Methods in Cell Biology Volume 52 Methods in Muscle Biology

Methods in Cell Biology VOLUME 52 Methods in Muscle Biology (ASCB( Series Editors Leslie Wilson Department of Biolo...

Author: Charles P. Emerson | H. Lee Sweeney

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Methods in Cell Biology VOLUME 52 Methods in Muscle Biology

(ASCB(

Series Editors

Leslie Wilson Department of Biological Sciences University of California, Santa Barbara Santa Barbara, California

Paul Matsudaira Whitehead Institute for Biomedical Research and Department of Biology Massachusetts Institute of Technology Cambridge, Massachusetts

Methods in Cell Biology Prepared under the Auspices of the American Society for Cell Biology

VOLUME 52 Methods in Muscle Biology

Edited by

Charles P. Emerson, Jr. Department of Cell and Developmental Biology University of Pennsylvania School of Medicine Philadelphia, Pennsylvania

H. Lee Sweeney Department of Physiology University of Pennsylvania School of Medicine Phdadelphia, Pennsylvania

ACADEMIC PRESS San Diego

London

Boston

New York

Sydney

Tokyo

Toronto

Front cover for comb-bound: Absence of nonmyogenic cells. After 21 days in culture, cells were stained with rhodamine-phalloidin to visualize actin filaments (red) and DAPI to visualize nuclei (blue). From Chapter 13 (Sweeney and Feng).

This book is printed on acid-free paper.

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Copyright Q 1997 by ACADEMIC PRESS All Rights Reserved. No part of this publication may be reproduced or transmitted in any form or by any means, electronic or mechanical, including photocopy, recording, or any information storage and retrieval system, without permission in writing from the Publisher. The appearance of the code at the bottom of the fmt page of a chapter in this book indicates the Publisher's consent that copies of the chapter may be made for personal or internal use of specific clients. This consent is given on the condition, however, that the copier pay the stated per copy fee through the Copyright Clearance Center, Inc. (222 Rosewood Drive, Danvers, Massachusetts 01923). for copying beyond that permitted by Sections 107 or 108 of the U.S.Copyright Law. This consent does not extend to other kinds of copying, such as copying for general distribution, for advertising or promotional purposes, for creating new collective works, or for resale. Copy fees for pre-1997 chapters are as shown on the title pages, if no fee code appears on the title page, the copy fee is the same as for current chapters. 0091-679XI97 $25.00

Academic Press a division of Harcourt Brace & Company 525 B Street, Suite 1900, San Diego, California 921014495, USA http://www.apnet.com

Academic Press Limited 24-28 Oval Road, London NWl 7DX, UK http://www.hbuk.co.uk/ap/ International Standard Book Number: 0-12-564154-0 (case.) International Standard Book Number: 0-12-238190-4 (comb) PRINTEDIN THE UNITED STATESOF AMERICA 97 98 9 9 0 0 01 0 2 E B 9 8 7 6

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3 2 1

CONTENTS

Contributors

xi

Preface

xv

PART I Embryological Analysis of Myogenesis 1. Avian Somite Transplantation: A Review of Basic Methods Charles P. Ordahl and Bod0 Christ I. 11. 111. IV.

Introduction Materials Methods Summary and Concluding Remarks References

4 5 11 25 26

2. Myogenesis in the Mouse Embryo Margaret Buckingham and Giulio Cossu I. 11. 111. IV. V.

Introduction Manipulation of Genes in the Mouse Analysis of Gene Expression in the Mouse Embryo Experimental Embryology Perspectives References

29 31 38 43 48 50

3. Myogenesis in Xenopus Embryos John B. Gurdon, Patrick Lemaire, and Timothy J. Mohun I. 11. 111. IV.

Fate Maps for Muscle Embryo Manipulations Molecular Markers of Muscle Differentiation Studies of Xenopus Heart Formation References

53 55 59 62 63

4. Zebrafish: Genetic and Embryological Methods in a Transparent Vertebrate Embryo Mark C . Fishman, Didier Y. R. Stainier, Roger E. Breitbart, and Monte Westerfield

I. Introduction 11. Genetics 111. Heart Development

68 68 70 V

vi

Contents

72 79

IV. Skeletal Muscle Development References

PART I1 Myogenesis in Cell Culture 5. Skeletal Muscle Cultures Craig Neville, Nadia Rosenthal, Michael McGrew, Natalia Bogdanova, and Stephen Hauschka

85 86 113 114

I. Introduction 11. Muscle Cell Cultures 111. Summary and Future Prospects

References 6. Avian Cardiac Progenitors: Methods for Isolation, Culture, and Analysis

of Differentiation Maureen Gannon and David Bader

118 119 119 120 121 122 124 125 126 127 131

I. Establishment of Embryonic Axis 11. Pregastrulation Location of Cardiac Progenitors 111. Postgastrulation Location of Cardiac Progenitors IV.Removal of Embryos from the Egg V. Removal of Cardiac Progenitors VI. Removal of Endoderm VII. Culturing Whole Embryos VIII. Culturing Explants of Cardiogenic Mesoderm IX. Culturing Primary Cardiomyocytes X. Analysis of Differentiation References

7. Vascular Smooth Muscle Cell Cultures Rebecca R. Pauly, Claudio Bilato, Linda Cheng, Robert Monticone, and Michael

T.Crow

I. Introduction 11. Establishment of Cell Cultures

111. IV. V. VI.

Characterization of Cultured Vascular Smooth Muscle Cells Specialized Methods for Human Tissues Specialized Methods to Maintain the Differentiated State Conclusions and Perspectives References

133 136 141 147 150 152 153

8. Skeletal Muscle Satellite Cell Cultures Ronald E. Allen, ConstanceJ. Temm-Grove, Shannon M. Sheehan, and Glenna Rice I. Introduction 11. Physiological or Developmental Background of Subjects

111. Monolayer Mass Cultures

155 157 158

vii

Contents

IV. Isolated Single Fiber Cultures V. Advantages and Disadvantages of Each Culture System VI. Conclusions References

170 173 174 175

PART I11 Viral and Cellular Gene Delivery Systems in Muscle 9. Retroviral Gene Delivery Mark J . Federspiel and Stephen H . Hughes I. 11. 111. IV. V.

Introduction Vector Construction and Virus Propagation Avian Model ALV/Receptor Mouse Model Summary/Reprise References

179 191 196 201 207 209

10. The Use of Replication-Defective Retroviruses for Cell Lineage Studies of Myogenic Cells Donald A. Fischman and Takashi Mikawa

I. Introduction 11. Basic Principles

111. Practical Procedures IV. Application of Retroviruses for Muscle Lineage Analyses in Chicken Embryos References

215 216 218 223 225

11. Adenoviral Gene Delivery 7'hierry Ragot, Paule Opolon, and Michel Pewicaudet I. Introduction to Research 11. Recombinant Adenovirus Design

111. IV. V. VI.

Practical Methods Applications of Adenoviral Gene Transfer to Muscle Current Problems and Future Prospects Conclusion References

230 234 238 24 1 248 256 256

12. Methods for Myoblast Transplantation Thomas A . Rando and Helen M . Blau I. Introduction 11. Preparation of Cells

111. Labeling of Myoblasu in Vitro by Retrovird Mediated Gene Transfer IV. Transplantation Techniques V. Evaluation of Transplant

261 262 264 266 268

Contents

viii VI. Concluding Remarks References

27 1 27 1

PART IV Molecular Analysis of Muscle Structure and Function 13. Structure-Function Analysis of Cytoskeletal/Contractile Proteins in Avian Myotubes H. Lee Sweeney and Huisheng Feng I. Introduction 11. Muscle Cultures 111. Other Considerations References

275 276 279 28 1

14. Functional and Structural Approaches to the Study of Excitation-Contraction Coupling Kurt G. Beam and Clara Franzini-Amstrong I. Functional and Structural Approaches to the Identification of e-c Coupling Components 11. The Dysgenic Muscle Model 111. Conclusions References

284 296 305 305

15. Adenovirus-Mediated Myofilament Gene Transfer into Adult

Cardiac Myocytes Margaret Wesijbll, Elizabeth Rust, Faris Albayya, and Joseph M . Metzger I. 11. 111. IV.

Introduction Isolation of Adult Cardiac Myocytes and Adenovirus-Mediated Gene Transfer Production of Recombinant Adenovirus Vectors Analysis of Myofilament Protein Expression and Incorporation in AdenovirusInfected Ventricular Myocytes V. Summary and Future Directions References

307 309 313 317 320 321

16. In Vivo Approaches to Neuromuscular Structure and Function Rita J. Balice-Gordon 1. Introduction 11. In Viuo Analysis of Skeletal Muscle and Neuromuscular Innervation 111. Manipulation of Motor Neurons, Muscle Fibers, and Synapses in Viuo IV. In Viuo Observations and Manipulations of Muscle Fibers and Neuromuscular

323 324 337 338

Innervation: Practical Considerations V. In Viuo Approaches: New Insights into Neuromuscular Junction Plasticity VI. summary References

339 346 346

ix

Contents

17. Molecular Diversity of Myofibrillar Proteins: Isofom Analysis at the Protein and mRNA Level Stefan0 Schiafino and Giovanni Salviati I. 11. 111. IV. V. VI. VII.

Myofibrillar Protein Isoforms in Muscle Tissues Multiple Approaches to Isoform Analysis Electrophoretic Techniques and Immunoblotting Immunohistochemistry and in Situ Hybridization Other Techniques for mRNA Analysis Isoform Analysis and Physiological Studies in Single Muscle Fibers Summary and Perspectives References

349 350 356 359 361 364 365 367

PART V Molecular Genetic Analysis of Muscle Gene Regulation 18. Transgenic Mice: Production and Analysis of Expression Alexander Faerman and Moshe Shani I. 11. 111. IV. V. VI.

Introduction Productipn of Stable Transgenic Mice Production of Transient Transgenics Analysis of Expression at the RNA Level Analysis of Expression at the Protein Level Future Directions References

374 375 383 384 397 400 401

19. D N A Transfection of Cultured Muscle Cells Craig Neville, Nadia Rosenthal, and Stephen Hauschka I. 11. 111. IV. V.

Introduction Transient Transfections Stable Transfections Identification of cis-Acting Control Regions with Reporter Assays Summary and Future Prospects References

405 406 412 415 42 1 422

20. DNA- and Adenovirus-Mediated Gene Transfer into Cardiac Muscle Alyson Kass-Eider and Leslie A . Loinwand I. Introduction 11. Methods for Cardiac Injection 111. Homogenization of Tissue and Assays IV. Experimental Design V. Future Directions References

423 426 428 430 433 435

Contents

X

21. Nuclear DNA-Binding Proteins Kristen L. Kucharczuk and DauidJ. Goldhamer I. Introduction 11. Nuclear Extract Preparation: Overview and Important Parameten 111. Nuclear Extract Protocol IV.Electromobility Shift Assays: Overview and Important Parameters V. EMSA Protocol VI. DNase I Protection Assays: Overview and Important Parameters VII. DNase I Protection Assay Protocol VIII. Relevance of the EMSA and DNase I Protection Assay to Muscle Research References

Index Volumes in 'Series

440 441 442 446 450 454 458 466 47 1

473 489

CONTRIBUTORS

Numbers in parentheses indicate the pages on which the authors’ contributions begin.

Faris Albayya (307), Department of Physiology, University of Michigan, Ann Arbor, Michigan 48109 RonaldE. Allen (155),Muscle Biology Group, Animal Sciences Department, University of Arizona, Tucson, Arizona 85721 David Bader (117), Department of Cell Biology, Vanderbilt University, Nashville, Tennessee 37232 Rita J. Balice-Gordon (323), Department of Neuroscience, University of Pennsylvania School of Medicine, Philadelphia, Pennsylvania 19104 Kurt G. Beam (283), Department of Anatomy and Neurobiology, Colorado State University, Fort Collins, Colorado 80523 Claudio Bilato (133), Laboratory of Cardiovascular Science, National Institute on Aging, National Institutes of Health, Baltimore, Maryland 21224 Helen M. Blau (261), Department of Molecular Pharmacology, Stanford University School of Medicine, Stanford, California 94305 Natalia Bogdanova (85), Cardiovascular Research Center, Massachusetts General Hospital-East, Charlestown, Massachusetts 02129 Roger E. Breitbart (67), Mdlenium Pharmaceuticals, Inc., Cambridge, Massachusetts 02139 Margaret Buckingham (29), CNRS, URS67, Department of Molecular Biology, Pasteur Institute, 75724 Pans Cedex, France Liida Cheng (133), Laboratory of Cardiovascular Science, National Institute on Aging, National Institutes of Health, Baltimore, Maryland 21224 Bod0 Christ (3), Anatomisches Institut, Albert Ludwigs Universitat Freiburg, D-7800 Freiburg, Germany Giulio Cossu (29), University of Rome “La Sapienza,” Institute of Histology and Embryology, 00161 Rome, Italy Michael T. Crow (133), Laboratory of Cardiovascular Science, National Institute on Aging, National Institutes of Health, Baltimore, Maryland 21224 Alexander Faerman (373), Institute of Animal Science, Agricultural Research Organization, The Volcani Center, Bet Dagan 50250, Israel Mark J. Federspiel(179),Molecular Medicine Program, Mayo Clinic and Mayo Foundation, Rochester, Minnesota 55905 Huisheng Feng (275), Department of Physiology, University of Pennsylvania School of Medicine, Philadelphia, Pennsylvania 19104 Donald A. Fischman (215),Department of CellBiology and Anatomy, Cornell University Medical College, New York, New York 10021 xi

xii

Contributora

Mark C. Fishman (67), Cardiovascular Research Center, Massachusetts General Hospital-East, Harvard Medical School, Charlestown, Massachusetts 02129 Clara Franzini-Armstrong (283), Department of Cell and Developmental Biology, University of Pennsylvania, Philadelphia, PA 19104 Maureen Gannon (117), Department ofCell Biology, Vanderbilt University, Nashville, Tennessee 37232 DavidJ. Goldhamer (439), Department of Cell and DevelopmentalBiology, University of Pennsylvania School of Medicine, Philadelphia, Pennsylvania 19104 John B. Gurdon (53), Wellcome/CRC Institute of Cancer and Developmental Biology, Cambridge CB2 lQR, United Kingdom Stephen Hauschka (85, 405), Biochemistry Department, University of Washington, Seattle, Washington 98195 Stephen H. Hughes (179), ABL-Basic Research Program, NCI-Frederick Cancer Research and Development Center, Frederick, Maryland 21702 Alyson Kass-Eisler (423), Cold Spring Harbor Laboratory, Demerec Building, Cold Spring Harbor, New York 11724 Kristen L. Kucharczuk (439),Department of Cell and DevelopmentalBiology, University of Pennsylvania School of Medicine, Philadelphia, Pennsylvania 19104 Leslie A. Leinwand (423), Department of Molecular, Cellular, and Developmental Biology, University of Colorado at Boulder, Boulder, Colorado 80309 Patrick Lemaire (53), Institut de Biologie du Developpment de Marseille, Marsedle, France Michael McGrew (85), Cardiovascular Research Center, Massachusetts General Hospital-East, Charleston, Massachusetts 02129 Joseph M. Metzger (307), Department of Physiology, University of Michigan, Ann Arbor, Michigan 48109 Takashi Mikawa (215), Department of Cell Biology and Anatomy, Comell University Medical College, New York, New York 10021 Timothy J. Mohun (53), National Institute for Medical Research, London, United Kingdom Robert Monticone (133), Laboratory of Cardiovascular Science, National Institute on Aging, National Institutes of Health, Baltimore, Maryland 21224 Craig Neville (85, 405), Cardiovascular Research Center, Massachusetts General Hospital-East, Charlestown, Massachusetts 02129 Paul Opolon (229), Unit6 de Pathologie Exphrimentale de 1’Institut Gustave Roussy, URA 1301 CNRS, Institut Gustave Roussy, PR2, 94805 Villejuif France Charles P. Ordahl(3),Department o f h a t o m y and Cardiovascular Research Institute, University of California at San Francisco, San Francisco, Cahfornia 19143 Rebecca R. Pauly (133),Laboratory of Cardiovascular Science, National Institute on Aging, National Institutes of Health, Baltimore, Maryland 21224 Michel Perricaudet (229), URA 1301 CNRS, Institut Gustave Roussy, PR2, 94805 Villejuif, France Thierry Ragot (229), URA 1301 CNRS, Institut Gustave Roussy, PR2, 94805 Villejd, France

Contributors

...

Xlll

Thomas A. Rando (261),Department of Molecular Pharmacology, Stanford University School of Medicine, Stanford, California 94305 Glenna Rice (155), Muscle Biology Group, Animal Sciences Department, University of Arizona, Tucson, AZ 85721 Nadia Rosenthal (85, 405), Cardiovascular Research Center, Massachusetts General Hospital-East, Charlestown, Massachusetts 02129 Elizabeth Rust (307), Department of Physiology, University of Michgan, Ann Arbor, Michigan 48109 Giovanni Salviaaf (349), Department of Biomedical Sciences and C N R Center of Muscle Biology and Physiopathology, University of Padua, 35121 Padua, Italy Moshe Shani (373), Institute of Animal Science, Agricultural Research Organization, The Volcani Center, Bet Dagan 50250, Israel Shannon M. Sheehan (155), Muscle Biology Group, Animal Sciences Department, University of Arizona, Tucson, AZ 85721 Stefan0 SchiafEno (349), Department of Biomedical Sciences and C N R Center of Muscle Biology and Physiopathology, University of Padua, 35121 Padua, Italy Didier Y. R. Stainier (67), Department of Biochemistry and Biophysics, University of Cahfornia at San Francisco, San Francisco, California 94143 H. Lee Sweeney (275), Department of Physiology, University of Pennsylvania School of Medicine, Phdadelpha, Pennsylvania 19104 Constance J. Temm-Grove (155), Muscle Biology Group, Animal Sciences Department, University of Arizona, Tucson, AZ 85721 Monte Westerfield (67), Institute of Neuroscience, University of Oregon, Eugene, Oregon 97404 Margaret Westfall (307), Department of Physiology, University of Michigan, Ann Arbor, Michigan 48109

This Page Intentionally Left Blank

PREFACE

The study of muscle has provided numerous fundamental biological insights during the past two centuries. Much of intermediate metabolism, including the citric acid cycle, was delineated during the biochemical study of muscle. Many of the fundamental insights into cellular energetics, including the discovery of ATP, were derived from investigations on muscle. The physics of muscle contraction led to the discovery of the first described molecular motor-myosin-as well as the discovery of actin. Although it is now understood that the myosin of muscle is simply one member of a large superfamily of motors and that actin is a key component of all eukaryotic cells, much of our understanding of the function of these proteins is derived from work on muscle. More recently, muscle research has proven seminal in unraveling the processes of determining and specifying a differentiated cell type. The discovery of the MyoD family of myogenic determination factors has greatly enhanced our understanding of the differentiation process not only in muscle, but in a variety of other cell types as well. The study of myofibrillogenesis has yielded and will continue to yield basic insights into the nature of macromolecular assembly processes as well as the interactions between the cell cytoplasm and the surrounding extracellular matrix. Differentiated muscle cells, as well as satellite cells, hold great promise as targets for gene and cell therapies not only for muscle diseases, but in situations where the protein export machinery of muscle can be used to provide circulating proteins. In this volume we have collected methods that allow both in vivo and in vitro application of muscle to fundamental problems in cell and developmental biology. After these chapters were written, a number of sigmficant technical advances occurred that beg mention. In the area of viral vectors for gene delivery into muscle, two new vectors hold immense promise for muscle. First, several groups have succeeded in constructing a recombinant adenovirus that contains no viral genes and is capable of packaging and expressing a large transgene with diminished immune response, compared to conventional adenoviral vectors. Second, recombinant adeno-associated virus (AAV), in the absence of adenovirus, appears selectively to target the postmitotic cells, such as striated muscle cells, and allows persistent expression, possibly via integration. In the areas of muscle development, physiology, and regeneration, mouse genetic technologies are being used to knock in genes for lineage tracing, to test isoform function, and for conditional expression of both structural and regulatory proteins. In addition, knockout mutant mice are being generated at a rapid pace, making available for the first time a large collection of genetic variant mouse strains, which will be valuable resources in all aspects of muscle biology research. The zebrafish offers

Preface

an additional genetic model for the study of many fundamental problems in muscle biology and offers a new complement to the chick and frog, which continue to play major roles as experimental models for studying developmental and cellular mechanisms. Finally, there have been recent applications of quantitative PCR to the analysis of gene expression at the single cell level in satellite cells and in embryonic cells undergoing developmental change and challenges. These applications will make possible the detailed description of gene regulatory processes occurring within the cellular complexity of the vertebrate embryo. Although this is not an exhaustive list, it is meant to underscore the rapidity with which new methodologic approaches are being developed to meet the challenges of muscle biology in what is now a rapidly evolving field. The chapters of this volume are meant to provide a practical guide to the range of fundamental techniques used by muscle biologists and by cell and developmental biologists working in related fields. Charles P. Emerson, Jr. H. Lee Sweeney

PART I

Embry010gica1 Analysis of Myogenesis


CHAPTER 1

Avian Somite Transplantation: A Review of Basic Methods Charles P. Ordahl*and Bod0 Christt ' Department of Anatomy and Cardiovascular Research Institute University of California at San Francisco San Francisco, California 94143 + Anatomisches Institut

Albert-Ludwigs-Universitat-Frieburg 79001 Freiburg, Germany

I. Introduction 11. Materials

A. Acquisition and Care of Avian Embryos for Surgery B. Candling C. Solutions D. Micropipets E. Microscalpels and Microneedles F. Microdissection Dishes G. Tissue Holding Dishes H. Tissue Markers I. Sealing Tape 111. Methods A. Isolating and Preparing Donor Embryos for Microsurgery B. Isolation of Quail (Donor) Somites C. Preparation of Chick (Host) Embryos for Somite Implantation D. Implantation of Donor Somites into Host E. Transplantation of Half-Somites F. Analysis of Chimeric Embryos IV. Summary and Concluding Remarks References

METHODS IN CELL BIOLOGY, VOL. 52 Copyright 0 1998 by A c d e m c Press. ALI nghD of reproduction in any form rescrvcd. 0091-679X/98 $25.00

3

Charles P. Ordahl and Bod0 Christ

I. Introduction The avian embryo has a long history of use in muscle research, from the earliest days of in vitro cell culture and the analysis of cytodifferentiation (Burrows, 1910; Lewis and Lewis, 1917). The surgical accessibility of the avian embryo (Hamburger, 1960; Hamilton, 1952; Rugh, 1962), moreover, provided a fertile ground for the study of embryonic development. Twenty-five years ago, the utility of the avian embryo increased dramatically with the development of quailchick grafting (Le Douarin, 1973), a method for tracing the developmental fate of embryonic anlage. The densely brilliant magenta nucleoli of the Japanese quail (Coturnh coarnix juponicu) in Feulgen-stained sections distinguish quail nuclei from those of chick in surgical chimeras. Since the quail nucleolar marker is heritable, the fate of cells descended from transplanted tissues can be traced indefinitely. This marking system has led to the current detailed understanding of vertebrate cell lineages. Inspired by a poster on quail-chick grafting by Nicole Le Douarin, one of us (BC) began a series of experiments in which the somites of a chick embryo were replaced with those of a quail. The original intention was to study the contribution of the somite to the vertebral column, but unexpectedly, the muscle tissue of the chick wing buds was found to be populated by quail cells (Christ et uL, 1974). This result supported early ideas about the somitic source of limb muscle while undermining many popular contemporaneous theories (Caplan, 1981; Zwilling, 1968). Subsequent employment of the basic somite transplantation technique established that all skeletal muscle in the body is derived from somites (Chevallier et ul., 1977,1978; Christ et aZ., 1974,1977,1978,1983). Moreover, this approach has also allowed a wide range of biological questions to be addressed, from the cellular localization of genetic defects in muscle (Kieny, 1983) to the molecular analyis of myogenic specification (Williams and Ordahl, 1994). Furthermore, by combining the quail-chick grafting technique with in situ localization of specific mRNAs or proteins, generalizations between avians and other vertebrate orders can be made. For example, this approach has been used to provide the first experimental evidence that the somite-to-limb muscle migration also occurs in mammalian embryos (Williams and Ordahl, 1994). Thus, the avian embryo has continuing potential for the analysis of all phases of vertebrate myogenic development. In recent years the development of increasingly discriminating antibody and nucleic acid probes enhances this potential by allowing the expression of specific macromolecules to be assigned to marked myogenic precursor cells within developing embryos. Moreover, the quail-chick grafting strategy can be utilized for “challenge assays” to analyze the acquisition and partitioning of developmental potential in marked myogenic precursor cells as they undergo the steps of specification. In such experiments quail embryo tissue fragments are transplanted into regions of the chick embryo that are permissive, nonpermissive, and/or inductive for specific developmental pathways.

1. Avian Somite Transplantation

5

The heritability of the quail nucleolus ensures that transplant-derived cells can be recognized regardless of their ultimate developmental fate. Such challenge assays are essential to assess the developmental plasticity of cells as they progress through embryonic cell specification and morphogenesis. Thus, the surgical accessibility of avian embryos make them an important experimental system for analysis of aspects of development that are difficult or impossible to address in other systems. Here we review the basic procedures used for transplantation of quail somites into chick embryos. Two basic methods are outlined: One is based upon techniques developed recently by one of us (CPO) while on sabbatical in Dr. Le Douarin’s Institut d’Embryologie outside of Paris. The other was developed independently in Germany by BC over the past 20 years. Both techniques give very satisfactory results and can be combined to yield increased flexibility in embryo surgery.

11. Materials A. Acquisition and Care of Avian Embryos for Surgery

Fertile chicken and quail eggs can be obtained from local hatcheries. In the United States, quail eggs can be ordered from Strickland Quail Farm ($0.55 per egg; telephone 912-748-5769). Eggs are stored at 4°C for up to 1 week and then incubated at 37-39°C in a humidified (60-80%) incubator. Chick eggs are incubated on their “sides” and are carefully maintained in their incubated orientation during windowing and surgery. Donor quail eggs may be incubated either with the narrow end down or on their sides. Commercial suppliers usually wash chicken eggs, but unwashed eggs give better viability. B. Candling

Candling is a method for visualizing the chick embryo within its shell using a strong light. Various boxes can be made for this purpose that have a light bulb in the bottom and an egg-sized hole in the top. It is also possible to candle an egg using a fiber-optic lamp held under the egg in a darkened room. In either method, the blastoderm appears as a reddish glow or spot, whereas the rest of the yolk appears yellow. The position of the blastoderm can be marked on the eggshell with a pencil to identify the site at which to cut the window in the shell for surgery. C. Solutions

Several isotonic buffers have been developed for work with avian embryos. The two formulas described below are equally suitable for the surgery described here.


