells. Changing the orientation of the pulses increases interstitial transport of plasmid DNA in the tissue and therefore gene transfer (35). 8. Square wave pulse generators are needed. As the tissue conductance is affected by the field-induced cell membrane permeabilization, the time constant of a capacitor discharge pulse generator is changing during the pulse. This leads to loss in control in the duration of the effective pulse. The pulse delivery must be controlled on line either on an oscilloscope or after digitization on a laptop. Because of the high current, one may observe that the voltage is slightly decreasing at the end of the millisecond pulse. Because of power limitation of the pulse generator in most cases, it is difficult to work a train frequency larger than

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1 Hz. This is a technical reason. There is no evidence that it is the optimized setting from a biophysical point of view. 9. One person should be responsible for holding the leg of the mouse in the same position under the macroscope to avoid lack of reproducibility in the exposition to the excitation beam.

Acknowledgments This work was supported by grants from the CNRS CEA “Imagerie du petit animal” program, the Region Midi Pyrenees (Therapie génique et cellulaire), and the Association française contre les Myopathies. References 1. Elbashir SM, Lendeckel W, Tuschl T (2001) RNA interference is mediated by 21 and 22 nucleotides RNAs. Genes Development 15:188–200 2. Elbashir SM, Harborth J, Lendeckel W, Yalcin A, Weber K, Tuschl T (2001) Duplexes of 21 nucleotides RNAs mediate RNA interference in cultured mammalian cells. Nature 411: 494–498 3. Golzio M, Mazzolini L, Moller P, Rols MP, Teissie J (2005) Inhibition of gene expression in mice muscle by in vivo electrically mediated siRNA delivery. Gene Therapy 12:246–251 4. Kishida T, Asada H, Gojo S, Ohashi S, Shin-Ya M, Yasutomi K, et al. (2004) Sequence-specific gene silencing in murine muscle induced by electroporation-mediated transfer of short interfering RNA. J Gene Med 6:105–110 5. Ryther RCC, Flynt AS, Philips JA, Patton JG (2005) siRNA therapeutics; big potential from small RNAs. Gene Therapy 12:5–11 6. Golzio M, Mazzolini L, Ledoux A, Paganin A, Izard M, Hellaudais L, et al. (2007) In vivo gene silencing in solid tumors by targeted electrically mediated siRNA delivery. Gene Ther 14:752–759 7. Tiscornia G, Singer, O, Ikawa M, Verna I (2003) A general method for gene knockdown in mice by using lentiviral vectors expressing small interfering RNA. Proc Natl Acad Sci USA 100:1844–1848 8. Uchida H, Tanaka T, Sasaki K, Kato K, Dehari H, Ito Y, et al. (2004) Adenovirus-mediated transfer of siRNA against surviving induced apoptosis and attenuated tumor cell growth in vitro and in vivo. Molecular Therapy 10: 162–171

9. Czauderna F, Santel A, Hinz M, Durieux B, Arnold W, Klippel A, et al. (2003) Inductible shRNA expression for application in a prostate cancer mouse model. Nucleic Acids Research 31:e127 10. Ill CR, Chiou HC (2005) Gene therapy progress and prospects: recent progress in transgene and RNAi expression cassettes. Gene Therapy 12:795–802 11. Scherr KJ, Morgan MA, Eder M (2003) Gene silencing mediated by small interfering RNAs in mammalian cells. Current Medicinal Chemistry 10:245–256 12. Wells DJ (2004) Gene therapy progress and prospects: electroporation and other physical methods. Gene Therapy 11:1363–1369 13. Vorhies JS, Nemunaitis J (2007) Nonviral delivery vehicles for use in short hairpin RNA-based cancer therapies. Expert Rev Anticancer Ther 7:373–82 14. Golzio M, Rols MP, Teissie J (2004) In vitro and in vivo electric field-mediated permeabilization, gene transfer and expression. Methods 32:126–135 15. Rols MP, Delteil C, Golzio M, Dumond P, Cros S, Teissie J (1998) In vivo electrically mediated protein and gene transfer in murine melanoma. Nature Biotechnology 16:168–171 16. Faria M, Spiller DG, Dubertret C, Nelson JS, White MR, Scherman D, et al. (2001) Phosphoramidate oligonucleotides as potent antisense molecules in cells and in vivo. Nature Biotechnology 19:40–44 17. Pedron-Mazoyer S, Plouët J, Hellaudais L, Teissie J, Golzio M (2007) New anti-angiogenesis developments through electro-immunization:

29 optimization by in vivo optical imaging of intradermal electrogenetransfer. Biochim Biophys Acta 1770:137–142 18. Heller R, Jaroszeski M, Atkin A, Moradpour D, Gilbert R, Wands J et al. (1996) In vivo gene electroinjection and expression in rat liver. FEBS Lett 389:225–228 19. Pringle IA, McLachlan G, Collie DD, SumnerJones SG, Lawton AE, Tennant P et al. (2007) Electroporation enhances reporter gene expression following delivery of naked plasmid DNA to the lung. J Gene Med 9:369–380 20. Aihara H, Miyazaki JI (1998) Gene transfer into muscle by electroporation in vivo. Nature Biotechnology 16:867–870 21. Mir LM, Bureau FB, Gehl J, Rangara R, Rouy D, Caillaud JM, et al. (1999) High-efficiency gene transfer into skeletal muscle mediated by electric pulses. Proc Natl Sci USA 96:4262–4267 22. Harrison RL, Byrne BJ, Tung L (1998) Electroporation-mediated gene transfer in cardiac tissue. FEBS Lett 435:1–5 23. Isaka Y, Yamada K, Takabatake Y, Mizui M, Miura-Tsujie M, et al. (2005) Electroporationmediated HGF gene transfection protected the kidney against graft injury. Gene Therapy 12:815–820 24. Khoury M, Bigey P, Louis-Plence P, Noel D, Rhinn H, Scherman D, et al. (2006) A comparative study on intra-articular versus systemic gene electrotransfer in experimental arthritis. Gene Therapy 8:1027–1036 25. Wang H, Ko CH, Koletar MM, Ralph MR, Yeomans J (2007) Casein kinase I epsilon gene transfer into the suprachiasmatic nucleus via electroporation lengthens circadian periods of tau mutant hamsters. Eur J Neuroscience 25:3359–66 26. Matsuda T, Cepko CL (2004) Electroporation and RNA interference in the rodent retina in vivo and in vitro. Proc Natl Acad Sci USA 101:16–22

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27. Cemazar M, Golzio M, Sersa G, Rols, MP, Teissié J (2006) Electrically-assisted nucleic acids delivery to tissues in vivo: where do we stand? Current Pharmaceutical Design 12:3817–3825 28. Heller LC, Heller R. (2006) In vivo electroporation for gene therapy. Hum Gene Ther 17:890–7 29. Golzio M, Rols MP, Gabriel B, Teissie J (2004) Optical imaging of in vivo gene expression: a critical assessment of the morphology and associated technologies. Gene Therapy 11: S85–S91 30. Yang M, Baranov E, Li XM, Wang JW, Jiang P, Li L, et al. (2001) Whole-body and intravital optical imaging of angiogenesis in orthotopically implanted tumors. Proc Natl Acad Sci USA 98:2616–2621 31. Bauer S, Kirschning CJ, Hacker H, Redecke V, Hausmann S, Akira S, et al. (2001) Human TLR9 confers responsiveness to bacterial DNA via species-specific CpG motif recognition. Proc Natl Acad Sci USA 98:9237–9242 32. Chevalier-Mariette C, Henry I, Montfort L, Capgras S, Forlani S, Muschler et al. (2003) CpG content gene silencing in mice : evidence from novel transgenes. Genome Biology 4:R53 33. Escoffre JM, Debin A, Reynes JP, Drocourt D, Tiraby G, Hellaudais L, Teissie J, Golzio M. (2008) Long-lasting in vivo gene silencing by electrotransfer of shRNA expressing plasmid. Technol Cancer Res Treat 7:109–16 34. André FM, Cournil-Henrionnet C, Vernerey D, Opolon P, Mir LM. (2006) Variability of naked DNA expression after direct local injection: the influence of the injection speed. Gene Ther 13:1619–27 35. Faurie C, Golzio M, Moller P, Teissié J, Rols MP. (2003) Cell and animal imaging of electrically mediated gene transfer. DNA Cell Biol 22:777–83

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Chapter 30 Detection of NF-kB Activity in Skeletal Muscle Cells by Electrophoretic Mobility Shift Analysis Jason M. Dahlman and Denis C. Guttridge Abstract An electrophoretic mobility shift assay (EMSA) is a common and invaluable technique which can be utilized to study the affinity of proteins to a specific DNA or RNA sequence. These assays are performed in vitro with protein extracts isolated from either cultured cells or isolated tissues. Here, we describe the methodology used to isolate the cytoplasmic and nuclear protein extracts from both cultured cells and tissues and utilize the nuclear protein fraction to assess NF-KB DNA-binding activity by EMSA analysis. Key words: NF-KB, Skeletal muscle, Transcription factors, Myogenesis, Differentiation, Muscular dystrophy, Atrophy

1. Introduction EMSAs are a common molecular biology technique used to detect the binding of proteins to a specific nucleotide sequence (1–3). This technique was originally described by Fried and Crothers (1) and Garner and Revzin (2) and more modern protocols have only differed slightly over the last 20 years. Some slight modifications have been made to make this technique more applicable for studying protein/nucleotide interaction from samples collected from both cultured cells as well as specific tissues (4–10). For the purpose of this review, we will go into detail how to isolate nuclear extracts from both cultured cells as well as skeletal muscle tissue and investigate NF-KB activity via EMSA analysis. Furthermore, we will highlight two techniques which will aid in identifying which visualized EMSA bands are NF-KB-specific through the use of a competitive “cold” EMSA probe (8) as well as supershift analysis (5–7).

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2. Materials Make all solutions using ultrapure H2O (prepared by purifying deionized H2O to attain a sensitivity of 18 M7 at 25°C) unless specified otherwise. 2.1. Nuclear Extraction from Cell Pellets

1. Cytoplasmic extract buffer: 10 mM HEPES pH 7.6, 60 mM KCl, 1 mM EDTA. For working solution, add 100 ML of 1 M HEPES pH 7.6, 200 ML of 3 M KCl, and 20 ML of 0.5 M EDTA to 9.68 mL of ultrapure H2O (see Note 1). 2. Nuclear extract buffer (NEB): 20 mM Tris pH 8.0, 420 mM NaCl, 1.5 mM MgCl2, 0.2 mM EDTA, 25% glycerol. For working solution, add 200 ML of 1 M Tris pH 8.0, 840 ML of 5 M NaCl, 15 ML of 1 M MgCl2, 50 ML of 0.5 M EDTA, and 2.5 mL of 100% glycerol (see Note 2) to 6.395 mL of ultrapure H2O (see Note 1). 3. Protease inhibitor cocktail: Either use Sigma’s cocktail (5 ML/mL) (P8340), or prepare your own mix: 2.5 Mg/mL leupeptin, 2.5 Mg/mL aprotinin, 2.5 Mg/mL Pepstatin, 1 mM PMSF, and 1 MM DTT. In both cases, store the cocktails at −20°C.

2.2. Nuclear Extraction from Skeletal Muscle

1. Low salt lysis buffer: 10 mM HEPES pH 7.6, 10 mM KCl, 1.5 mM MgCl2, 0.1 mM EDTA, 0.1 mM EGTA, 1 mM DTT, 0.5 mM PMSF, 50 ML of Protease inhibitor cocktail from Subheading 2.1, item 3, 0.5 mg/mL benzamidine. For working solution, add 100 ML of 1 M HEPES pH 7.6, 100 ML of 1 M KCl, 15 ML of 1 M MgCl2, 4 ML of 0.25 M EDTA, 10 ML of 0.1 M EGTA, 100 ML of 100 mM DTT, 100 ML of 50 mM PMSF, 50 ML of protease inhibitor cocktail from Subheading 2.1, item 3, and 5 mg of benzamidine in 9.5 mL of ultrapure H2O (see Note 1). 2. High salt nuclear buffer: 20 mM HEPES pH 7.6, 420 mM NaCl, 1 mM EDTA, 1 mM EGTA, 25% glycerol, 1 mM DTT, 5 ML of protease inhibitor cocktail from Subheading 2.1, item 3. For working solution, add 20 ML of 1 M HEPES pH 7.6, 84 ML of 5 M NaCl, 4 ML of 0.25 M EDTA, 10 ML of 0.1 M EGTA, 250 ML of 100% glycerol (see Note 2), 10 ML of 100 mM DTT, and 5 ML of protease inhibitor cocktail from Subheading 2.1, item 3 in 617 ML of ultrapure H2O (see Note 1).

2.3. Electrophoretic Mobility Shift Assay

1. 10× Tris–glycine–EDTA (TGE): 0.25 M Tris, 1.9 M glycine, 0.01 M EDTA. Make up a stock solution of 10× TGE by adding 121.12 g of Tris, 568 g of glycine, and 14.8 g of EDTA to 4 L of ultrapure H2O.

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2. 10% Ammonium persulfate (APS): For working solution, add 100 mg to 1 mL of ultrapure H2O (see Note 3). Store in −20°C. 0OLYD) D# sPOLYD) D# D)D# &ORNUCLEAREXTRACTSFROM cell pellets, add 0.5 ML of 50 mM PMSF, 0.5 ML of dIdC, and 6 ML of ultrapure H2O per assay sample. For nuclear extracts from muscle extracts, add 1 ML of 50 mM PMSF, 1 ML of dIdC, and 12 ML of ultrapure H2O per assay sample. 4. Normal 5× binding buffer: 50 mM Tris pH 7.6, 50% glycerol, 5 mM DTT, 2.5 mM EDTA. Add 50 ML of 1 M Tris pH 7.6, 500 ML of 100% glycerol (see Note 1), and 50 ML of 100 mM DTT, and 25 ML of 100 mM EDTA to 425 ML of ultrapure H2O. 5. Special 5× binding buffer: 125 mM HEPES pH 7.6, 2.5 mM EDTA, 2.5 mM DTT, 1% NP-40, 25% glycerol, 250 mM NaCl. Add 125 ML of 1 M HEPES pH 7.6, 10 ML of 0.25 M EDTA, 25 ML of 100 mM DTT, 100 ML of 10% NP-40 (see Note 4), 250 ML of 100% glycerol (see Note 1), 50 ML of 5 M NaCl to 445 ML of ultrapure H2O. 6. EMSA loading dye: 250 mM Tris–HCl pH 7.4, 0.2% bromophenol blue, 0.2% xylene cyanol, 40% sucrose.

3. Methods Perform all steps at room temperature unless otherwise specified. Volumes are determined for a preparation of 10 samples. All centrifugation was performed in a bench top microfuge. 3.1. Preparation of Nuclear Extracts from Cell Pellets

1. Aspirate off cell culture growth media from desired cells (for efficient nuclear extraction, a minimum of 1 × 106 cultured muscle cells should be used. Generally, this is easily achieved by growing cells in a 10 cm2 culture dish). 2. Wash the cells gently with 1 mL of nonsterile PBS. Aspirate off the PBS and repeat once. 3. Remove cultured cells from the polystyrene tissue culture dish by adding 1 mL of nonsterile PBS and scraping the cells gently with a sterile cell scraper. 4. Remove 1 mL of nonsterile PBS containing scraped cells with a 1 mL pipette and place in a 1.5 mL microcentrifuge tube. Centrifuge at 5,000 rpm for 5 min at 4°C. 5. While centrifuging your samples, prepare the following solutions in 1.5 mL microcentrifuge tubes: (a) Tube 1: Cytoplasmic extract buffer supplemented with NP-40: Mix 970 ML of cytoplasmic extract buffer with 25 ML of 10% NP-40 and 5 ML of protease inhibitor cocktail (see Note 5).

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(b) Tube 2: Cytoplasmic wash buffer: Mix 995 ML of cytoplasmic extract buffer with 5 ML of protease inhibitor cocktail. (c) Tube 3: NEB: Mix 995 ML of NEB with 5 ML of protease inhibitor cocktail. 6. After centrifugation, remove the supernatants and add 5 pellet volumes of Tube 1 to each sample. 7. Dissolve cellular pellets by flicking the tubes with your fingers. Try to minimize the amount of bubbles you generate while flicking. You may want to add buffer from Tube 1 to your samples first and let them stand on ice for 1–3 min before resuspending. Make sure cellular pellets are completely dissolved. 8. Incubate the resuspended pellets on ice for 5 min. 9. Centrifuge resuspended pellets at 2,600 rpm for 4 min at 4°C. 10. After centrifugation, carefully pipette off the supernatants and transfer them to new 1.5 mL microcentrifuge tubes. The supernatant contains the cytoplasmic proteins. While removing the supernatant, take special care not to disrupt the nuclear pellet, which now should take on an opaque appearance. The cytoplasmic extract can be either disposed of or stored at −80°C. 11. Add 100 ML of Tube 2 to the nuclear pellets to remove the excess NP-40 from the cell pellets. 12. Gently tap the 1.5 mL microcentrifuge tubes to dislodge the nuclear pellets. There is no need to resuspend the pellets. 13. Repeat centrifugation at 2,600 rpm for 4 min at 4°C. 14. Carefully pipette off and discard the supernatant, once again taking care not to disturb the nuclear pellet. 15. Add 1 pellet volume of Tube 3 to the 1.5 mL microcentrifuge tubes containing the nuclear pellets and resuspend by flicking the tubes until they are completely resuspended. Try to minimize the amount of bubbles you generate while flicking. NF-KB DNAbinding activity is sensitive to salt concentration. Therefore, for larger pellets such as those generated from cultured myotubes, it is important that the final concentration of NaCl remains at 420 mM. If necessary, adjust with addition of stock 5 M NaCl. 16. Incubate the resuspended nuclear pellets on ice for 10 min. During that 10 min incubation period, mix the 1.5 mL microcentrifuge tubes by flicking every 2 min. 17. After the 10 min incubation, centrifuge the resuspended nuclear pellets at 13,000 rpm for 10 min at 4°C. 18. After centrifugation, carefully pipette off the supernatants and transfer them to new prechilled 1.5 mL micro centrifuge tubes. The supernatants contain the nuclear proteins. 19. Store the nuclear extracts at −80°C.

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1. Remove desired skeletal muscles from euthanized animal and place the muscles in 1.5 mL microcentrifuge tubes. 2. Snap freeze the removed skeletal muscles by placing the 1.5 mL microcentrifuge tubes containing the muscles in a small styrofoam container filled with liquid nitrogen for 1 min. After this time, remove the microcentrifuge tubes from the liquid nitrogen, open the tubes briefly, reclose the tubes, and place them on dry ice. If the muscle samples are not going to be immediately used, they can be stored at −80°C indefinitely. 3. To begin the nuclear extract process, homogenize the desired amount of skeletal muscle in low salt lysis buffer in a 2 mL glass homogenizer at the ratio of 9 ML of Low Salt Lysis Buffer to 1 mg of muscle weight. We suggest using a minimum of 20 mg of tissue. 4. Grind the tissues on ice by inserting the glass pestle into the glass tube and rotate the glass pestle 360° 3–6 times until the tissue is fully homogenized. 5. Once homogenized, rinse the sides of the glass homogenizer with the same amount of low salt lysis buffer that was used for the homogenizing process. 6. Transfer the homogenized tissue from the glass homogenizer to a new prechilled 1.5 mL microcentrifuge tube via pipetting. 7. Allow the homogenized tissues in the 1.5 mL microcentrifuge tubes to incubate on ice for 5 min. 8. While the homogenized tissues are incubating, set up an ethanol/dry ice freeze bath by filling a styrofoam container with 0.5 in. of 100% ethanol and add one handful of dry ice. The ethanol will begin to “boil.” The freeze bath is ready to use once the ethanol begins to “simmer.” 9. Place the samples into a microcentrifuge tube floating rack. Freeze the samples by placing them into the ethanol/dry ice freeze bath for 5 min. 10. After 5 min, thaw the samples in a 37°C water bath for 1 min. 11. Vortex the samples at medium speed for 5 s and repeat steps 9 and 10 once. 12. Vortex the samples one last time at medium speed for 5 s and centrifuge the samples at 3,000 rpm for 3 min at 4°C. 13. After centrifugation, carefully pipette off the supernatants and transfer them to new 1.5 mL microcentrifuge tubes. The supernatant contains the cytoplasmic proteins. While removing the supernatant, take special care not to disrupt the nuclear pellet. The cytoplasmic extract can either be disposed of or stored at −80°C.

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14. Resuspend the nuclear pellets in ice cold high salt nuclear buffer at a ratio of 4 ML of high salt buffer per 1 mg of original skeletal muscle weight that was homogenized in Subheading 3.2, step 4. Resuspend the nuclear pellet by flicking the 1.5 mL microcentrifuge tubes. Try to minimize the amount of bubbles generated during the resuspension process. 15. Incubate the resuspended nuclear pellets on ice for 30 min. Flick the resuspended nuclear pellets every 5–10 min during this incubation period. 16. Centrifuge at 13,000 rpm for 5 min at 4°C. 17. After centrifugation, carefully pipette off the supernatants and transfer them to new prechilled 1.5 mL microcentrifuge tubes. The supernatant contains the nuclear proteins. While removing the supernatant, take special care not to disrupt the nuclear membrane and debris pellet. Store the nuclear extracts at −80°C. 3.3. Generating a Radiolabeled Probe for Detecting NF-kB Binding by EMSA

1. In order to detect NF-KB DNA-binding activity via EMSA analysis, we utilize a 37 base pair double-strand oligonucleotide probe containing an NF-KB consensus-binding site derived from the major histocompatibility complex (MHC) class I gene (10). The sequence of the probe is as follows (consensus site is underlined): 5c-CAGGGCTGGGGATTCCC CATCTCCACAGTTTCACTTC-3c. The complementary oligonucleotide strand for our purposes is a 15-mer with the following sequence: 5c-GAAGTGAAACTGTGG-3c. 2. To anneal the 37 and 15-mer oligonucleotides, dilute 236 pmol of the 37-mer, and 2,360 pmol of the 15-mer in a total volume of 98 ML in ultrapure H2O in a 1.5 mL microcentrifuge tube. Add 2 ML of 5 M NaCl to the diluted 37-mer and 15-mer mixture and place 1.5 mL microcentrifuge tubes into a boiling water bath for 2 min. After 2 min, turn off the heat source and allow the water bath to slowly cool down to room temperature (see Note 6). This usually takes 2–3 h. After the water bath has cooled down to room temperature, the reaction is now ready to be used to generate a radiolabeled probe specific to NF-KB binding. 3. To generate the NF-KB radiolabeled probe, add 4 ML of annealed oligonucleotide (see Subheading 3.3, step 2), 4 ML of 10 mM dATP, 4 ML of 10 mM dTTP, 4 ML of 10 mM dGTP, 1 ML of 0.1 mM dCTP, 5 ML of 10× klenow reaction buffer, 10 ML of A-32P dCTP (3,000 Ci/mmol), and 1 ML klenow DNA polymerase (5 units of activity) in 17 ML of ultrapure H2O to a 1.5 mL microcentrifuge tube for a total volume of 50 ML. Incubate labeling reaction at room temperature for 45 min. Quench the labeling reaction by adding 5 ML of 10 mM dCTP and incubate for an additional 15 min at room temperature. 4. During the probe quenching period, prepare a microspin Sephadex G-50 column. Resuspend the beads by gently flicking

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the column. Loosen the cap of the column and snap off the bottom. Place the column in a 1.5 mL microcentrifuge tube and spin for 1 min at 3,000 rpm. The Sephadex should be spun dry. After centrifugation, place the column in a new 1.5 mL microcentrifuge tube. Carefully dispense the labeled probe reaction from Subheading 3.3, step 3 in the center portion of the Sephadex beads. Centrifuge for 2 min at 3,000 rpm. After centrifugation, discard microspin column in an appropriate radioactive waste container and determine the counts per minute of the probe by using 1 ML in a scintillation counter. An efficiently labeled probe is one that has a count of 0.5– 1.0 × 106 cpm/ML. The probe should be stored at −20°C. 3.4. Electrophoretic Mobility Shift Assay 3.4.1. Cast a 5% Nondenaturing Acrylamide Gel

We recommend the Bio-Rad Protean II xi Electrophoresis system or an equivalent apparatus. 1. We utilize the following glass plate dimensions. The inner plate measures 20 × 20 cm × 3 mm, while the outside plate is 22 × 20 cm × 3 mm. The plates should be precleaned with 70% ethanol. 2. Place 1–1.5 mm spacers in between the two glass plates at the left and right most edges and insert the plates into the single screw clamps. Tighten the single screw clamps to create a leak proof seal along the edges of the glass plates and attach to the casting stand. 3. Add 25 mL of ultrapure H2O in between the two glass plates and check for any leaks. If the rig leaks go back to Subheading 3.4.1, step 1 and start again (see Note 7). If the rig shows no signs of leaking, dispense the H2O. 4. Mix 6 mL of 10× TGE, 10.5 mL of 30% acrylamide, 3 mL of 100% Glycerol (see Note 1), 450 ML of 10% APS, and 60 ML of TEMED in 40 mL of ultrapure H2O. 5. Using a 25 mL pipette, dispense the polyacrylamide solution in between the glass plates until the solution has reached the top edge of the shorter glass plate. Place a 1–1.5 mm comb into the casted gel being careful not to introduce air bubbles. 6. The gel will polymerize in approximately 15–20 min.

3.4.2. Preparation of Protein: DNA-Binding Reactions for Nuclear Extracts Isolated from Cellular Pellets

1. Add 5 Mg of nuclear protein to a 1.5 mL microcentrifuge tube. 2. Add NEB to a final volume 5 ML. 3. Add 7 mL of dIdC mix to the 1.5 mL microcentrifuge tubes containing nuclear extract protein and incubate for 10 min. 4. While samples are incubating, remove the labeled probe from the −20°C freezer. 5. After 10 min, add 5 ML of normal 5× binding buffer.

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6. Prepare and add the NF-KB radiolabeled probe by diluting the stock probe with ultrapure H2O to 6,667 cpm/ML. Add 3 ML of diluted probe to each sample and incubate for 20 min at room temperature (the final amount of probe should be 20,000 cpm/sample). 3.4.3. Preparation of Protein: DNA-Binding Reactions for Nuclear Extracts Isolated from Skeletal Muscle

1. Add 10 Mg of nuclear protein to a 1.5 mL microcentrifuge tube. 2. Add high salt nuclear buffer to a final volume of 10 ML. 3. Add 14 ML of dIdC mix to each 1.5 mL microcentrifuge tube containing nuclear extract protein and incubate for 10 min. 4. While the samples are incubating remove the labeled probe from the −20°C freezer. 5. Prepare and add the NF-KB radiolabeled probe by diluting the stock probe with ultrapure H2O to 13,334 cpm/ML. Add 3 ML of diluted probe to each sample and incubate for 20 min at room temperature (the final amount of probe should be 40,000 cpm/sample).

3.4.4. Prepare the Running Unit

1. Place casted 5% acrylamide gel in an appropriate lower and upper electrophoresis chamber (our laboratory uses the BioRad Protean II xi cell running unit). 2. Fill the inner and lower chamber of the unit with 1× TGE (dilute 10× stock solution in ultrapure H2O) running buffer. Carefully remove the comb from the gel and thoroughly clean each well of any remaining polyacrylamide solution.

3.4.5. Loading and Running the Samples

1. Add 2 ML of EMSA loading dye per sample. To verify the purity of the probe, we recommend also setting up a negative control sample containing loading dye, diluted probe, and normal 5× or special 5× binding buffer. 2. Use appropriate long loading tips that fit on a standard P20 pipetman and slowly load each sample into the wells of the casted 5% acrylamide gel. Run at 25 mA for 2–2.5 h. We have found that removing the bromophenol blue and xylene cyanol from the loading dye may improve the resolution of the bands. We will leave it to the researcher to decide which method is more suitable. 3. At the end of the run, separate the glass plates and place a piece of Whatman paper (we use 3 mm chromatography paper) that is slightly larger than the size of the gel onto the EMSA gel. 4. The gel will adhere to the Whatman paper. Starting at the bottom of the gel, carefully remove the gel from the glass plate by slowly pealing the Whatman paper away from the glass plate. 5. Place the gel and Whatman paper onto a gel dryer and dry the gel for approximately 45–60 min (see Note 8).

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Fig. 1. Measuring NF-KB DNA-binding activity by EMSA. (a) Proliferating C2C12 myoblasts were treated with 5 ng/mL of TNFA and nuclear extracts were isolated over a 60-min time course. Five micrograms of nuclear extract per sample were used for EMSA analysis. To verify the purity of the probe, prepare a negative control (NC) sample containing loading dye, diluted probe, and normal 5× binding buffer. The NF-KB and free probe bands are indicated. (b) EMSA analysis was performed with 10 Mg of nuclear extracts isolated from the skeletal muscles of postnatal day P5–P21 wild-type mice. Purity of the probe was verified as detailed in (a) above. 3.4.6. Develop the EMSA

1. After the gel is thoroughly dried, place it in a cassette and expose to either X-ray film at −80°C overnight or a blank phosphorous screen at room temperature overnight. Develop the following day (Fig. 1a, b).

3.5. Confirming Specificity of NF-kB Complexes

A competition EMSA allows you to address the specificity of your radiolabeled probe. To perform a competition EMSA:

3.5.1. Generate “Cold” Nonradiolabeled NF-kBSpecific Probe

1. Set up the following labeling reaction mix: 2. In a 1.5 mL microcentrifuge tube, add 4 ML of annealed oligonucleotide (see Subheading 3.3, step 2), 4 ML of 10 mM dATP, 4 ML of 10 mM dTTP, 4 ML of 10 mM dGTP, 4 ML of 10 mM dCTP, 5 ML of 10× klenow reaction buffer, and 1 ML klenow polymerase (5 units of activity) in 28 ML of ultrapure H2O. The total reaction volume will be 50 ML. 3. Incubate labeling reaction at room temperature for 45 min. 4. Place the “cold” probe in a prepared microspin Sephadex G-50 column (see Subheading 3.3, step 4). 5. Centrifuge for 2 min at 3,000 rpm. 6. After centrifugation, discard microspin column and store the “cold” probe at −20°C.

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Fig. 2. Specificity of NF-KB determined by a competition EMSA. Proliferating C2C12 myoblasts were left untreated (−) or treated (+) with 5 ng/mL of TNFA for 10 min. Nuclear extracts were isolated from cells and 5 Mg of nuclear protein was used to perform EMSA analysis. To verify the specificity of NF-KB binding, extracts were incubated prior to the addition of the radiolabeled probe with either no competitor, or the addition of competitor oligonucleotide, representing 500-fold molar excess of unlabeled wild-type (WT) or mutant (MUT) NF-KBbinding probe, and EMSA analysis was subsequently performed. NF-KB bands are indicated.

3.5.2. Perform a Competition EMSA

Follow the EMSA protocol as described in Subheading 3.4 with the following additions (8): 1. Prior to the addition of the “hot” radiolabeled probe, add 100– 1,000 fold molar excess of “cold” unlabeled probe, prepared in Subheading 3.5, step 1, to the 1.5 mL microcentrifuge tubes containing the 5–10 Mg of isolated nuclear extract samples. 2. Incubate the nuclear extract samples with the “cold” probe for 10 min prior to adding the “hot” probe. 3. Continue with EMSA protocol as described in Subheading 3.4 (Fig. 2).

3.5.3. Supershift Analysis

Supershift analysis allows you to specifically identify which NF-KB subunit complexes are bound to your radiolabeled EMSA probe by preincubating nuclear extracts with an antibody raised against individual NF-KB subunits. This preincubation step will cause your bound radiolabeled probe to run at a higher molecular weight resulting in an upward shift in the visualized band (5–7). A supershift reaction should be performed following the EMSA protocol as described in Subheading 3.4 with the added modifications (Fig. 3a, b): 1. Prior to the addition of the dIdC mixture, add 1.5 Mg of antibody to the 1.5 mL micro centrifuge tubes containing the 5–10 Mg of isolated nuclear extract samples. We have included a list of trusted commercially available NF-KB-specific antibodies,

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Fig. 3. Specificity of NF-KB determined by a supershift EMSA. (a) Supershift analysis was performed on 5 Mg of nuclear extract isolated from proliferating C2C12 myoblasts. The nuclear extract was incubated with antibodies raised against rabbit IgG, p65, or p50. N.C. denotes negative control lane. Shifted complexes along with NF-KB and free probe bands are indicated. (b) Supershift analysis was performed on 10 Mg of P5 skeletal muscle nuclear extract. Nuclear extract was incubated with antibodies raised against rabbit IgG, p65, or p50. N.C. denotes negative control lane. Shifted complexes along with NF-KB and free probe bands are labeled.

which in our hands work well for super shift analysis (see Note 9). As a control, set up an additional reaction sample, which contains nuclear extract incubated with equivalent amounts of isotype-specific IgG. 2. Incubate the nuclear extract samples containing 1.5 Mg of antibody for 10 min. 3. Proceed with the EMSA protocol as described in Subheading 3.4.

4. Notes 1. Make these solutions fresh to ensure optimal cytoplasmic and nuclear extract yields. 2. Use autoclaved 100% glycerol. 3. Sigma APS catalog number A3678. We advise that the 10% APS solution be made fresh on a routine basis. After mixing the APS in ultrapure H2O, you should hear a distinct “popping” noise in the solution. This noise indicates that the APS is of good quality which is important for proper gel polymerization.