6

Tyrode’s solution (Tyrode, 1910) is a balanced salt solution for avian embryos that is widely used in the United States and Europe. Its formula is as follows: 1.8 mM CaCl,; 1.1 mM MgC12; 2.7 mM KC1; 137 m M NaC1; 0.4 mM NaPO,; 5.6 mM D-glucose; 12 mMNaHC03;pH 7.2-7.6 adjusted with HC1 and/or NaOH. This formulation is adapted from Sigma Chemical Company’s instructions for Tyrode’s salts (Catalog No. T-2145). Another good balanced salt solution for avian embryos is Locke’s solution (Locke and Rosenheim, 1907): 2.2 mM CaC12; 5.4 mM KC1; 154 mMNaCl; 2.4 mM NaHC03 (NaHC03is often omitted without detriment). This formulation is adapted from Rugh (1962); see page 407. Pancreatin is a crude enzyme preparation (Sigma Cat. # P-3292) used to digest extracellular matrix in donor and host embryos. Pancreatin solution (4X) can be used straight from the bottle or diluted (1 :3) with either Tyrode’s or Locke’s solutions. It is particularly important to inactivate and remove any pancreatin used in host embryos. This is done by aspirating the enzyme solution and then flooding the area with fresh Tyrode’s solution.The Tyrode’s rinse is then aspirated and the rinsing-aspiration cycle repeated at least two more times. We typically apply fetal bovine serum (10%v/v in Tyrode’s) to the area treated with pancreatin prior to the rinses with Tyrode’s, but the efficacy of serum to quench pancreatin action has not been empirically determined. A 2%solution of serum is used to hold donor tissue fragmentsprior to implantation in hosts. The serum helps quench any pancreatin and also helps to prevent adherence of tissue fragments to glass or plastic surfaces. When moving tissue fragments from holding drop to host, it is advisable to draw the fragment into a transfer pipet charged with unadulterated Tyrode’s solution. This will ensure that tissue fragments do not float in the Tyrode’s solution within the surgical environment in ovo. D. Micropipets

Micropipets are used for delivering buffers, enzyme solutions, and tissue fragments. We prepare mouth-operated micropipets by pulling borosilicate (not flint) glass microcapillary tubes (100 pl size; Fisher No. 21-164-28) over a small flame. The tips of pulled pipets are broken off to achieve the desired diameter, typically about 50-100 microns, or occasionally larger (see Fig. 1A). Micropipets are connected to a mouthpiece using rubber or plastic hosing. The movement of solutions or tissue fragments within the micropipet is then controlled by mouth by connecting it to a rubber or plastic hose attached to a mouthpiece. A convenient and inexpensive mouthpiece can be fashioned using a pipe stem with room for a filter. The pipe stem allows the mouthpiece to be gripped with the teeth while the filter blocks saliva flow into the hosing. Inexpensive pipes and filters can be ordered from Medico (Department L, PO Box 789, Peekskill, NY 10566). Spemann pipets (Fig. 1B) allow the movement of solutions and/or tissue fragments to be controlled by hand rather than by mouth. To make a Spemann pipet, the small end of a Pasteur pipet is closed using a flame. Next, an area near the tapered region of the pipet is heated over a flame while pressure is

7


A

e -

\ opening

bulb for I coarse I

-1mm

Rubber membrane for fingertip cont ro I

Side view Fig. 1 Tools for moving embryos and tissues for microsurgery. (A) Mouth-operated pipet; (B) Spemann pipet; (C) perforated spoon.

applied by mouth at the large end of the pipet until a bubble forms. The bubble is then broken and the edges fire-polished, creating a smooth hole in the distal end of the wide portion of the pipet (see diagram). The tip of the pipet is then reopened and, if desired, bent to a convenient angle. A rubber bulb is attached to the wide end and a rubber sleeve (fashioned from tubing or a cut pipet bulb) is placed over the hole. The sleeve should be easily deformable. The rubber bulb is used to charge the pipet with buffer solution. Depressing the sleeve over the hole allows tissue fragments to be moved into and out of the pipet tip in a gentle, controlled manner. Cell injection electrodes are micropipets that can also be used to good advantage in embryo surgery. The very fine walls of these pipets allow tip bores of extremely small size. We pull FHC pipets (Cat. No.30-31-0; FHC, Brunswick,


8

ME 04011; telephone 207-729-1601)on a conventional electrode puller and then break the tips to the desired size (5-50 microns) by gently dragging them across soft tissue paper. When mouth-operated, these pipets are useful for precise, localized targeting of enzymes or for aspirating small tissue fragments that are difficult to remove cleanly with microneedles or microscalpels. E. Microscalpels and Microneedles

A variety of micro cutting tools have been used for embryo microsurgery, from cactus needles to finely drawn glass needles. Extremely fine microneedles (see Fig. 2A) can be conveniently prepared by electrolysis of tungsten wire (Davies, 1989). Tungsten wire can be ordered from Goodfellow (telephone 800821-2870; Cat. No. 005155; 0.38-mm dia, 99.95% purity). Tungsten microneedles have an extended thin taper with a tip that is often invisible, even under high stereoscopic magnification. The shape and thinness of the tungsten microneedle dictate two basic cutting strokes: First is the downward slash, in which the fine edge of the microneedle is drawn downwards through (or between) tissues. The second stroke, the upward slash, consists of first inserting the tungsten microneedle tip at an angle into, or between tissues, and then drawing the tip smoothly but sharply upwards. The upward slash requires care; the often invisible microneedle tip can easily be inserted more deeply than intended, causing unnecessary tearing of underlying tissues. As an alternative to microneedles, the Le Douarin lab developed microscalpels (Fig. 2B) in which the tips of stainless steel sewing needles were sculpted to fine blade shapes upon fine-grain Arkansas sharpening stones. A new method for creating tungsten microscalpels by electrolysis (Conrad et aL, 1993) now allows fine microscalpels to be prepared by those without the patience to learn to sculpt microneedles by hand. Microscalpels can be manipulated in a manner similar to that of a paring knife, because they are relatively rigid and have both cutting and flat edges. The cutting edges can be used to score, snip, or slash tissue, although the last is not recommended because it typically results in damaged tips. The flat edge, on the other hand, can be used to push or tease tissues apart. Because of the shape of the microscalpel, the tip is highly visible and is the only portion actually used for cutting. Other microtools useful for embryo surgery, such as forceps, microscissors, and perforated spoons (Fig. lC), can be obtained from Fine Science Tools (telephone 800-521-2109). F. Microdissection Dishes

Small glass concave embryological culture dishes (Cat. # 910B; Variety Glass (telephone 614-432-3643) are filled halfway with black Dow Corning Sylgard (KRAnderson Co., Cat. No. 184, with black pigment No. 1747; telephone 800672-1858). Using these dishes, isolated embryos can be held in place for surgery

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A Needle cross section-1

-

/

0

Tungsten needle

I I

The basic stroke

for a microneedie Is the slash

+d

slash Contact

Downward pressure

c3+ More pressure

s1a=4 +,+ +4

Upward

/

Upward

insert ion

More pressure

pressure

B Knife cross

knife

I Three types of strokes with microscalpel :

Score

Snip

Tease (with flat edge)

Fig. 2 (A) Microneedles and (B)microscalpels.


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or fixation using stainless steel insect pins. The black Sylgard provides good contrast for visualizing the white/translucent tissues of the embryo. Dishes exposed to fixatives should never be used for surgery. Microdissection dishes can also be made by filling plastic culture dishes with a 2-mm-thick layer of 2% agarose. Embryos are then spread out over the agarose and fixed in place by pressing the extraembryonic membranes into the agarose layer with a tungsten needle. Contrast is provided by placing a black background under the microdissection dish. G. Tissue Holding Dishes

Holding dishes are used to hold donor tissue fragments until needed. A cheap method is to use a plastic culture dish containing a droplet of holding solution (2% serum in Tyrode’s solution). The tissue fragment is transferred into the droplet where it can be visualized microscopically. To avoid evaporation, the dish lid should be replaced while the tissue is held. Tissue fragments can be held for up to 2 hr at room temperature. To transfer the tissue fragment to the host embryo, a micropipet charged with Tyrode’s solution (without serum) is inserted into the droplet and, under microscopic examination, a small amount of Tyrode’s solution is pipetted over the tissue fragment. The fragment is then gently aspirated into the tip of the pipet. These steps reduce the carryover of serum-containing medium, which can affect the buoyancy of the tissue fragment during implantation into the host embryo.

H.Tissue Markers Donor tissue is most easily marked for orientation purposes by the inclusion of asymmetric fragments of adventitious tissue with known orientation. For example, somites carrying either ectoderm or endoderm are easily oriented in the dorsoventral plane. Cranial-caudal orientation, on the other hand, can be established by including a fragment of an adjacent somite. This approach is outlined in the methods described here because it is easy to perform and highly reliable. As outlined next, however, the use of dyes and/or other markers can also be useful. Nile Blue sulfate is a vital dye that can be used to stain donor and host tissue. A glass rod is heated and pulled out to give a finely pointed end, which is then polished over a flame to give a small glass ball on its end. The glass ball is then dipped in melted (70°C) 2.5% agarose containing 1%Nile Blue sulfate and then removed to allow the agarose to cool. This is repeated 5-6 times until the glass ball is well coated with the Nile Blue sulfate/agarose. This staining instrument can be stored at 4°C for several weeks. To apply the dye, the tip of the glass ball is applied directly to tissue for a few seconds. The Nile Blue sulfate will stain the tissue.


11

Animal carbon (ash from animal tissue) can also be used for marking tissue (see, for example, Ordahl and Le Douarin, 1992). Animal carbon, however, is not easy to obtain in the United States. Moreover, carbon marking is difficult because of the nonuniformity of the carbon particles and their low adhesivity to tissue. I. Sealing Tape

Windows and other holes in eggshells can be sealed with tape. Scotch-type tape works well, but because of its stiffness, pleats are necessary to conform to the curved surface of the eggshell. Such pleats must be carefully sealed to prevent moisture loss. Leukosilk tape (Beiersdorf;Hamburg, Germany) is cloth tape that is easily conformable to eggshell contours that also provides a good moisture barrier.

III. Methods A. Isolating and Preparing Donor Embryos for Microsurgery

The shell of an incubated quail egg is fist disinfected with 70% ethanol. Then a small hole (5-10 mm) is cut into the shell at the extreme tapered end with curved scissors and the clear albumin decanted (see Fig. 3A). If the stringy white chalazae emerges through the hole, it can be cut with scissors to facilitate decantation. The removal of albumin lowers the yellow yolk so that the hole can be enlarged to allow the yolk to be gently decanted into a bowl containing Tyrode’s or Ca*+-freeLocke’s solution. The bath should be of sufficient depth to allow the yolk to float and the small white blastoderm (2-5 mm diameter) to rotate up. A forceps in one hand is used to steady the yolk, while an iridectomy scissors in the other hand is used to cut the yolk membrane around the blastoderm. Thus liberated, the blastoderm may be lifted away from the yolk using fine forceps, picked up with a perforated spoon, and transferred to a microdissection dish filled with Tyrode’s solution (Figs. 3B and 4A). If the milky vitelline membrane remains adhering to the upper surface of the blastoderm, it should be removed with a fine forceps and discarded. Removing yolk platelets with a Pasteur pipet will improve visibility. The embryo blastoderm is then pinned dorsal side up to the plastic bottom of the microdissection dish as described in Materials (Fig. 4a). 1. Staging Embryos and Counting Somites Chick embryos are staged according to a number of systems, the most widely used of which is the Hamburger and Hamilton (HH) system (Hamburger and Hamilton, 1951).Among the criteria for HH staging is the counting of the number

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Charles P. Ordohl and Bod0 Christ

A

0 Yolk

1

Cut shell

Cut around blastoderm with iridectomy scissors

Tyrode's or Locke's solution

B

C Method 1

Method 2 AI bum in level with

r

Window in

Albumin Collapsed Albumin removal to lower embryo under window

Buffer injection to raise embryo up to window

Fig. 3 Egg and embryo manipulation for surgery. (A) Removing yolk born quail (donor) egg. (B) Isolating blastoderm in dissection dish. (C) Positioning embryo for surgery in OVO.


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Fig. 4 Appearance of quail and chick embryo during surgery. (A) The 16-somite quail embryo has been removed from the egg, rinsed free of contaminating debris, and staked out in a blackbottomed Sylgard-coated dish with insect pins, as described in the text. The embryo is located centrally within the clear area pellucida and is surrounded by the densely white area opacu, the margins of which already contain rust-colored blood islands. Other features of the embryo are indicated. (B) The 17-somite chick embryo is viewed through a window in the eggshell after black ink has been injected under the blastoderm, as described in the text. Somite 3, the first clearly demarcated somite in an embryo of this age, is indicated. The surgical target area is schematically represented with the somite number and somite stages indicated. (See also Color Plates.)

of formed somites. Ten somites have formed in Hamburger Hamilton stage 10 (HH lo), although only eight somites are clearly visible owing to the complete disintegration of somite number one and partial disintegration of somite number two. Therefore, in embryos HHlO or older, the cranialmost, clearly delineated somite is somite number 3 (see Fig. 4B). Somites are numbered sequentially in a cranial-to-caudalsequence using arabic numerals. By HH 14, when the cranialmost somites are obscured by curvatures of the embryo, numbering of the visible somites can be deduced from the position of the omphalomesenteric artery, which typically exits the embryo body adjacent to somite number 21.

Charles P. Or-

14

and Bod0 Christ

2. Somite Stages Somites sequentiallybud off from the cranial end of the segmentalplate. Thus, in an embryo of any age, the caudalmost somites are the most recently formed and least mature developmentally. Somite staging provides an index of somite maturity with the most recently formed (or caudalmost) somite designated as a “stage I somite” and the next somite cranially as a “stage I1 somite,” and so on (Ordahl, 1993). Somite stages are diagrammatically illustrated in Figs. 4B and 5A.In general, somites at stage I or I1 are relatively plastic in their developmental potential, whereas at later stages of somite development progressive specification of specific somite lineages and structures occurs. The procedures outlined here target only the most recently formed (stage I and 11) somites. Finally, only brachial somites (somites number 15-20) are targeted in the experiments described here because they are highly accessible surgically and are well understood developmentally. Thus, because two adjacent somites are transplanted in the procedures outlined later, embryos with a total of 16 to 20 somites should be selected for these procedures. Surgery targeting somites at other axial levels, or other stages of somite development, would require some modifications of the techniques described here.

B. Isolation of Quail (Donor) Somites In this step, stage I and I1 somites, and a fragment from the caudal portion of somite I11 (as an orientation marker) will be cut free of surrounding tissue on the adjacent stage I11 somite the right-hand side of the quail embryo. The liberated somites are then teased out of the embryo, aspirated into a transfer pipe, and transferred to a holding dish. Two methods are outlined: the first employing microscalpels and enzymes, and the second employing microneedles without the use of enzymes. Both methods involve the same important sequence of incisions. Although both methods have advantages and disadvantages, they yield comparable results. In many cases the basic procedures can be recombined to give increased flexibility in surgery. Hand stability is crucial for embryo microsurgery under stereoscopic ma@cations reaching up to 1OOX. Most workers find it advantageous to have their hands in contact with each other during surgery, which increases stability. In a right-handed individual, for example, the left hand has the primary responsibility of embracing and stabilizing the holding dish (or host embryo eggshell). The right hand rests against both the microscope stage and the left hand, while wielding the microscalpel or microneedle between the thumb and forefinger. Some workers find that an elevated support for the wrists alleviates fatigue. 1. Method 1: Donor Somite Isolation using Microscalpels and Enzymes In this first method, target somites are removed in three steps. The first two steps are diagrammed in Figs. 5A and 5B. First, the ectoderm overlying the somites is removed. Second, four incisions are made medial, lateral, cranial, and


15

Fig. 5 Incisions used for somite surgery. (A) Incisions in superficial ectoderm for skin removal. (B) Deep incisions for somite removal. (C) Side views of isolated somites. In Method 1, tightly adherent ventral endoderm serves to hold somitestogether and aids in orientation during implantation in host. In Method 2, ventral endoderm may be dissected away, leaving tightly adherent dorsal ectoderm to hold somites together and aid in orientation. (D) Incisions for medial half-somite isolation. (E) Incisions for lateral half-somite isolation. (F) Isolated somites are used to separate dorsal and ventral half-somites with a fine microneedle.

caudal to the target somites. Finally, the somites are freed from the underlying endoderm either by teasing or by cutting out the patch of endoderm to which they adhere. Microscalpels are used for all but the final steps, and advantage is taken of the shape of a microscalpel to perform different types of surgical strokes. In addition, the enzyme pancreatin is used to digest extracellular matrix to augment surgical manipulations. a. Step 1: Removal of Skin Ectodem over Target Somites Four shallow incisions are made to circumscribe a rectangular patch of skin ectoderm over the target somites (Fig. 5A).The first incision is made in the skin

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Charles P. O r W and Bod0 Christ

ectoderm between the neural tube and the caudalmost somites and cranialmost segmental plate. A second parallel incision is made in the skin ectoderm lateral to the somites. The cranial and caudal ends of these first longitudinal incisions are then connected by two parallel transverse incisions. Each incision can be aided by first “scoring” the ectoderm repeatedly (3-10 times) by dragging the cutting tip of the microscalpel along the surface (Fig. 2B). Then, to complete the incision, the tip of the microscalpel is inserted into the scored ectoderm and “snipped” by sharply flicking it up and away (Fig. 2B). This is repeated along the line of incision until it is complete. Small tissue fragments can adhere to and blunt the microscalpel. These can be easily removed without damaging the microscalpel by submersing the tip in a sonicatingwater bath for 2-5 sec. Microscalpelsshould never be passed through a flame for cleaning because this will damage the tip and edge. After the four incisions are completed, the rectangular patch of skin ectoderm is “teased” away from the underlying tissues using the flat side of the microscalpel tip (see Fig. 2B). This can be facilitated by pipetting pancreatin (1X or 4X) directly over the incisions. The heavier enzyme solution will fall onto and cover the target area. The enzyme solution is left in place for 0.5-2 minutes to digest extracellular matrix material. During this period, the edges of skin ectoderm are teased up and away. After removal of the skin flap, the action of pancreatin is quenched by addition of 10% serum followed by liberal rinsing of the surgical area with fresh Tyrode’s solution. The exposed somites, segmental plate, and intermediate mesoderm are now more clearly revealed and accessible to surgery. b. Step 2: Excising Somites I and II and the Caudal Half of Somite ILZ This step involves 4 incisions, similar to those made in the skin ectoderm except that these incisions cut deeper, through the entire mesoderm layer. These incisions are diagrammed in Fig. 5B. The sequence of the incisions (medial first, then lateral, then two transverse) is important to exploit the mechanical advantages of the surrounding tissues. Once these four incisions are made, either the target somites can be teased away from the underlying endoderm, or the endoderm itself can be cut to liberate it with the somites still attached. Fimt Incision:Sepamtion of the Somitesjhm the Neuml Tube. Extracellular matrix binds the medial borders of the somites closely to the adjacent neural tube, and this association must be disrupted in order to remove the somites cleanly. First, the interval between the neural tube and somites is scored to break the fine strands of extracellular matrix that span this space. Next, the tip of the microscalpel is inserted between the tip of the segmental plate and neural tube and the flat side of the scalpel used alternately to push the lateral wall of the neural tube medially and the medial wall of the stage I somite laterally. The microscalpel is then moved incrementally cranially and these teasing motions repeated. This is continued along the entire interval between the target somites and the adjacent neural tube. Although the teasing action is the primary means


17

of separating the axial structures (neural tube + notochord) and somites, the sharp edges and tip of the scalpel can also be used to cut any recalcitrant strands of matrix to ensure that the separation extends ventrally to (but not through) the endoderm. When this step is completed, the scalpel should pass between the space between the somites and neural tube without resistance. Pancreatin can be used sparingZy to facilitate this separation. It must be kept in mind that it is desirable for adjacent donor somites to remain attached to each other and to the underlying endoderm. Excessive use of pancreatin can weaken these attachments. Second Incision: Incision betweenSomites and Intennediate Meso&m The Wolffian duct serves as a marker for the intermediate mesoderm because it lies on its surface, immediately beneath the skin ectoderm. A longitudinal incision through the mesoderm at the immediate lateral edge of the somite and medial to the Wolffian duct will therefore separate the somites from the intermediate mesoderm. The incision line should be first scored and then snipped, in a manner similar to that used for the ectoderm incisions. When fully completed, this incision should completely separate the paraxial and intermediate mesoderm, and extend ventrally all of the way to (but not through) the underlying aorta and endoderm. Since the cells of the lateral margins of the somites and medial margins of the intermediate mesoderm are somewhat intermixed, the actual line of this incision is less precise than that between neural tube and somites. It therefore may not be possible to remove somites without at least a little intermediate mesoderm attached. Third and Fourth Incisions: Two Tmnsveme Incisions Cranial and Caudal to Target Somitw: In this step, transverse incisions are made cranially and caudally to the the target somites to separate them from the paraxial mesoderm. These incisions can be made in the intersomitic clefts. It is advisable, however, to make the cranial incision through the the middle of the stage I11 somite. In this case, a fragment of extra somite tissue is left attached, thereby marking the cranial end. This can be a useful orientation marker when the somites are implanted into a host embryo. Other methods of marking involve application of dyes, such as Nile Blue sulfate, or carbon particles to one surface of target somites. Since the target somites are still firmly held in place prior to the transverse incisions, such marking strategies should be considered prior to making the transverse incisions. c. Step 3: Removal of Somites and Transfkr to Holding Dish Once the transverse incisions are completed the target somites are only held in place by strong attachments to the underlying aorta and endoderm. The somites can be removed by teasing them away from the endoderm by lifting with the flat edge of the microscalpel. Pancreatin can facilitate this, but runs the danger of disrupting attachments between adjacent somites. An alternative method is to slash through the underlying endoderm using the same medial-lateral-transverse sequence of incisions described before. The

18

Charles P. Ordohl and Bod0 Christ

slashing type of incision is most easily made using a microneedle (Fig. 2A) because a microscalpel is almost always damaged by this procedure. After the four incisions, the target somites with underlying endoderm attached are teased out of the host. The advantage of this second method is that the attached endoderm provides a good orientation mark for the ventral side of the donor somites (see Fig. 5C). After the somites are teased out of the donor, they are aspirated in a micropipet and transferred to a droplet of 2%calf serum (in Tyrode’s solution) contained in a holding dish. Isolated somites can be stored in this manner at room temperature for up to 2 hr.

2. Method 2: Donor Somite Isolation using Microneedles and without Enzymes This method differs from the previous one in two major ways. First, microneedles are used rather than microscalpels.Because of their shape, microneedles are highly bendable if they are long and, as a result, the most productive cutting stroke is a “slash.” Second, no enzymes are used. The sequence of microneedle incisions made to remove the somites is exactly as described for method 1and illustrated in Fig. 5B. A major difference,however, is that the incision stroke passes sequentially through the ectoderm, mesoderm, and endoderm. The tip of the microneedle is positioned on the surface ectoderm between the target somites and the neural tube. The microneedleis then smoothly and slowly pushed downward in a “slashing” motion between the neural tube and somites (Fig. 2A). The slashing motion cuts sequentially through (i) the surface ectoderm, (ii) the extracellular matrix between somites and neural tube, and (iii) the underlying endoderm. The second incision is made in a similar way between the Woman duct and the lateral edges of the target somites. Finally, parallel caudal and cranial incisions are made, taking into consideration the inclusion of excess tissue fragments for orientation. Microneedles also become contaminated with extracellular matrix material and tissue fragments that dull them. These microneedles can be cleaned by sonication or by passage through a small flame. The donor somites, with attached ectoderm and endoderm, are then teased out of the blastoderm and moved to a peripheral region of the dissection dish. When viewed from the side (see Fig. 5C), the whiter and thinner endoderm on the ventral surface of the somites can be distinguished from the thicker and more translucent ectoderm.The endoderm can be removed using slashing motions with the microneedle. The ectoderm should be left in place to help hold the somites together during transfer and implantation and to act as an orientation marker for the dorsal surface. The donor somites are then transferred by pipet to a holding dish containing Locke’s solution. C. Preparation of Chick (Host)Embryos for Somite Implantation Chick embryo surgery is performed through a small window cut in the side

of the eggshell. Two methods of doing this are outlined. In the first, the position


19

of the embryo within the eggshell is lowered by removing albumin. A large window is then cut in the shell over the embryo, giving a wide surgical field. Ink is injected under the embryo to provide contrast. The host is then operated upon using microscalpels and enzymes. In the second method, a relatively small window is cut in the shell and the embryo raised into it. Low-angle light, rather than ink, is used to visualize the embryo in this method. The surgery is then performed using microneedles without the aid of enzymes.

A. Method 1: Large Window with Lowered Embryo a. Step f :Lowering of Embryo by Removal of Albumin Chick eggs are washed with 70% ethanol. A small hole is punched in the eggshell at the narrow end using the tips of forceps or scissors. An 18- to 20-gauge needle fitted to a 10-cc syringe is then inserted through the hole and 0.5 to 2.0 ml albumin aspirated. This lowers the yolk a few millimeters from the shell membrane (Fig. 3C). b. Step 2: Windowing the Embryo Using a small curved pair of scissors, a 1-2 cm diameter hole is cut in the eggshell and the subjacent eggshell membrane directly over the position of the embryo. Particles of eggshell that fall onto the egg will lie on the vitelline membrane, a tough, acellular membrane covering the embryo. A few drops of Tyrode’s solution are applied to moisten the vitelline membrane over the embryo and the larger shell particles removed using a fine forceps. The embryo appears white or translucent within the urea pellucidu (Fig. 4B), which is surrounded by the whiter area opacu where orange-colored blood islands and vessels are beginning to form (Fig. 4A). Inking the Embryo Pelikan (Hannover, Germany) Drawing Ink A No. 17 black mixed 1:1 with Tyrode’s solution is drawn up into a fine glass micropipet and 10-20 p1 injected under the blastoderm. The needle is inserted into the area opucu and then angled under the midgut of the embryo, and ink is gently deposited there by gentle mouth pressure. As the ink spreads, the embryo appears white against the blackened background (Fig. 4B). Deposition of excessive volumes of ink should be avoided as it will lead to siphoning of the ink out of the urea pellucidu. The vitelline membrane can also be more easily seen now, particularly at the site of ink injection. The injection hole in the vitelline membrane, or a new hole created with a microscalpel, can be widened using a fine forceps and the entire vitelline membrane removed and discarded. This will give a much clearer view of the embryo allowing the number of somites to be counted and the target area for surgery to be identified. Considerations regarding the number of formed c. Step 3:

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Charles P. Ordaht and Bod0 Chrirt

somites and target area are essentially identical to those outlined earlier for the quail donor embryo. d. Step 4: Skin Incision A singre incision only is made in the skin ectoderm between the target somites and neural tube (see inset, Fig. 4B)using the same scoring and snipping methods as described for the donor embryo. Pancreatin is applied to the incision for 0.52 min, while the tip of a scalpel is used to tease the skin flap away from the underlying somites and other tissue. The skin flap should be teased away enough to allow complete exposure of the somites. It will quickly re-attach and grow back after surgery. e. Step 5: Somite Incisions With enzyme still in place, and working with deliberate speed, four incisions are made around the target somites in the same sequence and manner as employed for the donor embryo (see Fig. 5B),taking care not to cut the endoderm. The enzyme aids in the separation of the somites from the neural tube and from the endoderm and aorta. The liberated somites are then teased out of the host, aspirated using a micropipet, and discarded. The surgical area is then flooded with 1-10 pl of 10% serum in Tyrode’s solution, which is then aspirated and discarded. Three flooding/aspiration washes of Tyrode’s solution follow. Tyrode’s solution is then applied and left in place. The host embyro should be used for transplantation as quickly as possible (within 30 min) to prevent healing of the incisions prior to implantation.