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4. Sigma NP-40 catalog number I7771. Dilute stock NP-40 in ultrapure H2O to a final concentration of 10%. This 10% solution should be made up fresh and is stable at room temperature for approximately 1 week. We have noticed that using 10% NP-40 after this time results in lower protein yields. 5. Add freshly made 10% NP-40 to fresh cytoplasmic extract buffer to ensure optimal protein isolation. 6. This 2–3-h cooling down step is essential for proper and efficient annealing of the oligonucleotides. 7. If you are having trouble with the casting rig leaking, place a thin layer of gasket grease on the bottom edge of the glass plates that rest on the rubber gasket of the casting stand. 8. Once the gel is thoroughly dry, it will have a tendency of turning a shade of yellow due to the glycerol. 9. The following is a list of trusted antibodies for supershift REACTIONS AVAILABLE FROM 3ANTA #RUZ "IOTECHNOLOGY  P RelA (SC-109), p50 (SC-114), c-Rel (SC-70 or SC-6955), and Bcl-3 (SC-185). For c-Rel and Bcl-3, we recommend incubating 3 Mg of antibody with 5–10 Mg of NEB for 45 min prior to the addition of the dIdC mixture. We have yet to identify a commercially available antibody capable of shifting p52 or Rel-B. References 1. Fried M, Crothers DM (1981) Equilibria and kinetics of lac repressor-operator interactions by polyacrylamide gel electrophoresis. Nucleic Acids Res 9:6505–6525 2. Garner MM, Revzin A (1981) A gel electrophoresis method for quantifying the binding of proteins to specific DNA regions: application to components of the Escherichia coli lactose operon regulatory system. Nucleic Acids Res 9:3047–3060 3. Revzin A, Ceglarek JA, Garner MM (1986) Comparison of ncleic acid-protein interactions in solution and in polyacrylamide gels. Anal Biochem 153:172–177 4. Chaturvedi MM, Kumar A, Darnay BG, Chainy GB, Agarwal S, Aggarwal BB (1997) Sanguinarine (pseudochelerythrine) is a potent inhibitor of NF-kappaB activation, IkappaBalpha phosphorylation, and degradation. J Biol Chem 272:30129–30134 5. Cheshire JL, Baldwin AS, Jr (1997) Synergistic activation of NF-kappaB by tumor necrosis factor alpha and gamma interferon via enhanced I kappaB alpha degradation and de novo I kappaBbeta degradation. Mol Cell Biol 17:6746–6754

6. Dahlman JM, Bakkar N, He W, Guttridge DC (2010) NF-kappaB functions in stromal fibroblasts to regulate early postnatal muscle development. J Biol Chem 285:5479–5487 7. Dahlman JM, Wang J, Bakkar N, Guttridge DC (2009) The RelA/p65 subunit of NF-kappaB specifically regulates cyclin D1 protein stability: implications for cell cycle withdrawal and skeletal myogenesis. J Cell Biochem 106:42–51 8. Guttridge DC, Albanese C, Reuther JY, Pestell RG, Baldwin AS, Jr (1999) NF-kappaB controls cell growth and differentiation through transcriptional regulation of cyclin D1. Mol Cell Biol 19:5785–5799 9. Kumar A, Lnu S, Malya R, Barron D, Moore J, Corry DB, Boriek AM (2003) Mechanical stretch activates nuclear factor-kappaB, activator protein-1, and mitogen-activated protein kinases in lung parenchyma: implications in asthma. Faseb J 17:1800–1811 10. Mayo MW, Norris JL, Baldwin AS (2001) Ras regulation of NF-kappaB and apoptosis. Methods Enzymol 333:73–87

Chapter 31 Isolation of Nuclei from Skeletal Muscle Satellite Cells and Myofibers for Use in Chromatin Immunoprecipitation Assays Yasuyuki Ohkawa, Chandrashekara Mallappa, Caroline S. Dacwag Vallaster, and Anthony N. Imbalzano Abstract Studies investigating mechanisms controlling gene regulation frequently examine specific DNA sequences using chromatin immunoprecipitation (ChIP) assays to determine whether specific regulatory factors or modified histones are present. While use of primary cells or cell line models for differentiating or differentiated tissue is widespread, the ability to assess factor binding and histone modification in tissue defines the events that occur in vivo and provides corroboration for studies in cultured cells. Many tissues can be analyzed with minimal modification to existing ChIP protocols that are designed for cultured cells; however, some tissues, such as skeletal muscle, are problematic in that accessibility of the cross-linking agent is limited. We describe a method to isolate skeletal muscle tissue nuclei suitable for use in ChIP protocols. Furthermore, we utilize a simple fractionation of digested skeletal muscle tissue that can separate mature myofibers from satellite cells, which are responsible for postnatal skeletal muscle regeneration, thereby allowing simultaneous preparation of nuclei from both cell types. Key words: Skeletal muscle, Satellite cells, Nuclei isolation, Chromatin immunoprecipitation, Brg1, Brahma

1. Introduction In general, the chromatin structure that exists in eukaryotic cells is refractory to processes that utilize the DNA, including transcription, replication, and recombination. In response to this problem, two large families of enzymes that can alter chromatin structure in manners that facilitate access of regulatory factors to the DNA have evolved. These include the ATP-dependent chromatin remodeling enzymes, which utilize the hydrolysis of Joseph X. DiMario (ed.), Myogenesis: Methods and Protocols, Methods in Molecular Biology, vol. 798, DOI 10.1007/978-1-61779-343-1_31, © Springer Science+Business Media, LLC 2012

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ATP to alter histone–DNA contacts (1, 2), and histone modifying enzymes, which posttranslationally modify specific histone residues by acetylation, methylation, phosphorylation, ubiquitylation, ADP-ribosylation, and sumoylation (3–5). The conventional chromatin immunoprecipitation (ChIP) assay is a powerful tool that has allowed investigators to cross-link proteins to DNA, immuno-purify complexes of protein bound to fragmented DNA, and recover the DNA to identify specific DNA sequences that were interacting with the protein of interest in the cell at the time of cross-linking. This assay has been widely used to identify sequences bound by regulatory factors that directly bind DNA, regulatory factors that interact with DNA indirectly through protein–protein interactions, including chromatin remodeling and modifying enzymes, and the structural components of chromatin, including specifically modified histones (6, 7). Typically, the crosslinking agent can be added directly to cell culture media to cross-link proteins to DNA in cultured cells. Protein–DNA interactions in many tissues can be similarly cross-linked by immersion of minced tissue pieces in media containing the cross-linking agent (8). While some investigators have successfully performed ChIP experiments from cross-linked skeletal muscle (9), cross-linking of skeletal muscle tissue is relatively inefficient compared to most other tissues, presumably due to the myofiber structure presenting a physical barrier that limits access of the cross-linking agent to the nuclei. As an alternative, we have determined that brief enzymatic digestion of skeletal muscle and subsequent purification of nuclei from the tissue allow for efficient cross-linking and highly reproducible ChIP results. A further advantage of this methodology is that the enzymatic digestion of skeletal muscle tissue releases satellite cells from between the basal lamina and sarcolemma of myofibers. Satellite cells are the resident muscle stem cell population present in muscle fibers. Upon activation by injury, they exit the microenvironment provided by the myofiber, enter a proliferative stage, migrate to the site of injury, and differentiate (10, 11). Separation of satellite cells from mature myofibers permits analysis of protein–DNA interactions in both populations, which will increase our understanding of similarities and differences in gene activation that occur in these distinct cell types. It is important to note that while satellite cells in noninjured tissue are quiescent, the act of isolating the tissue will cause activation of some of the cells, thus the pool of nuclei that is obtained represents a mixture of activated and quiescent cells. While myogenic regulators of differentiation can therefore be detected in the satellite cell sample as well as in the myofiber sample, assessment of the separation of satellite cells from myofibers can still be monitored by quantification of specific mRNAs from the isolated cells or nuclei, as will be discussed.

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The reader should also note that although the satellite cell pool is devoid of myofibers, it is not a homogenous population of satellite cells. Fibroblasts, in particular, are present in large numbers. Thus, investigations are necessarily restricted to the activation and maintenance of skeletal muscle-specific gene expression. This is because the presence of nuclei from other cell types does not impact the results since the relevant myogenic genes are silent in those nuclei.

2. Materials Prepare reagents using double-distilled deionized water and analytical or molecular biology grade chemicals. 2.1. Isolation and Digestion of Tissue

1. 95% ethanol in a spray bottle. 2. Tray of ice. 3. Plastic wrap. 4. Scalpel, forceps, and scissors suitable for animal dissection. 5. Phosphate-buffered saline (PBS): For 1 L of 1× PBS, add 8 g sodium chloride, 0.2 g potassium chloride, 1.44 g disodium hydrogen phosphate, and 0.24 g potassium dihydrogen phosphate to 800 mL of deionized water. Adjust pH to 7.4 with hydrochloric acid and make up the volume to 1 L with deionized water. 6. Plastic 10 cm petri dishes.

2.2. Separation of Satellite Cells and Myofibers

1. Collagenase type II (Invitrogen). 2. 1 M calcium chloride (CaCl2) stock solution: Autoclave and store at room temperature. 3. Collagenase type II in PBS supplemented with 1 mM CaCl2: Dissolve 10,000 units of collagenase type II in 100 mL of 1× PBS and filter sterilize by passing through a 0.22 Mm filter to obtain 100 units/mL. Store in 5 mL aliquots in −20°C. Add 5 ML of 1 M CaCl2 stock solution to the 5 mL aliquot of collagenase type II before use to generate a final concentration of 1 mM CaCl2. 4. Temperature-controlled incubator/shaker. 5. 70 Mm cell strainer (Becton Dickinson). 6. 50 mL plastic tubes. 7. 14 mL round bottom polypropylene tubes (Falcon). 8. 1.5 mL microcentrifuge tubes. 9. Liquid nitrogen. 10. Dewar flask.

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11. Forceps. 12. 1 M 4-(2-hydroxyethyl)-1-piperazineethanesulfonic acid (HEPES-KOH) (pH 7.3) stock solution: To 700 mL of deionized water, add 238.3 g of HEPES, stir to dissolve, and set the pH to 7.3 using potassium hydroxide crystals. Make up the volume to 1,000 mL using deionized water. Sterilize using a 0.22 Mm filter and store at 4°C. 13. 1 M potassium chloride (KCl) stock solution. 14. 1 M magnesium chloride (MgCl2) stock solution. 15. 1 M dithiothreitol (DTT) stock solution: Store in 0.5 or 1 mL aliquots at −20°C. 16. 0.1 M phenylmethanesulfonylfluoride (PMSF) stock solution: Add 174 mg of PMSF to 10 mL of isopropanol and dissolve by vortexing or rotation on a rotating platform. 17. 10% Nonidet P-40 (NP-40) stock solution. 18. 1 mg/mL leupeptin stock solution: Dissolve 10 mg of leupeptin into 10 mL of deionized water and store in aliquots at −20°C (stable up to 6 months). 19. 3 mg/mL cytochalasin B (Calbiochem) stock solution: Dissolve 30 mg in 10 mL of dimethyl sulfoxide (DMSO) and store as aliquots at −20°C. 20. Lysis buffer (10 mM HEPES-KOH (pH 7.3), 10 mM KCl, 5 mM MgCl2, 0.5 mM DTT, 0.2 mM PMSF, and 10 Mg/mL leupeptin): Prepare 25 mL of lysis buffer by adding 250 ML of 1 M HEPES-KOH, 250 ML of 1 M KCl, 125 ML of 1 M MgCl2, 12.5 ML of 1 M DTT, 50 ML of 100 mM PMSF, 25 ML of 1 mg/mL of leupeptin in 15 mL deionized water and make up the volume to 25 mL with deionized water. Add the DTT, PMSF, and Leupeptin just before use. 21. Lysis buffer plus cytochalsin B (10 mM HEPES-KOH (pH 7.3), 10 mM KCl, 5 mM MgCl2, 0.5 mM DTT, 0.2 mM PMSF, 10 Mg/mL leupeptin, 3 Mg/mL cytochalasin B): Prepare 25 mL of lysis buffer as described in Subheading 2.2, item 20, then add 25 ML of 3 mg/mL cytochalasin B just before use. 22. Lysis buffer +0.1% NP-40: Prepare 25 mL of lysis buffer as described in Subheading 2.2, item 20, except that 250 ML of 10% NP-40 is added prior to the addition of DTT, PMST, and leupeptin. 23. Tissue homogenizer. 24. 10 mL Dounce with pestle A. 25. Hoechst 33258 dye. 26. 1 M Triethanolamine pH 7.5: For 100 mL, add 13.3 mL of triethanolamine (Sigma; the stock is 7.5 M) to 80 mL of deionized

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water and adjust the pH to 7.5 with HCl. Add deionized water to raise the volume to 100 mL. 27. 10-STM buffer (10% sucrose, 10 mM triethanolamine pH 7.5, 5 mM MgCl2, 10 Mg/mL leupeptin): For 500 mL, add 5 mL of 1 M triethanolamine pH 7.5 stock solution, 2.5 mL of 1 M MgCl2, and 50 g sucrose to 300 mL of deionized water and dissolve by stirring. Raise the volume to 500 mL using deionized water. Sterilize using a 0.22 Mm filter and store at 4°C. Just before use, add leupeptin to a final concentration of 10 Mg/mL. 28. 1 M Tris–hydrochloride (Tris–HCl), pH 7.4: For 100 mL, add 12.11 g Tris to 80 mL of deionized water and dissolve using a magnetic stirrer. Adjust the pH to 7.4 with HCl and make up the volume to 100 mL using deionized water. Autoclave and store at room temperature. 29. 2.0 M Sucrose/10 mM Tris–HCl/5 mM MgCl2 buffer: For 500 mL, add 342.3 g of sucrose to 300 mL of water. Dissolve using a magnetic stirrer. Add 5 mL of 1 M Tris–HCl pH 7.4 and 2.5 mL of 1 M MgCl2. Make up the volume to 500 mL using deionized water. Sterilize using a 0.22 Mm filter and store at 4°. 30. Ultracentrifuge with hanging bucket rotor that holds 5 mL ultracentrifuge tubes. 31. 5 mL ultracentrifuge tubes. 2.3. Preparation of Cross-linked Nuclei for Subsequent Use in a Chromatin Immunoprecipitation Assay

1. Formaldehyde. 2. 1 M glycine stock solution. 3. Lysis buffer (see Subheading 2.2, item 20).

3. Methods 3.1. Isolation and Digestion of Tissue

A schematic of the protocol is presented in Fig. 1. The procedure described below is for isolation of upper hindlimb skeletal muscle from young adult (4–8 week old) mice. This procedure should be applicable to any skeletal muscle from mice of any age; however, the number of satellite cells present varies depending upon the specific muscle and the age of the organism (12). 1. Wash the forceps, scissors, and scalpel with detergent. Wrap in aluminum foil or suitable autoclave bag. Autoclave to sterilize. 2. Sacrifice the mice in accordance with your institutionally approved protocol (see Note 1).

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Fig. 1. Schematic of protocol.

3. Spray the mouse with 95% ethanol and place back up on Styrofoam tray on ice to slow autolysis. 4. Lift the dermis with the forceps above the base of the tail (Fig. 2a). 5. Make a small subcutaneous incision with the scissors above the base of the tail (Fig. 2b). 6. Using the scissors, cut along the midline toward the head (Fig. 2c). 7. Use the scissors to cut the dermis perpendicular to the midline and to separate the dermis from the underlying musculature (Fig. 2d). 8. Use the forceps to grasp the dermis and pull away from the back and the limbs to reveal the musculature (Fig. 2e, f). 9. Use the scalpel to dissect skeletal muscle from both hindlimbs. 10. Use the forceps to place skeletal muscle in cold 1× PBS in a petri dish on ice.

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Fig. 2. (a-f) Sequential images of the exposure of hindlimb musculature to facilitate isolation of skeletal muscle.

11. To isolate liver tissue for use as a control, create a V-shaped incision in the parietal peritoneum to reveal the underlying visceral peritoneum (see Note 2). 12. Make an additional V-shaped incision in the visceral peritoneum to expose the liver. 13. Dissect several lobes of the liver and place in a different petri dish containing cold 1× PBS on ice. 14. Using the scalpel and forceps, remove contaminating tissues such as adipose and connective tissues from the skeletal muscle. 15. Using the scalpel and forceps, mince the skeletal muscle to approximately 1 mm3 pieces and transfer to a prechilled 50 mL conical tube on ice. 16. Repeat 3.1 steps 14 and 15 with the control tissue. Transfer to a prechilled 50 mL conical tube and place on ice. Control tissues are not subjected to the collagenase digestion or filtration steps described below (see Subheading 3.2). Mincing is sufficient for subsequent steps. 3.2. Separation of Satellite Cells and Myofibers (see Note 3)

1. Partially digest the minced skeletal muscle tissue by resuspension in 100 units/mL collagenase type II in PBS supplemented with 1 mM CaCl2. 2. Incubate the tissue with agitation at 37°C for 1 h (13, 14) in a temperature-controlled incubator/shaker (see Note 4). 3. Following the collagenase treatment, separate the satellite cells from the mature myofibers by filtration by gravity through a 70 Mm nylon cell strainer into a 50 mL conical tube at room temperature (15). The flow through material is enriched for satellite cells.

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4. To isolate the fraction retained on the membrane (myofibers), turn the filtration unit upside-down and tap over a 50 mL conical tube to facilitate transfer of the myofibers into the tube. 5. Place both tubes on ice along with the control tissue samples. 6. Centrifuge at 300 × g at 4oC for 5 min. 7. Remove the supernatant above the pellet by aspiration or with a pipette. 8. If some of the tissue is to be used for RNA analysis, transfer small aliquots of each tissue sample into 1.5 mL microcentrifuge tubes using a wide-mouth P1000 pipetman tip. Freeze by placing the tubes into a Dewar flask (or other suitable container) with liquid nitrogen (see Notes 5 and 6). Remove the tubes from the flask with forceps and place samples in a −70°C freezer. 9. Estimate the volume of the pellet in each 50 mL conical tube. 10. Resuspend the pellet in 7 volumes of lysis buffer (10 mM HEPES-KOH (pH 7.3), 10 mM KCl, 5 mM MgCl2, 0.5 mM DTT) containing freshly added protease inhibitors (0.2 mM PMSF, and 10 Mg/mL leupeptin). The lysis buffer for the myofiber fraction, but not the satellite cell or control tissue samples, also includes 3 Mg/mL cytochalasin B (see Note 7). 11. Incubate the samples for 30 min on ice. 12. Transfer the samples to 14 mL round bottom tubes. 13. Homogenize the samples using a tissue homogenizer (see Note 8) to disrupt larger fragments of tissue and to create a more homogenous single cell suspension to ease subsequent douncing. Place homogenized sample on ice. 14. Between samples, clean the homogenizer with 95% ethanol, then sonicate in a fresh tube of deionized water. Repeat the water sonications twice more, each time with fresh water. 15. Centrifuge the samples at 3,000 × g for 5 min at 4°C. 16. Estimate the volume of the pellet. 17. Resuspend the pellet in 2.5 volumes of 10-STM buffer (10% sucrose, 10 mM triethanolamine pH 7.5, 5 mM MgCl2, 10 Mg/mL leupeptin). 18. Add twice the original volume of the pellet of 2.0 M sucrose/10 mM Tris–HCl/5 mM MgCl2 and mix the sample by gentle pipeting. 19. Transfer the sample to a prechilled 10 mL Dounce homogenizer by pouring. 20. Dounce homogenize each sample on ice for 7 slow strokes using pestle A to release the nuclei. Move the pestle slowly enough to avoid foaming of the sample solution. Rinse the Dounce homogenizer well with cold 1× PBS between samples.

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21. Check the release and integrity of nuclei under a light microscope after staining of a small aliquot (1–2 ML) with an equal volume of Hoechst 33258 dye; nuclei will stain blue while any remaining unlysed cells will clear the dye and appear unstained. If lysis is insufficient (<90% cells lysed), repeat Subheading 3.2, step 20. 22. Transfer the sample from the Dounce homogenizer to a prechilled 14 mL tube. 23. Prepare the samples for ultracentrifugation. Aliquot 750 ML of the 2.0 M sucrose/10 mM Tris–HCl/5 mM MgCl2 buffer into the bottom of a 5 mL ultracentrifuge tube, and gently overlay with the Dounce homogenized nuclei mixture from Subheading 3.2, step 22. 24. Ultracentrifuge the samples at 116,100 × g for 1 h at 4°C. 25. Aspirate or pipet off the supernatant. 26. Resuspend the pellet in 500 ML of lysis buffer +0.1% NP-40. 27. If desired, an aliquot can be taken for RNA preparation (see Notes 5 and 6). 28. Use 1–2 ML to determine the A260 reading using a spectrophotometer or spectrofluorometer (see Note 1). 3.3. Preparation of Cross-linked Nuclei for Subsequent Use in a Chromatin Immunoprecipitation Assay

1. Cross-link the sample by adding formaldehyde at a final concentration of 1% for 5 min at room temperature (see Note 9). 2. Quench the cross-linking reaction by the addition of glycine to a final concentration of 0.125 M for 5 min at room temperature. 3. Pellet the cross-linked nuclei by centrifugation for 1 min at room temperature at 13,500 × g. 4. Aspirate or remove the supernatant. 5. Freeze the pellet in liquid nitrogen. Store the samples at −70°C for later use or thaw on ice for immediate use. 6. Resuspend the cross-linked nuclei in 400 ML of lysis buffer. 7. We have made use of the Millipore ChIP protocol (http:// www.millipore.com/immunodetection/id3/immunoprecipitationprotocols) to detect cross-linked protein–DNA interactions.

3.4. Examples of Results Achieved with Isolated Satellite Cell, Myofiber, and Liver (Control) Nuclei

Subheading 3.2, steps 8 and 27 indicate that aliquots of sample can be reserved for RNA isolation. We have utilized the Trizol reagent (Sigma) and followed the manufacturer’s directions to isolate RNA, although any standard RNA isolation method can be utilized. We looked for the presence or absence of specific mRNAs to examine the quality of the separation of satellite cells and myofibers.

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Fig. 3. mRNA analysis indicates clean separation of satellite cells from myofibers. mRNA quantification of (a) Pax3, a satellite cell marker, and (b) dystrophin, a marker of mature skeletal muscle tissue, by real-time PCR in satellite cells, myofibers, and liver tissue. Expression levels were normalized to EF-1a mRNA levels, and the relative expression in liver cells was set at 1. A representative experiment is shown.

Pax3 mRNA is expressed in satellite cells but not mature myofibers (16), while the dystrophin gene is inactive in myoblasts but expressed in mature myofibers (17). Experimental analysis of Pax3 and dystrophin mRNA expression in the separated tissue prior to isolation of nuclei (Fig. 3) or following isolation of nuclei (data not shown) revealed clean tissue separation with little to no cross-contamination. We utilized cross-linked satellite cell, myofiber, and liver nuclei for ChIP experiments to examine the interaction of Brahma (Brm) and Brg1 (Brahma-related gene 1) with the promoter sequences controlling the expression of the myogenin gene, which encodes a myogenic regulatory protein that is expressed early in the differentiation process (18, 19). Brg1 and Brm are highly related DNAdependent ATPases that act as the catalytic subunit for the SWI/ SNF family of mammalian chromatin remodeling enzymes (20, 21). These enzymes alter histone–DNA contacts to permit structural alterations in local chromatin structure and are essential for most cellular differentiation programs (2, 22). Previous work has demonstrated that the Brg1 protein is essential for activation of the myogenin gene and for skeletal muscle differentiation (23–25), and that it is likely targeted to the myogenin promoter by a DNA-bound complex containing Pbx/Meis proteins and MyoD (24, 26). Affinity-purified antibodies isolated from polyclonal antisera generated against either the Brg1 or Brm protein (27) were used for the immunoprecipitation of cross-linked chromatin fragments derived from all three nuclei preps (see Note 10). Real-time PCR was used with primers to the myogenin promoter to amplify the

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Fig. 4. Brg1 and Brm bind to the myogenin promoter in skeletal muscle tissues. Representative ChIP experiments performed on nuclei isolated from satellite cells, myofibers, and liver. The data reflect relative Brg1 or Brm recruitment to the myogenin promoter and normalized to input as determined by real-time PCR. Primers (31) spanned −143 to −5 relative to the start site of transcription; this region includes binding sites for TFIID, MyoD, Mef2, and Pbx. Recruitment is relative to that in liver cells, which was assigned a value of 1. Negative controls included immunoprecipitation with preimmune antisera or purified IgG and amplification of the IgH enhancer, where no binding of SWI/SNF proteins is evident (24). Background signals were obtained for all control experiments.

DNA isolated by the ChIP procedure. The results from this experiment are presented in Fig. 4. Brg1 was associated with the myogenin promoter in both satellite cells and in myofibers, which correlates with transcriptional expression of myogenin in these tissues. In contrast, Brg1 was not associated with the myogenin promoter in liver cells, where the gene is transcriptionally inactive. The results for the related ATPase, Brm, were different. Brm was present at the myogenin promoter in myofibers, but not in satellite cells or the control liver cells. These experiments indicate that only Brg1 is present at the myogenin locus in satellite cells and suggest that Brg1 is the relevant SWI/SNF ATPase associated with myogenin activation in these cells. Both Brg1 and Brm were present at the myogenin locus in mature myofiber tissue. Since Brg1 and Brm are mutually exclusive in formation of SWI/SNF enzyme complexes (28); these data suggest that the myogenin promoter in myofiber nuclei is either occupied by multiple SWI/SNF complexes or that individual loci can be occupied by SWI/SNF complexes containing either ATPase. In conclusion, the data clearly demonstrate the feasibility of identifying protein–DNA interactions that occur at specific regions of DNA in satellite cell and myofibers isolated from skeletal muscle tissue.

4. Notes 1. Typically, we have pooled tissue from four mice per isolation to isolate nuclei. We do not count the number of isolated nuclei; instead, we quantify recovery based on a simple A260 reading. An average preparation yields approximately 100 Mg of satellite cell DNA and approximately 600 Mg of myofiber DNA.

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2. Liver tissue is suggested as a control because it is a large tissue and is easy to isolate. We have also utilized brain tissue as well as adipose from various depots as control tissues. Any nonmuscle tissue should act as an adequate control. 3. Separating the satellite cells from the myofibers is an option, but not a requirement. Total skeletal muscle nuclei can be obtained by omitting the separation step. 4. Do not overdigest. If the solution becomes sticky and very viscous, it means that overdigestion has occurred and the isolation should be abandoned. 5. As an alternative to reserving aliquots of tissue for subsequent RNA isolation, aliquots of the purified nuclei (see Subheading 3.2, steps 27) can be utilized for RNA isolation. 6. RNA can be prepared subsequently using any standard RNA isolation procedure. 7. Inclusion of cytochalasin B is important because it inhibits actin polymerization, which promotes actin depolymerization, instability of actin-based filaments, and nuclei release. 8. Settings for the homogenizer will vary by manufacturer and by tissue, so the investigator will need to empirically determine the minimum amount of homogenization required. The objective is to create a single cell suspension. 9. Cross-linking conditions may be optimized by the investigator. Typical ranges for formaldehyde concentration are 0.5–2%. Typical time ranges are 4–20 min, and some investigators perform the cross-linking at 37°C. 10. The nature of ChIP experiments makes the procedure largely dependent upon the quality of the antibody being used. Thus, there is no absolute amount of nuclei necessary for a particular ChIP experiment. We routinely use 10–20 Mg of nuclei for experiments. We suspect that even less can be used; prior experiments with embryonic tissue (29, 30) or nuclei isolated from embryonic tissue (data not shown) has yielded such small quantities that we did not sacrifice any for quantification purposes and instead simply used the samples for the desired ChIP experiments. However, antibodies that work poorly in ChIP experiments where tissue culture cells are the starting material usually also work poorly, if at all, with tissue nuclei, even with larger amounts of starting material. Therefore, we recommend that the investigator determine the minimal amount of starting material empirically.

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Acknowledgments We thank C. Baron for assistance with the figures and OH Cho for manuscript comments. This work was supported by NIH grant GM56244 to ANI and by grants from the Ministry of Education, Culture, Sports, Science and Technology of Japan to YO. References 1. Racki, L. R., and Narlikar, G. J. (2008) ATPdependent chromatin remodeling enzymes: two heads are not better, just different. Curr Opin Genet Dev 18, 137–144. 2. Saha, A., Wittmeyer, J., and Cairns, B. R. (2006) Mechanisms for nucleosome movement by ATP-dependent chromatin remodeling complexes. Results Probl Cell Differ 41, 127–148. 3. Berger, S. L. (2007) The complex language of chromatin regulation during transcription. Nature 447, 407–412. 4. Bhaumik, S. R., Smith, E., and Shilatifard, A. (2007) Covalent modifications of histones during development and disease pathogenesis. Nat Struct Mol Biol 14, 1008–1016. 5. Kouzarides, T. (2007) Chromatin modifications and their function. Cell 128, 693-705. 6. Massie, C. E., and Mills, I. G. (2008) ChIPping away at gene regulation. EMBO Rep 9, 337–343. 7. Minard, M. E., Jain, A. K., and Barton, M. C. (2009) Analysis of epigenetic alterations to chromatin during development. Genesis 47, 559–572. 8. Wells, J., and Farnham, P. J. (2002) Characterizing transcription factor binding sites using formaldehyde crosslinking and immunoprecipitation. Methods 26, 48–56. 9. Mejat, A., Ramond, F., Bassel-Duby, R., Khochbin, S., Olson, E. N., and Schaeffer, L. (2005) Histone deacetylase 9 couples neuronal activity to muscle chromatin acetylation and gene expression. Nat Neurosci 8, 313–321. 10. Le Grand, F., and Rudnicki, M. A. (2007) Skeletal muscle satellite cells and adult myogenesis. Curr Opin Cell Biol 19, 628–633. 11. Shi, X., and Garry, D. J. (2006) Muscle stem cells in development, regeneration, and disease. Genes Dev 20, 1692–1708. 12. Conboy, I. M., and Rando, T. A. (2005) Aging, stem cells and tissue regeneration: lessons from muscle. Cell Cycle 4, 407–410.

13. Asakura, A., Komaki, M., and Rudnicki, M. (2001) Muscle satellite cells are multipotential stem cells that exhibit myogenic, osteogenic, and adipogenic differentiation. Differentiation 68, 245–253. 14. Cornelison, D. D., and Wold, B. J. (1997) Single-cell analysis of regulatory gene expression in quiescent and activated mouse skeletal muscle satellite cells. Dev Biol 191, 270–283. 15. Olguin, H. C., and Olwin, B. B. (2004) Pax-7 up-regulation inhibits myogenesis and cell cycle progression in satellite cells: a potential mechanism for self-renewal. Dev Biol 275, 375–388. 16. Conboy, I. M., and Rando, T. A. (2002) The regulation of Notch signaling controls satellite cell activation and cell fate determination in postnatal myogenesis. Dev Cell 3, 397–409. 17. Lev, A. A., Feener, C. C., Kunkel, L. M., and Brown, R. H., Jr. (1987) Expression of the Duchenne’s muscular dystrophy gene in cultured muscle cells. J Biol Chem 262, 15817–15820. 18. Edmondson, D. G., and Olson, E. N. (1989) A gene with homology to the myc similarity region of MyoD1 is expressed during myogenesis and is sufficient to activate the muscle differentiation program. Genes Dev 3, 628–640. 19. Wright, W. E., Sassoon, D. A., and Lin, V. K. (1989) Myogenin, a factor regulating myogenesis, has a domain homologous to MyoD. Cell 56, 607–617. 20. Khavari, P. A., Peterson, C. L., Tamkun, J. W., Mendel, D. B., and Crabtree, G. R. (1993) BRG1 contains a conserved domain of the SWI2/SNF2 family necessary for normal mitotic growth and transcription. Nature 366, 170–174. 21. Muchardt, C., and Yaniv, M. (1993) A human homologue of Saccharomyces cerevisiae SNF2/ SWI2 and Drosophila brm genes potentiates transcriptional activation by the glucocorticoid receptor. Embo J 12, 4279–4290. 22. de la Serna, I. L., Ohkawa, Y., and Imbalzano, A. N. (2006) Chromatin remodelling in

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Kingston, R. E., and Imbalzano, A. N. (2000) Mammalian SWI-SNF complexes contribute to activation of the hsp70 gene. Mol Cell Biol 20, 2839–2851. Wang, W., Cote, J., Xue, Y., Zhou, S., Khavari, P. A., Biggar, S. R., Muchardt, C., Kalpana, G. V., Goff, S. P., Yaniv, M., Workman, J. L., and Crabtree, G. R. (1996) Purification and biochemical heterogeneity of the mammalian SWI-SNF complex. Embo J 15, 5370–5382. Ohkawa, Y., Marfella, C. G., and Imbalzano, A. N. (2006) Skeletal muscle specification by myogenin and Mef2D via the SWI/SNF ATPase Brg1. Embo J 25, 490–501. Ohkawa, Y., Yoshimura, S., Higashi, C., Marfella, C. G., Dacwag, C. S., Tachibana, T., and Imbalzano, A. N. (2007) Myogenin and the SWI/SNF ATPase Brg1 maintain myogenic gene expression at different stages of skeletal myogenesis. J Biol Chem 282, 6564–6570. Lu, J., McKinsey, T. A., Zhang, C. L., and Olson, E. N. (2000) Regulation of skeletal myogenesis by association of the MEF2 transcription factor with class II histone deacetylases. Mol Cell 6, 233–244.