2. Method 2: Small Window with Raised Embryo The initial steps of this method are diagrammed in Fig. 3C. First, the chick egg is candled and the position of the embryo marked by pencil on the shell. The surface of the shell is then disinfected with 70% ethanol. A he-toothed saw (such as a he-toothed hacksaw blade) is used to saw a small rectangle in the shell over the embryo, taking care not to extend the saw cut into the underlying eggshell membrane. A second saw cut in the shell is made at the wide end, over the air sac, again without cutting the shell membrane. The square piece of eggshell over the embryo is then removed with forceps, again taking care not to cut or disrupt the underlying shell membrane. The shell membrane is then covered with Locke’s solution. Under the stereomicroscope a small hole is made in the shell membrane with a forceps or needle, and the Locke’s solution can be seen to fall between the shell membrane and the vitelline membrane directly over the embryo. The air sac is next opened by pushing a needle through the shell membrane at the cut previously made in the wide end of the eggshell (Fig. 3C). As the air sac collapses, the embryo falls away from the shell membrane, and the latter can be removed from the square hole without damaging the embryo. After the


21

shell membrane is removed, the embryo is re-elevated by dripping Locke’s solution into the square hole in the shell. When the embryo reaches the level of the shell, it is easily accessible for surgery. A dome of Locke’s solution should always cover the embryo to maintain hydration, but addition of excessive amounts of Locke’s solution should be avoided to prevent distortion of the embryo in the window. To provide good contrast, the lighting of the embryo should be from a low angle rather than from above. By this method, all regions of the embryo can be easily visualized without the necessity of injecting India ink under the blastoderm. The vitelline membrane is then labeled with Nile Blue sulfate and removed over the surgical target area. Too large a hole in the vitelline membrane is not advisable to avoid distortion of the embryo body due to bulging upwards. The ectoderm over the target somite area is then labeled with Nile Blue sulfate to make it more easily visible. The ectoderm between the neural tube and somites is cut using the upward slash of a microneedle (Fig. 2A). The tip of the needle is inserted through the ectoderm and into the potential space between the somites and neural tube. The needle is then slashed sharply upwards in a cranial direction, creating a cut between neural tube and somite. This is repeated until an incision in the ectoderm spans the two target somites. The ectoderm is then teased away in a lateral direction to expose the dorsal surface of the somites. If necessary to improve visualization, the dorsal surfaces of the somites can also be labeled with Nile Blue sulfate. The same sequence of cuts is made in the host as was made in the quail donor. However, the downward slashing motion used for the quail cannot be used for the chick host to avoid cutting underlying structures. Therefore, the fine tungsten needle is inserted at an angle and slashed in an upwards direction, as illustrated in Fig. 2A: first, between the neural tube and the medial boundary of the somite; second, between the lateral boundary of the somite and the intermediate mesoderm; and finally, transversely, at the cranial and caudal intersomitic clefts. The surgically isolated pair of somites is then carefully teased away from the underlying endoderm and aorta using a stiff tungsten needle. This teasing should proceed from medial to lateral. In cases where the embryo is bleeding from a tom aorta, the embryo can be cooled in a refrigerator for 5 min, which will reduce the beating of the heart and consequent bleeding. D. Implantation of Donor S o d t e s into Host

1. Method 1 Using a micropipet charged with Tyrode’s solution, the donor somites are aspirated from the droplet in the holding dish and pipetted onto the blastoderm of the host embryo. The deposition of the somites onto the blastoderm should be followed visually under the stereomicroscope to avoid losing them as they emerge from the micropipet.


22

The somites are then moved into position above the implant site using the flat edge of a microscalpel and then manipulated into the host site using a scalpel or a fine needle. Markings are noted at this time so that the orientation of the donor somites can be noted and/or altered. At this point, the quality of the graft should be assessed and recorded in terms of (i) orientation; (ii) alignment relative to contralateral somites; (iii) size and fit as compared to host somites; (iv) whether the skin flap closes over the transplant; and (v) any other damage or changes in the host or donor tissue. The window in the shell is then closed with tape and carefully sealed to prevent moisture loss. The embryo can now be incubated in a conventional egg incubator until time to harvest.

2. Method 2 Using a Spemann pipet, the graft previously labeled with Nile Blue sulfate is transferred onto the surface of the host embryo and teased into position using a tungsten needle. The markings are used to place the graft in the correct orientation. Once the graft is in place, the Locke’s solution meniscus covering the embryo is carefully removed using the comer of a small piece of filter paper. The skin ectoderm flap is then replaced over the graft site to help hold the donor tissue in place. Finally, the embryo is lowered by removing 2-3 ml of albumin by aspiration with a syringe via the small hole made in the wide end of the eggshell. Both holes are then covered with tape, to prevent evaporation and contamination, and the egg placed in an incubator.

E. Transplantation of Half-Somites Tranplantation of somite fragments can be used to elucidate the fate of different portions of the developing somite. Half-somite transplantation has proven particularly useful in this regard because the somite can easily be divided in half along each of its three axes. The techniques for performing these transplants are modifications of those outlined earlier. The connections between half-somite fragments, however, are somewhat more fragile than those between whole somites, so more care must be taken in their isolation and transfer.

1. Medial-Lateral Half-Somite Transplants The procedures outlined above can be modified to isolate either medial or lateral half-somite fragments by a method adapted from Ordahl and Le Douarin (1992). For medial half-somites, the same sequence of incisions is made except that the lateralmost incision 2 is made through the somite midline (see Fig. 5D). For lateral half-somites, the first incision is made in the somite midline with subsquent incisions made as outlined earlier for whole somites (see Fig. 5E). The sequence and position of these incisions is the same regardless of the method employed.


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2. Dorsal-Ventral Half-Somites An incision through the horizontal plane of the somite is best made on isolated whole somites that have the ectoderm and endoderm in place to provide stability and orientation. Thus, the no-enzyme method is preferable for this procedure. Isolated somites are manipulated onto their sides and the ectoderm and endoderm identified (see Fig. 5F). A fine microneedle is then used to divide the dorsal and ventral halves of the somites (Christ et al., 1978).

3. Cranial-Caudal Half-Somites Cranial and caudal somite halves can only be isolated as individual half-somite units (see Kalcheim and Teillet, 1989). The sequence of incisions is identical to that described earlier, that is, the final two incisions divide the target half-somite from its half-somite partner and the adjacent whole somite.

F. Analysis of Chimeric Embryos 1. Harvesting Chimeric Embryos The reincubation period (hours to days) depends upon the experimental goals. In general, embryos to be harvested are first irrigated with Tyrode’s or Locke’s solution, and then cut away from the yolk using iridectomy scissors. Using a perforated spoon inserted under the blastoderm, the embryo is elevated away from the yolk and transferred to a dish containing Tyrode’s solution. After detritus is washed away and excess tissue or membranes are cut away, the Tyrode’s solution is replaced with fixative. The choice of fixative can be important depending upon the type of assay to be performed. If the quail nucleolar marker is to be visualized by the Feulgen reaction in wax sections (Le Douarin, 1973), ice-cold Carnoy’s fixative (60% absolute ethanol, 30% chloroform, 10% glacial acetic acid) or Serra’s fixative (same as Carnoy’s, except chloroformis replaced with formalin) is recommended. However, if the embryos are to be subjected to antibody staining or in situ hybridization, or are to be cryosectioned,4% paraformaldehyde may be a more appropriate fixative. Other fixatives can also be valuable, and their suitability is usually determined empirically. After gentle washing with Tyrode’s, young embryos (2-3 days) are initially staked out in a dedicated fixation dissection dish in a manner similar to that described earlier for quail embryos. The Tyrode’s solution is aspirated and replaced with fixative and the embryos allowed to fix for 2-3 min before excess body tissue and extraembryonicmembranes are trimmed with iridectomy scissors. Embryos should be fixed for 30-60 minutes, but can be removed from the dissection dish after 10-15 minutes. Older embryos (4 days or older) can be washed in culture dishes and extraembryonic tissues removed and discarded. These embryos are then immersed di-

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Charles P. Ordnhl and Bod0 Christ

Fig. 6 Histologicaldiscriminationbetween quail and chick nuclei in chimericembryos. (A) Feulgenstained cross-section through a chimeric chick embryo 1 day after surgically replacing the lateral halves of Wing level somites with those of quail as described in the text. The neural tube and wing bud are out of the field to the left and right, respectively. Under light microscopy, the quail nuclei contain bright, magenta-colored nucleoli (black arrows labeled Q-1, Q-2, and Q-3), whereas the staining of chick nuclei is pale and diffuse (white arrows labeled C-1, C-2, and C-3). Nuclei marked C-1 and C-2 reside in cells of the dermomyotome and myotome, respectively. C-3 marks the nucleus of a sclerotome cell. Q-1 marks the nucleus of a cell in the lateral part of the dermomyotome. Q2 marks the nucleus of a dermomyotome-derived cell that has dissociated from the lateral edge of the dermomyotome at the onset of its migration to the limb bud (out of field on right). Q-3 marks the nucleus of a cell in the lateral sclerotome. Note that the medial-lateral boundary between chick and quail cells extends from the dermomyotome through the entire sclerotome. Note also that the cells in the lateral m a t margin of the myotome are chick derived, whereas all of the cells of the lateral dermomyotome and migratory cells are quail derived. (B) Feulgen-stained cross-section through a chimeric chick embryo 1 day after surgically replacing the dorsal half of a wing level somite with that of a quail as described in the text. The neural tube (nt) can be seen on the lefthand margin, whereas the Wing bud is out of the field to the right. Nuclei marked C-1, C-2, and C3 are those of chick cells in the dorsomedial, ventromedial, and lateral sclerotome, respectively. Nuclei marked Q-1 and Q-2 reside in the medial and lateral myotome regions, respectively. The nucleus marked Q-3 resides within a cell of the dermomyotome. Note that quail nuclei are restricted to dorsal somite derivatives, dermomyotome and myotome, and that chick nuclei are restricted to sclerotome, a derivative of the ventral somite half. (See also Color Plates.)

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rectly in fixative. The timing of fixation can be important. For fixation of larger embryos in Carnoy’s, a good rule of thumb is to incubate the embryo in a refrigerator until it falls to the bottom of the fixation vessel, or overnight, whichever comes first. For embedding, the orientation of the embryo is important to obtain the desired section planes. If the embryo is very small, the embedding should be performed under stereo microscopy.

2. Histology Typically, chimeric embryos are paraffin embedded, sectioned at 5-7 pm, and stained by the Feulgen method (Le Douarin, 1973). Under light microscopy, quail nuclei are distinguished by the presence of a large, luminously magenta nucleus (Fig. 6A). Black-and-white photography of the quail nucleolar marker is best captured using Agfa ortho film. Quail nuclei can also be visualized immunohistochemicallyusing monoclonal antibodies (Carlson, 1992, personal communication). The anti-quail monoclonal antibody “QCPN’ is not published but is available from the Developmental Studies Hybridoma Bank maintained by the Department of Pharmacology and Molecular Sciences at Johns Hopkins University School of Medicine, Baltimore, Maryland, and the Department of Biology at the University of Iowa, Iowa City, Iowa, under contract number NOl-HD-23144 from the NICHD.

IV. Summary and Concluding Remarks We have outlined two basic procedures for the transplantation of quail embryo somites into chick embryos. Although the two methods were developed independently, both employ a remarkably similar surgical strategy, even down to the sequence of incisions. The similarity in incision sequence to a large degree reflects mechanical constraints inherent to the tissues within the avian embryo. Nevertheless, combining materials or techniques from both procedures can exploit advantages and minimize disadvantages. In addition, these basic procedures can also be used to perform modified somite transplantations or transplantation of somite fragments. For example, we oulined half-somite (dp; d/v; d)transplantation strategies that have proven useful in revealing novel aspects of early development. In addition, experiments involving transplantation of fragments of more mature somites (e.g., sclerotome, dermomyotome) can be developed using these basic strategies. Undoubtedly, procedures with increased power and precision will continue to be developed. Acknowledgments CPO would like to acknowledge the patient mentorship of Drs. Nicole Le Douarin and MarieAimee Teillet of the Institut d’Embryologie in Nogent-sur-Mame,France, in the development of

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Charles P. Ordahl and Bod0 Christ methods described herein. The authors also thank Brian Williams for helpful suggestions and critical reading of the manuscript. This work was supported by grants from the Muscular Dystrophy Association and NIH (to CPO) and the Deutsche Forschungsgemeinschaft (Ch 44/9-2,44/12-1 to BC). We also thank the UCSF Cardiovascular Research Institute for partial support of one of us (BC) during his sabbatical through funds from its NHLBI Program of Excellence in Cardiac Molecular Biology (HL43821).

References Burrows, M. (1910). The cultivation of tissues of the chick embryo outside the body. J. Amer. Med. ASSOC. 55,2057-2058. Caplan, A. (1981). The molecular control of muscle and cartilage development. In “Levels of Genetic Control in Development” (S. Subtelny andh U. Abbott, eds.), pp. 37-68. New York Alan R. Liss. Chevallier, A., Kieny, M., and Mauger, A. (1977). Limb-somite relationship: Origin of the limb musculature. J. Embryol. Exp. Morph. 41,245-258. Chevallier, A., Kieny, M., and Mauger, A. (1978). Limb-somite relationship: Effect of removal of somitic mesoderm on the wing musculature. J. Embryol. Exp. Morph. 43,263-278. Christ, B., Jacob, H., and Jacob, M. (1974). Ober den U r s p m g der Flugelmuskulature. Experienria 30,1446-1448. Christ, B., Jacob, H., and Jacob, M. (1977). Experimental analysisof the origin of the wing musculature in avian embryos. Anat. Embryol. WO, 171-186. Christ, B., Jacob, H., and Jacob, M. (1978). On the formation of the myotomes in avian embryos. An experimental and scanning electron microscope study. Experientio 34,514-516. Christ, B., Jacob, M., and Jacob, H. (1983). On the origin and development of the ventrolateral abdominal muscles in the avian embryo. Anat. Embryol. 166,87-107. Conrad, G., Bee, J., Roche, S., and Teillet, M. (1993). Fabrication of microscalpels by electrolysis of tungsten wire in a meniscus. J. Neuroscience Merhods SO, 123-127. Davies, A. (1989).Neurotrophic factor bioassay using dissociated neurons. In “Nerve Growth Factors” (R. Rush, ed.), pp. 95-109. New York John Wiley. Hamburger, V. (1960). “A Manual of Experimental Embryology.” Chicago: University of Chicago Press. Hamburger, V.,and Hamilton, H. (1951). A series of normal stages in the development of the chick embryo. J. Morph. a49-92. Hamilton, H. L. (1952). In Lillie’s Development of the Chick An Introduction to Embryology” (B. H. Wilber, ed.). New York Henry Holt & Co. Kalcheim, C., and Teillet, M. A. (1989). Consequences of somite manipulation on the pattern of dorsal root ganglion development. Development 1685-93. Kieny, M. A. (1983). Cell and tissue interactions in the organogenesis of the avian limb musculature. In “Limb Development and Regeneration, Part B” (R. 0. Kelley, P. F. Goetinck, and J. A. MacCabe, eds), pp. 293-302. New York Alan R. Liss, Inc. Le Douarin, N. (1973). A Feulgen-positive nucleolus. Exp. Cell Res. R,459-468. Lewis, M., and Lewis,W. (1917). Behaviour of cross-striated muscle in tissue cultures. Am. 1.Anat. 22,169-194. Locke, F., and Rosenheim, 0. (1907).Contributions to the physiology of the isolated heart. The consumption of dextrose by mammalian cardiac muscle. J. Physiol. (London) 36,205-220. Ordahl, C., and Le Douarin, N. M. (1992). Two myogenic lineages within the developing somite. Development 114,339-353. Ordahl, C. (1993). Myogenic lineages within the somite. In “Molecular Basis of Morphogenesis” (M. Berdeld, ed.), pp. 165-176. New York John Wdey & Sons. Rugh, R. (1%2). “Experimental Embryology: Techniques and P r d u r e s , ” 3rd ed. Minneapolis: Burgess Publishing Co.

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Tyrode, M. (1910). The mode of action of some purgative salts. Arch. Intern. Phumucodyn. 17, 205-209. Williams, B., and Ordahl, C. (1994). Pax-3expression in segmental mesoderm marks early stages in myogenic cell specification. Development UO,785-796. Zwilling, E. (1968). Morphogenetic phases of development. Dev. Biol. Supplement 2,184-207.


CHAPTER 2

Myogenesis in the Mouse Embryo Margaret Buckingham’ and Giulio COSSU**~ CNRS, URA1947 Department of Molecular Biology Pasteur Institute 75724 Pans Cedex, France University of Rome “La Sapienza” Department of Histology and Medical Embryology 00161 Rome, Italy t

I. Introduction 11. Manipulation of Genes in the Mouse A, Transgenes B. Homologous Recombination 111. Analysis of Gene Expression in the Mouse Embryo A. In Situ Hybridization B. Immunocytochemistry IV. Experimental Embryology A. Culture of Somites and Somitic Cells V. Perspectives References

I. Introduction Myogenesis became one of the favorite model systems for the study of differentiation in the 197Os,largely because skeletal-muscle myoblasts could be grown in culture, where they will fuse to form myotubes that accumulatemuscle proteins. The myoblast-to-myotube transition is normally accompanied by the activation of muscle-specific genes. This process can be followed in primary cultures, often derived from late fetal or neonatal rat or chick muscle or in permanent muscle cell lines, most of which are of rat or mouse origin. The exploitation of muscle cell lines led to the isolation of the first myogenic transcription factors, MyoD METHODS IN CELL BIOLOGY, VOL. 52 Copyright 0 1998 by Academic Press. AU righa of reproduction in my form rrscwcd 0091-679X/98$25.00

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and myogenin, in the 1980s (see Weintraub et aZ., 1991). As far as in vivo studies of muscle are concerned, these have classically centered on the study of the sarcomere, its proteins, and their organization; of fiber types, their contractility, and their isoform content; and of innervation, with the associated electrophysiology of muscle. The species of choice here have tended to be the rabbit and rat, although some aspects of avian and even fish muscles have also been extensively analyzed. The other major line of research on muscle concerned its embryology. Most of the classic work on the embryological origin of muscle in vertebrates was based on observations with Amphibia and birds, largely because of the accessibility of the embryo and consequent ease of manipulation. Manipulations with birds, in particular, where chick/quail chimeras provide a powerful experimental tool, has led to most of the detailed information that is available for higher vertebrates (see Wachtler and Christ, 1992). In recent years, molecular biologists have become increasingly interested in the process of myogenesis in vivo. Molecular tools are now available to distinguish between the products of related genes as muscles mature, and the striking property of myogenic conversion exhibited in vitro by the MyoD family (see Weintraub et al., 1991) provides an opening to the definition of myoblast precursor cell populations in the embryo. Although mammalian embryos are more difficult to work with, the mouse has become a key experimental material for muscle research, despite the fact that most classical work in vivo has been with other species. This is partly because of the work with mammalian muscle cell lines and the fact that mouse embryology is relatively sophisticated compared with that of other mammals. Mouse genetics is also a potentially powerful tool, not just for localizing muscle genes (e.g., Robert et aZ., 1985), but also for identifying mutations, such as “splotch,” that affect muscle development (reviewed in Olson and Rosenthal, 1994). However, the most important single factor has been the technical progress in gene manipulation in the mouse. Transgenic mice can be obtained routinely by injecting gene constructs into fertilized eggs in vitro, followed by reimplantation into foster mothers. In this way transgenic animals are obtained and transgenic lines can be bred, where all the cells derived from the fertilized egg carry one or more copies of the transgene integrated into the genome at a random site. In order to extend classical genetics into an area where specific mutations in specific genes can be created at will, as for bacteria and yeast, the endogenous gene must undergo homologous recombination with an exogenous DNA, and the cells that have a modified genotype must subsequently be selected and then reintroduced into an embyro. After passage of these cells through the germ line, heterozygotes and then homozygotes for the mutation can be obtained. This technology has been developed for mice. Murine embryonic stem cells can be cultivated in vitro and maintained in a dividing pluripotent state, while undergoing DNA electroporation and subsequent clonal selection. Such cells, when injected into an embryo, will participate in subsequent development to form ES/chimeric mice. If colonization of the germ line by the ES cells occurs, mutant mouse lines can be generated. This approach, previously not

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available for vertebrates, has now made it possible to manipulate genes and to ask questions about function and regulation in vivo. Increasing numbers of new mouse mutants are being generated, and this technology has largely contributed to the fact that mdecular developmental biology has become one of the rapidly expanding fields of the 1990s. Furthermore, mice carrying marked genes provide a novel material for embryological manipulations that previously were restricted to birds and Amphibia.

11. Manipulation of Genes in the Mouse A. Transgenes

Transgenic mice have been employed by a number of laboratories in the muscle field, to examine transcriptional regulation in vivo. Regulatory elements, usually including the proximal promoter and enhancer sequences, initially identified in cell culture systems,have been shown to reproduce,partially or completely, the expression pattern of the endogenous gene. Tissue specific expression has been the first criterion, followed by more detailed analysis of the onset of transgene transcription in the embryo (e.g., Donoghue et aL, 1991), and restriction of expression later to specificmuscle fiber types (e.g., Banarjee-Basu and Buonnano, 1993). In some cases, genes normally expressed in both cardiac and skeletal muscle show expression in only one striated tissue with the sequence elements used (e.g., Li et al,, 1993); fiber type distribution in mature skeletal muscle may also be anomalous (Hallauer et al., 1993),pointing to additional regulatory motifs lacking in the transgene. The identification of sequences that are involved in fiber specification or in developmental timing will be of major interest in the future. These kinds of questions can only be addressed in vivo. In addition to regulatory studies on muscle genes, transgenicanimals also introduce the possibility of overproducinga gene product or a mutated gene product and of deliberately producing it ectopically, at a different site andor with different timing. Transgenes encoding muscle protein growth factors or transcriptional factors, under the control of specilic muscle regulatory sequences, will no doubt be used in the future as an experimental approach to dissecting myogenesis in vivo. Making transgenic mouse lines is a long and expensive procedure. For a detailed description the reader is referred to Section X of Wasserman and DePamphilis (1993); a brief outline of the procedure, stressing points that, in our experience, are important, is given here. Preparation of the DNA fragment to be injected is critical. We purify the plasmid DNA using a Qiagen column, before restriction digestion to remove as much vector sequence as possible, and purification through an elutip (Schleicher and Schiill) column. Some laboratories use high-pressure liquid chromatography to ensure DNA purity. One-cell-stage fertilized eggs are obtained by flushing the oviducts of superovulated females that had been mated the previous night, and the DNA is injected into the larger male pronucleus at the two pronuclei

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stage. This step requires DNA microinjection equipment and specilized technical skill. Surviving zygotes, after several hours of incubation, are reimplanted at the one- or two-cell stage into the oviducts of pseudopregnant foster mothers. Litters are tested for presence of the transgene by extracting tail DNA and assaying by PCR or on a Southern blot, the latter providing useful additional information about transgene DNA configuration. A realistic estimate of yields obtained by a skilled manipulator would be as follows: for 100 eggs, 90 are injectable, 70 survive the injection process, and 35 still look all right after incubation and are reimplanted, leading to about 15 transgenic animals, that is, an overall yield of 15%. Transgenic founder (Fo)animals are then bred and the progeny analyzed for transmission and expression of the transgene. In the case of skeletal muscle, this can be more rapidly checked, without sacrificing the animal, by taking a piece of tail, from a neonate, for example, and assaying for transgene product, since skeletal muscle forms along the tail. If the transgene is ZucZ, this is immediately obvious from P-galactosidase activity. Once a transgenic line is established, animals are of course readily available for analysis. However, the process takes several months to a year. To short-circuit the time factor, particularly where expression in the embryo is the main interest, so-called “transitory” transgenics can also be used. In this case, embryos are sacrificed immediately from the first litter, tested for the transgene, and analyzed for expression at the same time. The main problem with this approach is that it is necessary to analyze a large number of transgenic embryos to obtain an estimate of the expression pattern during development. This means many microinjections. Potential problems arising from DNA integration are a major consideration. Although the site of integration may not be completely random, it is different each time and the surrounding sequences may exert an effect on transgene expression, resulting in either no expression or additional ectopic sites. This can be particularly striking when the transgene does not contain strong musclespecificenhancer type sequences. Such context effects make it essential to analyze more than one transgenic line, or a number of Foembryos at each time point. Another feature of transgene integration is copy number, which should be monitored on Southern blots. A few copies integrated in tandem reduce possible end effects on the transgene, but it is not uncommon to see reports of animals with 50-100 copies inserted together, making quantitativeinterpretation very difficult. So-called “end” effects may involve rearrangements of the 5’ and 3‘ ends of the transgene, again making interpretation of results on expression difficult. Major rearrangements should be detectable by Southern blotting, and mice can be eliminated. Another potential problem in founder mice is mosaicism, which can be detected by quantitative Southern analysis. In this case not all cells contain the transgene, and transmission depends on the presence of the transgene in germ cells. Integration effects make detailed quantitative analysis between lines difficult. This becomes a major problem when the relative contribution of different regulatory elements is assessed.Very few such experimentshave been carried

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out with muscle genes; the need to analyze several mouse lines with each construct, in order to obtain a sound quantitative estimate, makes this a formidable task. Where there is an all-or-none effect, as reported for the MEF'2 and E-box motifs in the myogenin promoter (Cheng et aZ., 1993; Yee and Rigby, 1993), this can be immediately visualized with a lac2 reporter, but for many regulatory elements, transfection experiments already point to combinatorial mechanisms for optimal expression. Direct injection of DNA into muscle is a very promising alternative approach to problems that can be examined with postnatal animals and is not necessarily limited to mice. It has already proved valuable in experiments on gene regulation in the heart (e.g., Buttrick et aZ., 1992), and with the finding that regenerating skeletal muscle will also take up DNA efficiently (Vittadello et aZ., 1994), experiments where DNA constructs are injected into skeletal fibers are now feasible, with quantitation of transgene products. An alternative approach involves the use of adenoviral vectors, which give much more widespread expression of the transgene (Bessereau et aZ., 1994), although artifacts due to viral sequences are a potential hazard. The choice of reporter gene for regulatory studies is an ongoing subject of discussion. In many of the transfection experiments with cultured muscle cells CAT was used, because activity measurements are sensitive and rapid. It is therefore convenient to use the same constructions in transgenic mice. The disadvantage is that tissue has to be dissected and extracted for CAT activity measurements. This is also the case for luciferase, the reporter of choice if detection sensitivity is a major issue. In the case of CAT, in situ hybridization and antibodies (see Cui et aZ., 1994) have been used to examine regionalized muscle expression, notably in the case of the myosin (MLClF) transgene. The P-galactosidase assay is less sensitive, but ZacZ transgenes offer the great advantage of immediate visualization of expression on whole mounts, in addition to sections that can be cut subsequently if necessary. As an in situ marker pgalactosidase is very sensitive, comparing favorably with antibodies, and has the added advantage over radioactive RNA probes that it gives cellular-level resolution. This is enhanced if a nuclear localization sequence is added to the ZacZ (nZac2) (see Chapter 27, Wasserman and DePamphilis, 1993) and indeed, for studies on multinucleated muscle fibres, the nlacZ transgene has obvious advantages; the protein tends to relocalize to the adjacent nucleus. There has been considerable debate about possible artifacts produced by the reporter gene, and some anomalies in ZacZ expression have been reported (see Cui et aZ., 1994). In our experience, the nZac2 reporter shows different expression patterns in embryonic and adult muscle (Kelly et al., 1994), depending on the regulatory sequences of the transgene, without any evident bias introduced by the reporter. The convenience of having immediate visualization of transgene expression has resulted in widespread use of this reporter by mouse embryologists, and the accumulated comparative data are also an important consideration. Other transgenes also permit whole-mount visualization, but background reactivity has

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tended to be a problem. The human placental alkaline phosphatase gene is a potentially interesting reporter (see Chapter 56, Wasserman and DePamphilis, 1993), and its use in double transgenics with lac2 may be of future interest. B. Homologous Recombination Although integration of transgenes into the mouse genome has been a widely used technical approach for over a decade now, directed modification of a specific endogenous gene by homologous recombination in embryonic stem cells is only beginning to become standard technology (see Section XI, Wasserman and DePamphilis, 1993). The question of function is particularly relevant to muscle research, where multigene families often coding for closely related isoforms are frequent, both for the contractile proteins and for their transcriptional regulators. The recent “knockout” experiments with the MyoD family of myogenic factor genes were of major importance for our understanding of vertebrate myogenesis, providing the first functional data in vivo (see Buckingham, 1994). It is now clear that myogenin plays a critical role in muscle cell differentiation, whereas MyoD and myf-5 are essential for establishing or maintaining the myoblast precursor population. Experiments such as these involved disrupting the coding sequence of the gene. The first consideration here is the type of DNA construction to be used for electroporation into ES cells. An example of such a construct is given in Fig. 1.

I

0

0

I t

nlecZpolyA

Pol Ii neo ply A

0HSV-TK

w exons

-

intmnic and flanking genomic DNA

-1 kilobase

Fig. 1 Diagrammatic representation of a construct for homologous recombination, as discussed in the text (e.g., see Tajbakhsh et al., 19%a).