Chapter 32 An Improved Restriction Enzyme Accessibility Assay for Analyzing Changes in Chromatin Structure in Samples of Limited Cell Number Yasuyuki Ohkawa, Chandrashekara Mallappa, Caroline S. Dacwag Vallaster, and Anthony N. Imbalzano Abstract Studies investigating mechanisms that control gene regulation frequently examine the accessibility of specific DNA sequences to nuclease cleavage. In general, sequences that are sensitive to nuclease cleavage are considered to be in an “open” chromatin conformation that is associated with regulatory factor binding, while sequences resistant to nuclease cleavage are considered to be in a “closed” conformation commonly associated with chromatin that is neither poised for transcription nor being actively transcribed. Changes in nuclease accessibility at specific genomic sequences reflect changes in the local chromatin structure that can occur as a result of signaling cues in the extracellular environment. These changes in chromatin structure usually precede or are coincident with changes in gene expression patterns and are therefore a useful marker of regulatory events controlling transcription. We describe a method to perform restriction enzyme accessibility assays (REAA) that utilizes ligationmediated polymerase chain reaction (LM-PCR) technology and that permits assessment of samples from any source containing as few as 1,000 cells. Use of this modified REAA protocol will enhance analysis of chromatin structural changes at specific DNA sequences of interest by making it possible to analyze samples where unrestricted amounts of sample are not readily available. Key words: Restriction enzyme accessibility assay, Chromatin accessibility, Chromatin remodeling enzymes, Skeletal muscle, Satellite cells

1. Introduction Eukaryotic genomic DNA is packaged around histone octamers to form nucleosomes. Nucleosomes form arrays on the DNA and fold and condense to form filaments and fibers of chromatin, which is the natural state of the genomic DNA in a eukaryotic cell nucleus. However, chromatin structure is not uniform across the nucleus; Joseph X. DiMario (ed.), Myogenesis: Methods and Protocols, Methods in Molecular Biology, vol. 798, DOI 10.1007/978-1-61779-343-1_32, © Springer Science+Business Media, LLC 2012

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indeed it is dynamic and can change as a function of input from a multitude of signaling pathways regulating nearly every biological process and response. In particular, it was recognized many years ago that the structure of those portions of the genome encoding actively transcribed genes was inherently different from genomic regions devoid of genes or containing transcriptionally silent genes. This conclusion was based on the observation that different genomic regions exhibited differential sensitivity to digestion with DNAse I. Specifically, this study demonstrated that the globin genes in nuclei isolated from chicken erythrocytes were more accessible to cleavage by DNAse I than were the globin sequences in nuclei isolated from other cell types in which globin is not expressed (1). Subsequent studies expanded the concept of differential DNAse I accessibility by demonstrating that specific sequences within DNAse I accessible chromatin at active genes were hypersensitive to enzyme cleavage (2, 3) and that these sequences likely corresponded to regulatory regions controlling the expression of the gene being analyzed (4). Work from many laboratories over the following years clearly established the idea that DNAse I hypersensitivity frequently correlated with the location of cis-acting regulatory sequences (reviewed in refs. (5–7)). Restriction endonuclease enzymes have also been used to probe accessibility of genomic DNA, first to analyze satellite DNA sequences (8), and later to also demonstrate that actively transcribed genes are more accessible than nontranscribed genes (9). A significant advantage to the use of restriction enzymes as opposed to DNAse I is that the sequence specificity with which they cleave DNA clearly delineates the site of DNA cleavage. However, the sequence specificity also limits use of the assay to genomic regions near where the restriction enzyme recognition sequence exists. Typically, if the restriction enzyme cleavage site is incorporated into a nucleosome, cleavage is inhibited. In contrast, if nucleosome positioning is altered or nucleosomes are remodeled or otherwise structurally altered, the restriction enzyme cleavage site usually becomes more accessible to the enzyme and results in an increased frequency of cleavage. For these reasons, restriction enzyme accessibility is a useful assay to generally measure changes in chromatin structure at specific gene loci and has been particularly useful in monitoring structural changes caused by chromatin remodeling enzymes. It is important to note that changes in nuclease accessibility do not structurally define the changes that are occurring to genomic chromatin. Instead, they simply indicate that a structural change has occurred. Although some of the initial studies utilized solution hybridization to detect cleaved genomic DNA, the field quickly turned to detection of nuclease cleavage by Southern blot. In this technique, the nuclease-digested chromatin is deproteinated and purified, then subjected to restriction enzyme cleavage at sites flanking the sequence of interest to create a restriction fragment of known size.

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If cleavage of genomic DNA at the sequence of interest occurred, some fraction of the specific restriction fragment will be reduced in size. When the sample is separated on an agarose gel, transferred to a membrane and then hybridized with a suitably labeled probe to the sequences of interest; detection of the cleaved restriction fragment reflects the extent of nuclease accessibility, and by extension, the extent of change to the local chromatin structure. The advent of PCR-based technologies led to the development of ligation-mediated polymerase chain reaction (LM-PCR) as a means to greatly increase the sensitivity of detecting cleaved genomic DNA (10). Genomic DNA cleaved by enzymatic or chemical methods is deproteinated and processed as necessary to prepare double-stranded fragments suitable for ligating to a defined linker DNA (11). Amplification of the cleaved genomic fragment can then be accomplished by PCR using one primer to the linker sequence and a second primer that hybridizes to the genomic DNA of interest near the cleavage site. Because of the ability to PCR amplify from small amounts of starting material, this method greatly increases the sensitivity of detection. Thus, it is ideal for analyzing chromatin accessibility when starting with limiting amounts of material. More detailed explanation of LM-PCR use to measure regions of genomic DNA cleaved by restriction endonucleases is presented below.

2. Materials 2.1. Isolation of Nuclei

1. Phosphate-Buffered Saline (PBS) pH 7.4: For 1 L of 1× PBS, add 8 g sodium chloride, 0.2 g potassium chloride, 1.44 g disodium hydrogen phosphate, and 0.24 g potassium dihydrogen phosphate to 800 mL deionized water. Adjust pH to 7.4 with hydrochloric acid and make up the volume to 1 L with deionized water. 2. Rubber policeman or cell scraper. 3. 1 M Piperazine-N,N c-bis(2-ethane-sulfonic acid) (PIPES) pH 8.0 stock solution. 4. 1 M Potassium chloride (KCl) stock solution. 5. 1 M Calcium chloride (CaCl2) stock solution. 6. 10% Nonidet P-40 (NP-40) stock solution. 7. 1 mg/mL Leupeptin stock solution: Dissolve 10 mg of leupeptin into 10 mL of deionized water and store in aliquots at −20°C (stable up to 6 months). 8. 0.1 M Phenylmethanesulfonylfluoride (PMSF) stock solution: Add 174 mg of PMSF to 10 mL of isopropanol and dissolve by vortexing or rotation on a rotating platform.

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9. Tissue culture cell (TCC) lysis buffer (10 mM PIPES pH 8, 85 mM KCl, 1 mM CaCl2, 5% sucrose, 0.5% NP-40, plus protease inhibitors added immediately before use, including 1 Pg/mL leupeptin, and 1 mM PMSF): Prepare 500 mL of tcc buffer by dissolving 25 g sucrose in 400 mL of deionized water using a magnetic stirrer. Then add 5 mL of 1 M PIPES pH 8.0 stock solution, 42.5 mL of 1 M KCl stock solution, 0.5 mL of 1 M CaCl2 stock solution, and 25 mL of 10% NP-40 stock solution. Bring the volume to 500 mL. Immediately before use, aliquot the desired amount and add the protease inhibitors. 10. 10 mL Dounce homogenizer with Pestle A. 11. Hoechst 33258 dye. 2.2. Restriction Enzyme Digestion and Ligation of Linker to Cleaved DNA

1. Restriction enzyme.

2.3. Preparation of Linker and Ligation of Linker to Cleaved DNA

1. 1 M Tris–hydrochloride (Tris–HCl), pH 7.5 stock solution: For 1 L, add 121.1 g Tris to 800 mL of deionized water and dissolve using a magnetic stirrer. Adjust the pH to 7.5 with HCl and make up the volume to 1,000 mL using deionized water. Autoclave and store at room temperature.

2. Appropriate 10× restriction enzyme reaction buffer. 3. DNeasy purification kit (Qiagen).

2. 0.5 M Ethylenediaminetetraacetic acid (EDTA) stock solution: Add 186.12 g EDTA to 800 mL of deionized water. Stir with a magnetic stirrer to begin to dissolve. Add NaOH pellets slowly to adjust the pH to 8.0. Note that EDTA will not fully dissolve unless the pH is near 8.0. Make up the volume to 1,000 mL using deionized water, autoclave and store at room temperature. 3. 10× Annealing buffer (100 mM Tris, pH 7.5, 500 mM NaCl, 10 mM EDTA): For 50 mL, dissolve 0.58 g NaCl in 40 mL of deionized water. Add 5 mL of 1 M Tris–HCl pH 7.5 stock solution and 1 mL of 0.5 M EDTA stock solution. Bring the volume to 50 mL with deionized water. Filter sterilize through a 0.22 Pm filter. 4. Mighty Mix Ligation kit (Takara). 5. RNAse/DNAse-free water.

3. Methods 3.1. Isolation of Nuclei

A detailed method for the isolation of nuclei from primary tissues is presented in the accompanying report (see Chapter 31). The same protocol can be used for isolation of nuclei from tissue culture

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cells, but we have found that gradient purification of the nuclei is not generally necessary. A slightly different method to isolate nuclei from a 100 mm plate of adherent tissue culture cells follows (see Notes 1 and 2). 1. Place the cell culture dishes containing the cells to be isolated on a tray of ice. 2. Aspirate or remove media covering the cells. 3. Wash the cells twice by slowly pouring or pipeting 5–10 mL of cold PBS onto the cells (see Note 3). 4. Add 1 mL of cold PBS to the cells. 5. Scrape cells off the plate with a rubber policeman or with a cell scraper. Hold the plate on an angle and use the scraper to push the cells and media to the bottom of the dish to facilitate collection. 6. Use a pipet or Pipetman to collect the cells in PBS and transfer to a prechilled 1.5 mL microcentrifuge tube. 7. Pellet by centrifugation at 300 × g for 2 min at 4°C. 8. Resuspend pellet in 500 PL of tcc lysis buffer. 9. Transfer suspension to a prechilled 10 mL Dounce homogenizer. 10. Lyse the cells by using a Dounce homogenizer with pestle A for seven to eight slow strokes on ice, taking care not to cause foaming. The optimum number of strokes may need to be determined empirically. 11. Check the release and integrity of nuclei under a light microscope after staining of a small aliquot (1–2 PL) with an equal volume of Hoechst 33258 dye; nuclei will stain blue while any remaining unlysed cells will clear the dye and appear unstained. If lysis is insufficient, repeat step 10. 12. Collect the resulting nuclei solution in a fresh, prechilled microcentrifuge tube. 13. Use 1–2 PL to determine the A260 reading using a spectrophotometer or spectrofluorometer. 3.2. Restriction Enzyme Digestion

The choice of restriction enzyme will be specific to the sequences of interest. Generally, a site at or within a few hundred base pairs of the sequence of interest should provide an accurate reflection of changes in local chromatin accessibility. Restriction enzymes inhibited by CpG methylation should be avoided. If multiple genomic sequences are of interest but do not share common restriction enzyme sites, multiple enzymes can be combined for genomic DNA digestion. The same considerations that exist for combining restriction enzymes to cleave naked DNA apply to genomic DNA digestion, including reaction components and total amount of

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enzyme added to the reaction. In practice, we have had success combining different restriction enzymes that will work in similar reaction conditions. Sequential digests are an alternate option. 1. Aliquot 100 Pg of nuclei into a 1.5 mL microcentrifuge tube (see Notes 4 and 5). 2. Add the appropriate restriction enzyme 10× buffer, water, and the restriction enzyme of choice at 0.5 Units/Pg nuclei. The reaction size should be approximately 100 PL and should not exceed 150 PL. 3. Digest for no more than 1 h at 37°C or the suggested optimum temperature for the restriction enzyme of choice (see Note 6). 4. Stop the digestion using the conditions recommended by the restriction enzyme manufacturer (see Note 7). 5. Purify the DNA using the DNeasy purification kit (Qiagen) in accordance with the manufacturer’s instructions. Elute the samples in 40 PL of elution buffer (supplied as part of the kit). 3.3. Preparation of Linker and Ligation of Linker to Cleaved DNA

We have frequently utilized the linkers LM-PCR1 and LM-PCR2 (11), which when annealed provide a linker suitable for ligation to genomic DNA cleaved with a blunt end cutter (see Fig. 1a; includes linker sequence). If the genomic DNA was cleaved with an enzyme leaving a 5c or 3c recessed end, the linker design can be modified to incorporate the bases complimentary to the existing overhang (see Fig. 1b). This strategy permits use of restriction enzymes that leave either a 3c or 5c recessed end.

Fig. 1. Schematic of alternate possibilities for linker ligation to restriction enzyme-cleaved genomic DNA. (a) ligation to blunt-ended genomic DNA. (b) ligation to genomic DNA with 3`- or 5`-recessed ends.

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1. To prepare the linker, mix 20 PL of each single stranded oligonucleotide from 100 pmol/PL stock in 1× annealing buffer in a 50 PL reaction volume, heat to 95°C for 5 min, incubate at 75°C for 15 min, followed by incubation at 35°C for 45 min, and cooling to 4°C to obtain 40 PM of double-stranded linker (see Notes 8 and 9). 2. Ligate 1 Pg of purified digested DNA to 2.5 PL of 40 PM double-stranded linker oligonucleotide using the Mighty Mix Ligation kit (Takara) reagents as instructed by the manufacturer. The ligation reaction volume ideally should be 10 PL and should not exceed 20 PL (see Note 10). 3. Dilute the ligation reaction to 40–60 PL using RNAse/ DNAse-free water. The sample may be stored at −20°C. 3.4. PCR Detection of Cleaved DNA

The standard amplification protocol utilizes a primer to the linker and a second primer upstream or downstream of the anticipated cleavage site (see Fig. 2; P1a and P2). This permits detection of the cleaved product, thereby providing direct measurement of increased enzyme accessibility at the genomic site examined. However, because the linker has been ligated onto every cleaved genomic DNA fragment, there is potential for increased background signal in the PCR reaction. One method to decrease background and improve specificity is to design the linker primer to span the linker sequence and the genomic sequence that is predicted to adjoin the linker (see Fig. 2; P1b and P2). Theoretically, this decreases the number of genomic fragments to which the primer can hybridize to the single fragment of interest. We have found this to be particularly helpful when performing experiments with limited amounts of starting material. Real-time PCR as a detection method also greatly improves the sensitivity of the assay as compared to

Fig. 2. Schematic of ligation-mediated polymerase chain reaction (LM-PCR) protocol. P1a, P1b, P2, P3, and P4 represent primers that are described in the text.

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conventional PCR, since this method allows quantification of the PCR product during the amplification stages of the reaction. Typically, a sequence upstream or downstream of the cleavage site of interest is amplified as a control for quantitative comparison of different samples in a particular experiment (see Fig. 2; P3 and P4). Alternatively, the quantity of a cleaved genomic fragment can be compared to input by using primers that flank the cleavage site of interest for amplification of undigested genomic DNA. 1. Prepare the following reaction mixture on ice for real-time q-PCR using an ABI StepOne Plus (see Note 11). For a 20 PL reaction volume add; L

L

L

2 PL of Ligated DNA from Subheading 3.3, step 3. 1 PL of Forward primer from 4 pmol/PL stock (200 nM final concentration). 1 PL of Reverse primer from 4 pmol/PL stock (200 nM final concentration) (see Note 12).

L

10 PL of ABI Fast SYBR green master mix.

L

6 PL of Sterile deionized water.

2. Analyze the data using delta delta Ct method (12). Most realtime PCR machine manuals will also have detailed explanations for the analysis of raw data by this method. For normalization, use the amplification values obtained from the upstream primer pairs, P3 and P4 (see Fig. 2). 3.5. Detection of Changes in Restriction Enzyme Accessibility in as Few as 1,000 Cells

An example of this methodology is presented in Fig. 3. Our laboratories have a longstanding interest in the mammalian SWI/SNF complexes, which are ATP-dependent chromatin remodeling enzymes that break histone:DNA contacts that facilitate both nucleosome mobility as well as regulatory factor interactions with chromatin (13, 14). Mammalian SWI/SNF complexes contain one of two highly related ATPase subunits called Brahma (Brm) or Brahma-related gene 1 (Brg1) (15). To identify biological roles for SWI/SNF enzymes, we created fibroblast cell lines that express ATPase-deficient, dominant negative Brm or ATPase-deficient, dominant negative Brg1 in a tetracycline repressible manner (16). The dominant negative form of the protein interacts with the other subunits of the complex, but is functionally inactive (16, 17). Thus, the expression of genes that require functional SWI/SNF enzymes is inhibited in cells expressing the mutant ATPase. Fibroblasts can be reprogrammed along numerous different lineages, including skeletal muscle. Introduction of the MyoD regulator followed by serum withdrawal promotes skeletal muscle differentiation (18). We previously used this experimental system to show that expression of dominant negative Brm or dominant negative Brg1 inhibits skeletal muscle-specific gene expression and differentiation because the chromatin at muscle-specific gene promoters

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Fig. 3. Restriction enzyme accessibility assays (REAA) using as few as 1,000 cells. Restriction enzyme accessibility of Pvu II at the promoter of a muscle-specific (desmin) or fat-specific (aP2) gene in differentiated muscle cells in the presence (−tet) or absence (+tet) of a dominant negative version of Brahma-related gene 1 (Brg1) was measured using tenfold serial dilutions of isolated nuclei. Accessibility was observed in the muscle-, but not the fat-specific promoter in a Brg1 dependent manner, as expected in a muscle cell. The amount of cleavage at each promoter site in the cells expressing dominant negative Brg1 was set to 1; the amount of cleavage in cells not expressing dominant negative Brg1 is expressed relative to that number. Primers to amplify desmin regulatory sequences were described (19). Primers for the aP2 promoter were 5c caaaatgtgtgatgcctttgtgg 3c and 5c tccatgcgggaagcagcattttc 3c.

is not remodeled (19–22). In the experiment presented in Fig. 3, cells were cultured in the presence or absence of tetracycline, infected with a retrovirus expressing MyoD, and subsequently differentiated. Nuclei were isolated from these cells, and side-byside enzymatic digestion reactions containing 105, 104, or 103 nuclei were performed. We used Pvu II for digestion because its recognition sequence matches the consensus sequence for E boxes, which are binding sites for MyoD- and MyoD-related myogenic factors. Thus, this experiment measures accessibility at muscle-specific genes in the regions where MyoD may be binding. As shown in Fig. 3, cells maintained in tetracycline, which do not express dominant negative Brg1, show a robust increase in Pvu II accessibility in the promoter of the muscle-specific desmin gene. In contrast, in cells cultured in the absence of tetracycline, where the dominant negative Brg1 enzyme was expressed, there was much less accessibility, resulting in greatly reduced levels of cleavage. This implicates functional SWI/SNF enzyme in the creation of the open chromatin structure that could be cleaved by the restriction enzyme. As a control, there were no changes in chromatin accessibility at the adipogenic aP2 gene, which contains a Pvu II site in its promoter but is not expressed in the MyoD differentiated cells. Importantly, the same results were obtained whether the reaction contained 100,000 nuclei or 1,000 nuclei.

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This demonstrates the exquisite sensitivity of this assay and validates use of this approach with samples where large quantities of starting material are not available. We have made use of this approach for measuring gene-specific changes in chromatin structure in embryonic mouse limb buds and embryonic mouse limb skeletal muscle (19, 23). This technique should be applicable to any tissue from which the nuclei can be isolated.

4. Notes 1. Volumes should be scaled proportionally for larger or smaller size samples. 2. For tissue culture cells grown in suspension, the first step is to pellet the cells by centrifugation. Cells should be poured or pipeted into 15 mL conical tubes and centrifuged at 300 × g. Modifications to the centrifugation may be necessary based on the cell type being used. 3. For cell types that do not adhere strongly to the culture dish surface, do not pour the PBS onto the cells. Instead, tilt the dish with one hand, and with the other, slowly expel 5–10 mL of cold PBS from a pipet along the side of the culture dish, not directly on top of the cells. 4. When cell lines or commonly available primary tissue are used as starting materials, 100 Pg of nuclei is a reasonable starting point for restriction enzyme digestion. However, if sample size is limiting, 10–20 Pg of nuclei can be utilized. 5. We recommend starting with trial experiments using commonly available nuclei and making dilutions of this material prior to initiating work with samples where availability and/or amount are restricted. 6. Limited digestion to optimize differences between accessible and inaccessible chromatin is essential; overdigestion will minimize accessibility differences. Some empirical determination of enzyme concentration and/or time of digestion may be necessary. 7. Although the digestion reactions can be stopped using conditions recommended by the enzyme supplier for inactivation, we do not find this to be a necessary step. Instead, we proceed directly to DNA purification as described in Subheading 3.2, step 5. 8. Preparation of the double-stranded linker should occur well in advance of the procedure. Large quantities can be prepared, aliquoted, and stored at −20°C. 9. An alternative annealing method is to place the linker mixture on the bench following the 95°C incubation and allow to cool to room temperature.

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10. We have successfully utilized different ligation times at different temperatures. Ligations can be performed in as little as 30 min at 25°C. Alternatively, 4 h at 16°C or overnight at 4°C can be used. 11. The suggested protocol is optimized for the ABI StepOne Plus real-time q-PCR machine. Any real-time q-PCR machine is adequate; the reader will need to modify the protocol and reagents in accordance with the requirements of the machine to be used. 12. One of the primers should amplify the linker sequence or the linker sequence and adjacent sequence as discussed in Subheading 3.4.

Acknowledgments We thank C. Baron for assistance with the figures. This work was supported by NIH grant GM56244 to ANI and by grants from the Ministry of Education, Culture, Sports, Science and Technology of Japan to YO. References 1. Weintraub, H., and Groudine, M. (1976) Chromosomal subunits in active genes have an altered conformation. Science 193, 848–856. 2. Wu, C., Wong, Y. C., and Elgin, S. C. (1979) The chromatin structure of specific genes: II. Disruption of chromatin structure during gene activity. Cell 16, 807–814. 3. Wu, C., Bingham, P. M., Livak, K. J., Holmgren, R., and Elgin, S. C. (1979) The chromatin structure of specific genes: I. Evidence for higher order domains of defined DNA sequence. Cell 16, 797–806. 4. Wu, C. (1980) The 5c ends of Drosophila heat shock genes in chromatin are hypersensitive to DNase I. Nature 286, 854–860. 5. Reeves, R. (1984) Transcriptionally active chromatin. Biochim Biophys Acta 782, 343–393. 6. Gross, D. S., and Garrard, W. T. (1988) Nuclease hypersensitive sites in chromatin. Annu Rev Biochem 57, 159–197. 7. Elgin, S. C. (1981) DNAase I-hypersensitive sites of chromatin. Cell 27, 413–415. 8. Lipchitz, L., and Axel, R. (1976) Restriction endonuclease cleavage of satellite DNA in intact bovine nuclei. Cell 9, 355–364. 9. Pfeiffer, W., and Zachau, H. G. (1980) Accessibility of expressed and non-expressed genes to a restriction nuclease. Nucleic Acids Res 8, 4621–4638.

10. Mueller, P. R., Salser, S. J., and Wold, B. (1988) Constitutive and metal-inducible protein:DNA interactions at the mouse metallothionein I promoter examined by in vivo and in vitro footprinting. Genes Dev 2, 412–427. 11. Carey, M., and Smale, S. T. (2000) Transcriptional Regulation in Eukaryotes. Cold Spring Harbor Laboratory Press, Cold Spring Harbor, NY. 12. Livak, K. J., and Schmittgen, T. D. (2001) Analysis of relative gene expression data using real-time quantitative PCR and the 2(-Delta Delta C(T)) Method. Methods 25, 402–408. 13. Imbalzano, A. N., Kwon, H., Green, M. R., and Kingston, R. E. (1994) Facilitated binding of TATA-binding protein to nucleosomal DNA. Nature 370, 481–485. 14. Kwon, H., Imbalzano, A. N., Khavari, P. A., Kingston, R. E., and Green, M. R. (1994) Nucleosome disruption and enhancement of activator binding by a human SW1/SNF complex. Nature 370, 477–481. 15. Wang, W., Cote, J., Xue, Y., Zhou, S., Khavari, P. A., Biggar, S. R., Muchardt, C., Kalpana, G. V., Goff, S. P., Yaniv, M., Workman, J. L., and Crabtree, G. R. (1996) Purification and biochemical heterogeneity of the mammalian SWISNF complex. EMBO J 15, 5370–5382. 16. de La Serna, I. L., Carlson, K. A., Hill, D. A., Guidi, C. J., Stephenson, R. O., Sif, S.,

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Kingston, R. E., and Imbalzano, A. N. (2000) Mammalian SWI-SNF complexes contribute to activation of the hsp70 gene. Mol Cell Biol 20, 2839–2851. 17. Sif, S., Saurin, A. J., Imbalzano, A. N., and Kingston, R. E. (2001) Purification and characterization of mSin3A-containing Brg1 and hBrm chromatin remodeling complexes. Genes Dev 15, 603–618. 18. Davis, R. L., Weintraub, H., and Lassar, A. B. (1987) Expression of a single transfected cDNA converts fibroblasts to myoblasts. Cell 51, 987–1000. 19. Ohkawa, Y., Marfella, C. G., and Imbalzano, A. N. (2006) Skeletal muscle specification by myogenin and Mef2D via the SWI/SNF ATPase Brg1. EMBO J 25, 490–501. 20. de la Serna, I. L., Ohkawa, Y., Berkes, C. A., Bergstrom, D. A., Dacwag, C. S., Tapscott, S. J., and Imbalzano, A. N. (2005) MyoD targets

chromatin remodeling complexes to the myogenin locus prior to forming a stable DNA-bound complex. Mol Cell Biol 25, 3997–4009. 21. de la Serna, I. L., Carlson, K. A., and Imbalzano, A. N. (2001) Mammalian SWI/SNF complexes promote MyoD-mediated muscle differentiation. Nat Genet 27, 187–190. 22. Mallappa, C., Nasipak, B. T., Etheridge, L., Androphy, E. J., Jones, S. N., Sagerstrom, C. G., Ohkawa, Y., and Imbalzano, A. N. (2010) Myogenic microRNA expression requires ATPdependent chromatin remodeling enzyme function. Mol Cell Biol 30, 3176–3186. 23. Ohkawa, Y., Yoshimura, S., Higashi, C., Marfella, C. G., Dacwag, C. S., Tachibana, T., and Imbalzano, A. N. (2007) Myogenin and the SWI/SNF ATPase Brg1 maintain myogenic gene expression at different stages of skeletal myogenesis. J Biol Chem 282, 6564–6570.

Chapter 33 ChIP-Enriched in Silico Targets (ChEST), a ChIP-on-Chip Approach Applied to Analyzing Skeletal Muscle Genes Guillaume Junion and Krzysztof Jagla Abstract Mapping the cis-regulatory modules (CRMs) to which bind myogenic transcription factors is an obligatory step towards understanding gene regulatory networks governing muscle development and function. This can be achieved in silico or by chromatin immunoprecipitation (ChIP) approaches. We have developed a ChIP-enriched in silico targets (ChEST) strategy designed for mapping the CRMs by combining in silico and ChIP methods. ChEST involves a software-assisted prediction of transcription factor (TF) – specific CRMs, which are spotted to produce a computed genomic CRM microarray. In parallel, the in vivo pool of targets of a given TF is isolated by ChIP and used as a probe for hybridization with the array generated. Here we describe ChEST strategy applied to identify direct targets of Myogenic Enhancer Factor, Dmef2 in Drosophila embryos. Key words: ChIP, ChEST, Transcription factor, Cis regulatory module, Microarray, Drosophila

1. Introduction Gene activity is controlled by the dynamic association of transcription factors and chromatin modifiers with target DNA sequences. Over the last decade, different chromatin immunoprecipitation-based methods have been developed and widely used to map the interactions of regulatory proteins with their cognate target sequences (for review see (1)). In the last few years, an important effort has been made to reduce the amount of starting material necessary to perform chromatin immunoprecipitation (2, 3) and in particular to shorten the protocols (4). Fast ChIP assay has introduced two modifications. First, incubation of antibodies with chromatin in an ultrasonic bath, which increases the rate of antibody-protein binding and allows to shorten incubation time from several hours to 15 min.

Joseph X. Dimario (ed.), Myogenesis: Methods and protocols, Methods in Molecular Biology, vol. 798, DOI 10.1007/978-1-61779-343-1_33, © Springer Science+Business Media, LLC 2012

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Second, applying a cation-chelating resin (Chelex 100)-based DNA instead of traditional reversal of cross-linking and proteinase K digestion, which reduces the total time for preparation of PCRready templates from 1 day to 1 h. These modifications can be applied to the protocol we present below. Independently, a set of computational tools has been made available to predict cis-regulatory modules (CRMs) in large and complex genomes (5–9). In the ChIPenriched in silico target (ChEST) strategy described here, we took advantage of genome scanning tools and annotations available for the fruit fly genome and demonstrate that this cost-effective method allows rapid identification of in vivo TF targets in Drosophila. Using the ChEST approach, we uncovered a large set of motifs to which binds the key myogenic factor Dmef2 (10), but also a set of functional enhancers recognized by the transcription factor Ladybird required for specification of a subset of Drosophila muscles (11). We anticipate that the ChEST strategy originally designed to identify regulatory modules in Drosophila can be adapted for any sequenced and annotated genome. Actually, ChIP followed by the secondgeneration sequencing (ChIPseq) (12) has become the most sensitive and unbiased approach for mapping regulatory modules. However, ChEST remains an attractive alternative to ChIP followed by the Q-PCR and an interesting approach for a pilot CRM screen and/or for optimization of the ChIP-on-chip protocols. As our knowledge about the bona fide enhancers remains still limited, the ChIP-identified as well as in silico defined CRMs need to be validated by reporter gene transgenesis in an in vivo context.

2. Materials All solutions are prepared using ultrapure water. Analytic grade reagents are used as solution’s components. Population cages with about 2,000 flies are used to collect Drosophila embryos. Prediction of regulatory modules to which bind a given TF is done using FlyEnhancer (http://www.opengenomics.org) (see Note 1) and filtered with the use of custom-made SCAFI software (available via http://www.soluscience.fr/dup_soft/). For each in silico selected CRM, at least two different long oligonucleotides (60 nt) are designed, synthesized, and printed to produce a computed CRM’s microarray (see Note 2). 2.1. Cross-linking and ChIP Components

1. Embryo Wash Buffer (PBT): 1 × PBS containing 0.01% Triton X100 (see Note 3). Prepare up to 500 mL and store at room temperature. 2. Cross-linking Buffer: 1.8% formaldehyde, 50 mM Hepes, 1 mM EDTA, 0.5 mM EGTA, 100 mM NaCl pH8.0, and 30 mL of n-heptane. Prepare up to 100 mL. Store at 4°C. Make sure it is at room temperature before use.

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3. Stop Cross-link Buffer: PBT containing 0.125 M glycine. Prepare up to 100 mL immediately before use (see Note 4). 4. Homogenization Solution 1: 0.3 M sucrose, 10 mM Hepes, 10 mM KCl, 1.5 mM MgCl2, 0.1 mM EGTA, 0.5 mM DTT, 2 mM AEBSF, 1 mg/mL leupeptin. Prepare 50 mL and keep at 4°C. 5. Homogenization Solution 2: 10 mM Hepes, 10 mM KCl, 2 mM MgCl2, 0.5 mM DTT, 2 mM AEBSF, 1 mg/mL leupeptin. Prepare 10 mL and store at 4°C. 6. Sonication buffer: 50 mM Hepes, 140 mM NaCl, 1 mM EDTA, 1% Triton X100, 0.1% sodium deoxycholate, 0.1% SDS. Prepare 20 mL and store at 4°C. 7. IP buffer: 140 mM NaCl, 10 mM Tris–HCl pH8.0, 1 mM EDTA, 1% Triton X100, 0.1% SDS, 0.1% sodium deoxycholate, 1 mM PMSF (see Note 5). Prepare up to 20 mL and store at room temperature. 8. IP washing solution 1: IP buffer with 500 mM NaCl instead of 140 mM NaCl. Prepare up to 20 mL and store at room temperature. 9. IP washing solution 2: 250 mM LiCl, 10 mM Tris–HCl pH 8.0, 1 mM EDTA, 0.5% NP-40, 0.5% sodium deoxycholate. Prepare up to 20 mL and store at room temperature. 10. TE buffer: 10 mM Tris–HCl pH 8.0, 1 mM EDTA. Prepare up to 100 mL and store at room temperature. 11. Sepharose A beads: Use 30 mL of Protein A Sepharose CL4B (Sigma) for each immunoprecipitation reaction (see Note 6). 2.2. Purification and Amplification of Immunoprecipitated DNA

1. 5 mg/mL RNAse A (DNase free). 2. 10% SDS. 3. 10 mg/mL proteinase K (stored at −20°C). 4. 1.5 mL Phase Lock tubes. 5. Two primers: Phosphoprimer 1: phosphorylated at 5¢-end (see Note 7) 24-mer AGA AGC TTG AAT TCG AGC AGT CAG Primer 2: 20-mer CTG CTC GAA TTC AAG CTT CT These primers are used for preparing PCR linkers (see Note 8). 6. 10× ligation buffer: 0.5 M Tris–HCl pH 7.6, 125 mM MgCl2, 250 mM DTT, 12.5 mM ATP 7. T4 DNA ligase, 4 U/mL 8. Amplitaq Polymerase and buffer (Roche). 9. 10 mM dGTP, dATP, dCTP, dTTP (diluted from commercially available stocks in 10 mM Tris–HCl pH 8.0, and stored in small aliquots at −20°C).

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10. Qiagen PCR purification columns. 11. Nanodrop spectrometer. 2.3. Probe Synthesis and Hybridization Solutions

1. Random prime labeling kit (Invitrogen). 2. Cy3 and/or Cy5-dUTP. 3. Corning hybridization chambers. 4. Hybridization solutions: 20 × SSC, 10% SDS, 50× Denhardt’s solution, 10 mg/mL polyA, formamide (see Note 9). 5. Chamber humidifying solution: 3 × SSC. Prepare 50 mL. 6. Hybridization wash solution 1: 1 × SSC, 0.03% SDS. Prepare 250 mL. 7. Hybridization wash solution 2: 0.2 × SSC. Prepare 250 mL. 8. Hybridization wash solution 3: 0.05 × SSC. Prepare 250 mL.