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To achieve homologous recombination, several kilobases of 5’ and 3’ flanking sequencesshould be included for optimal efficiency. Although keeping one flanking sequence short facilitates analysis of recombinant pools of clones and then single clones by PCR, in practice this is only advantageous for genes where recombination frequencies are low. In addition, PCR analysis tends to be more problematic, and it is necessary, anyway, to verify the recombination event by Southern blotting. A second important consideration is the nature of the mutation introduced into the gene. It is necessary to target a known functional motif, such as the basic or helix-loop-helix region of a myogenic factor. Even so, a straightforward null mutation may not be generated. Partial transcripts derived from the mutated locus, either as a result of transcription initiated at the normal Cap site or at cryptic initiation sites within the gene, or as a result of readthrough transcripts from the promoter of the selectable marker gene, may result in the presence of truncated proteins or fusion proteins that retain some biological activity. Different “null” mutant mice can potentially show different phenotypes. Once a mutation has been made, it is essential to check for partial transcripts and for the presence of protein. In the case of the myf-5 knockout (Braun et aZ., 1992), for example, such transcripts were present, although there is no evidence that partially functional protein was produced. To avoid read-through it is important to verify that insertion of the selectable marker (neo) disrupts the reading frame. Since partial transcripts are often generated by read-through from the neo promoter, it is also important to try to control this, for example, by introducing strong “stop” signals with a poly A polyadenylation signal. The most frequently used selection gene is that coding for neomycin resistance. In our experience, it is worth attempting to insert a reporter gene upstream of the neo gene and under transcriptional regulation of the endogenous locus. The potential disadvantage of this is a reduction in recombination frequency. Despite the theoretical claim that only the flanking sequences influence recombination efficiencies and that the size of insertion is neutral, evidence from a number of laboratories, where the same gene has been targeted with or without an additional lac2 sequence, indicates that a longer insertion results in lower recombination efficiency. However, lac2 or nZacZ inserted in this way provides a valuable marker of endogenousgene expression. This is useful for studies on heterozygotes where unexpected features of endogenous gene expression may be revealed that were not detected with exogenous probes. This was the case for myf-5 expression in the central nervous system (Fig. 2), for example (Tajbakhsh et uZ., 1994, and Tajbakhsh and Buckingham, 1996). Such lac2 mice also provide a baseline for comparison with lines where a ZucZ transgene is under the control of fragments of regulatory sequences. The lac2 reporter inserted in the endogenous gene is also very valuable for marking the cell population where the endogenous gene would normally be expressed in homozygote mutants. Again in the case of myf5, the presence of the lac2 marker clearly shows that the precursor muscle cells are formed in the absence of this early myogenic factor (Tajbakhsh et aZ., 1996b). In order not to interfere with the regulation of the locus, which is essential if a

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Fig. 2 Microinjection of ES cells into morula. A to D show successive phases of the injection. The holding pipet is on the left and the injecting pipet with a few ES cells is on the right. Bar: 30 pm.

lac2 marker is used, the neo gene should be under the control of a neutral promoter; enhancer sequences (used in the past with the thymidine kinase promoter) should be avoided. One possibility, using the RNA polymerase I1 promoter, gives encouraging results (M. Capecchi, personal communication; see Tajbakhsh et al., 1996b). Another important consideration where the construct is concerned is whether or not to include a second selection gene, which, in contrast to positive selection by neo, will permit negative selection for clones that have not undergone homologous recombination. Here the most frequently used gene is thymidine kinase (see Fig. 1).Gancyclovir should potentially eliminate ES cell clones that carry this gene. The efficiency of this negative selection can vary from 1-2-fold to 50-fold enrichment for clones that have undergone homologous recombination and eliminated the TK gene. The reasons for this variability are not clear, although inactivation of TK after random integration of the construct (either by rearrangement or by integration site effects) probably plays a major role. An alternative second selection system is provided by the introduction of a gene conferring sensitivity to diphtheria toxin, which has given very promising results (Yagi et aZ., 1990), and in our hands has proved satisfactory (Tajbakhsh et al., 1996b).How much effort is worth putting into a second negative selection system will depend on how difficult it proves to be to target the gene. This is empirical, depending no doubt on factors that remain difficult to predict,

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such as DNA sequence composition and chromatin composition. In principle, a gene that is transcribed in dividing ES cells should be easier to target by using a promoterless neo gene, although there are many examples now where an inactive gene has been successfully mutated. Although many ES cells (usually from 129 mice) have been targeted with DNA constructs from other mouse strains, it is clear that targeting efficiencies are higher with isogenic DNA. If no clones that have undergone homologous recombination are obtained, the first thing to do is to go back and isolate the gene from 129 mice or indeed, for optimal results, from the 129 ES cell that is to be used. The second major technical hurdle in gene targeting experiments is the choice of ES cell line. There are two aspects to this. The most important concerns the potential of the cells for germ line transmission, after culturing and selection. The first ES lines, like D3, did not give consistent transmission, probably depending partly on the passage number from the original embryonic isolate. Ideally, each lab should isolate its own primary lines, but in practice this is a great deal of work. A second generation of ES lines, such as HM1 (Selfridge et aZ., 1992), in our hands gives germ-line transmission frequently. If a targeted clone does not give heterozygotes rapidly-even if it gives a high percentage of chimerism, generally estimated in terms of coat colour, such as agouti (129 ES cells) on a dark (C57BL recipient embryo) background-it is not worth pursuing. It is of course also feasible to target both alleles in the ES cell. Increasing the level of neomycin in order to select for cells in which both chromosomes are targeted has proved satisfactory in a number of laboratories (e.g., Braun and Arnold, 1994) and avoids subjecting the ES cells to a second round of selection with a different drug. A second aspect of the ES cell line that is worth considering is its potential for myogenic differentiation. For embyro injection, ES cells are maintained as a dividing undifferentiated population grown in the presence of LIF and/or on feeder layers of embryonicfibroblastsderived from transgenic mice carrying the neo-resistance gene. Clones in this state are selected for blastocyst injection. Already, however, as a criterion for successful targeting, the capacity of a clone that carries the lac2 marker gene to express is a useful index. Secondly, targeted ES cells can be used for in vitro studies on myogenesis. Most ES cell lines, when grown under conditionswhere they form aggregates and differentiate (see Chapter 54, Wasserman and DePamphilis, 1993), produce relatively large numbers of nascent cardiocytes, in keeping with the early onset of cardiac myogenesis in mice (Miller-Hance et al, 1993). Skeletal myocytes tend to form later and more rarely. ES cells have mainly been used as vehicles for gene targeting. However, ES clones in which an exogenous DNA construct has been randomly inserted into the genome are also potentially useful for generating transgenic Foembryos that can be examined immediately for transgene expression. The advantage is that the same ES clone, with the same site of integration, can be used for repeated injections and that, again, in vitro expression in differentiating ES cells can be examined in parallel with that in vivo. Embryos can be monitored for expression

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Margaret Buckin+

and Giulio Corsu

in this way and a transgenic line subsequently produced. The disadvantage is that the transgenic embryos generated in this way are chimeras. If mutant mice are the object of the exercise, blastocyst injection is a standard procedure. Embryonic injection is also a tricky step and requires specialized technical skill. Depending on the ES line, a number of cells varying from 5 to 20 per blastocyst is usually optimal. In our hands, morula (eight-cell embyro) injection (see Fig. 3) gives a reproducibly high percentage of chimeras (Tajbakhsh et al., 1996a), although most of the mice do not survive long enough to breed. Injection of ES cells at the morula stage, before compaction, is also technically easier than blastocyst injection. This approach is useful for obtaining rapid in vivo information on the expression of a targeted gene, carrying the lacZ marker in the locus, in addition to its application for nontargeted transgenics. Once germ-line transmission is obtained and homozygote mutant mice are available (this requires a minimum of several months), it is important to bear in mind variations in phenotype that may result from variation in genetic background (e.g., 129 on C57 BL). This problem is illustrated by the 10-fold variation in a-actin mRNA levels documented for different inbred mouse lines (Alonso et aL, 1990). Compensatory phenomena may well be influenced by differences in transcriptional and processing efficiencies of related sequences. The solution is to try to breed the mutation into a homogeneous genetic background.

111. Analysis of Gene Expression in the Mouse Embryo The expression patterns of a number of muscle genes (see Buckingham et aZ., 1992), including those encoding contractile protein isoforms, muscle enzymes,

Fig. 3 A 9.5-day heterozygoteembryo containing nhcZ in one allele of the myogenic factor gene myf-5, fixed in 4% paraformaldehyde for 1.5 hr, stained for /3-galactosidase activity. In addition to myotomes, /3-gal-positive cells are detected in the midbrain (arrow).


39

membrane associated components, and transcriptional factors, have been described by in situ hybridization on sections of mouse embryos. Where antibodies have been available, immunocytochemistry has also been used on sections to look at muscle protein accumulation. The advantage of such in situ analyses is that they can be carried out on sections of the whole embryo and therefore give an immediate overall picture of the sites of expression of a gene. Different probes and antibodies can be used on the same or adjacent sections, covering different transverse, parasagittal, or frontal planes of the embryo, taken at different stages of development, to build up a complete regional and temporal map for the relative expression patterns of different genes. Underlying the in situ approach is the crucial importance of identifying the embryonic structures seen on a section. This can be implemented by careful orientation of the embryo before sectioning, together with frequent reference to published sources (e.g., Kaufman, 1992). Precise staging of embryos is essential and, for early stages of muscle formation, the number of somites is the best criterion. Embryos taken at the same time (the morning of the vaginal plug is counted as 0.5 day) may vary in age by as much as half a day, and, even within the same litter, some embyros are usually more advanced than others. It is essential to do a somite count on every embryo before sectioning. At later fetal stages, when somites are no longer present in the trunk, the morphology of structures such as limb buds provides a developmental marker. Antibodies, cold probes, and P-galactosidase staining can also be used for whole-mount studies on the complete embryo, providing rapid information on the overall expression pattern of a gene, which can then be investigated in more detail on sections. One of the disadvantages of the in situ approach is that quantitation is difficult. Extraction of RNA and analysis by Northern or slot blotting, or, better, by nuclease protection, is one solution. PCR-based methods have the great advantage that they can be applied to very small quantities of material, although quantitation is tricky (see Chapters 19 and 20, Wasserman and DePamphilis, 1993). PCR analysis becomes important, for example, in the context of low-level expression of muscle transcription factors prior to somitogenesis (Kopan et al., 1994). In such experiments, pooled material is used, and it is essential that the staging of all embryos, based on morphology, should be correct; the inclusion of negative controls for downstream muscle genes is also essential. A range of PCR cycles (e.g., 15-25) helps in appreciating relative expression levels. The significance of transcripts only detectable at 1 3 0 cycles is perhaps less evident. A major difficulty in comparative studies and quantitation is the differential efficiencies of different oligonucleotide primers, as well as the fact that they may interfere with each other. In an extreme situation some primers just do not work, probably because of secondary structure or because they recognize other sequences, and the only solution is to use another sequence. When RNA preparations from different tissues, for example, different compartments of the heart (Kelly et al., 1994), are compared, it is essential to include a constant positive control to adjust for minor differences in RNA concentration. Choosing such a

40

Margaret Buckinghun and Giulio Corru

sequence is not easy; a ribosomal protein sequence is one option if the division status of the tissues is similar. Similar considerations apply at the protein level, provided that antibodies are available. With labeled antibodies, quantitation even of very small quantities of extracted protein is feasible. Any method that involves extraction loses, of course, the distinction between a few cells producing a lot of material and many cells producing less within a tissue.

A. I n Situ Hybridization Although DNA probes have also been used successfully for maximum resolution, we would recommend in situ hybridization with 35S-labeledRNA probes, on paraffin-embedded sections mounted on subbed slides. A detailed description of the standard protocol can be found in Chapters 22 and 24 of Wasserman and DePamphilis (1993) and in Wilkinson (1992). A number of technical points should be emphasized.First, the selection of the probe is critical. Muscle proteins frequently belong to multigene families, which by definition contain sequence homologies. If a coding sequence probe is used, the risk of cross-hybridization should be checked. Noncoding sequences from the 3‘ or 5’ ends of the mRNA avoid this difficulty, although the use of a shorter sequence may pose problems for detecting rare messengers. A gene that gives rise to several transcripts is also a common phenomenon in muscle, and to follow individual mRNAs, exonspecific probes should be selected. Again, this may mean using a short probe. In the case of cardiac actin mRNA, for example, a probe containing only 50 nucleotides from the 5’ noncoding sequence gave a good signal (see Sassoon ef aZ., 1988). Fewer than 30 nucleotides is probably tricky, since this begins to approach the limit size for hybridization. Longer labeled probes require alkali digestion to reduce them to fragments approaching the 50-150 nucleotides size, to facilitatetissue penetration.The signal, of course, does not depend on fragment size, but on the total length of sequence recognized by the probe. The sequence selected should be cloned into a bluescript type vector that permits sense and antisense RNA transcription. One or two 35S-labelednucleotides can be used, according to the sensitivity required. In order to obtain low background in the hybridization, correct probe preparation, with removal of contaminating free labeled nucleotides by passage through G50 Sephadex after alkali digestion, is crucial. Correct orientation of the embryo when it is embedded in paraffin, to obtain clear parasaggital, frontal, or transverse sections later, is also important, as is the sectioning itself. One of the advantages of modem paraffin (e.g., Paraplast) is that it contains wax and plastic polymers that give high-quality sections, facilitating the identification of embryonic morphology. Sections are usually cut at 6-8 pm thickness, although thinner sections (e.g., 3 pm) can be obtained. If different probes are to be compared, it is essential to hybridize them to adjacent sections and to ensure that probe (a), used first, is rehybridized to the last section in the series. Particularly with younger embryos, muscle masses, such as the myotome, are small, and it is important to check that a section at the end of a


’

41

series is still within the muscle, rather than at the surface. A negative result may be misleading. Prehybridization,hybridization,and washing conditionshave been adapted to give maximum signalhackground, as described in Chapters 22 and 24, Wasserman and De Pamphilis (1993). The addition of dithiourea to the wash is helpful in further reducing background (G. Lyons, personal communication). Some antibodies will work on paraffin sections and can therefore be used in conjunction with labeled probes. However, often this is not the case. Frozen sections can also be used for in situ hybridization, following a similar protocol to that used for paraffin. The morphology tends to be less good, but antibodies can be used without problem. As with paraffin sections, they should be used before hybridization in situ, which involves a proteinase K treatment, and visualized before application of the emulsion to the slide. The alternative to 35S-labeledprobes are so-called “cold probes.” These avoid the use of radioactivity and also have the advantage that they work on whole mounts. In addition, more than one cold probe can be applied to the same whole mount or embyro, if different labels are used. The most commonly used label at the moment is digoxigenin. The procedure (Chapter 22, Wasserman and DePamphilis, 1993) is very similar to that used for radioactive probes, with the major difference that the signal is visualized by immunochemical detection of the probe, usually with an alkaline phosphatase-coupled anti-digoxigenin antibody. An alternative probe labeling system is provided by biotinylated nucleotides (Chapter 23, Wasserman and DePamphilis, 1993), which are visualized by streptavidin, coupled to an immunochemical detection system, or conjugated in a P-galactosidase complex. Cold probes are convenient and their use is becoming more widespread. In our experience, however, they remain less sensitive than 35S-labeledprobes, which may be an important issue when low levels of transcript are being monitored on sections. In the case of mice where a lacZ sequence has been introduced into the genome, it is frequently desirable to carry out in situ hybridization in parallel with p-galactosidase staining, either to compare it with the expression pattern of the endogenous gene, or to look at the expression of other genes that may have compensated for, or been perturbed by, a targeted mutation. In fact, in our experience, embryos that have been fixed in 4% paraformaldehyde and stained for P-galactosidase activity overnight, if refixed, can then be successfully hybridized with radioactive or cold probes, either after subsequent embedding in paraffin and sectioning, or in the latter case also as whole-mount embryos (Tajbakhsh and Houzelstein, 1994). For mRNAs such as that encoding MyoD, enough survive the P-galactosidase staining to give a good hybridization signal, contrary to what one might predict (Fig. 4; see Color Plates).

B. Immunocytochemistry A detailed description of immunocytochemical methods would be beyond the topic of this chapter. For this, the reader is referred to extensive general reviews

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Margaret Buckinghom and Giulio Cossu

(Harlow and Lane, 1988; Catty, 1988) and, more specifically, to an excellent description of immunocytochemical methods adapted for embryonic material (Stem, 1993). More pertinent to this chapter is probably a brief description of the use of antibodies on myogenic cells, tissue sections, or embryonic explants that have been previously stained for 8-galactosidase activity. Somites or somitic cells (see later discussion) are usually cultured on matrixcoated (collagen, fibronectin, or laminin) plastic or on feeder layers. They adhere very poorly to glass coverslips (even if precoated), and therefore we perform immunocytochemistrydirectly on the plastic dish; we find no special background fluorescence due to plastic. The product of the P-galactosidase reaction by itself quenches the signals of fluorochromes and also interferes with color development after peroxidase or alkaline phosphatase activity; there are, however, several possibilities for localizing one or two antigens in a P-galactosidase-positive cell. One crucial point is the cellular localization of the 8-galactosidase compared with the antigen(s) to be localized. If the construct contains, in addition to 8-galactosidase, a nuclear localization signal (n), this helps the identification of single expressing cells. However, after staining for P-galactosidase, the nucleus of positive cells will become dark blue and other nuclear antigens will be difficult to localize. In this case it is still possible to know whether a given cell coexpresses P-galactosidase and a nuclear antigen (such as a transcription factor) by using antibodies directed against the antigen under investigation and against P-galactosidase. In this case the advantages of the histochemical stain for P-galactosidase will be lost. If, however, the antigen is either cytoplasmic or membrane-bound, it will be easily localized in 8-galactosidase positive cells. The use of the fixative depends on the specific requirements of the antibodies to be used later. In most cases, 4%paraformaldehyde in PBS can be satisfactorily used. After normal staining of the sample, the X-gal solution is removed, and after two rapid washes with PBS, the sample is incubated with a blocking solution (for example, 1%bovine serum albumin, 0.05% TritonX-100, and 1%heatinactivated goat serum in PBS) before addition of the first antibodies (at the appropriate dilution in the same solution). The conditions of incubation have to be optimized empiricallyfor the antibodies used. We find convenient an overnight incubation at 4"C, but shorter times and higher temperatures (up to 38°C for antibodies with weak binding affinities) can be used. To use small aliquots of antibodies whose availability may be limited, we grow the sample in the center of a 35-mm 0 dish, and dry the dish all around the central spot with a paper tissue so that the incubation is carried out in 20 pl of the final dilution of antibody in a humidified chamber for the required time. After incubation, the samples are washed twice with PBS (1 min for each wash) and once with the blocking solution for 30 min at room temperature (RT) with gentle agitation and then incubated with second antibodies (labeled with either fluorochromes or horseradish peroxidase or alkaline phophatase). For double-labeling we prefer fluorochromes because they are more sensitive, and


43

moreover there is no interference with color development, which may occur after the first staining for P-galactosidase. We use fluorescein-labeled anti-rabbit Ig and rhodamine-labeled anti-mouse Ig (according to manufacturer’s instructions) for 1 hr at RT. After the incubation, the samples are washed again as described earlier, and then mounted with Gelvatol (Stem, 1993) or with 80% glycerol in 0.15 M Tris-C1, pH 8. For cells and tissue sections, samples can be observed directly under an epifluorescence microscope (Fig. 5; see Color Plates). When possible it is convenient to take photographic records of the samples by using two cameras (most modem microscopesare equipped with two), one loaded with a sensitive film for fluorescence (800 ASA or more) and another loaded with a less sensitive film for light photography of the same field (if the 8galactosidase signal is not strong, it is better to take a first picture under transmitted light and a second one under phase contrast, as the latter gives a nicer picture but may mask a weak P-galactosidase positive signal). For explants that are about 100 pm thick it may help to use a confocal microscope or, otherwise, embed and cut frozen sections to be labeled later with antibodies. For this, the method described in Stem (1993) gives excellent morphology, when compared with other methods. Briefly, the sample is fixed in 4% PAF, stained for flgalactosidase, and then washed in PBS. The sample is later transferred to 7% sucrose in PBS, then 15 (or 20)% sucrose in PBS (each incubation is continued until the sample sinks), and finally to 15 (or 20)% sucrose, 7.5% (final w/v) gelatin in PBS (this last solution has to be warm because it becomes a gel at RT). After the sample is impregnated with gelatin, the solution is cooled to gel (if this is done under a dissecting microscope, it is possible to orient the sample using warm needles or forceps), trimmed into a small cube with a razor blade, mounted on a cryostat block with OTC (tissue-tek) and quickly frozen for 20-30 sec in N2-cooledisopentane (although liquid N2 itself is usually satisfactory), and then sectioned in a cryostat. Whole-mount immunocytochemistry can also be performed after Pgalactosidase staining. We tried several published and unpublished protocols with no striking differences among different methods. We suggest the method described by Stem (1993) and find that the quality of the reaction is much more linked to the size of the embryo or to the presence of thick epithelium on the surface of the explant than to the previous /3-galactosidase staining.

IV.Experimental Embryology For decades, postimplantation mammalian embryos have been a difficult if not an impossiblesystem for experimental embryology. Embryos can be obtained in small numbers only after the sacrifice of the mother; they are small and do not grow well ex utero, certainly not to the point at which terminal differentiation and morphogenesis of most organs occur.

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Margaret Bucldngham and Giulio Corsu

However, gene manipulation in the mouse, both for generating new mutants and for introducing a marker sequence, such as lacZ, into the genome, offers a powerful and as yet largely unexplored new tool for experimental embryology. If the reporter gene is inserted into the locus of a tissue-specific gene, then all and only the cells expressingsuch a gene will carry a cell-autonomous,inheritable, undilutable marker that can be unambiguously identified by a fast and highly sensitive staining technique. This technique works easily and quickly in whole mount and, more importantly, will allow identification of a single cell (or its progeny) in an explant, after reconstitution with other tissues from non labeled embryos. In addition to cells from targeted heterozygotes or homozygotes where a particular gene has been targeted with lacZ, transgenic mice, which express a reporter under the regulatory sequences of a gene of interest, can also be analyzed. Furthermore, as discussed earlier, immunocytochemistry or in situ hybridization can be performed after staining for 8-galactosidase,thus allowing further phenotypic characterization within the same experimental sample. In principle this method constitutes the equivalent of the quail-chick transplantation system for birds, with the clear advantage of intraspecific transplantation or reconstitution of different embryonic cells or tissues, but with the problems related to the difficulty of studying mammalian embryos ex ucero. However, thanks to recently developed methods for culturing early postimplantation mammalian embryos, and also for culturing tissue explants and germ-layer derivatives from early or more developed embryos (Chapter 10, Wasserman and DePamphilis, 1993), many experiments previously restricted to Amphibia and birds can now be performed in mammals. In principle it is possible to study postimplantation development with embryo cultures up to 10 or 11 days post coitum (dpc). Afterwards, in vitro culture of the organ anlagen of interest makes it possible to follow terminal differentiation and, to a lesser extent, morphogenesis in an ex utero situation. Experiments such as, for example, those on migration (Serbedzija et af., 1990), interactions (Vivarelli and Cossu, 1986; Hashimoto et al., 1987), and developmental potential of different embryonic cells should be greatly facilitated by using cells that carry P-galactosidase as an endogenous lineage marker. Similarly, cell-lineage and cell-transplantation analyses (Beddington and Lawson, 1990), which are now technically possible in mammals with the use of embryo cultures, still present some limitations (such as duration of the label); this may be overcome by the use of embryos carrying P-galactosidase or green fluorescent protein as an endogenous lineage marker (Zernicka-Goetz etal. 1997). As an example of the use of manipulated lines of mice for experimental embryology and their potential development for cell-lineage studies, we will describe mice that carry P-galactosidase under the control of the promoter of the fast skeletal myosin heavy chain 3 (Kelly et al., 1995) and mice that have lac2 inserted into the myf-5locus (Tajbakhsh et al., 1996a). Lac2 has become a reporter of choice because of the easy and sensitive assay required to detect it, but other reporter genes can be used. Gene regulatory sequences other than those that are tissue specific may also be used for example, in the case of mice


45

where p-galactosidase is under the control of homeotic gene sequences where the transgene is being employed to address the question of whether and to what extent their expression can be modified by the rostrocaudal level and/or by the neighboring tissues (reviewed by Olson and Rosenthal, 1994). By taking advantage of embryos where only myogenic cells will become pgalactosidase-positiveat the time of terminal differentiation (such as mice where the enzyme is under the control of a muscle promoter), it is possible to isolate a somitic cell, or a group of cells, or one or more somites, and challenge their differentiation potential under different experimental conditions. Among these, reaggregation of the sample with other embryonic cells or organs from nontransgenic siblings will allow easy, fast, and sensitive analysis of the extent of terminal differentiation. It will also have the distinct advantage of being unbiased by possible cross-contamination between samples. For example, we reported years ago that cells from the neural tube were necessary to promote muscle differentiation in cultures from early somites (Vivarelli and Cossu, 1986). When we attempted to reconstitute neural tubes with somites in vitro, we observed that the former invariably contained differentiated mononucleated muscle cells. This fact, which led us, later, to identify a population of myogenic cells in the embryonic neural tube (Tajbakhsh et al., 1994), made it impossible initially to continue the experiments, which have been reapproached now that MLC3F-pgalactosidase embryos are available. To study neural induction of myogenesis in vitro,we use neural tube from nontransgenic siblings and somites or segmental plates from transgenic ones. As far as cell lineage studies are concerned, cells (and, in principle, a single cell) from a transgenic embryo may be injected ortho- or heterotopically and ortho- or heterochronically into nontransgenic embryos in utero (Muneoka et af., 1990) and their clonal progeny followed. Since not all the cells of somites will be myogenic, a certain number of white embryos will have to be taken into account, but dozens of injected embryos (injection itself being the limiting step) can be stained easily and only those containing a positive clone will be further analyzed. Finally, the simple anatomical examination of these transgenic embryos will immediately reveal information that could only be obtained after much work and with less certainty on normal embryos. For example, counting the number nuclei in whole mount or (at later stages) in of MLC3F-~-galactosidase-positive thick sections of myotomes will reveal the exact number of terminally differentiated myocytes per myotome at any given time. This has led us to the unpredicted observation that cervical myotomes contain fewer differentiated myocytes, and the number increases in a rostro-caudal gradient up to the first thoracic ones (Cossu et al., unpublished observations). A. Culture of Somites and Somitic Cells

To study the phenotype, lineage, or developmental potential of early myogenic cells, it is necessary first to isolate the tissues where these cells are originally

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Margaret Buckinghpm and Giulio Corsu

located. Somites are usually dissected with fine needles (either sharpened tungsten or glass) from embryos or embryo fragments that have been digested previously with enzymes. The following protocol has been adapted for isolation and culture of somites or single somitic cells from several published methods to which the reader is referred for more detail and discussion for other tissues. Pregnant mice are sacrificed at a developmental stage ranging from 8.5 to 10.5 days post coitum (dpc), the morning of vaginal plug detection being counted as 0.5 dpc. Usually, if embryos ranging from 0 to 6-8 somites are required, experiments should be performed in the early morning of day 8.5, then later in the day as more somites develop, up to 12-18 somites for experiments performed in the evening. In the morning of day 9.5 dpc, most embryos have developed to the stage of approximately 20-26 somites, whereas by the morning of 10.5 dpc, approximately 38-46 somites have formed. When a precise stage of development is needed, timing must be determined empirically, since there may be variations from strain to strain of mice and even in the same strain maintained in different animal houses. Embryos are isolated from the decidua under a dissecting microscope in saline such as PBS or M 2 (Chapter 10, Wasserman and DePamphilis, 1993). If the dissection takes some time, as in the case of small embryos and/or inexperienced investigators, it may be wise to keep embyros in a richer medium, such as a Hepes-buffered tissue culture medium (which contains, in addition to glucose, a variety of nutrients). 10.5-dpc embryos are easily dissected because they are relatively large; 9.5-dpc embryos usually pop out from their membranes during the dissection. This would not be a problem (as it is for whole-embryo culture) if it was not for the fact that the tail (where the newly formed somites are) is often lost. At 8.5 dpc, isolation of the embryo is not easy (for details, see Chapter 10, Wasserman and DePamphilis, 1993). Briefly, it is necessary first to remove each individual decidua from the uterine wall by gently cutting it on the antimesometrial side and then peeling out the decidua without breaking it. The isolated decidua is then gently transferred to a new dish with fresh medium (blood makes it very difficult to see the small embryos). The decidua looks like a pear with the embryo located in the smaller portion. It can be removed by gently opening the lower, larger part until the amnion is reached. Alternatively, it is possible to hold the decidum by the larger part, and to cut the upper fourth with fine forceps. This will normally expose the amnion, which then can be gently scooped out of the decidua. Somites or presomitic segmental plates can be isolated either mechanically or enzymatically. If there is no reason for culturing pure somites, mechanical dissection is simpler and faster, but the tissues are invariably contaminated by neural tissues (shown by later immunostaining of the culture for neural antigens) and by fragments of ectoderm and lateral structuressuch as the dorsal aorta. Because embryos older than 8.5 dpc are curved, it is simpler first to cut a segment of the trunk of the embryo (approximately 10-15 somites long). After gentle removal of ventral structures and with the embryonic fragment held with one needle on


47

the cranial edge of the neural tube, a first cut is made along the lateral edge of somites (or segmental plate), and then a second cut is made between somites and neural tube (Cossu et aL, 1996). If relatively pure somites are required, enzymatic dissection is necessary. Usually pancreatin, trypsin, or a mixture of both is used for short periods on ice (we use 0.25% pancreatin-0.1% trypsin for 5 min at 4°C). With this or any similar protocol, it is essential to test empirically each new batch of enzymes to find optimal conditions (concentration and period of incubation), then store the rest in frozen aliquots, which, however, can be thawed a few times. Embryonic tissues are extremely sensitive to proteolysis, and it is easy to over-digest tissues, which then become very sticky, leading to high levels of embryonic cell death. If, on the other hand, the digestion is not completed, it will be difficult later to dissect the different structures. After proteolysis, tissues are transferred to a serum-rich medium to stop the action of proteases. If somites are to be grown as explants, one or more somites can be cultured under different conditions, again depending on the goal of the experiment. Better morphogenesis is attained by rotating suspension cultures. Cell differentiation (see Fig. 6) or tissue interactions can be studied best after adhesion to, or inclusion in, extracellular matrix (for details of the various methods, see Chapter 10, Wasserman and DePamphilis, 1993). If a single-cell suspension is required, it can be obtained by gentle pipetting of the digested structures. For this purpose siliconized capillaries of progressively smaller diameter, obtained after flaming Pasteur pipets, can be used, but sterile yellow tips of a Gilson pipet can also be employed and are easier to prepare and to store. Pipetting must be very gentle and prolonged, since foaming and vortexing will increase the number of dead cells dramatically. Usually, about 80% of isolated cells are viable at the time of plating, as judged by Trypan Blue exclusion. When a single-cell suspension is required from tissues that have been

Fig. 6 3-day-old organ culture of the neural tube with adjacent segmental plates from MLC3FI LacZ embryos, stained for P-galactosidase activity. Note well-developed myotomes (arrowheads). Bar: 40 pm.