3. Methods We present the ChEST protocol based on previously published experiments (10) designed to identify direct targets of muscle differentiation factor Dmef2. 3.1. In Silico Prediction of CRMs and Generation of Computed CRM Array

1. Computer-assisted genome scanning. Custom-developed scanning/filtering SCAFI software (available via http://www.soluscience.fr/dup_soft/) was used for in silico prediction of CRMs to which bind transcription factor Dmef2 and bHLH factor Twist (see Note 1). SCAFI uses flyenhancer (http:// www.flyenhancer.org) as the scanning engine. Scanning parameters and the number of identified CRMs in each scanning condition (see Note 10) are presented below in Table 1.

Table 1 Scanning parameters and in silico identified Dmef2-bound CRMs Flyenhancer scanning Dmef2 sites

E-box/E-box like sites

Window size (bp)

1X TTATTTTTAG

4X CANNTR

100

383

1X CTATTTTTAG

2X CANNTR

100

616

1X CTATATTTAG

3X CANNTR

150

244

Total

Number of CRMs

1,243

The three previously reported in vivo Dmef2-binding sites were combined with different numbers of E-box/E-box-related sequences to which bind key mesoderm factor Twist in a short genomic window ranging from 100 to 150 bp. This led to the prediction of more than 12,000 Dmef2-dependent CRMs

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2. Filtering of predicted CRMs. The subsequent filtering steps were performed using SCAFI and were based on the distance of predicted CRMs in relation to adjacent genes (available via http://www.flyenhancer.org) and on the annotations of potential target genes (available via Flybase and BDGP in situ database). Only CRMs located within 12 kb of genes annotated as having mesodermal function and/or expressed in the mesoderm or in a subset of mesodermal cells were selected (see Note 10). The CRMs present in the vicinity of genes of unknown function and expression were rejected. An example of in silico prediction, filtering, and subsequent ChEST detection on computed arrays is shown in Fig. 1.

Fig. 1. An overview of CRM selection from the predicted Dmef2 CRM’s pool and an example of ChEST detection. (a) A scheme illustrating selection steps during filtering process. (b) A sample from printed 60 nt oligos microarrays hybridized with Dmef2 ChIPed DNA probe (left panel) or with a mock DNA probe (right panel). Enriched sequences in left compared to the right panel are indicated.

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3. Generation of custom CRM microarrays. Selected CRMs are used to design 60-mer oligonucleotides (one per CRM, see Note 11). Design, oligonucleotide synthesis, and microarray printing are done by Eurogenetec (see Note 2). Oligonucleotides are printed in replicates (2 times each oligonucleotide). 3.2. ChIP from Drosophila Embryos

1. Cross-linking: Dechorionate about 1 g of age-matched embryos (stage 11–15 for Dmef2 ChIP) (see Note 12), wash using Embryo Wash Buffer, and cross-link for 15 min at room temperature in 40 mL of Cross-linking Buffer (see Note 13). After spinning down at 500 × g for 1 min and decanting the supernatant, wash embryos briefly with 50 mL of Stop Crosslinking Buffer. Pellet embryos again at 500 × g for 1 min, decant supernatant, and wash briefly with 50 mL of Embryo Wash Buffer. Remove supernatant and pour embryos onto Nytex membrane lying on the filter paper to blot the liquid and to get dry embryos (see Note 14). 2. Homogenization: Resuspend dechorionated and cross-linked embryos (thaw when using frozen embryos) in 10 mL of Homogenization Solution 1. Homogenize 20× on ice using a Dounce homogenizer with B type pestle (tight). Transfer to a suitable tube and centrifuge at 500 × g for 1 min at 4°C. Collect supernatant and centrifuge at 1,000 × g for 10 min at 4°C to pellet the nuclei. Resuspend the nuclei in 2 mL of Homogenization solution 1 and purify on a sucrose gradient at 11,000 × g for 30 min at 4°C using a SW41 rotor (Beckman). Wash the pellet containing nuclei with 2 mL of Homogenization Solution 2 and centrifuge at 1,000 × g for 10 min at 4°C. 3. Sonication: Dissolve the purified nuclei in 3 mL of Sonication buffer. Sonication is performed on ice in conditions adapted to obtain DNA fragments of 200–1,500 bp (Branson Sonifier B15, output control: 4, duty cycle%: 60 with 15 pulses of 1s repeated 12 times) (see Note 15). Transfer sonicated chromatin to 1.5 mL eppendorf tubes and centrifuge for 15 min at 11,000 × g at 4°C. Pool clarified supernatant containing fragmented chromatin and use for immunoprecipitation (see Note 16). 4. Immunoprecipitation first day: Use 30 mL of Protein A Sepharose beads (see Note 6) resuspended in 100 mL of IP buffer for each immunoprecipitation reaction (including the mock reactions) for chromatin preclearing. Dilute an aliquot of chromatin (200–300 mL) by adding 100 mL of 10% Triton X100, 100 mL of 1% deoxycholate, 100 mL of 1% SDS, 100 mL of 1.4 M NaCl, and 10 mL of 100 mM PMSF. Preincubate chromatin with Protein A Sepharose for 2 h at 4°C to eliminate nonspecific binding to the resin. Transfer precleared chromatin to a new tube. Take 10 mL as an input sample and keep it at 4°C. Add 3 mL of Dmef2 antibody or nonimmune serum

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(mock reaction) to the preincubated chromatin and incubate overnight at 4°C with gentle mixing. In parallel for each immunoprecipitation (including mock reactions), prepare 30 mL of preblocked Protein A Sepharose beads by washing beads with 1 mL of IP buffer containing 1 mg/mL BSA and then by incubating with another 1 mL of IP buffer with 1 mg/mL BSA overnight at 4°C. 5. Immunoprecipitation second day: Spin down preblocked Protein A Sepharose beads and resuspend in 100 mL of IP buffer per immunoprecipitation reaction. Purify protein-DNA complexes from chromatin samples by adding 100 mL of preblocked beads and incubating for 4 h at 4°C. Wash beads for 10 min at 4°C with 1 mL of IP buffer, 3 times with IP washing solution 1, once with IP washing solution 2 and 2 times with TE buffer (see Note 17). Resuspend beads with protein-DNA complexes in 100 mL of TE and incubate for 30 min at 37°C with 50 mg/mL RNase. Adjust sample to 0.5% SDS and 0.5 mg/mL proteinase K and incubate overnight at 37°C. 6. Immunoprecipitation third day: Reverse formaldehyde crosslinking by incubating samples for 6 h at 65°C. Eliminate antibody/antigen complexes by phenol/chloroform extraction using phase lock tubes and precipitate DNA fragments. Resuspend pellet in 30 mL of TE and store at −20°C (see Note 18). 3.3. DNA Amplification, Probe Synthesis, and Hybridization

1. Linker ligation: Ligate 7 mL of blunted DNA fragments (see Note 19) from ChIP reaction with 1 mL of 1 mM annealed linker (see Notes 7 and 8) by adding 1 mL of 10× ligation buffer and 1 mL of T4 DNA ligase. Incubate overnight at 16°C. 2. Amplification: Use the ligation mixture directly for PCR amplification. Add to the tube with the ligation reaction 68 mL of H2O (PCR grade), 10 mL of 10× Amplitaq buffer (Roche), 10 mL of 2 mM dNTP mixture, 1 mL of Amplitaq polymerase (Roche), 1 mL of ChIP 20-mer primer. The PCR amplification scheme is as follows. 1 cycle of: 2 min at 94°C; 34 cycles of: 1 min at 94°C/1 min at 55°C/3 min at 72°C; 1 cycle of: 1 min at 94°C/1 min at 55°C/10 min at 72°C. Purify PCR products on Qiagen PCR purification columns (following manufacturer’s instructions). Final elution of the DNA is in 30 mL of Qiagen “EB buffer” (10 mM Tris-Cl, pH 8.5). Determine the concentration of the eluted DNA with a Nanodrop spectrophotometer (OD260). Further check the size and yield of DNA by agarose gel electrophoresis. 3. Probe labeling: Adjust 0.5 mg of PCR amplified DNA to a volume of 10.5 mL with H2O and mix with 10 mL of 2.5× random prime reaction buffer (Invitrogen). Heat the sample for 5 min at 100°C to denature DNA and place on ice. Add 2.5 mL of 10× UTP-mix, 1.5 mL Cy3 or Cy5-dUTP, and 0.5 mL Klenow

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polymerase 40 U/mL (Invitrogen Bioprime kit). Incubate overnight at 37°C. Protect from light. Purify with Qiagen minElute columns. Elute with 20 mL of EB buffer into a brown Eppendorf tube and quantify the DNA content and the specific activity with the Nanodrop spectrometer. 4. Hybridization in Corning chambers: Resuspend 1 mg of speedvac-dried DNA probe in 35 mL of hybridization mix (see Note 9). To humidify the chamber, pipet 15 mL of 3× SSC into each humidifier well and 10 mL of 3 × SSC to each side of the slide. Clean a suitable coverslip with compressed air. Boil the probe 2 min at 100°C in a heat block. Spin down and cool for 30 s at room temperature. Load the probe onto the center of the coverslip and immediately lay down the slide with printed CRMs over the coverslip. Take care to avoid bubbles. Put the slide into a hybridization chamber, attach the lid of the chamber, seal and submerge it in a 42°C water bath. Hybridize for 16 h in the dark.

4. Notes 1. Other genome scanning tools such as SECOMOD (http:// crfb.univ-mrs.fr/secomod/public/) or Enhancer Element Locator (http://www.cs.helsinki.fi/u/kpalin/EEL/) can be used for identifying potential TF-binding regulatory modules in other species. 2. The custom-made 60 nt computed CRM microarrays were originally designed and printed using Eurogenetec services. As genomics microarrays printing is no longer proposed by Eurogenetec, a comparable service can be purchased from Agilent. 3. 1× PBS (Phosphate-Buffered Saline) Buffer consists of: 137 mM NaCl, 2.7 mM KCl, 100 mM Na2HPO4, 2 mM KH2PO4. Routinely, it is prepared from autoclaved 10 × PBS, pH 7.4. 4. To prepare 100 mL of stop cross-link buffer, dissolve 0.93 g of glycine in 1 × PBS and then add 0.1 mL of Triton X100. 5. Add PMSF immediately before use from a 100 mM stock in isopropanol. Protease inhibitors and 1 mg/mL BSA are only added to the IP buffer for immunoprecipitation but not for washing. 6. To prepare Sepharose A beads, take for each IP reaction 30 mL of Protein A Sepharose, wash with 1 mL IP buffer, spin down at 1,000 × g, and resuspend in 100 mL of IP buffer. 7. To phosphorylate the Primer 1, mix 1 mL of 1 mM 24-mer oligonucleotide, 16 mL H2O, 2 mL of 10× T4 kinase buffer,

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1 mL of T4 polynucleotide kinase (10 U/mL). Incubate for 1 h at 37°C and then for 10 min at 68°C (heat inactivation). Add 40 mL of H2O. Mix and add 240 mL of 5 M ammonium acetate. Mix and then add 750 mL of 80% ethanol. Precipitate at −20°C. Centrifuge for 30 min, remove supernatant, dry in air, and resuspend in 100 mL of TE to obtain a 10 mM solution of Phosphoprimer 1. 8. To generate the PCR linker, mix 100 mL of 10 mM Phosphoprimer 1 with 100 mL of 10 mM Primer 2. Incubate for 5 min at 70°C (to remove secondary structures) and then for 5 min at 55°C (annealing). Let the solution cool down slowly at room temperature to obtain 200 mL of 5 mM blunt PCR linker. 9. To prepare hybridization, mix for each labeled probe (speedvac dried), mix 6 mL of 20 × SSC, 4 mL of 10% SDS, 4 mL of 50× Denhardt’s, 3.2 mL of 10 mg/mL polyA, 20 mL of 100% formamide. 10. A wide range of scanning and filtering conditions can be applied. A large number of CRMs will be predicted by increasing the size of the genomic window in which the search for regulatory motifs is performed and by reducing the number and the complexity of motifs. Conversely, searching for relatively long motifs in a genomic window of restricted size will result in the prediction of a comparatively lower number of CRMs. Filtering parameters are also fully flexible, enabling a defined combination of functional criteria to be used for the final CRM selection. This is possible because the relevant information about the CRMs and the adjacent genes is stored in a database that is automatically generated by SCAFI software during the initial scanning step. Below (Table 2) is an example of ChEST efficacy scoring for three different TFs tested in our laboratory, which takes into account both in silico and in vivo parameters. 11. The size of selected Dmef2 CRMs was 100–150 bp. In this case, it was difficult to design more than one 60 nt oligonucleotide per CRM, especially for 100 bp CRMs. In case the size of CRMs is larger, it is advised to design more than one oligonucleotide per CRM. A library of long oligonucleotides for genomic arrays is actually available via Agilent so that it is possible to choose oligonucleotides from the existing catalog. 12. Wash the embryos from the agar plate and transfer them into a sieve. Dechorionate for 2.5 min in 100 mL of 50% bleach at room temperature in a beaker. Use a stir bar to keep embryos in suspension. Wash embryos with water and transfer them into a preweighted eppendorf tube to calculate their weight. 13. Gentle shaking of embryos during cross-link is advised.

***

*

Lmd

Lb

*

–

*

Other TF sites expected to regulate the same CRMs (***max)

In silico and in vivo parameters have been scored

***

Dmef2

TF

Complexity of DNA-binding sites (***max)

In silico ChEST parameters

* *

–

*

Stringency of filtering (***max)

*

*

Restricted genomic window used for scanning (***max)

*

*

*

Quality and specificity of antibodies (***max)

–

*

**

Proportion of TF-expressing cells (***max)

In vivo ChEST parameters

Table 2 Estimation of ChIP-enriched in silico target (ChEST) efficacy for three different TFs tested

4

7

9

Max score 10

ChEST efficacy score

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14. At this step, cross-linked embryos can be frozen in liquid nitrogen and stored at −80°C. There is a possibility to pool several samples of frozen embryos for ChIP reaction. 15. Sonication can also be done in the Bioruptor sonicator water bath (Diagenode, Liège, Belgium) in 15 mL falcon tubes 10× (15s on/15s off). The water bath should contain some ice. 16. At this step, sonicated chromatin can be subdivided into aliquots (200–300 mL) and frozen in cryotubes by flash freezing in liquid nitrogen. Keep chromatin at −80°C. 17. Spin at 1,000 × g for 2 min to pellet Protein A beads, then remove supernatant. 18. 1 mL of resuspended DNA fragments can be used for real-time PCR reaction to test enrichment of specific fragments. 19. Perform blunting reaction for 30 min at 37°C. Mix 29.5 mL of H2O, 10 mL of 5× T4 polymerase buffer, 2.5 mL of dNTPs (each 2 mM), 1 mL of T4 DNA polymerase and add 7 mL of ChIP-material. Stop the reaction by heating for 5 min at 65°C and purify DNA with the Qiagen Mini-Elute kit. Elute with 20 mL of H2O and reduce the volume to about 7 mL using a Speedvac. References 1. Collas, P. (2010) The current state of chromatin immunoprecipitation. Mol Biotechnol 45, 87–100. 2. Acevedo, L.G., Iniguez, A.L., Holster, H.L., Zhang, X., Green, R. and Farnham, P.J. (2007) Genome-scale ChIP-chip analysis using 10,000 human cells. Biotechniques 43, 791–797. 3. Dahl, J.A. and Collas, P. (2008) A rapid micro chromatin immunoprecipitation assay (microChIP). Nat Protoc. 3, 1032–1045. 4. Nelson, J.D., Denisenko, O., Sova, P. and Bomsztyk, K. (2006) Fast chromatin immunoprecipitation assay. Nucleic Acids Res 34 :e2. 5. Bergman, C.M., Carlson, J.W. and Celniker, S.E. (2005) Drosophila DNase I footprint database: a systematic genome annotation of transcription factor binding sites in the fruitfly, Drosophila melanogaster. Bioinformatics 21, 1747–1749. 6. Hallikas, O., Palin, K., Sinjushina, N., Rautiainen, R., Partanen, J., Ukkonen, E. and Taipale, J. (2006) Genome-wide prediction of mammalian enhancers based on analysis of transcriptionfactor binding affinity. Cell 124, 47–59. 7. Palin, K., Taipale, J. and Ukkonen, E. (2006) Locating potential enhancer elements by

comparative genomics using the EEL software. Nat Protoc 1, 368–74. 8. Halfon, M.S., Gallo, S.M. and Bergman, C.M. (2008) REDfly 2.0: an integrated database of cis-regulatory modules and transcription factor binding sites in Drosophila. Nucleic Acids Res 36 (Database issue), D594–8. 9. Khoueiry, P., Rothbächer, U., Ohtsuka, Y., Daian, F., Frangulian, E., Roure, A., Dubchak, I. and Lemaire, P. (2010) A cis-regulatory signature in ascidians and flies, independent of transcription factor binding sites. Curr Biol 20, 792–802. 10. Junion, G., Jagla, T., Duplant, S., Tapin, R., Da Ponte, J.P. and Jagla, K. (2005) Mapping Dmef2-binding regulatory modules by using a ChIP-enriched in silico targets approach. Proc Natl Acad Sci USA 102, 18479–84. 11. Junion, G., Bataillé, L., Jagla, T., Da Ponte, J.P., Tapin, R. and Jagla, K. (2007) Genome-wide view of cell fate specification: ladybird acts at multiple levels during diversification of muscle and heart precursors. Genes Dev 21, 3163–80. 12. Johnson, D.S., Mortazavi, A., Myers, R.M. and Wold, B. (2007) Genome-wide mapping of in vivo protein-DNA interactions. Science 316, 1497–502.