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Margaret Buckingham and Giulio Corru

isolated mechanically, we prefer to digest them with 0.1% collagenase-0.1% dispase for 5 min at 30"C, gently wash them in complete medium (see later discussion), and then gently pipette through a siliconized capillary to obtain a single-cell suspension. In the case of single-cell cultures, density is crucial. Only high-density cultures will grow and differentiate satisfactorily. We usually plate 5 X 10" cells in 30 pl of medium in a spot in the center of a 30-mm 0 dish. When the cells have settled, they should appear already confluent or slightly overcoduent (given that the suspension contains a certain fraction of dead cells) under the microscope. Cells are allowed to adhere to the substrate for 1-2 hr at 37°C in a humidified atmosphere (or evaporation will concentrate the medium and kill the cells); then, 2 ml of complete medium are gently added to the dish, which is returned to the incubator. If a lower density is needed or if clonal cultures have to be performed, feeder layers can be used. We have tried many different cell lines (3T3, 10T1/2, BC3H, C2C12) and also primary embryonic fibroblasts, with no striking differences among the different cells. Usually we use 1OT1/2, since these cells grow slowly and do not overgrow after confluence, and thus need not be treated with mitomycin C. For low-density or cloning experiments, cells from somites older than the fifth newly formed one should be used, since younger somitic cells are still dependent upon a community effect, at least for myogenic differentiation (Cossu et al., 1995). Cultures are grown in rich media. We usually use RPMI medium (Gibco) supplemented with 15%fetal calf serum, 300 pM P-mercaptoethanol, and 50 pg/ ml gentamycin (complete medium). Other media, different concentrations of FCS, or even rat serum, can be used, but for organ or cell culture, we do not see a distinct advantage in using rat serum, and therefore FCS is preferred for practical reasons.

V. Perspectives An increasing amount of information is already being generated by different laboratories on expression patterns of genes during mouse embryogenesis, both at the protein level with antibodies and at the transcript level with radioactive and "cold" nucleic acid probes. Although not yet fully operational,image analysis programs are being developed that should make it possible to store and relate all this information from different laboratories in a computer bank (Ringwald et al., 1994). The need for this has become pressing, as increasing numbers of transgenicmice are produced, many with ectopic expression patterns, which may be of particular interest to scientists working in another field. Without easy access to this information, its potential use is lost. The endogenous expression of many genes has also been analyzed in greater detail, as a result of the resolution provided by the j?-galactosidaseassay, either in transgenics with a lac2 reporter gene or on mice where lac2 is introduced into the locus by homologous recombination. Again, this extensive information requires collating and storing. Particu-

2. Myogeneiis in the Mouse Embryo

49

larly in the case of mutant mice, the effects of the mutation on the expression of other genes necessitate comparative analyses. Transgenes can be used not just to test regulatory sequences with a reporter, but also to manipulate expression of the gene itself. Overexpressionin the correct context, ectopic expression, or incorrect temporal expression of the protein, together with expression of a mutated version of the protein to give a dominant negative variant, are all ways in which function can be tested. For example, directing the expression of a fast myosin to slow fibers as a transgene with slow myosin regulatory sequences is a way to change isoform composition. Such an experiment has recently been reported for myosin isoforms in the heart (Gulick et al., 1997). This kind of experiment can also be envisaged in a null background, if, for example, a fast myosin gene has been “knocked out.” The first generation of homologous recombination experiments has been aimed at targeting a gene to produce a null mutant. Future experiments of this kind will no doubt also aim at modifying the endogenous gene itself, either its coding sequence or its regulatory sequences, by recombining in a different sequence. The presence of a lac2 sequence already in the locus facilitates this approach, since lac2 can be replaced in the targeting vector by another gene. A MyoD coding sequence, for example, could be introduced to look at the effect of expressing this myogenic factor in place of myf-5. Indeed, introduction of myogenin into the myf-5 locus corrects at least part of the mutant phenotype (Wang et al., 1996). An alternative type of manipulation would be to introduce a killer gene to eliminate all cells that would normally express the myf-5 myogenic factor. This provides information complementary to that obtained by knocking out the gene while retaining the cell, although killer gene technology in mammalian embryos is not yet fully operational. One potential approach is to use the thymidine kinase gene and kill cells expressing it with gancyclovir, perfused via the maternal bloodstream. Manipulation of the diphtheria toxin receptor gene (Yagi et al., 1990) provides another approach. In vivo manipulation of endogenous genes is beginning to become feasible with the development of the Cre-lox recombinase system (see Chapter 53, Wasserman and DePamphilis, 1993). As more and more mutant mice are generated, many interesting experiments can be camed out by crossing mutants. A first example of this was provided by the myf-5MyoD crosses that established the essential role of these two factors in myogenesis (Rudnichi et af., 1993). The limitation of the gene targeting approach is that it is restricted to known genes. Random targeting of lac2 into ES cells, with subsequent screening for mutant phenotypes and interesting pgalactosidase expression patterns, is beginning to be exploited to isolate, via the lac2 marker sequence, potentially important novel genes. The ES cell system provides an intermediate step that may be useful for screening out potentially interesting clones, before investing in time-consuming and expensive in vivo experiments. Genes of interest for myogenesis can be identified, at least where muscle differentiation is concerned, by lac2 expression in cardiocytes or myotubes found in embryoid bodies. Genome sequencing projects are also going to

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Margaret Buckingham and Gidio Corsu

lead to the identification of new genes. Here most of the emphasis is on the human genome, but because of the already detailed genetic map in the mouse, and also because of its experimental potential for gene manipulation, new genes are likely to be tested in this species. The localization of new genes may well lead to the identification of previously known mouse mutants, as in the case of Pax-3 and the splotch mutation, which provided several ready-made mutant alleles. Mutagenesis with chemical mutagens is likely to lead to the generation of new muscle mutations; a number of such mutations that have arisen spontaneously await further analysis. The difficulty in these cases, of course, is to clone out an untagged gene, based on a genetic localization. Manipulation of genes in the mouse is certainly going to have a major impact on our understanding of myogenesis in the next decade. However, generating mutants is very labor intensive, and although it can give invaluable information about gene function and molecular regulation, many questions will remain that can only be answered by embryological manipulation. The inductive effect of one tissue on another is a classicexample. The possibility of mutating and marking genes does not exist to the same extent in other systems, and therefore developing embryological techniques for the mouse will be very important. It is already possible to culture mouse embryos for up to 48 hr in vitro. Surgery on older embryos is also possible in utero. However, a potentially easier and more flexible approach is provided by explant experiments. Recent results on the manipulation of material from mouse embryos carrying an Otx2 gene marked with lac2 provide an example of the potential of this approach (Ang et al., 1994). Acknowledgments We should like to thank Denis Houzelstein and Shahragim Tajbakhsh for the photograph in Fig. 4 and Shahragim Tajbakhsh for Figs. 2 and 3 and for helpful discussions. M. B.’s laboratory is

supported by grants from the A.F.M. and the MENESR. G. C.’s laboratory is supported by grants from Telethon (A.67), Cenci-Bolognetti,and MURST. M.B. and G.C. have an EC grant from the HCM program.

References Alonso, S., Gamer, I., Vandekerckhove, J., and Buckingham, M. (1990).Genetic analysis of the interaction between cardiac and skeletal actin gene expression in striated muscles of the mouse. J. Mol. Biol. 211,727-738. Ang, S.-L.,Conlon, R. A., Jin, O., and Rossant, J. (1994).Positive and negative signalsfrom mesoderm regulate the expression of mouse Otx2 in ectoderm explants. Development 1u),2979-2989. Banejee-Basu, S.,and Buonanno, A. (1993).cis-Acting sequences of the rat troponin-I slow gene confer tissue-specific and development-specific transcription in cultured muscle cells as well as fiber type specificity in transgenic mice. Mol. Cell. Biol. l3,7019-7028. Beddington, R. S. P., and Lawson, K.A. (1990).Clonal analysis of cell lineages. I n “Post-implantation Mammalian Development” (A. J. Copp and D. L. Cockoft, eds.), pp. 267-292. Oxford University Press.


51

Bessereau, J. L., Stratford-Pemcaudet, L. D., Piette, J., Le Poupon, C., and Changeux, J. P. (1994). I n vivo and in vitro analysis of electrical activity-dependent expression of muscle acetylcholine receptor genes using adenovirus. Proc. Natl. Acad. Sci. USA 91,1304-1308. Braun, T., and Arnold, H. H. (1994).ES-cells carrying two inactivated myf-5 alleles form skeletal muscle cells: Activation of an alternative myf-5-independent differentiation pathway. Dev. Bio. 164924-36. Braun, T., Rudnicki, M. A., Arnold, H. H., and Jaenisch, R. (1992). Targeted inactivation of the muscle regulatory gene myf-5 results in abnormal rib development and perinatal death. Cell 71,369-382. Buckingham, M. (1994). Which myogenic factors make muscle? Curr. Biol. 461-63. Buckingham, M. E., Lyons, G. E., Ott, M. O., and Sassoon, D. A. (1992). Myogenesis in the mouse. Ciba Found. Symp. 165,111-124. Buttrick, P. M., Kass, A., Kitsis, R. N., Kaplan, M. L., and Leinwand, L. A. (1992). Behavior of genes directly injected into the rat heart in vivo. Circ. Rex 70,193-198. Catty, D. (1988). “Antibodies a practical approach.” IRL Oxford University Press. Cheng, T. C., Wallace, M. C., Merlie, J. P.,and Olson, E. N. (1993). Separable regulatory elements governing myogenin transcription in mouse embyrogenesis. Science 261,215-218. Cossu, G.. Kelly, R., Di Donna, S.,Vivarelli, E., and Buckingham, M. (1995).Myoblast differentiation during mammalian somitogenesis is dependent upon a community effect. Proc. Nut. Acad. Sci. US 92,2254-2258. Cossu, G., Kelly, R., Tajbakhsh, S., Di Donna, S., Vivarelli, E., and Buckingham,M. (1996).Activation of different myogenic pathways: myf-5 is induced by the neural tube and MyoD by the dorsal ectoderm in mouse paraxial mesoderm. Development l22,429-437. Cui, C., Wani, M. A., Wight, D., Kopchick, J., and Stambrook, P. (1994).Reporter genes in transgenic mice. Transgenic Reseurch 3, 182-194. Donoghue, M., Merlie, J. P., Rosenthal, N., and Sanes, J. R. (1991). A rostrocaudal gradient of transgene expression in adult skeletal muscle. Proc. Natl. Acud. Sci. USA 88, 5847-5851. Gulick, J., Hewett, T. E.,Klevitsky, R., Buck, S. H., Moss, R. L., and Robbins, J. (1997).Transgenic remodelling of the regulatory myosin light chains in the mammalian heart. Circ. Res. 80,655-664. Hallauer, P. L.,Bradshaw, H. L., and Hastings, K. E. M. (1993).Complex fiber-type-specificexpression of fast skeletal muscle troponin I gene constructs in transgenic mice. Development 119,691-701. Harlow, E., and Lane, D. (1988). “Antibodies: A Laboratory Manual.” New York Cold Spring Harbor. Hashimoto, K., Fujimoto, H., and Nakatsuji, N. (1987). An ECM substratum allows mouse mesodermal cells isolated from the primitive streak to exhibit motility similar to that inside the embyro and reveals a deficiency in the TIT mutant cells. Development 100,587-598. Kaufman, M. H. (1992). “The Atlas of Mouse Development.” London: Academic Press. Kelly, R., Alonso, S.,Tajbakhsh, S., Cossu, G., and Buckingham, M. (1995). Myosin light chain 3F regulatory sequences confer regionalised cardiac and skeletal muscle expression in transgenic mice. J Cell Biol. 129, 383-396. Kopan, R., Nye, J. S., and Weintraub, H. (1994). The intracellular domain of mouse Notch: A constitutively activated repressor of myogenesis directed at the basic helix-loop-helix region of MyoD. Development lzo, 2385-2396. Li, Z., Marchand, P., Humbert, J., Babinet, C., and Paulin, D. (1993). Desmin sequence elements regulating skeletal muscle-specific expression in transgenic mice. Development ll7,947-959. Miller-Hance, W. C., LaCorbiere, M., Fuller, S. J., Evans, S. M., Lyons, G., Schmidt, C., Robbins, J., and Chien, K. R. (1993).I n vitro chamber specification during embryonic stem cell cardiogenesis. J. Biol. Chem. 268,24244-25252. Muneoka, K., Wanek N.,Trevino, C., and Bryant, S.V. (1990).Ex utero surgery. I n “Postimplantation Mammalian Development” (A. J. Copp and D. L. Cockroft, eds.) pp. 41-60. IRL Oxford University Press. Olson, E. N., and Rosenthal, N. (1994). Homeobox genes and muscle patterning. Cell 79,9-12.

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Ringwald, M., Baldock, R., Bard, J., Kaufman, M., Eppig, J. T., Richardson, J. E., Nadeau, J. H., and Davidson, D. (1994). A database for mouse development. Science 265,2033-2034. Robert, B., Barton, P., Minty, A., Daubas, P.,Weydert, A., Bonhomme, F., Catalan, J., Chazottes, D., Guhet, J. L., and Buckingham, M. (1985). Investigation of genetic linkage between myosin and actin genes using an interspeciflcmouse back-cross. Nature 314,181-183. Rudnicki, M. A., Schneglesberg, P. N. J., Stead, R. H., Braun, T., Amold, H. H., and Jaenisch, R. (1993). MyoD or myf-5 is required for the formation of skeletal muscle. Cell 75,1351-1359. Sassoon, D., Gamer, I., and Buckingham, M. (1988). Transcripts of acardiac and a-skeletal actins are early markers for myogenesis in the mouse embyro. Development 104,155-164. Selfridge, J., Pow, A. M., McWhir, J., Magin, T. M., and Melton. D. W. (1992). Gene targeting using a mouse HPRT minigene/HPRT-deficient embryonic stem cell system: Inactivation of the mouse ERCC-I gene. Somatic Cell and MoL Genet. 18,325-336. Serbedzija,G., Fraser, S. E., and Bonner-Fraser, M. (1990). Pathways of trunk neural crest migration in the mouse embryo revealed by vital dye analysis. Development 108,605-612. Stem, C . (1993).Immunocytochemistry of embronic material. In “Essential Developmental Biology” (C. D. Stem and P. W. Holland, eds.), pp. 193-212. IRL Oxford University Press. Tajbakhsh, S., and Buckingham, M. (19%). Lineage restriction of the myogenic conversion factor myf5 in the brain. Development 121,4077-4083. Tajbakhsh, S., Bober, E., Babinet, C., Pournin, S., Arnold, H. H., and Buckingham, M. (1996a). Gene targeting the myf-5 locus with nlacZ reveals expression of this myogenic factor in mature skeletal muscle fibres as well as early embryonic muscle. Dev. Dynamics 206,291-300. Tajbakhsh, S., Rocancourt, D., and Buckingham, M. (1996b). Muscle progenitor cells failing to respond to positional cues adopt nonmyogenic fates in myf-5 null mice. Nature 384,266-270. Tajbakhsh, S., Vivarelli, G., Cusella-De Angelis, G., Rocancourt, D., Buckingham, M., and Cossu, G. (1994).A population of myogeniccells derived from the mouse neural tube. Neuron l3,813-821. Tajbakhsh, S., and Houzelstein, D. (1995). In situ hybridization and P-galactosidase: A powerful combination for analysing transgenics, TIC 11,42. Tajbakhsh, S., Vivarelli, G., Cusella-De Angelis, G., Rocancourt, D., Buckingham, M., and Cossu, G. (1994).A population of myogeniccells derived from the mouse neural tube. Neuron -813-821. Vittadello, M., Schiaffino,M. V., Picard, A., Scarpa, M., and Schiaffino, S. (1994). Gene transfer in regenerating muscle. Human Gene Therapy 5,ll-18. Vivarelli, E., and Cossu, G. (1986). Neural control of early myogenic differentiation in cultures of mouse somites. Develop. Biol. 117,319-325. Wachtler, F., and Christ, B. (1992).The basic embryology of skeletal muscle formation in vertebrates: The avian model. Seminars in Dev. Bid. 3,217-227 Wang, Y., Schnegelsberg, P.N. J., Dausman, J., and Jaenisch, R. (19%). Functional redundancy of the muscle-specific transcription factors myf-5 and myogenin. Nature 379,823-825. Wasserman, P. M., and DePamphilis, M. L., eds. (1993). “Guide to Techniques in Mouse Development,” Methods in Enzymology, Vol. 225. New York Academic Press. Weintraub, H., Davis, R., Tapscott, S., Thayer, M., Krause, M., Benezra, R., Blackwell,T. K., Turner. D., Rupp, R., Hollenberg, S., Zhuang, Y., and Lassar, A. (1991). The MyoD gene family: Nodal point during specification of the muscle cell lineage. Science 25% 761-766. W W s o n , D. G., (1992). “In Sim Hybridization.” IRL Oxford University Press. Yagi, T., Ikawa, Y., Yoshida, K.,Shigetani,Y., Takeda, N., Mabuchi, I., Yamamoto, T., and Aizawa, S. (1990). Homologous recombination at c-fyn locus of mouse embryonic stem cells with use of diphtheria toxin A-fragment gene in negative selection. Proc. Natl. Acad. Sci. USA 87,9918-9922. Yee, S.-P., and Rigby, P. W. J. (1993). The regulation of myogenin gene expression during the embryonic development of the mouse. Genes Dev. 7,1277-1289. Zemicka-Gomez,M., Pines, J., McLean Hunter, S., Dixon, J. P. C., Siemering, K.R., Haseloff, J. and Evans, M. J. (1997). Following cell fate in the living mouse embryo. Development -1133-1137.

CHAPTER 3

Myogenesis in Xenopus Embryos John B. Gurdon,* Patrick Lemaire; and Timothy J. Mohun* Wellcome/CRC Institute of Cancer and Developmental Biology Cambridge CB2 1QR United Kingdom t Institut de Biologie du Developpment de Marseille

Marseille, France

*

National Institute for Medical Research The Ridgeway, Mill Hill London "7 1AA United Kingdom

I. Fate Maps for Muscle A. Limb and Head Muscle B. Axial Muscle C. Cardiac Muscle 11. Embryo Manipulations A. Methods of Dissection B. W and Lithium Treatment of Early Embryos C. Animal Caps and Conjugates D. Signaling Sources E. Signaling Molecules Meeting Mesoderm Formation and Patterning 111. Molecular Markers of Muscle DiEerentiation A. Transcript Markers B. Methods of Analysis C. DNA-Binding Proteins (EMSA) IV. Studies of Xenopus Heart Formation References

I. Fate Maps for Muscle A. Limb and Head Muscle It is assumed that Xenopus is similar to three other vertebrates where it has been shown that limb muscles arise by migration of muscle progenitor cells from METHODS IN CELL BIOLOGY, VOL. 52 Copynght 0 1998 by Academic Pres. AU rightr of repduction in m y form reserved. 0091-679X/98 $25.00

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John B. Gurdon ct al.

54

axial somites. The origin of head muscle has not been traced in detail, but derives from the most anterior part of the invaginating mesoderm, which is located in postgastrular stages in front of the notochord. B. Axial Muscle

This derives from the somites, which in turn come from mesoderm tissue that is invaginated during gastrulation. The creation of different regions of the mesoderm, and its changing shape during gastrulation, are illustrated in color by Keller (1991). Somites are formed progressively at the rate of about 1.5 per hr from stage 17 onwards (Nieuwkoop and Faber, 1956). In Xenopus, by far the greatest part of a somite consists of myotome cells that will form muscle. A small dorsolateral part of each somite will form skin (dermatome), and only the smallest amount nearest the notochord is thought to form sclerotome. The location of dermatome and myotome regions are indicated by Hausen and Riebesell (1991; see plates 35 and 38). The location of the mesoderm as seen during gastrulation is more complicated in Xenopus than in other amphibians such as urodeles. Dale and Slack (1987b) show that most axial muscle derives from the dorsal marginal zone at stages 8-10. It is significant that mesoderm cells are located inside the embryo, and that surface equatorial (or future marginal) cells at the late blastula and early gastrula stages do not contribute to the mesoderm (Keller, 1991). The position of muscle progenitor cells at the early gastrula stage has been determined by culturing explants. Kato and Gurdon (1993) find that 40-60% of the cells contained in an internal dorsolateral region of an early gastrula will form muscle cells. The lineage tracing of mesoderm cells has been carried out at the 32-cell stage by Dale and Slack (1987a). They find that most (83%)of the somite, and therefore most of the muscle, derives from blastomeres B2+3 and (2-4 (corresponding to the lateral region), though no more than 20% of the daughters of any one of these blastomeres give rise to muscle (Fig. 1). Their results show that cell lineage is markedly imprecise at these early stages, and muscle can occasionally be contributed by daughters of nearly all other 32-cell blastomeres. Similar results have been described by Moody (1987). For some experiments, it is important to know the origin of experimentally treated cells that may form muscle. For this purpose, 10-15 nl of the fixable (by formaldehyde) lineage markers rhodamine or fluorescein lysinated dextran can be injected into eggs or early embryo blastomeres at 2 m g / d in distilled water. Alternatively, mRNA encoding P-galactosidase can similarly be injected. The fluorescent markers and the encoded P-galactosidase persist in cells until at least stage 40, by which time the majority of muscle markers have been expressed. C. Cardiac Muscle

Lineage labeling studies have shown that two blastomeres of the 32-cell stage contribute to the embryonic heart (Dale and Slack, 1987a). These correspond

3. Myogenesis in Xeropur Embryos

55

Fig. 1 Designation of blastomeres in 32-cell stage Xenopus embryo. Those shaded dark gray contribute 40-508 of their progeny to somites. Those shaded light gray contribute 30-408 of their progeny to somites. Most of the somite becomes muscle (from Dale and Slack, 1987a). The future dorsal side of the embryo lies to the right.

to blastomeres C1 and C2 (Fig. 1). During gastrulation, the regions of mesoderm destined to form the heart tube are found on either side of the dorsal lip and are distinct from the somitic mesoderm (Keller, 1976). By the end of gastrulation, these are arranged bilaterally, at the anterior edge of the newly forming neural plate. During neurulation, the two regions of cardiac mesoderm are found in progressively more ventral locations, until by stage 16, they begin to fuse along the ventral midline (Nieuwkoop and Faber, 1956). By stage 19, a single presumptive heart anlage has been formed, which lies caudal to the cement gland. Little further morphological differentiation can be detected until stage 28, when the endocardial tube begins to form. The characteristic S-shaped looping of the tube occurs between stages 33 and 36. Heartbeat is initiated at stage 34 and distinct atrial and ventricular chambers can be distinguished by stage 3516.

11. Embryo Manipulations

A. Methods of Dissection After gastrulation, it is sometimes useful to be able to dissect embryos into different tissues. Xenopus embryos are large enough that direct biochemical analyses (not RT.PCR) can be carried out on tissues from only a few embryos. Interestingly, tissues between which an induction is taking place tend to be tightly adherent to each other and therefore difficult to separate. We have found that a 0.3%solution of commercial collagenase (e.g., Sigma (2-2139, which like other sources is impure) in 1X MBS (Gurdon, 1977) is very helpful in enabling tissues to be separated by gentle action with needles. However, it is essential first to break the very impermeable surface coat of embryos, for example, by removing

John B. Gurdon et al.

56

the endoderm. Incubation in the collagenase solution for 10-20 min enables tissues to be gently peeled apart, without dissection of the component cells, as long as calcium ions are retained in the MBS-collagenase solution. Once separated, tissues can be cultured indefinitely in normal l X MBS, with no apparent ill effects from the collagenase treatment. This procedure was used to prepare reasonably pure samples of muscle, notochord, neurectoderm, ventral mesoderm, and ectoderm from tail-bud stages (Mohun et aL, 1984). Between stages 18 and about 35, complete strips of somite can be dissected from each side of any embryo, the anterior part consisting of segmented somites, and the posterior part of unsegmented mesoderm. Isolated somites can be dissociated into single muscle cells by incubation in calcium-free MBS, with 0.5 mM EDTA, at pH 8.0. In Xenopus, muscle cells are mononucleate for the whole of embryonic life. B. UV and Lithium Treatment of Early Embryos Xenopus embryos can be epigenetically manipulated to obtain ventralized or dorsalized phenotypes. These phenotypes can be described using the dorsoanterior index (DAI) (Kao and Elinson, 1988). Embryos with a DAI of 0 to 4 are ventralized; normal embryos have a DAI of 5; and dorsalized embryos have a DAI of 6 to 10. As muscle is an intermediate tissue, it is not found in severely ventralized embryos (DAI 0 to 2) and is probably absent from dorsalized embryos (DAI 9 to 10). Therefore, these embryos can be a useful source of nonmuscle mesoderm for recombination experiments. Severalprocedures have been used to ventralize embryos. The most convenient is the vegetal irradiation of embryos with ultraviolet light during the first cell cycle. A good discussion of the important parameters to obtain reproducible UV ventralization can be found in Kao and Danilchik (1991). The UV source can be a handheld illuminator (UV Light Products, San Gabriel, California, USA), a transilluminatorused for viewing ethidium bromide-stained gels, or an inverted UV oven (Stratalinker, Stratagene). The last allows a more careful and reproducible dosage of the irradiation and is therefore recommended. We find it convenient to place a small opaque shelf across the UV oven. This shelf should have two holes: one over which the fused quartz dish containing the embryos is placed, and a second one to allow the meter to receive some ultraviolet light. The optimal energy required to ventralize a given batch of embryos must be determinedempirically. Dorsalization of embryos is obtained by placing 32-128-cell embryos in a solution containing 0.3 it4 lithium chloride for 5 to 10min. The important parameters for an efficient dorsalization are discussed in Kao and Danilchik (1991). C. Animal Caps and Conjugates

The Nieuwkoop conjugate, fist used by Nieuwkoop (1969), is the basis of the animal cap assay widely used in current embryological work with amphibia. An

3. Myogenesis in Xmopur Embryos

57

animal cap is that part of a blastula that, if isolated, will form ectoderm. If an animal cap is exposed to the appropriate dose of a mesoderm inducer such as activin, it will form mesodermal cell types, including muscle. Similarly, an animal cap isolated from embryo injected at the two-cell stage with mRNA that encodes an inducer protein will also form muscle. This animal cap assay has been extensively used to determine the identity and mode of action of growth factors. To prepare animal caps, dejellied blastulas should be placed in 1 X MBS (or in equivalent medium) and the vitelline membrane removed with forceps, taking care not to damage the animal hemisphere of the embryo. Using forceps, the one-third of the embryo nearest the animal pole is cut off and placed upside down (surface coat and pigmented region downwards) in a glass dish. Forceps or needles are then used to cut away all of the thicker, less pigmented region on the periphery. The remaining thin layer of animal pole ectoderm can then be cultured. It is often helpful to combine two animal caps face-to-face to obtain survival to late stages (equivalent of stage 40 or beyond). Animal caps not treated with growth factor, and not injected as eggs with mRNA, should form ectoderm, with no trace of muscle. It is necessary to include controls of this kind in all experimental series since different batches of embryos differ in how cleanly all mesoderm (equatorial) tissue can be removed. D. Signaling Sources The differentiation of part of the marginal zone tissue from a blastula into axial muscle involves a cascade of cell-cell interactions. The first is the induction of mesoderm by one or several signals emitted by vegetal cells. In addition, a dorsalizing signal acts in the late blastula and gastrula stages. This signal has the ability to convert ventral mesoderm into more dorsal types such as muscle or notochord and is secreted from the organizer. Also, Gurdon and colleagues (1993b) have shown that muscle precursors cultured in vitro during the gastrula stages differentiate into muscle only if they are in communication with a large number of like cells, an effect named the community effect. Little is known about the community-effectfactor, but it is assumed that it is a soluble polypeptide secreted by future muscle cells. A useful discussion of the roles of these three types of factors can be found in Gurdon et al. (1993a). Other factors, acting later in development, also have an influence on muscle development. For example, signals emanating from the notochord inhibit the differentiation of dorsal somitic derivatives (Pourquie et al., 1993). E. Signaling Molecules Affecting Mesoderm Formation and Patterning

Several secreted polypeptides can have an effect on mesoderm differentiation and the formation of axial muscle during the blastula and early gastrula stages. Different assays exist to test the activity of these factors. The most common ones are the animal cap assay and perturbation of the body axis following mRNA


58

injections. The former reveals the mesoderm-inducingability of a factor, whereas the latter reveals the ability of a factor to modify the fate of induced mesodermal cells. In the animal cap assay, a dissected blastula animal cap is treated with a dilute solution of a secreted polypeptide. If the factor used is a mesoderm-inducing factor, the animal pole ectoderm is converted into mesoderm, a conversion that can be demonstrated by histological analysis of the explants. This assay is described in detail in Smith (1993) and Slack (1993). Numerous variants of this assay exist. Rather than treating the explant with a soluble polypeptide, it is possible to overexpress the polypeptide by injecting its mRNA into fertilized eggs (reviewed in Vize et al., 1991). The factor is then synthesised in all blastomeres, including those that form the animal pole of the embryo. Using animal caps isolated from such embryos it is possible to test the activity of factors that are not available in a soluble form. Table I shows the effects of some known secreted factors in this assay. The ability of a polypeptide to affect the formation of the body axis can be assayed by injecting mRNA or DNA for this factor into vegetal blastomeres of early cleavage stage embryos (reviewed in Vize et al., 1991). Injection of mFWA for ventralizing factors into a dorsal blastomere leads to the truncation of the axis, whereas injection of mRNA for dorsalizing factors into a ventral blastomere leads to the formation of a secondary axis (Table 11). For example, a dorsal injection of BMP4 mRNA leads to ventralized embryos (Dale et aL, 1992), Table I Conversion of Ectoderm into Mesoderm by Secreted Polypeptide Factors Molecule Mesoderm inducers

Vg-1"

RNA injection

Activin

Soluble factor or RNA injection Soluble factor RNA injection

TGFIP;! BMP4 bFGF Modifiers

Administration

eFGF Noggin XWnt-8 Wnt-1 XWnt-11

Soluble factor or RNA injection RNA injection Soluble factor or RNA injection RNA injection RNA injection RNAS injection

Mesoderm

Muscle

+

+ + +

+

+ + +

+ -c

Hybrid mRNA producing a mature form of the Vg-1 polypeptide. At high concentrations only. Conversion of the injected ectodermal tissue into neurectoderm.

-

+ -

Reference

Dale er al. (1993); Thomsen and Melton (1993) Smith et al. (1990) Rosa et al. (1988) Dale et al. (1992); Jones et al. (1992) Kimelman and Kirschner (1987); Kimelman and Maas (1992) Isaacs et al. (1994) Smith and Harland (1992); Smith er al. (1992) Christian et al. (1992) McMahon and Moon (1989) Ku and Melton (1993)

3. Myogenesis in Xmopw Embryos

59

Table I1 Conversion of Ventral Mesoderm into Muscle Ventral to axial mesoderm

Rescue of UV ventralized embryos

AXiS duplication

ND*

Complete

ND

Activin BMP4

+

Partial

Partial

bFGF Noggin XWnt-8

ND

+ +

None Complete Complete

Wnt-1 XWnt-11

ND ND

ND Partial

Molecule Mesoderm

Modifiers

a

Vg-1"

-

ND

-c

ND ND Complete Complete

ND

Reference Dale ef al. (1993); Thomsen and Melton (1993) Sokol and Melton (1991) Dale et al. (1992); Jones et af. (1992) Kimelman and Maas (1992) Smith and Harland (1992) Smith and Harland (1991); Sokol et al. (1991) McMahon and Moon (1989) Ku and Melton (1993)

Hybrid mRNA producing a mature form of the Vg-1 polypeptide. ND: not done. Ventralization of injected embryos.

whereas ventral injection of mRNA for several members of the wnt family leads to embryos with two axes (Moon, 1993). A variant of this approach is the injection of mRNA into a vegetal blastomere of a UV-ventralized embryo. One then looks for the rescue of the axis (described in Smith and Harland, 1991; Sokol et al., 1991). It is assumed that in all cases, the perturbation of the axis foIlowing vegetal injections of mRNAs results from the alteration of the fate of the mesodermal precursor cells located just above the progeny of the injected vegetal cells. Indeed, whenever tested, mRNA that could perturb the axis when injected vegetally could also respecify the fate of mesodermal cells when injected in the marginal zone (Table 11).

111. Molecular Markers of Muscle Differentiation A. Transcript Markers A number of genes, normally expressed from the late blastula stages onwards, are activated as a direct response to mesoderm induction. Although their expression is not limited to muscle, they have proved useful markers for studying the inductive interactions that are responsible for specifying muscle progenitor cells (Table 111). Muscle-specific genes are first activated during gastrulation and subsequently identify differentiating myotomal muscle cells. Genes encoding cardiac muscle-specific isoforms are only expressed much later in development, at the tail-bud stage (Table IV).


60

Table 111 Some Useful Early Markers for Mesodermal Differentiation Specificity at gastrula stage

Specificity at neurula stage

Goosecoid

Invaginating mesoderm Nondorsal mesoderm and vegetal Cells Mesoderm and vegetal cells Organizer

Noggin

Probe

Onset

Comments

Posterior mesoderm and notochord Ventral and lateral mesoderm

MBT”

Easy to detect by in situ hybridization Low abundance declines in gastrula

Smith et al. (1991)

Not detectable

MBT

Rosa et al. (1988)

Prechordal plate

MBT

Organizer

Prechordal plate and notochord

MBT

Transient expression peaks St. 10 Easy to detect by in situ hybridization Variable amount of maternal transcripts

XLiml

Organizer

MBT

Taira et al. (1992)

XFHKl

Organizer

Notochord, pronephros, and pineal body Notochord

MBT

mot2

Organizer

MBT

eFGF

Invaginating mesoderm

Notochord, floor plate, and pineal body Posterior mesoderm

Dirksen and Jamrich (1992) von Dassow et al. (1993)

Xbra XWnt-8

MiXl

MBT

MBT

Reference

Christian et al. (1991); Lemaire and Gurdon (1994)

Cho et al. (1991) Smith and Harland (1992)

Isaacs et al. (1994)

Mid-blastula transition.

B. Methods of Analysis Analysis of the expression of the markers mentioned earlier can be performed using biochemical or histological methods. RNase protection and Northern blots are useful for the quantitation of transcript amounts and can be performed using well-established protocols. In situ hybridizations (ISH) are preferred if information about the spatial distribution of transcripts is required. Depending on the application, ISH can be performed on paraffin sections using radioactive (O’Keefe et d.,1991) or digoxygenin-labeled probes (Lemaire and Gurdon, 1994), or on whole embryos using digoxygenin-labeled probes (Harland, 1991). Protocols using digoxygenin probes are faster and give a much better resolution, but may fail to detect rare transcripts. Whole-embryo protocols offer the advantage of showing a three-dimensional view of the expression pattern of a gene, but penetration problems may prevent visualization of transcripts in the yolky vegetal cells of early embryos (Lemaire and Gurdon, 1994).

3. Myogenesis in Xenopur Embryos

61

Table IV Some Useftl Markers for Muscle Differentiation Probe

Specificity

Onset

Comment

Reference

Cardiac actin Skeletal actin Femoral actin XMyoD

Striated muscle Striated muscle Striated muscle

Stage 10.5" Stage 11" Stage 12"

Highly abundant Highly abundant Highly abundant

Mohun et al. (1988, 1994) Mohun et al. (1988,1994) Mohun et al. (1988, 1994)

Myotomal muscle

Stage 1 O S b

XMyfS

Myotomal muscle

Stage 9.5

Hopwood et al. (1989); Harvey (1991) Hopwood et al. (1991)

xMRF4 XNKX2.5 XMHCa XMLC;! Troponin I

Myotomal muscle Cardiac mesoderm Cardiac muscle Cardiac muscle Cardiac muscle

Stage 18 Stage 15 Stage 27 Stage 27 Stage 28

Low levels-decline sharply in tail bud Low levels-decline sharply in tail bud Low abundance Low abundancec Abundant Abundant Abundant

~

Jennings (1992) Tonissen et al. (1994) Logan and Mohun (1993) Chambers et 01. (1994) Drysdale et al. (1994)

~~

" Detected by PCR. Maternal transcripts detected. Transient zygotic expression throughout the mesoderm (Frank and Harland, 1991; Harvey, 1991). Detected throughout heart field and at low levels in other regions of the embryo.

C. DNA-Binding Proteins (EMSA) Analysis of DNA-binding activities in Xenopus embryos is most conveniently camed out using whole cell extracts. Because of the large size of amphibian embryos, extracts from single embryos or explants can be analyzed successfully. The presence of large amounts of yolk proteins in such extracts does not appear to interfere with binding assays or with the subsequent electrophoresis step. A convenient protocol for electrophoretic mobility shift assays (EMSA), which can be scaled up or down as appropriate, is the following: 1. Homogenize 12 embryodoocytes in 100 p1 ice-cold modified Dignam C (MDC) buffer: 50 mM Tris-HC1, pH 8.0 50 mM KC1 0.1 mM EDTA 2mMDTT 25% glycerol

A cocktail of ihhibitors may improve the extract quality. For example: Pepstatin OSpg/ml Leupeptin OSpg/ml Chymostatin 0.5pg/ml Antipain OSpg/ml


62

Benzamidine 2 mM PMSF 2 mM For extracts enriched in muscle-specific binding activities, dorsal posterior embryo fragments (stages 28-32) or tail pieces from early tadpoles (stages 34-38) are convenient to use (Taylor el aL, 1991; Chambers et aL, 1992). 2. Immediately microfuge (14,000K) at 4°C for 2minutes and remove the aqueous layer, avoiding as much of the floating lipid as possible. Flash freeze the extract in aliquotsin dry icelethanol and store at -70°C. (Note: Extracts deteriorate with freezekhaw cycles, so discard unused extract after thawing.) Procedures for electrophoretic mobility shift assays vary widely and frequently need to be optimized for the binding activity of interest. The following has provided a useful starting point: 3. Combine the following: 4 pl embryo extract 5 pl MDC buffer 1 p1 15 mM spermidine 1 p1 nonspecific competitor DNA (e.g., pUC, salmon sperm DNA, polydI : dC) 1pl1OX MglEDTA mix (15 mM MgC12; 7.5 mM EDTA; 1 mM D"; 1 mM PMSF) 6 p1 water (including any specific competitor DNA) Preincubate for 15 min on ice and then add 2 pl radiolabeled probe oligo or fragment (approx. 1 ng or 1020,OOo cpm) 4. Incubate for 15 min on ice, add bromophenoYglycero1 loading dye (optional), and load onto a 29 :lacrylamide :bisacrylamide gel (4-696) prerun at 100 V/30 mA for 2-3 hr. Gels can be run at room temperature or 4°C.

This procedure has been successfully used to study binding activities containing the myogenic factors (Taylor et af., 1991) and members of the MEF'2 family of transcription factors (Chambers et af., 1992).When comparing embryo fragments, a ubiquitous binding activity such as SP1 can be used to normalize results. (Note: SP1 levels vary widely in adult tissues.)

W.Studies of Xenopus Heart Formation A series of studies from Jacobson and Sater summarize the experimental approaches that can conveniently be used to study heart formation in Xenopus embryos (Sater and Jacobson, 1989,1990a,1990b).At the gastrula stage, explants

3. Myogenesis in Xmopvs Embryos

63

containing the cardiac mesoderm can be obtained by removing the region between 30 and 45 degrees with respect to the newly formed dorsal lip on each side of the embryo (Sater and Jacobson, 1989). Such fragments have been studied using hanging drop cultures, but can also be successfully maintained in agarlined dishes or microtiter plates containing amphibian saline (e.g., 3/4 NAM). They differentiate to form a beating heart-tube structure if maintained for several days. Recent studies have demonstrated that the ability of such early explants to differentiate depends upon signals from the underlying endoderm, which is required until stage 10.5 (Nascone and Mercola, 1995). By stage 18, the two regions have fused on the ventral midline and can be isolated as a single fragment immediately behind the cement gland (see Sater and Jacobson, 1990a). Again, explants differentiate to form a vesicle containing a heart tube showing some degree of S-shaped looping and undergoing spontaneous, rhythmic contractions. Differentiation can occur in the absence of associated endodermal tissue and will also occur in each half of a subdivided explant. Conjugates prepared from pairs of explants from which the endoderm has been removed will form a single heart structure. Heart-forming capacity is not restricted to the anterior ventral region of the embryo in early tail-bud embryos. Adjacent regions of lateral mesoderm will also form beating hearts, and this capacity is retained to some extent in subdivided fragments. During tail-bud stages of development, the capacity of lateral fragments to undergo heart morphogenesis declines and is lost by stage 28 (Sater and Jacobson, 1990a). Little is currently known about how the “heart morphogenetic field” is established, nor do we understand why heart-forming potential is subsequently lost by ventrolateral mesoderm. Cardiac muscle differentiation can also be induced in animal cap explants by treatment with high doses of activin A. Under these conditions, a small proportion of cells within the explant form cardiac muscle, while a much larger number form skeletal muscle or notochord (Logan and Mohun, 1993). The time course of cardiac muscle differentiation in such explants approximates that of normal development, and a characteristic rhythmic “heartbeat” is frequently observed. Only weak induction is obtained with individual explants, whereas fused aggregates contain proportionally far more cardiac muscle tissue. References Chambers, A. E., Kotecha, S., Towers, N., and Mohun, T. J. (1992). Muscle-specific expression of SRF-related genes in the early embryo of Xenopus luevis. EMBO J. 11,4981-4991. Chambers, A. E., Logan, M., Kotecha, S., Towers, N., Sparrow, D., and Mohun, T. J. (1994). The RSRFMEF2 protein SLl regulates cardiac muscle-specific transcription of a myosin light-chain gene in embryos. Genes Dev. 8,1324-1334. Cho, K. W., Blumberg, B., Steinbeisser,H., and De, R. E. M. (1991). Molecular nature of Spemann’s organizer: The role of the Xenopus homeobox gene goosecoid. Cell 67,1111-1120. Christian, J. L., McMahon, J. A., McMahon, A. P., and Moon, R. T. (1991). Xwnt-8, a Xenopus Wnt-l/int-l-related gene responsive to mesoderm-inducing growth factors, may play a role in ventral mesodermal patterning during embryogenesis.Development 11%1045-1055.

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Christian, J. L., Olson, D. J., and Moon, R. T. (1992). Xwnt-8 modifies the character of mesoderm induced by bFGF in isolated Xenopus ectoderm. EMBO J. ll,33-41. Dale, L., and Slack, J. M. W. (1987a). Fate map for the 32-cell stage of Xenopus laevis. Development 99,527-551. Dale, L., and Slack, J. M. W. (1987b). Regional specification within the mesoderm of early embryos of Xenopus laevis. Development 100, 274-295. Dale, L., Howes, G., Price, B. M., and Smith, J. C. (1992). Bone morphogenetic protein 4 A ventralizing factor in early Xenopus development. Development 115,573-585. Dale, L., Matthews, G., and Colman, A. (1993). Secretion and mesoderm-inducing activity of the TGF-beta-related domain of Xenopus Vgl. EMBO J. 12,4471-4480. Dirksen, M. L., and Jamrich, M. (1992). A novel, activin-inducible, blastopore lip-specific gene of Xenopus laevis contains a fork head DNA-binding domain. Genes Dev. 6,599-608. Drysdale, T. A., Tonissen, K. F., Patterson, K. D., Crawford, M. J., and Krieg, P. A. (1994). Cardiac troponin-I is a heart-specific marker in the Xenopus embryo. Expression during abnormal heart morphogenesis. Dev. Biol. 165,432-441. Frank, D., and Harland, R. M. (1991). Transient expression of XMyoD in non-somitic mesoderm of Xenopus gastrulae. Development ll3,1387-1393. Gurdon, J. B. (1977). Methods for nuclear transplantation in Amphibia. Methods Cell Biol. 16, 125-139. Gurdon, J. B., Kato, K., and Lemaire, P. (1993a). The community effect, dorsalization and mesoderm induction. Curr. Opin. Genet. Dev. 3, 662-667. Gurdon, J. B., Tiller, E., Roberts, J., and Kato, K. (1993b). A community effect in muscle development. Curr. BioL 3,1-11. Harland, R. M. (1991). In situ hybridisation: An improved whole mount method for Xenopus embryos. I n “Xenopus laevis: Practical Uses in Cell and Molecular Biology” (B. K. Kay and H. B. Peng, eds.), Vol. 36, pp. 685-695. London: Academic Press. Harvey, R. P. (1991). Widespread expression of MyoD genes in Xenopus embryos is amplified in presumptive muscle as a delayed response to mesoderm induction. Proc. Natl. Acad. Sci. USA 88,9198-9202. Hausen, P., and Riebesell, M. (1991). “The Early Development of Xenopus Laevis.” London: Springer-Verlag. Hopwood, N.D., Pluck, A., and Gurdon, J. B. (1989). MyoD expression in the forming somites is an early response to mesoderm induction in Xenopus embryos. EMBO J. 8,3409-3417. Hopwood, N.D., Pluck, A., and Gurdon, J. B. (1991). Xenopus Myf-5marks early muscle cells and can activate muscle genes ectopically in early embryos. Development 111,551-560. Isaacs, H. V., Pownall, M. E., and Slack, J. M. W. (1994). eFGF regulates XBra expression during Xenopus gastrulation. EMBO J. l3,4469-4481. Jennings, C.G. (1992). Expression of the myogenic gene MRF4 during Xenopus development. Dev. BioL Wl, 319-332. Jones, C. M., Lyons, K. M., Lapan, P. M., Wright, C. V., and Hogan, B. L. (1992). DVR-4 (bone morphogeneticprotein-4) as a posterior-ventralizingfactor in Xenopus mesoderm induction.Development 115,639-647. Kao, K., and Danilchik, M. (1991). Generation of body plan phenotypes in early embryogenesis. Methods Cell BWl. 36,271-284. Kao, K.R., and Eliinson, R. P. (1988). The entire mesodermal mantle behaves as Spemann’s organiser in dorsoanterior enhanced Xenopus laevis embryos. Dev. BWL l27,64-77. Kato, K., and Gurdon, J. B. (1993). Singlecell transplantation determines the time when Xenopus muscle precursor cells aquire a capacity for autonomous differentiation. Proc. Natl. Acad. Sci. USA 90,1310-1314. Keller, R. E.(1976). Vital dye mapping of the gastrula and newula of Xenopus laevis. I. Prospective areas and morphogenetic movements of the deep layer. Dev. BioL Sl, 118-137. Keller, R. (1991). Early embryonic development of Xenopus laevis. Methods Cell Biol. 36,61-113.

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Kimelman, D., and Kirschner, M. (1987). Synergistic induction of mesoderm by FGF and TGF-fi and the identification of an mRNA coding for FGF in the early Xenopus embryo. Cell 51,118-137. Kimelman, D., and Maas, A. (1992). Induction of dorsal and ventral mesoderm by ectopically expressed Xenopus basic fibroblast growth factor. Development 114,261-269. Ku, M., and Melton, D. A. (1993). Xwnt-11:A maternally expressed Xenopus wnt gene. Development 119,1161-1173. Lemaire, P., and Gurdon, J. B. (1994). A role for cytoplasmic determinants in mesoderm patterning: Cell-autonomous activation of the goosecoid and Xwnt-8 genes along the dorsoventral axis of early Xenopus embryos. Development l20,1191-1199. Logan, M., and Mohun, T. (1993). Induction of cardiac muscle differentiation in isolated animal pole explants of Xenopus laevis embryos. Development 118,865-875. McMahon, A. P., and Moon, R. T. (1989). Ectopic expression of the proto-oncogene int-Z in Xenopus embryos teads to duplication of the embryonic axis. Cell 58, 1075-1084. Mohun, T., Brennan, S., Dathan, N., Fairman, S., and Gurdon, J. (1984). Cell type-specificactivation of actin genes in the early amphibian embryo. Nature 311, 716-721. Mohun, T., Garrett, N., Stutz, F., and Spohr, G. (1988). A third striated muscle actin gene is expressed during early development in the amphibian, Xenopus laevis. J. Mol. Biol. 202,67-76. Mohun, T., Wilson, R., Gionti, E., and Logan, M. (1994). Myogenesis in Xenopus laevis. Trenh in Card. Vasc. Med. 4,146-151. Moody, S . A. (1987). Fates of the blastomeres of the 32-cell-stage Xenopus embryo. Dev. Biol. 122, 300-319. Moon, R. T. (1993). In pursuit of the functions of the Wnt family of developmental regulators: Insights from Xenopus laevis. Bioessays 15,91-97. Nascone, N., and Mercola, M. (1995). An inductive role for the endoderm in Xenopus cardiogenesis. Development, 121,515-523. Nieuwkoop, P. D. (1969). The formation of mesoderm in Urodelean amphibians. I. Induction by the endoderm. Wilhelm Roux’s Arch. Entw. Mech. Org. 162,341-373. Nieuwkoop, P., and Faber, J. (1956). “Normal Table of Xenopus laevis (Daudin).” Amsterdam: North-Holland. O’Keefe, H. P., Melton, D. A., Ferreiro, B., and Kintner, C. (1991). In situ hybridization. MethoL Cell Biol. 36,443-463. Pourquie, O., Coltey, M., Teillet, M. A., Ordahl, C., and Le Douarin, N. M. (1993). Control of dorsoventral patterning of somitic derivatives by notochord and floor plate. Proc. Natl. Acad. Sci. USA 90,5242-5246. Rosa, F., Roberts, A. B., Danielpour, D., Dart, L. L., Sporn, M. B., and Dawid, I. B. (1988). Mesoderni induction in amphibians: The role of TGFbeta2-like factors. Science 239,783-785. Sater, A. K., and Jacobson, A. G. (1989). The specification of heart mesoderm occurs during gastrulation in Xenopus laevis. Development 105,821-830. Sater, A. K., and Jacobson, A. G. (1990a).The restriction of the heart morphogenetic field in Xenopus laevis. Dev. Biol. 140,328-336. Sater, A. K., and Jacobson, A. G. (1990b). The role of the dorsal lip in the induction of heart mesoderm in Xenopus laevis. Development 108,461-470. Slack, J. (1993). In “Growth Factors: A Practical Approach” (I. A. McKay and I. Leigh, eds.), pp. 73-84. Oxford IRL Press. Smith, J. C. (1993). I n “Cellular Interactions in Development: A Practical Approach” (D. A. Hartley, ed.), pp. 181-204. Oxford IRL Press. Smith, W. C., and Harland, R. M. (1991). Injected Xwnt-8 RNA acts early in Xenopus embryos to promote formation of a vegetal dorsalizing center. Cell 67,753-765. Smith, W.C.. and Harland, R. M. (19%). Expression cloning of noggin, a new dorsalising factor localized to the Spemann organizer in Xenopus embryos. Cell 70,829-840. Smith, J. C., Price, B. M., Van Nimmen, K., and Huylebroeck, D. (1990). Identification of a potent Xenopus mesoderm-inducing factor as a homologue of activin A. Nature 345,729-731.

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Smith, J. C., Price, B. M., Green, J. B., Weigel, D., and Herrmann, B. G. (1991).Expression of a Xenopus homolog of Brachyury (T) is an immediate-early response to mesoderm induction. Cell 67,7947. Smith, W. C., Knecht, A. K.,Wu, M., and Harland, R. M. (1992).Secreted noggin protein mimics the Spemann organizer in dorsalizing Xenopus mesoderm. Nature 36% 547-549. Sokol, S.,and Melton, D. A. (1991). Preexistent pattern in Xenopus animal pole cells revealed by induction with activin. Nature 35% 409411. Sokol, S., Christian, J. L., Moon, R. T., and Melton, D. A. (1991). Injected Wnt RNA induces a complete body axis in Xenopus embryos. Cell 67,741-752. Taka, M., Jamrich, M., Good, P. J., and Dawid, I. B. (1992). The LIM domain-containing homeo box gene Xlim-1is expressed specifically in the organizer region of Xenopus gastrula embryos. Genes Dev. 6,356-366. Taylor, M. V., Gurdon, J. B., Hopwood, N. D., Towers, N., and Mohun, T. J. (1991). Xenopus embryos contain a somite-specific,MyoD-like protein that binds to a promoter site required for muscle actin expression. Genes Dev. 5,1149-1160. Thomsen, G. H., and Melton, D. A. (1993).Processed Vgl protein is an axial mesoderm inducer in Xenopus. Cell 74,433-441. Tonissen, K.F., Drysdale, T. A., Lints, T. J., Harvey, R. P., and Krieg, P. A. (1994).XNkx 2.5: A Xenopus homologue of Nkr-2.5 and tinman. Evidence for a conserved role in cardiac development. Dev. Bwl. 162,325-328. Vue, P.D.,Melton, D. A., Hemmati Brivanlou, A., and Harland, R. M. (1991). Assays for gene function in developing Xenopus embryos. Methods Cell Biol. 36,367-387. von Dassow, G., Schmidt, J. E., and Kimelman, D. (1993). Induction of the Xenopus organizer: Expression and regulation of Xnot, a novel FGF and activin-regulated homeo box gene. Genes Dev. 7,355-366.