sdfsdf

INDEX A Adeno-associated virus ............................................ 267, 270 Amino acid tracer ............................................ 246–248, 259 Anesthesia ......................................... 66, 188, 193, 194, 214, 236, 259, 456, 466, 495 Antibodies ............................6, 12, 18, 22, 27–29, 32, 33, 39, 40, 47, 54, 57, 60–62, 74, 79, 81, 82, 129, 133–135, 138, 146–150, 155–159, 165, 166, 172, 175, 180, 218, 219, 224, 374, 377, 430–432, 438, 446, 450, 453–454, 514–516, 526, 528, 543, 552 Atrophy ............................ 215, 227, 232, 233, 237, 311, 475

B B-galactosidase (B-gal) .........................23, 26, 143, 445–458

C Caenorhabditis elegans (C.elegans) ..................... 171–180, 476 Calcium ............................................. 75, 100, 270, 272, 273, 280, 373–381, 383–391, 395–408, 411–420 ATPase .............................................................. 411–420 dyes .................................................................... 398, 399 release channels.......................................... 373, 374, 395 spark .................................................................. 395–408 Cardiac myocytes..................................................... 411–420 Catalase ............................................192, 193, 204–206, 208 C2C12. ..................... 26, 66, 71, 75, 239, 312, 314, 316, 428, 431, 437, 476–478, 480–482, 485, 486, 513–515 C.elegans. See Caenorhabditis elegans Cell culture .......................... 7, 14, 25–28, 31–34, 38–40, 43, 66, 80, 82, 85, 86, 91, 270, 286, 288, 312–313, 321, 378, 380, 386, 397, 413–417, 426, 428, 431–432, 439, 469, 485, 507, 518, 528, 534, 535, 540 Cell isolation ..............................................21, 22, 26–38, 54 Chicken ....................................23, 25, 28, 29, 31, 34, 42, 43, 80, 106, 117, 157, 158, 532 ChIP enriched in silico targets ................................ 543–553 Chromatin accessibility ................................... 533, 535, 539 Chromatin immunoprecipitation............................ 431–434, 517–528, 543 Chromatin remodelling .................... 517, 518, 526, 532, 538 cis-regulatory modules (CRM) .......................544, 546–548, 550–552

Collagenase .....................................................5, 8, 9, 27, 30, 55, 56, 86, 87, 90, 94, 96, 98, 99, 384, 385, 387, 397–399, 406, 419, 519, 523 Cre recombinase ........................................................ 27, 445 Cross-sectional area ............ 24, 216–224, 226, 232, 237, 238 Cryosectioning .........................................146, 449–453, 457

D Depolarization ......................................................... 384, 412 Dermomyotome .............................................................. 161 Development ........................................... 3, 4, 29, 32, 47, 54, 103–107, 109, 110, 115–117, 127–150, 153, 154, 200, 216, 223, 277, 312, 326, 335, 358, 390, 396, 433, 446, 475, 477, 491, 533 Difference in-gel electrophoresis ..................................... 358 Differentiation ............................... 3, 4, 7, 15, 16, 18, 24–26, 28, 29, 31, 32, 42, 43, 66, 67, 71–73, 75, 78, 85, 114, 128, 130, 132, 153, 257, 289, 294, 313, 314, 384–387, 390, 425–428, 431, 437, 476–478, 481, 482, 486, 518, 526, 538, 546 Digestion ......................................... 9, 26, 27, 29, 30, 33–37, 47, 54, 56, 86, 87, 92, 94, 99, 155, 162, 166, 271, 274, 275, 361, 366, 368, 387, 397–399, 401, 406, 414–416, 419, 488, 518, 519, 521–523, 532, 534–536, 539, 540, 544 Drosophila .........127–150, 154, 157, 433, 476, 544, 548–549 Dynamometer ......................................... 188–190, 193–195 Dystrophin ......................... 22, 157, 294, 357, 358, 477, 526

E Eccentric contractions ..................................... 197–198, 214 EDL. See Extensor digitorum longus (EDL) Electrical stimulation.......................................... 77–83, 190, 207, 214, 240, 383 Electrophoresis .................200, 234, 235, 261, 323, 358–361, 363, 367, 380, 479, 481, 486, 487, 511, 512, 549 Electrophoretic mobility shift assay .................430, 506–507, 511–513 Electroporation................................ 233, 235–236, 240, 241, 300, 302, 462, 464, 466, 470 Electrotransfer ...232, 236–238, 240, 462, 492, 494–495, 497 Embryo................ 4, 22, 80, 89, 103, 128, 153, 311, 426, 544 Estrogen receptor ............................................................ 297

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MYOGENESIS 556 Index Evoked contractions ........................................................ 391 Excitation-contraction coupling ...............383, 391, 412, 418 Exercise .....................185–208, 213, 215, 216, 311, 326, 396 Extensor digitorum longus (EDL) ...... 24, 87, 214, 215, 221, 225, 396–399, 401–404, 406, 407

F Fast fiber .................................................................... 78, 219 Fast twitch ......................................................24, 30, 77, 446 Fatty acids..................326, 328, 340, 341, 348–350, 352, 353 FDB fibers. See Flexor digitorum brevis (FDB) fibers Fiber type .............................. 77–83, 85, 199, 219, 220, 222, 224, 225, 426, 445–458, 477 Fixation ......................16, 32–33, 38–40, 101, 130, 133–134, 136–137, 140, 143–146, 148–150, 154, 160, 165, 172, 173, 175–179, 363, 380, 432, 447, 449–453 Flexor digitorum brevis (FDB) fibers ................87, 387, 398, 400–402, 404, 406, 407 Flooding dose .......................................................... 245–263 Fluorescence ............................................ 7, 8, 13–14, 16, 18, 23, 27, 28, 33, 53–63, 67, 70, 129, 138, 144, 165, 166, 172, 202, 208, 291, 358, 363, 376, 386–390, 398, 399, 402–404, 413, 415–418, 420, 449, 454, 455, 471, 492–496, 498, 499 Fluorescence-activated cell sorting ...................... 4, 7, 13–14 Founder cell ..................................................... 128–130, 132 Functional overload ................................................. 214, 215

G Gas chromatography ........................326, 328, 329, 333–334 Gene expression............................... 4, 25, 54, 60, 61, 78, 79, 130, 139, 142–144, 268, 269, 297, 307, 312, 326, 373, 391, 426–428, 439, 445–458, 461, 462, 477, 478, 491–495, 519, 538 Gene profiling ......................................................... 311–323 Gene promoter ......................... 425–439, 446, 461–471, 538 Gene silencing ................................................. 418, 493, 494 Gene therapy ........................................................... 268, 492 Gene transfer ............................ 231–241, 268, 418, 495, 499 Glutathione ..............................................191, 199, 200, 375

H Homologous recombination ............................ 298, 304–306 Hybridization ................................ 89, 92, 96, 100, 101, 129, 131, 139–142, 271, 275, 305, 312, 479, 482, 532, 546, 549–551 Hypertrophy ..................... 130, 186, 213–227, 311, 418, 477

I Immunocytochemistry............................16, 82–83, 153–166 Immunohistochemistry ................... 129, 142, 146, 154, 234, 236–238, 453

Immunostaining ...................... 22, 23, 29, 32–33, 35, 38–40, 60, 79, 87, 95, 97, 144, 148, 172, 177–178, 432, 446–448, 451, 454, 458, 471 Inositol triphosphate receptor.......................... 373, 383–391 In vitro ..................................3, 4, 7–8, 14–17, 23, 78, 79, 86, 140, 190, 214, 221, 225, 312, 438, 445, 477, 491 In vivo.. .................................23, 78, 79, 85–87, 98, 186, 195, 218, 232, 240, 245–263, 428, 430, 433, 438, 439, 445, 446, 449, 461–471, 477, 478, 491–495, 499, 544, 546, 551, 552 Isometric contractions ..............................186, 194, 196, 221 Isometric maximal force .................................................. 198

K Knock-in ................................................................. 298, 301 Knock-out ....................................................................... 298

L Larva... ................................ 130, 132, 136, 137, 139–140, 148 Lentivirus ........................................................ 285–294, 477 Ligation-mediated PCR .......................................... 533, 537 Lineages ..................................... 4, 65, 72, 85, 426, 445, 538 Lipidome ......................................................................... 328 Lipids.................154, 325, 326, 328–330, 337, 344, 351, 352 Localization ........................ 96, 171–180, 232, 374–375, 377 Luciferase ........................ 429, 437, 462–464, 466–471, 478, 480, 483–485

M Mass spectrometry........................... 256, 357, 358, 361, 363, 429, 430, 436 MEF......... .............2, 128, 129, 161, 163, 426–428, 433, 434 Mesoangioblasts .......................................................... 65–75 Mesoderm ................................ 103–120, 128, 130, 546, 547 Microarray ...............4, 60, 312, 384, 478, 544, 547, 548, 550 MicroRNA (miRNA)...............................306, 476, 479, 483 Mouse.............................6, 23, 25–33, 36, 40–42, 65–75, 87, 89, 90, 92–93, 98, 101, 154, 157–159, 188–190, 192, 194–198, 218, 219, 225, 232, 233, 235, 236, 238, 240, 241, 248, 249, 251–253, 259, 261, 278, 293, 297, 304–307, 358, 374, 380, 384, 387, 389, 396, 399, 405, 414, 418, 431, 446–449, 458, 476–478, 480, 483, 485, 495, 500, 522, 540 Muscle. .............3–19, 21–47, 53–63, 65–75, 77–83, 85–101, 127–150, 153, 154, 157–159, 161, 163, 171–180, 185–208, 213–227, 231–241, 245–263, 268, 285–294, 311–323, 325–354, 357–368, 373–381, 383–391, 395–408, 425–439, 445–458, 461–471, 475–488, 491–500, 505–528, 538–540, 543–553 Muscle fiber....................... 22, 65, 77–83, 85–101, 128, 129, 153, 159, 217, 219–221, 224, 225, 231, 232, 237, 238, 241, 325, 326, 357, 383–391, 395–408, 426, 446–448, 451, 470, 495, 499, 518

MYOGENESIS 557 Index Muscular dystrophy ..................... 24, 47, 150, 312, 357, 358, 396, 426, 476, 477 Myoblast .........................3–19, 22, 24–26, 29, 42, 43, 60, 66, 71, 79–82, 128–130, 133, 139, 288–290, 292–294, 312, 314, 315, 384–386, 426–428, 430, 431, 433, 437, 476–478, 482, 485, 486, 513–515, 526 Myocyte............................ 411–420, 425, 430–433, 436, 438 MyoD.......................22, 25, 26, 29, 158, 426–429, 431–434, 476, 526, 527, 538, 539 Myofiber ....................22–26, 53, 86, 186, 311, 391, 517–528 Myogenesis ............................ 25, 26, 28, 127, 128, 153–166, 297–307, 311, 358, 425, 432 Myogenin ................25, 29, 72, 158, 426, 431–433, 526, 527 Myosin heavy chain ...........29, 72, 77, 79, 128, 219, 323, 446 Myotome ...................................... 4, 113, 157–159, 166, 426 Myotubes ............................16, 22, 25, 29, 32, 71, 72, 75, 78, 79, 82, 83, 86, 128, 294, 312, 314, 315, 384–391, 397, 431, 508

N NF-KB ............................................................. 239, 505–516 Northern blot .......................... 312, 461, 478–479, 481, 482, 485, 486, 488 Nuclear extract ......................... 430, 435, 436, 438, 505–515 Nuclei isolation........................................ 517–529, 533–535

O Osmotic shock ................................................................. 499 Osmotic stress ................................................. 396, 402, 403 Oxidative stress................................................ 185–208, 396

P Pax............7, 22, 23, 25–29, 87, 158, 298, 300–303, 306, 307 PCR. See Polymerase chain reaction Pericytes ............................................................................23 Phospholipids ...........................................326–350, 353, 354 Plasmid DNA...................140, 220, 231–235, 239, 240, 279, 462–466, 468–470, 492, 498, 499 Polymerase chain reaction (PCR) ......................60, 132, 140, 269, 271, 274, 275, 281, 304–306, 312, 313, 317, 320, 323, 384, 431, 432, 479, 483, 485, 526, 533, 537–538, 545, 546, 549, 551, 553 Proliferation....................9, 23–25, 28, 29, 31, 32, 42, 43, 68, 71, 72, 85, 89, 93, 97–99, 293, 384–386, 476–478 Pronase .......................................................27, 29, 30, 35–37 Protein degradation ................................................. 246, 257 Protein synthesis ...................................................... 245–263 Proteomics ........................ 357, 358, 361, 428, 430, 434–436 Pupa......................................................................... 136, 137

Q Quantitative real-time PCR (qRT-PCR) ........................ 312

R Rat........ ............ 23, 25, 26, 28, 40, 86, 89, 91, 158, 188–190, 193–199, 207, 215, 216, 220, 224–227, 232, 233, 235–241, 247, 384, 396, 404, 413–415, 464–467, 469–471 Reactive oxygen species (ROS)................................ 187, 204 Reporter gene .................... 27, 142–144, 280, 291, 293, 297, 307, 428, 437, 439, 445–458, 462–464, 466, 468, 492, 494, 495, 544 Resistance exercise ....................................213, 215, 216, 311 Resistance loading ................................................... 185–208 Restriction enzyme accessibility assay ..................... 531–541 Ribonucleic acid (RNA) .................... 60, 100, 140, 268, 277, 285–287, 312–317, 322, 476, 482, 483, 485, 487, 488, 491, 493, 524, 525, 528 interference ................................................ 268, 418, 491 RNA. See Ribonucleic acid (RNA) Ryanodine receptors ................................................ 373–381

S Sarcoplasmic reticulum ............. 225, 325, 373, 395, 411–420 Satellite cells ...................... 21–47, 53, 54, 57–62, 65, 85–90, 94–100, 517–528 Segmentation................................................... 104, 107–116 Serotype................................................................... 268, 278 Short hairpin RNA ......................................................... 491 Single fiber isolation .................................................... 89–91 Skeletal muscle .................... 3–19, 21–47, 65–75, 78, 86, 89, 127–150, 185–208, 213–227, 231–241, 246, 247, 250, 285–294, 311–323, 325–354, 357–361, 367, 373–381, 383–391, 395–408, 425–439, 446, 461–471, 475–488, 491–500, 505–528, 538, 540, 543–553 Slow fiber .................................................................. 77, 426 Slow twitch ...................................................24, 77, 426, 446 Small interfering RNA .................................................... 491 Somites ..................... 104, 105, 107, 109, 111–114, 116–119 Somitogenesis ...........................................107–109, 114, 120 Specific tension................................. 217, 222, 224, 225, 227 Stem cell ............................... 24, 43, 47, 53–63, 68, 298, 518 Stimulation .................................... 77–83, 95, 193–195, 197, 198, 207, 214, 221, 232, 240, 383, 388, 389, 396, 413, 416–418 Stretch-shortening............................186, 187, 194–196, 198 Synergist ablation ............................................................ 214

T Tamoxifen................................................................ 297–307 Thin-layer chromatography..................................... 326–327 Transcription ..................23, 25–26, 128, 140, 232, 240, 268, 286, 312, 313, 317, 384, 412, 425–439, 461–463, 476, 479, 483, 488, 492, 517, 527, 543, 544, 546

MYOGENESIS 558 Index Transcription factors ............................ 25–26, 128, 232, 240, 312, 384, 426–431, 434–439, 476, 543 Transduction.....................227, 232, 268, 269, 278, 285–289, 291–294, 326, 439 Transfection ......................232, 237, 240, 241, 270, 272, 273, 279, 280, 286, 287, 289–293, 300, 437, 439, 462, 465, 466, 468, 484–486, 488, 499 Transgene ........................ 232, 268, 275, 287–289, 291–294, 446, 447, 458 Transgenic ....................23, 27, 128, 148, 172, 232, 267–282, 298, 304, 305, 406, 418, 426, 439, 445–458, 477 Transients ......................... 376, 378–379, 384, 390, 411–420 Translation............................................................... 246, 247

Triacylglycerols ......................... 326–332, 334–350, 353, 354 Two-dimensional gel electrophoresis............... 358, 361, 367

V Viral vector ...............................................267–282, 491, 492

W Whole mount .................................. 129, 130, 136–139, 154, 160–162, 164, 165

Z Zebrafish ........................................89–91, 96, 116, 153–166