CHAPTER 4

Zebrafish: Genetic and Embryological Methods in a Transparent Vertebrate Embryo Mark C. Fishman,' Didier Y. R. Stainier: Roger E. Breitbart,*and Monte Westerfields ' Cardiovascular Research Center Massachusetts General Hospital-East Harvard Medical School Charlestown. Massachusetts 02129 t Depaxtment of Biochemistry and Biophysics

University of California at San Francisco San Francisco, California 94143

* h4illenium Pharmaceuticals, Inc. Cambridge, Massachusetts 02139 Institute of Neuroscience University of Oregon Eugene, Oregon 97404

I. Introduction 11. Genetics

A. Mutagenesis B. Transgenic Fish 111. Heart Development A. Morphology B. Cellular Embryology C. Genetics IV. Skeletal Muscle Development A. Morphology B. Cellular Embryology C. Innervation D. Genetics E. Molecular Biology References METHODS IN CELL BIOLOGY, VOL. 52 Copyright 0 1998 by Academic Press. AU righn of reproduction in any form m w c d

0091-679X/98 S25.W

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Mark C. Pishman ct al.

I. Introduction The powers of the zebrafish are its embryological accessibility combined with tractable genetics. It is amenable to large-scale mutagenesis of the germ line and to visual screening for defects. The fish are hardy, lay hundreds of eggs on a regular basis, and can be raised by the thousands in reasonably sized facilities. Fertilization is external and the embryo is nearly transparent, so cells are accessible for injection or transplantation, and organs can be directly visualized. The cells of the blastula or gastrula can be transplanted and will incorporate into new hosts, permitting determination of whether mutations affect specific cells in a cell-autonomousmanner. Current major drawbacks include the lack of efficient techniques for insertional mutagenesis and for homologous recombination. All relevant techniques, including aquaculture, microscopy, mutagenesis, and histology, are described in detail in The Zebrafish Book (Westerfield, 1995) and so will not be elaborated here. Rather, we will briefly introduce several facets of this fish relevant to the genetic and cellular study of cardiac and skeletal muscle, focusing on working strategies and methodologies under active investigation.

11. Genetics For simple embryological manipulations, zebrafish can be obtained from pet stores or directly from fish farms. For genetics, it is preferable to begin mutagenesis with lines that lack lethal or other deleterious mutations. It had been hoped that heterozygosity could be eliminated by parthenogenesis. Eggs activated by UV-inactivated sperm bear only the maternal chromosomes. Hydrostatic pressure, which disrupts microtubules and inhibits the second meiotic division, can convert these to the diploid state, but such fish are not homozygous at every locus because of recombination between sister chromatids during meiosis. Heat shock of haploid embryos does produce homozygous embryos (Streisinger et al., 1981),but for unclear reasons this is lost over generations. Therefore, inbreeding and heat shock together have been utilized to generate lethal-free strains with minimal heterozygosity, although with concomitant loss of hybrid vigor.

A. Mutagenesis The timing and dose of mutagen are crucial (Grunwald and Streisinger, 1992; Mullins and Nusslein-Volhard, 1993; Solnica-Krezel et al., 1994). In practice, quantitation of effectiveness of a mutagen is achieved using pigmentation mutants. For example, a mutagenized male is crossed with an albino/albino hornozygous female and the F1 progeny examined to assess the frequency of new mutations at the albino locus (Grunwald and Streisinger, 1992; Mullins and Nusslein-Volhard, 1993;Solnica-Krezelet al., 1994).The pigmentation also serves

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to assay mosaicism of the mutation in the germ line. The commonly utilized agent, ENU, alkylates only one strand of DNA, and this modification is fixed only after replication or repair. Therefore, if the fish is bred shortly after mutagenesis, the ENU-affected germ cells will derive from mature spermatids (where the mutation has not yet been fixed) so some cells of the F1embryo will bear the mutation and others will not. With time after mutagenesis, the directly mutagenized spermatids are replaced by progeny of mutated spermatogonia that have divided several times. Because both DNA strands are affected, the F1 will be nonmosaic. In practice, this is achieved by waiting 1month or so after ENU before breeding the mutated male. The specific locus rate, of course, varies with the extent of DNA damage caused by the mutagen. For example, ENU is an alkylating agent believed to cause primarily point mutations. Used at just sublethal levels, it causes specific locus mutations at average frequencies of 1/1000. Gamma rays, which usually cause deletions, have specific locus frequencies of MOO. Therefore, for ensuring that single genes are affected, ENU is preferred. Once the F1 offspring of the mutagenized fish have been raised (3-4 months to sexual maturity), there are two strategies for screening (Kimmel, 1989;Weinberg, 1992; Driever et al., 1994). Haploid screening takes advantage of the parthenogenetic development that occurs if the eggs are activated with UV-irradiated sperm. These eggs develop with the maternal haploid chromosomal content, and generally die after five days. Although the embryos are always abnormal, distinctive mutant phenotypes can be visible over this background. The advantage of haploid screening is that it saves one generation of breeding prior to screening. The alternative is to generate FZfamilies, by crossing F1 fish with either wild-type or other F1 fish, and then screening the F3 progeny embryos obtained by sibling crosses. Even for flsh screened using haploid screening, study of the effect of the mutation without the other embryonic deficiencies inherent to haploids requires generation of F2families. In reality, isolation of single mutations by outcrossing of the identified F2fish is done, usually twice, prior to detailed study, to “ensure” that effects are due to single gene defects.

B. Transgenic Fish DNA injected into the early embryonic zebrafish is expressed in a mosaic pattern. Stable integration occurs and provides a reproducible pattern from animal (Westefield et al., 1992) to animal (Stuart et al., 1990). It is relatively easy to inject hundreds of 1- to 2-cell embyros at a sitting. Standard techniques use micropipets to introduce DNA. Although some have advocated electric fieldmediated transfer (Powers et al., 1992; Zhao et al., 1993) or microprojectiles (Zelenin et al., 1991) these have not been rigorously assessed. Transgenic rates vary between 0 and 25% of injected fish done by standard methods (Stuart et al., 1988, 1990; Culp et al., 1991; Lin et al., 1992). Unlike the mouse pronucleus (or that of the teleost medaka), the zebrafish pronucleus is not readily identifiable,

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Mark C. Fishman rt al.

so there may be an inherent limit to transgenic efficiency. Less than 50% of F1 offspring of the injected Go inherit the transgene, presumably because it is incorporated in a mosaic fashion into the germ line, but it is passed on to 50% of the FZ progeny. Both lac2 and chloramphenicol acetyl transferase (CAT) have been used as marker genes for expression and are expressed infrequently (and in some reports, not at all) after germ-line passage. Various promoters regulate cell-specific expression (Westerfield et al., 1992; Reinhard et al., 1994), but there are also likely to be position effects dependent upon the site of chromosomal integration and the strength of the promoter. Stable expression patterns through the Flo generation have been obtained (Bayer and Campos-Ortega, 1992). One reason to improve the efficiency of trangenesis is to make it usable for insertional mutagenesis. For this purpose it will be crucial to develop dominant markers for rapid screening for the rare transgenic among injected animals; otherwise, the aquaculture and genomic analyses required for screening transgenics are foreboding. One such approach is prescreening of animals expressing lac2 using a fluorescent P-galactosidase substrate (FDG; Westerfield et al., 1992) rather than staining of fixed specimens, as is standard, with X-Gal (5-bromo-4chloro-3-indolyl-~-~-galactoside). FDG in DMSO permeates the living fish (with some toxicity, unfortunately) and is cleaved by P-galactosidase, releasing fluorescein. More powerful and less toxic is green fluorescent protein (GFP), originally isolated from the jellyfish Aequorea victoria, which fluoresces on exposure to UV light without the need for exogenous substrate (Chalfi et al., 1994). Mutant GFPs have been generated that are stable and highly fluorescent (ZernickaGoetz et al., 1996). In theory, even more facile screening could be achieved using a dominant pigment marker, or a secreted protein that modifies pigmentation, because this would not require embryonic manipulation for prescreening. Another crucial advance toward efficient DNA integration is the use of viral infection rather than exogenous DNA injection. The first steps in this direction have been taken. To include fish in the viral host range, this virus contains the Moloney murine leukemia virus (MoMLV) genome in a vesicular stomatitis virus (VSV) envelope (Bums et al., 1993; Lin et al., 1994a). Injected into the blastula, this pseudotyped virus can integrate into the germ line (Lin et al., 1994a). To be useable for an insertional mutagenesis screen, the major remaining step is to increase the efficiency of transgenesis. 111. Heart Development A. Morphology

The zebrafish heart has not been studied in detail, but clearly is similar to those of other teleosts. There are four chambers to the fish heart: the sinus venosus, atrium, ventricle, and bulbus arteriosus, from venous to arterial end (Randall, 1968; Santer, 1985). The sinus venosus acts as a reservoir for venous


71

return, and the bulbus arteriosus serves to dampen pulsatility to cardiac output (a “windkessel” function), presumably to protect the gills. The ventricle is thickerwalled than the atrium and is trabeculated. Although not specifically studied in the zebrafish, the fish heart has a coronary circulation, which is of interest because many species (including the zebrafish; MCF, unpublished) evidence focal intimal proliferations of smooth muscle that resemble atheroscleotic lesions. The sinoatrial junction is the site of nodal tissue, which is believed to serve as the cardiac pacemaker, and is heavily innervated, especially by parasympathetic nerves. It is not clear whether there is a specialized conduction system in the hearts of most fish, but there is clearly a coordinated sequential activation of contraction that, along with valves between the chambers, serves to ensure that blood flows from the venous to arterial end without regurgitation. The mature teleost myocardial cell does not appear remarkably different from those of other species at an ultrastructural level (Santer, 1985) and contains myofibrils, sarcoplasmic reticulum, and gap junctions. The last is responsible for electrical coupling between cardiocytes, although the distribution and precise patterning varies between species and has not been studied in the zebrafish. The external surface of the myocardium is covered by the epicardium and the inner surface by the endocardium, a single layer of endothelial cells on a basal laminum. Careful ion current analysis has not been performed on the fish heart, although the action potential is similar to that of other vertebrates. Heart rate increases with catecholamines,high temperature, and activity (Randall, 1968). Contractility obeys Starling’s law, that is, increased filling enhances contractility.

B. Cellular Embryology The general features of zebrafish heart development (Stainier and Fishman, 1992, 1993, 1994) are similar to those classically described in the shad (Senior, 1909). The heart arises from lateral plate mesoderm in apposition to the yolk syncytial layer. Just medial to the lateral plates bilaterally are cords of cells, referred to as the portion moyenne, which appear to contain the precursors to the endocardium. By the 15-somitestage the myocardial progenitors have formed bilateral tubular primordia ventral to the CNS, extending in an arterior-posterior position, with the rostra1 extent reaching to the trigeminal ganglia and the caudal end to the level of the otic vesicle. By the 18-somite stage these cells express tropomyosin. The two tubes move medially into apposition and generate a single midline tube by the 21-somite stage. The portion moyenne sits between the two tubes and then is incorporated within the fusing heart tube and differentiates into the endocardium. The heart tube is actually fist a cone, the base (the future venous end) resting on the yolk and the apex (the future arterial end) continuous with the aorta. Contractions begin by the 22-somite stage, but are irregular until the 26-somite stage. The heart rate increases progressively thereafter as the pacemaker shifts from arterial to venous end, and shortly thereafter (about 24 hr after fertilization) circulation begins. Although the heart appears microscopically

Mark C. Fishman ct al.

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homogeneous at this stage, different myosin heavy chains are expressed by the 26-somite stage in the atrial and ventricular primordia (Stainier and Fishman, 1992). The heart loops to the right between 30 and 36 hr after fertilization. Morphological differences between chambers and cardiac cushions (valvular precursors) are evident by 36 hr after fertilization. The precursors of the heart arise from a discrete region of the blastula. Singlecell injections with fluorescently tagged dextrans are used to track the fates of individual blastomeres and their progeny (Kimmel and Warga, 1986, 1987b; Kimmel et aZ., 1990, Warga and Kimmel, 1990). Tissue restriction begins only around the time of gastrulation. However, there is a predictable cardiogenic zone even in the 512- or 1OOO- celled blastula, as determined by single-cell injection (Stainier et aZ., 1993). This region includes the few tiers of cells adjacent to the margin and extends around the ventral 180"of the embryo. Cell transplantation reveals apparently complete plasticity until commitment begins around midgastrulation. Endocardia1precursors in the 512-celled embryo are more prevalent in the most ventral region of this zone (Lee et aZ., 1994). Myocardial precursors at the 512-cell stage contribute to both atrium and ventricle. The progeny of cells injected at the 1OOO-cell stage are chamber-restricted. C. Genetics

The visibility of the heart and of contractions and circulation make it amenable to genetic analysis, the results of which have recently been published, including a subset that perturb heart function (Chen et aZ., 1996; Stainier et aZ., 1996).

IV. Skeletal Muscle Development A. Morphology The axial muscles are the best-studied skeletal muscle cells in zebrafish, although there have also been a few studies of head muscles (Hatta et aZ., 1990, Schilling and Kimmel, 1994) and fin muscles (Dane and Tucker, 1985). The axial muscles are composed of muscle fibers that arise from the myotomes of the somites. Direct observations of developing embryos show that somites begin to form at the end of epiboly, around 10 hr after fertilization, and new somites are added at a rate of about 2 per hr, at 28.5"C (Hanneman and Westerfield, 1989). A myoseptum forms in the borders between adjacent myotomes and between dorsal and ventral muscles in each myotome. Muscle-cell precursors elongate along the anterio-posterior body axis to form muscle fibers that extend between the borders of the somite. After their initial appearance, the myotomes are block shaped, but within an hour they begin to take on a chevron shape, with the points of the chevronspointed anteriorly. By about 24 hr after fertilization, all of the somites have formed and have acquired the chevron shape. Some time

4. Zebrafish Genetic and Embryological Methods

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later, a horizontal septum forms separating the myotome into dorsal and ventral muscles. By two weeks after hatching, the dorsal and ventral tips of the chevrons fold anteriorly to form the W shape of adult axial muscles (Van Raamsdonk et al., 1974). Muscle cell differentiation is first seen at the medial surface of the somite, which is nearest the notochord (Waterman, 1969; Van Raamsdonk et al., 1978). These first muscle cells may derive from cells in the unsegmented, paraxial mesoderm adjacent to the notochord, termed adaxial cells, that express the snail1 gene (Thisse et aL, 1993). Within an hour of somite formation, a few of the medial cells elongate and express contractile properties (Felsenfeld et aL, 1991). The medial, early-differentiating cells are also the first to express clusters of acetylcholine receptors (Liu and Westerfield, 1992) and acetylcholinesterase (Hanneman and Westerfield, 1989). These first cells to elongate can also be distinguished from the rest of the cells in the myotome by their expression of two of the engrailed homeobox genes (Hatta et al., 1991a; Ekker et al., 1992) and an antigen recognized by the zn5 antibody (Trevarrow et al., 1990). Because of their precocious differentiation, they have been termed muscle pioneers (Felsenfeld et al., 1991). Myogenesis progresses through the myotome from medial to lateral (Waterman, 1969). At 24 hr after fertilization, the majority of muscle cells are mononucleate (Waterman, 1969; Kimmel and Warga, 1987a). The skeletal muscles of fish, like those of most vertebrates, contain multiple types of muscle fibers that have been divided into three major types: red, intermediate, and white muscle, based on their metabolic, morphological, and immunohistochemical properties (for review, see Van Raamsdonk et al., 1982a,b). Unlike those in avian and mammalian muscles, the fiber types in fish axial muscles are largely segregated into homogeneous groups. In adults, the most superficial cells of the myotomes are red, slow-contracting, and fatigue-resistant; the deep cells are fast-contracting, white, and fatigue rapidly; and fibers of the intermediate type are located between the red and white (Van Raamsdonk et al., 1987). The early-differentiating,medial muscle cells express some of the properties of adult red muscle shortly after they elongate. By the end of somitogenesis, only the most medial muscle cells express white-muscle properties, and more lateral cells express red-muscle markers. At hatching, only the lateral cells in the somite express red-muscle markers, while the deep cells, which now make up the majority of fibers, have white-muscle characteristics (Van Raamsdonk et al., 1982b). Adult-type red and intermediate muscle fiber properties first appear at the lateral surface of the myotomes several weeks after hatching (Van Raamsdonk et al., 1982b). B. Cellular Embryology

In zebrafish embryos, the patterns of cell divisions that give rise to muscles can be followed by direct observation in live developing embryos using lineage

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tracer techniques. From this type of analysis, Kimmel and Warga (1986,1987b) showed that tissue-specific cell lineages first arise during gastrulation. A single labeled cell at this stage can give rise exclusively to muscle, whereas single cells at earlier stages typically contribute progeny to more than one tissue. At this stage, the beginning of gastrulation (50% epiboly), the precursors of axial muscle cells are located toward the ventral side of the embryo, near the blastoderm margin, a fate-map position topologically similar to that occupied by cells that give rise to axial muscles in other chordates (Kimmel et al., 1990). During the subsequent few hours of development, as epiboly progresses, the cells that will later give rise to axial muscle converge towards the dorsal side of the embryo where the axis is forming. At the same time, they extend along the axis as the process of epiboly sweeps the blastoderm margin posteriorly over the yolk cell (Warga and Kimmel, 1990). The cells that later give rise to axial muscles in the one day embryo typically have their terminal cell divisions during the late stages of gastrulation at 810 hr (Kimmel and Warga, 1987a). This terminal cell division produces daughter cells that show no particular spatial compartmentation within the muscle lineage; daughters can be found later in different muscle segments, on opposite sides of the embryo, or in different muscles (dorsal or ventral) within the same segment (Kimmel and Warga, 1987a). C. Innervation

Intracellular recording and injection of dyes into spinal-cord cells have revealed aspects of their differentiation and interaction with muscle cells during the time of innervation. The axial muscles are innervated first by primary motoneurons. The growth cones of these motoneurons initially extend from the spinal cord to the region occupied by the muscle pioneers beginning around 16-17 hr after fertilization (Eisen et aZ., 1986; Myers et al., 1986). At this time, clusters of acetylcholine receptors appear on the muscle pioneers (Liu and Westerfield, 1992) and the first localized contractions begin (Hanneman and Westerfield, 1989). Very shortly thereafter, slow side-to-side movements of the embryo can be seen, generated by coordinated contractions of muscle cells on each side of the embryo. Subsequently, the growth cones follow cell-specific pathways extending dorsally and ventrally first along the medial surface of the myotome and then later through the muscle to the lateral surface (Liu and Westerfield, 1990). Most neuromuscular junctions form under side branches that sprout from varicosities along the main trunks of the motor axons beginning after the first day. With few exceptions (Liu and Westerfield, 1990), the initial pattern of innervation is appropriate for the final adult pattern (Westerfield et al., 1986). Moreover, the initial patterning appears independently of electrical activity; it develops normally in nic mutants that lack neuromuscular transmission (Westerfield et aL, 1990) and in embryos paralyzed with neuromuscular blocking agents (Liu and Westerfield, 1990).The growth cones of secondary motoneurons extend into the muscles, beginning several hours after the primary motor growth cones


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(Myers et al., 1986),and follow the pathways established by the primary motoneurons (Pike et al., 1992). Intracellular recordings from adult muscle fibers have shown that each fiber is innervated by a single primary motoneuron and several secondary motoneurons (Westerfield et al., 1986). Recordings from swimming fish demonstrated that primary and secondary motoneurons are sometimes coactivated and that primary motoneurons function primarily, although not exclusively, during fast swimming, struggling, and the startle response, whereas secondary motoneurons function primarily during slower swimming (Liu and Westerfield, 1988). D. Genetics

Several mutations have been isolated that have revealed various features of muscle development in zebrafish. These mutations fall into three broad categories: mutations that affect cell movements and mesoderm patterning during gastrulation, mutations that perturb muscle cell differentiation, and mutations that affect later muscle cell functions. The spadetail mutation blocks the convergence of trunk muscle precursors in a cell-autonomous manner (Ho and Kane. 1990) without severe effects on extension (Kimmel et al., 1989). This suggests that these two morphological movements may be regulated by genetically distinct mechanisms. Trunk somites in spadetail mutants are poorly formed, if at all, and at first are virtually devoid of muscle cells, whereas there is an abnormal accumulation of cells in the tail, including cells that would have given rise to trunk muscles. A mutation in the no tail gene, the zebrafish homologue of the mouse T (Brachyury) gene (Schulte-Merker et al., 1994), acts autonomously in notochord precursor cells to block the differentiation of a mature notochord (Halpern et al., 1993). This mutation leads to a loss of the engrailed-expressing early medial muscle fibers, the muscle pioneers. Rescue of notochord by transplantation of wild-type notochord precursor cells into mutant embryos locally rescues these engrailed-expressing fibers, which can be derived from mutant host cells. This suggests that the notochord may induce engruiled expression in these cells. The myotomes in no tail mutants fail to maintain their chevron shape, and also fail to form a horizontal myoseptum. Thefibers unbundled mutation blocks myofibril formation, perhaps by preventing the initial bundling of actin filaments (Felsenfeld et al., 1990, 1991). Muscle cells elongate normally infibers unbundled mutants, suggesting that actin filament bundling is unnecessary for this to occur. Finally, the nic mutation acts autonomously in muscle cells to block expression of functional acetylcholine receptors (Westerfield et al., 1990; Sepich et al., 1994). This mutation results in muscle paralysis, but has no effect on the initial pattern of innervation.

E. Molecular Biology The zebrafish is exceptionally well suited to molecular approaches in the study of somite and muscle formation, both because of the rapid development, optical

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et

al.

clarity, and ease of manipulation of its embryos, and because of the availability of mutant strains in which mesoderm development and somitogenesis are disturbed. Detailed examination of these embryos using in situ hybridization,immunohistochemistry, and embryo microinjection provides critical information about the molecular determinants of skeletal myogenesis, expanding on the substantial body of knowledge about these factors in a variety of organisms (reviewed in this volume, and Olson and Klein, 1994). Evidence obtained thus far indicates that vertebrate myogenic regulatory mechanisms are largely, but not wholly, conserved in the zebrafish. Here we consider the regulatory molecules that have been described in zebrafish muscle development, taken in the order in which they appear in the embryo. Among the factors known to participate in early patterning of the zebrafish mesoderm, snaill is distinguished by its expression in the forming somites (Hammerschmidt and Nusslein-Volhard, 1993;Thisse et al., 1993).snaill encodes a zinc finger protein homologous to Drosophila snail, which is required for mesoderm formation in the fly (Alberga et al., 1991). Patterned zygotic transcription of fish snaill begins at about 4 hr after fertilization in a ring of cells around the margin of the blastula, just prior to the onset of gastrulation. It persists in the elongating embryonicshield, extending anteriorly in the paraxial hypoblast of the segmental plate that will give rise to the somites. Preceding somitogenesis,the linear array of adaxial cells along either side of the notochord, among which are the presumptive precursors of the muscle pioneer cells (Felsenfeld et al., 1991, and see earlier discussion) expresses snaill. As segmentation occurs, beginning at about 10 hr, snaill transcripts appear just anterior to each new somite furrow and accumulate progressively from medial to lateral, remaining confined to the posterior of each somite. From the 14-somite stage onward, however, snaill expression disappears progressively from the most mature (rostral) somites, again medial to lateral, while it accumulates in the newly forming ones. A new zebrafish snail-family gene, snail2, is also expressed in the adaxial cells along the notochord and persists longer than snaill in the maturing somites (C. Thisse, B. Thisse, and J. H. Postlethwait, personal communication). Thus, the timing and location of snaill expression is consistent with a key role in patterning the somitic mesoderm. snaill expression has also been studied in two mutant zebrafish lines in which somitogenesis is disturbed. In spadetail, paraxial mesoderm fails to converge normally toward the dorsal axis of the embryo during gastrulation, collecting instead in the tailbud (Kimmel et aL, 1989, and see earlier discussion). snaillexpressing cells are still present in this mutant, but they are improperly localized to the tail (Thisse et al., 1993). Activation of snail, therefore, must be independent of the spadetail locus. Furthermore, snaill expression is evidently insufficient for normal somitogenesis. In the nu tail mutant, the zebrafish homologue of the mouse T (Brachyury) gene, differentiated notochord, muscle pioneer cells, and the caudalmost somites are absent (Halpern el al., 1993; Schulte-Merker et d.,1994, and see earlier discussion). Here, snaill expression occurs in its normal distribution but at consid-


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erably reduced levels, indicating that although no tail is not required for snaill activation, it might participate in snail1 maintenance or enhancement (HammerSchmidt and Nusslein-Volhard, 1993; Thisse et al., 1993). Consistent with this hypothesis is the finding that overexpression of no tail injected into pregastrula embryos produces locally enhanced snaill levels (Hammerschmidt and NussleinVolhard, 1993). Animal cap cells can be experimentally induced to express snaill by the protein growth factor activin A (Hammerschmidt and Nusslein-Volhard, 1993); however, the actual inducers of snaill in the fish embryo, whether activin or possibly a zebrafish dorsal-related factor, remain to be elucidated. The requirement for activin in fish mesoderm formation is supported by experiments in another teleost, the Japanese medaka, where injection of dominant negative activin mutants disrupts the axis and mesoderm (Wittbrodt and Rosa, 1994). The temporal and positional expression pattern of snail1 in the presomitic mesoderm is largely recapitulated by a cascade of myogenic regulatory factors that are members of the bHLH and MEF;! gene families. Zebrafish myoD transcripts are first detected at 7 hr after fertilization in two small clusters of cells flanking the embryonic shield (Weinberg et al., 1996). As the embryonic axis elongates anteriorly, this expression extends in a pair of narrow stripes of adaxial cells flanking the notochord, again assumed to include precursors of the muscle pioneer cells. Caudally, these stripes are connected across the midline by a group of cells that encircle the posterior notochord. These myoD-expressing cells in the gastrula appear to be a subset of those expressingsnaill, although this remains to be determined. As with snaill, the onset of somitogenesis is marked by lateral extension of myoD just anterior to each new furrow in a rostral-caudal progression, occupying the posterior region of the somite. But although snaill ultimately disappears, myoD expression persists and extends to the anterior myotome as well, and by 30 hr appears in the fin bud and other nonaxial muscles. The expression of zebrafish myogenin follows essentially the same pattern as that of myoD, but its onset in the adaxial cells is delayed by 3 hr and its lateral extension in the somites by 1hr (R. Riggleman and E. S. Weinberg, personal communication). Zebrafish homologues of the other myogenic bHLH factors Myf5 and MRF4/Myf6 have not been reported. Zebrafish my00 has also been examined in the mesodermal mutants (E. S. Weinberg, personal communication). In spadetail, there is no myoD expression through 14 hr of development; thereafter, transcripts are detected in scattered cells in the poorly organized paraxial mesenchyme. In no tail, the initial appearance of myoD is delayed until 10.5 hr, at which point there is limited expression in an apparently normal pattern in the most rostra1 somites, reminiscent of snaill in this mutant. Thus, it seems that normal expression of myoD also depends, presumably indirectly, on the no tail gene. Three zebrafish MEF2 genes, mef2A, mef2C, and mef2D, have been identified and characterized (Ticho et al., 1996).These are the highly conserved homologues of the previously described mammalian MEF~s,all members of the MADS family of transcription factor and homeotic genes (see Breitbart et al., 1993, and

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references therein). The zebrafish MEF2s are sequence-specific DNA binding proteins and functional transactivators of muscle promoters. mef2D is the first of these genes expressed in the zebrafish embryo, appearing just 0.5 to 1hr after myoD in small cell clusters flanking the embryonic shield. mef2D expression then extends anteriorly in the adaxial cells of the presomitic mesoderm along the notochord, and later extends laterally within each somite upon segmentation. This pattern is indistinguishable from that described for myoD. Similarly,mef2A and mef2Cfollow 2 hr later, approximatelycoincident with the onset of segmentation and the appearance of myogenin. Later, as the development of the axial muscles proceeds, the expression patterns of the three zebrafish MEF2 genes diverge, such that by 36 hr mef2A is present throughout the myotome, mef2C persists only in the ventral myotome, and rnef2D has disappeared altogether. Whether these differences indicate a refinement of muscle gene regulation in distinct muscle groups remains to be determined. It should be noted that the order of appearance of the myogenic bHLH and MEF2 factors in skeletal muscle development in the zebrafish (myoDmef2D~myogenin/mef2A-tmef2C)is decidedly different from that recently reported for the mouse (Myogenin-+Mef2CiMef2~Mef2D-*MyoD; Edmondson et al., 1994), despite the relatively strong phylogenetic sequence conservation among these factors. Although both the myogenic bHLH and MEF2 multigene families must have emerged prior to the divergence of fish and mammals, it appears that the individual family members have fallen under different developmental regulatory programs in the two phylogenetic classes. The possibility warrents investigation that individual factors within each of these gene families might be functionally equivalent, for example, that rnef2D serves the same function in zebrafish as Mef2C serves in the mouse. The molecular regulation of somitic mesoderm development, therefore, is characterized by successive waves of expression of a series of key regulatory factors, each of which largely repeats the pattern established initially by snaill. Taken together, these findings point to a cascade of transcriptional regulatory activity in the skeletal myogenic cell lineage. Not surprisingly, the appearance of differentiated muscle cells in the early zebrafish embryo follows much the same progression. The first of these, the muscle pioneers, arise in the adaxial position adjacent to the notochord at 13 hr and express, in addition to contractile proteins, Engruiled-related homeodomain proteins with as yet unknown functions in muscle (Felsenfeld et d.,1991; Hatta et al., 1991b). Myotomal cell differentiation proceeds as the somites mature in a rostral-caudal progression, and within each myotome in a medial-to-lateral progression (Waterman, 1969), again recapitulating the patterns established by the myogenic factors. Acknowledgments Work in the MCF laboratory was supported by NIH R01 SROlHL49579-03,NIH T32 HL.07208, NIH R01 RR08888,and a sponsored research agreement with Bristol-Myers Squibb. Work in the


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RB laboratory was supported by NIH 5ROlHL48544 and grants from the Council for Tobacco Research and the Charles H. Hood Foundation. Work in the MW laboratory was supported by NIH NS21132 and HD 22486. We thank colleagues for communicating results prior to publication.

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Hatta, K., Schilling, T. F., B r e w e r , R. A., and Kimmel, C. B. (1990). Specification of jaw muscle identity in zebrafish Correlation with engrailed-homeoprotein expression. Science 250,802-805. Hatta, K., Bremiller, R., Westerfield, M., and Kimmel, C. B. (1991a). Diversity of expression of engrailed-like antigens in zebrafish. Development 112,821-832. Hatta, K., Kimmel, C. B., Ho, R. K.,and Walker, C. (1991b).The cyclopsmutation blocks specification of the floor plate of the zebrafish central nervous system. Nature 350,339-341. Ho, R. K., and Kane, D. A. (1990). Cell-autonomous action of zebrafish spt-1 mutation in specific mesodermal precursors. Nature 348,728-730. Kimmel, C.B. (1989). Genetics and early development of zebrafish. TZG 5,283-288. Kimmel, C. B., and Warga, R. M. (1986). Tissue-specific cell lineages originate in the gastrula of the zebrafish. Science 231,365-368. Kimmel, C. B., and Warga, R. M. (1987a). Cell lineages generating axial muscle in the zebrafish embryo. Nature 327,234-237. Kimmel, C. B., and Warga, R. M. (1987b). Indeterminate cell lineage of the zebrafish embryo. Developmental Biology W , 269-280. Kimmel, C.B., Kane, D. A., Walker, C., Warga, R. M., and Rothman, M. B. (1989). A mutation that changes cell movement and cell fate in the zebrafish embryo. Nature 337,358-362. Kimmel, C. B., Warga, R. M., and Schilling, T.F.(1990). Origin and organization of the zebrafish fate map. Development 108,581-594. Lee, R. R. K., Stainier, D. Y. R., Weinstein, B. M., and Fishman, M. C. (1994). Cardiovascular development in the zebrafish. II. Endocardial progenitors are sequenced within the heart field. Development, 1u)(12), 3361-3366. Lin, S., Long, W., Chen, J., and Hopkins, N. (1992). Production of germ-line chimeras in zebrafish by cell transplants from genetically pigmented to albino embryos. Proc. Natl. Acad. Sci. USA SS, 4519-4523. Lin, S., Gaiano, N., Culp, P., Burns, J. C., Friedmann T., Yee J. K.,and Hopkins, N. (1994a). Integration and germ-line transmission of a pseudotyped retroviral vector in zebrafish. Science 265,666-669. Lm,S., Yang, S.,and Hopkins, N. (1994b). IacZ expression in germline transgenic zebrafish can be detected in living embryos. Dev. Biol. 161, 77-83. Liu, D. W., and Westerfield, M. (1988). Function of identified motoneurons and coordination of primary and secondary motor systems during zebrafish swimming. J. Physiol. 403,73-90. Liu, D. W.C., and Westerfield, M. (1990). The formation of terminal fields in the absence of competitiveinteractions among primary motoneurons in the zebrafish. J. Neurosci. l2,3947-3959. Liu,D. W., and Westerfield,M. (1992). Clusteringof muscle acetylcholine receptors requires motoneurons in live embryos but not in cell culture. J. Neurosci. 12,1859-1866. Mullins, M. C., and Nusslein-Volhard, C. (1993). Mutational approaches to studying embryonic pattern formation in the zebrafish. Current Opinion in Genetics and Development 3,648-654. Myers, P. Z., Eisen, J. S.,and Westerfield, M. (1986). Development and axonal outgrowth of identified motoneurons in the zebrafish. The Journal of Neuroscience 6,2278-2289. Olson, E. N., and Klein, W. H. (1994). bHLH factors in muscle development: Dead lines and commitments, what to leave in and what to leave out. Genes Dev 8,l-8. Pike, S. H., Melancon, E. F., and Eisen, J. S. (1992). Pathfinding by zebrafish motoneurons in the absence of normal pioneer axons. Development 114,825-831. Powers, D. A., Hereford, L., and Cole T.(1992). Electroporation: A method for transferring genes into the gametes of zebrafish (Brachydanio rerio), channel catfish (Zctalurus punctatus), and common carp (Cyprinus carpio). Molecular Marine Biotechnology 1,301-308. Randall, D. J. (1968). Functional morphology of the heart in fishes. Am. Zoologist 8,179-189. Reinhard, E., Nedivi, E., Wegner, J., Skene, J. H. P.,and Westerfield, M. (1994). Neural selective activation and temporal regulation of a mammalian GAP43 promoter in zebrafish. Development UO, 1767-1775.


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Santer, R. M. (1985). Morphology and innervation of the fish heart. I n “Advances in Anatomy. Embryology, and Cell Biology,” Vol. 89, pp. 1-102. Berlin: Springer-Verlag. Schilling,T. F., and Kimmel, C. B. (1994). Segment and cell type lineage restrictions duringpharyngeal arch development in the zebrafish embryo. Dev UO, 483-494. Schulte-Merker, S., van Eeden, F. J. M., Halpern, M. E., Kimmel, C. B., and Nusslein-Volhard, C. (1994). no tail (ntl) is the zebrafish homologue of the mouse T (Brachyury) gene. Development 120,1009-1015. Senior, H. D. (1909). The development of the heart in shad. The American Journal of Anatomy IX,212-262. Sepich,D. S., Ho, R. K., and Westerfield, M. (1994). Autonomous expression of the nic 1acetylcholine receptor mutation in zebrafish muscle cells. Dev. Biol. 16l,84-90. Solnica-Krezel,L., Schier,A. F.,and Driever, W. (1994). Efficient recovery of ENU-induced mutations from the zebrafish germline. Genetics l36,1401-1420. Stainier, D. Y.R., and Fishman, M. C. (1992). Patterning the zebrafish heart tube: Acquisition of anteroposterior polarity. Developmental Biology l53,91-101. Stainier, D. Y.R., and Fishman, M. C. (1993). Cardiac morphogenesis in the zebrafish, patterning the heart tube along the anteroposterior axis. I n “Molecular Basis of Morphogenesis,” pp. 79-91. New York Wiley-Liss. Stainier, D. Y. R., and Fishman, M. C. (1994). The zebrafish as a model system to study cardiovascular development. Trends in Cardiovascular Medicine 4,207-212. Stainier, D. Y. R., Lee, R. K., and Fishman, M. C. (1993). Cardiovascular development in the zebrafish I. Myocardial fate map and heart tube formation. Development 119,31-40. Stainier, D. Y.R., Fouquet, B., Chen, J.-N., Warren, K. S., Weinstein, B. M., Meiler, S. E., Mohideen, M.-A. P. K., Neuhauss, S. C. F., Solnica-Krezel,L., Schier, A. F., Zwartkruis, F., Stemple, D. L., Malicki, J., Driever, W., and Fishman, M. C. (1996). Mutations affecting the formation and function of the cardiovascular system in the zebrafish embryo. Development lZ3,285-292. Streisinger et al. (1981) p. 100 Streisinger, G., Walker, C., Dower, N., Knauber, D., and Dower, N. (1981). Production of clones of homozygous diploid zebrafish (Brachydanio rerio). Nature 291,293-296. Stuart, G . W., McMurray, J. V., and Westerfield, M. (1988). Replication, integration and stable germ-line transmission of foreign sequences injected into early zebrafish embryos. Development 103,403-412. Stuart, G. W., Vielkind, J. R., McMurray, J. V., and Westerfield, M. (1990). Stable lines of transgenic zebrafish exhibit reproducible patterns of transgene expression. Development 109,577-584. Thisse, C., Thisse, B., Schilling, T. F., and Postlethwait, J. H. (1993). Structure of the zebrafish snail I gene and its expression in wild-type, spadetail and no tail mutant embryos. Development 119,1203-1215. Ticho, B. S., Stainier, D. Y. R., Fishman, M. C., and Breitbart, R. E. (1996). Three zebrafish MEF2 genes delineate somitic and cardiac muscle development in wild-type and mutant embryos. Mech. Dev. 59,205-218. Trevarrow, B., Marks, D. L., and Kimmel, C. B. (1990). Organization of hindbrain segments in the zebrafish embryo. Neuron 4,669-679. Van Raamsdonk, W., Van der Stelt, A., Diegenbach, P. C., Van der Berg, W., DeBruyn, H., Van Dijk, J., and Mijzen, P. (1974). Differentiation of the musculature of the teleost Brachydanio rerio. I. Myotome shape and movements in the embryo. Z. Anat. Enfwickl. Gesch. 145,321-342. Van Raamsdonk, W., Pool, C. W., and te Kronnie, G. (1978). Differentiation of the muscle fiber types in the teleost Brachydanio rerio. Anat. Embryol. 153,137-155. Van Raamsdonk, W., Van Veer, L., Veeken, K., Heyting, C., and Pool, C. W.(1982a). Differentiation of muscle fiber types in the teleost, Brachydanio rerio, the zebrafish. Anat. Embryol. 164,51-62. Van Raamsdonk, W., Van Veer, L., Veeken, K., te Kronnie, G., and de Jager, S. (1982b). Fiber type differentiation in fish. Mol. Physiol. 2,31-47.

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Van Raamsdonk, W., Smit-Onel, M., Scholten, G., Hemrika, W., and Robbe, B. (1987). Metabolic specialization of spinal neurons and the myotomal muscle in post-hatching stages of the zebrafish, Brachydanio rerio. A histochemical study. 2.Mikrosk-Anat. Forsch. 101,318-330. Warga, R. M., and Kimmel, C. B. (1990).Cell movements during epiboly and gastrulation in zebrafish. Development 108,569-580. Waterman, R. E. (1969).Development of the lateral musculature in the teleost, Brachydanio rerio: A fine structural study. Am. J. Anat. 125,457-493. Weinberg, E. S.(1992).3 analysisof early development in the zebrafish embryo. Results and Problems of Differentiation 18,91-150. Weinberg, E. S.,Allende, M. L., Kelly, C. S.,Abdelhamid, A., Murakami, T.,Andermann, P., Doerre, 0. G., Grunwald, D. J., and Riggleman, B. (1996).Developmental regulation of zebrafish MyoD in wild-type, no tail and spadetail embryos. Development l22(1), 271-280. Westerfield, M. (1995).“The Zebrafish Book.” Eugene, Oregon: University of Oregon Press. Westerfield,M., McMurray,J. V.,and Eisen, J. S.(1986).Identilied motoneurons and their innervation of axial muscles in the zebrafish. The Journal of Neuroscience 6,2267-2277. Westerfield, M.,Liu, D. W., Kimmel, C. B., and Walker, C. (1990).Pathfindingand synapse formation in a zebrafish mutant lacking functional acetylcholine receptors. Neuron 4,867-874. Westerfield, M., Wegner, J., Jegalian, B. G., DeRobertis, E. M., and Puschel, A. W. (1992).Specific activation of mammalian Hox promoters in mosaic transgenic zebrafish. Genes Dev. 6,591-598. Wittbrodt, J., and Rosa, F. M. (1994).Disruption of mesoderm and axis formation in fish by ectopic expression of activin variants: The role of maternal activin. Genes Dev. 8,1448-1462. Zelenin, A. V., Alimov, A. A., Barmintzev, V. A,, Beniumov, A. O., Zelenina, I. A., Krasnov, A. M., and Kolesnikov, V. A. (1991).The delivery of foreign genes into fertilized fish eggs using high-velocity microprojectiles. FEES Letters 287,118-120. Zernicka-Goetz, M., Pines, J., Ryan, K.,Siemering, K.R., Haseloff, J., Evans, M. J., and Gurdon, J. B. (19%). An indelible lineage marker for Xenopus using a mutated green fluorescent protein. Development 122,3719-3724. Zhao, X., Zhang, P. J., and Wong, T. K. (1993).Application of baekonization: A new approach to produce transgenic fish. Molecular Marine Biology and Biotechnology 2,63-69.

PART I1

Myogenesis in Cell Culture


CHAPTER 5

Skeletal Muscle Cultures Craig Neville,* Nadia Rosenthal,' Michael McGrew,* Natalia Bogdanova,* and Stephen Hauschkat Cardiovascular Research Center Massachusetts General Hospital-East Charlestown, Massachusetts 02129 t Department of Biochemistry

University of Washington Seattle, Washington 98195

I. Introduction 11. Muscle Cell Cultures A. Primary Muscle Cell Cultures

B. Established Muscle Cell Lines 111. Summary and Future Prospects

References

I. Introduction In the past decade, the field of vertebrate skeletal myogenesis has been revolutionized by the discovery that a muscle phenotype can be induced in cultured nonmuscle cells by the transfection of specific regulatory genes. This breakthrough, and the ease which differentiation can be induced and visualized in myogenic cell cultures, has allowed a molecular dissection of the differentiation process and the characterization of multiple, muscle-specific DNA control elements associated with differentiation-specific gene expression. From an experimental standpoint, cell transfection has been central in establishing muscle culture as a model system for studying the transcriptional regulation of differentiation. Skeletal muscle has proven particularly amenable to molecular analysis because primary myoblast cultures can be prepared fairly easily, and they spontaneously differentiate as observed by the fusing of mononucleate myoblasts into multinucleate syncytia. These morphological changes are accompanied by the METHODS IN CELL BIOLOGY, VOL. 52 Copynghr Q 199R by Arademc Prcs. All nghu of reproduction tn any form reserved. 0091-679X/9H $25 00

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rapid activation of a battery of genes, including those that encode the components of the muscle contractile apparatus. Well-characterized stable cell lines have been established that retain various aspects of the muscle phenotype seen in primary cells. These unique features of cultured muscle cells have been exploited to establish important paradigms in myogenesis, and have also facilitated the identification of a family of transcription factors whose expression in nonmuscle cells induces the myogenic phenotype. In transgenic mouse studies involving gene knockouts, several of these factors have been shown to be critical components of the genetic pathways leading to muscle determination and differentiation in the embryo. The myogenic determination factors (MDFs), a small family of related transcription factors, are expressed exclusively in skeletal muscle lineages. The four MDFs (MyoD, myogenin, myf5, and MRF4) were first characterized as potent regulators of myogenesis by forced expression in transfected nonmuscle cell cultures. Transfected cells were converted into myoblasts, which could then be differentiated into multinucleate myotubes by mitogen withdrawal (Davis et al., 1987; Edmondson and Olson, 1989; Wright et al., 1989; Rhodes and Konieczny, 1989; Braun et al., 1989b,1990,Miner and Wold, 1990). Forces expression of MDFs in nonmuscle cell culture also induces transcription of endogenous MDF genes (Braun et al., 1989a;Thayer et al., 1989), suggesting that continuous, autoregulatory activation of MDF gene expression may be an important component of the commitment to a myogenic phenotype. With the characterization of the MDFs, the means to control many steps along the pathway from an uncommitted mesodermal cell to a fully differentiated myotube in cell culture are now in hand. This chapter will review the preparation of primary muscle cell cultures, and the characteristics of different established muscle cell lines and the special culture conditions and handling they require. In the accompanying chapter, various approaches for the analysis of gene function in myogenic cultures will be described, with detailed protocols for muscle cell transfection and reporter gene assays.

11. Muscle Cell Cultures In mature muscle tissue, terminally differentiated myocytes form multinucleate syncytia in which structural genes are expressed and the contractile apparatus is assembled. Lying within the same basal lamina, undifferentiated mononucleate myoblasts, called satellite cells, form a reservoir of stem cells for regeneration and repair (Fig. 1). These cells can be induced to proliferate by muscle injury and make up an enriched source of myoblasts that can be extracted from the muscle tissue and propagated in culture. Numerous muscle cell lines have been established from these stem cell populations, all of which maintain the ability to undergo myogenic differentiation in serum-deficient media (Fig. 2). When

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Fig. 1 Intracellular structure of a differentiated muscle cell. The majority of the cytoplasm is Wed with myofibrils, comprising the contractile apparatus. Myonuclei are located in the periphery under the cell membrane sarcolemma. Satellite cells are situated between the sarcolemma and the outer membrane (basal lamina) that surrounds the cell. (Drawing courtesy of B. Carlson.)

differentiated these cell lines recapitulate to a greater or lesser degree the phenotype of the intact muscle tissue from which they were derived. A. Primary Muscle Cell Cultures

Primary muscle cell cultures are usually prepared from fetal or neonatal muscles, since large quantities of proliferative myoblasts are difficult to isolate from

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B

C

Fig. 2 Myogenic differentiation of the C#21zmuscle cell line. (A) subconfluent proliferative myoblasts. (B) myoblasts soon after serum withdrawal. Note the elongated morphology of some cells. (C) fully differentiated myotubes after several days in low serum conditions.

muscle tissue at later stages. Primary myoblasts require special conditions for optimal growth and differentiation. First, the initial isolate invariably contains a mixture of other cell types, mainly fibroblasts, which may overgrow the culture if they are not removed. Myoblast enrichment protocols often take advantage of the fact that myoblasts adhere to plastic much less avidly than do fibroblasts, and therefore fibroblasts can be removed from the culture by preabsorption on plastic tissue-culture plates. Second, the very characteristics that allow myoblast enrichment require that the plates they ultimately are grown on be coated with gelatin or collagen to facilitate their adherence. Once these steps are taken, primary myoblasts are relatively easy to propagate, and they spontaneously recapitulate the myogenic differentiation process upon reaching confluence or after sufficient time in culture.

1. Mouse Primary Cultures The procedure presented here is based largely on the Powell laboratory protocol (Powell and Fambrough, 1973). A modification of procedures for obtaining large numbers of primary mouse myoblasts that can be used for cell transplantation and cell-mediated gene therapy purposes has been described by Rando and Blau (1994).


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Muscle tissue from the hindlimbs of one neonatal mouse pup will yield sufficient cells to be plated on one to two 35-mm tissue culture dishes. More care is needed in preparation of mouse muscle cultures in comparison to that of rat or chick because of mouse muscle cultures’ increased sensitivity to media conditions and cell density. Mouse primary cultures grow best with chick embryo extract (CEE) included in the plating media. This should be prepared by the accompanying protocol. If isolated mouse cells are initially plated at too low a density, the cultures will never produce a healthy and prolific population. Furthermore, mouse myoblasts also experience a longer lag period than rat muscle cultures before attaching to the tissue culture plate and extending processes. For this reason it is important not to change the medium for the first two days to allow the cells time to adhere to the substrate. Afterwards, the medium should be changed every two days. Mouse myoblasts cultures are not normally passaged and are usually transfected on day five of culture. This should always be followed by a glycerol shock on the following morning before switching to differentiation media. Cultures are normally harvested after two or three days in differentiation media. Addition of 0.3 mg/100 ml cytosine /3-D-arabinofuranoside (AraC; Sigma #6645) to the differentiation media will select against proliferating cell types (i.e., fibroblasts) and generate pure myotube cultures. This should be added after the first day in differentiation media and left on for 24-hr. Sterile techniques should be utilized at all times in this protocol. a. Dissections

1. Rat and mouse dissections are performed in a similar manner. Mouse pups are euthanized under C02, 2 min for neonatal day 1 mouse pups and 4 min for rat pups. Pups are placed in a beaker containing 95% ethanol for 2 min, then transferred to a second beaker of 95% ethanol for an additional 3 min. Subsequently, pups are placed in a sterilized beaker containing Hanks balanced salt solution (Cat, Mg2+free) (HBSS) on ice for a minimum of 10 min to remove ethanol and wash tissues. Pups are removed one at a time for the following dissections. Dissecting tools should be laid on a paper towel and constantly rinsed with 95% ethanol. Allow instruments to air dry before touching tissue. 2. Transfer pups to a 100-mm Petri dish and remove head and hind limbs. Be sure to cut sagittally up the side of the animal to include the upper thigh with the dissection. Collect legs in a Petri dish containing HBSS on ice. 3. Transfer hindlimbs to a clean dry Petri dish and deskin and depaw. Start at the top of the leg and pull skin down over the paw. Any visible fat deposits must be removed at this time. Also be sure not to remove the looser muscle tissue, which will tend to separate with the dermal skin layer instead of sticking to the underlying muscle tissue. Place skinned limbs in a Petri dish containing HBSS on ice. 4. Transfer limbs individually to a dry Petri dish. Remove bones by using two tweezers to pull muscle tissue away from bone. Care must be used with mouse

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cultures to ensure that muscle tissue is not left behind. Place the isolated muscle tissue in a new Petri dish containing HBSS on ice.

b. Isolation of Myoblust All subsequent manipulations are done in a tissue culture hood. The protocol described below is for 10 newborn mouse or rat pups. 1. Remove HBSS with Pasteur pipet. Mince tissue with small surgical scissors (curved iris variety) for three min. Add a few milliliters of prewarmed enzyme solution to dish and transfer minced tissue to enzyme tube. Triturate several times and incubate at 37°C for 7 min. 2. Triturate again about 20 times. When tissue begins to pass freely through the pipet tip, flame-polish a Pasteur pipet to 4 of the original diameter and triturate 20 more times. Return tube to water bath and incubate 6 more min. 3. Triturate solution again for about 20 times. Flame polish tip to 4 diameter and triturate 20 times. The tissue should now be dispersed with no visible clumps. 4. Add an equal volume of plating media (7 ml) and pipet into a 100-pm nylon mesh cell strainer (Falcon #2360), using fresh strainers as necessary. Collect flow-through in a 50-ml conical tube. 5. Centrifuge in a tabletop centrifuge at 650g for 5 min. Aspirate supernatant and resuspend pellet gently in 10 ml plating media. 6. Pour solution into a 100-mm Petri dish and preplate in incubator for 1 hr to remove fibroblasts. 7. Carefully remove solution from Petri dish by tilting it to one side. This solution contains the enriched myoblast population and can be plated in either Falcon Primaria tissue culture dishes or tissue culture dishes coated with 1% gelatin, bloom ,175 (Sigma #2625) (see also Section 1I.B.l.a pertaining to MM14 cultures). Mouse plating medium 20 ml fetal calf serum 2 ml200 mM glutamine 5.0 units/ml penicillin-streptomycin (PS) 2mlCEE 76 ml Dulbecco’s modified Eagle’s medium (DMEM) Enzyme solution 3.5 ml 0.25%trypsin 1.4 ml 2.5%pancreath 2.1 ml HBSS

Mouse differentiation medium 10 ml horse serum 2 ml 200 mM glutamine 5.0 unitdm1 PS 88 ml DMEM

c. Preparation of Chick Embryo Extract

Addition of CEE supplies mitogens, which results in a significant increase in the number and the size of the myotubes formed. CEE is commerciallyavailable; however, its quality is highly variable, and thus the following protocol is supplied. It is carried out using sterile techniques even though the final product is filter sterilized by passage through a 0.45-pm filter.

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1. Clean day 9-10 chicken eggs by wiping with ethanol. Harvest the embryos into a large Petri dish. Sacrifice the embryos and place in a 10-ml syringe. 2. Pass embryos twice through the 10-ml syringe. Collect the processed embryos in a 50-ml conical centrifuge tube. 3. Add an equal volume of HBSS. Triturate solution with a wide-bore 25-ml pipet. Gradually reduce pipet size until solution is able to be drawn into a 5-ml pipet. 4. Freeze solution overnight at -20". 5. Thaw solution and continue trituration until solution can be passed through a Pasteur pipet. 6. Centrifuge solution in a tabletop centrifuge at l500g for 30 min. 7. Remove supernatant and freeze this solution in 2-ml aliquots. When preparing plating medium add CEE to DMEM and passage through 0.45 pm filter before adding additional ingredients. An alternative CEE protocol that works well for rodent, avian, and human muscle cultures has been devised by Konigsberg (Konigsberg, 1968; Linkhart et al., 1981; Seed and Hauschka, 1988; Hauschka, 1974a).

2. Derivation of Permanent Clonal Lines of Mouse Myoblasts The availability of numerous mouse strains that lack or overexpress regulatory and structural genes, or that serve as models for human neuromuscular disease, provides the opportunity for studying these phenomena in vitro via the use of permanent myogenic cell lines derived from the mice. Clonal lines of skeletal muscle myoblasts can be readily derived from primary mouse myogenic cultures by taking advantage of the natural propensity of mouse cells for spontaneous transformation (Hauschka et al., 1979; Linkhart et al., 1981). Although it is possible that the derivation of muscle cell lines is facilitated by using muscle from mouse strains that also carry transgenic oncogens such as the SV40 T antigen (Bradley et al., 1993), this practice is not required. Permanent mouse muscle cell lines can be derived from any developmental stage and anatomical muscle from which one can obtain healthy primary cultures. The procedure described next has been used routinely by the Hauschka lab for obtaining permanent satellite muscle cell lines from adult mouse muscles, but it has also worked well with fetal muscle as young as E16. Attempts to derive lines from earlier mouse stages were unsuccessful, but this may have been due to culture problems as trivial as a suboptimal serum lot. Depending on the use to be made of the muscle cell lines, one has the option of deriving them from original primary clones, from pooled primary clones, or from clones obtained from mass cultures that have been taken through several passages. The basic strategy in all of these cases is similar: namely, to expand a pure population of myoblasts until most cells undergo proliferative senescence, while a few rare spontaneous transformants continue to proliferate.

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A typical derivation protocol starting from single primary muscle clones is as follows:

1. Media, passaging, and freezing procedures are as described later for the handling of MM14 cells. Several modifications that seem beneficial, although not absolute requirements, are increasing the basic fibroblast growth factor (bFGF) levels to 6 ng/ml and supplementing the media with 1%chick embryo extract. Both of these changes increase the yield of primary myogenic clones. The use of higher bFGF levels also permits primary clones to be grown higher cell numbers without loss of cells to spontaneous differentiation. It is also beneficial to use Ham’s FlOC as the nutrient medium rather than the many other nutrient media that work well with established myogenic cell lines. The advantage of FlOC is that it provides a slight selective advantage of myoblast growth over that of fibroblasts; this is particularly useful when the experimental plan is to grow mass cultures for several passages prior to the cloning steps. Finally, although mouse primary muscle cells can be grown in media containing fetal calf serum (FCS) rather than horse serum, it may be wise not to use FCS since it facilitates the growth of fibroblasts. It is also possible that the use of FCS rather than the HS plus FGF combination will potentiate the derivation of FGF-independent myogenic cell lines rather than the FDF-dependent lines typically derived via this protocol. 2. Primary mouse muscle cultures should be initiated as described earlier, or via virtually any alternative technique that a lab has found successful. Following cell dissociationand an optional preplating step to reduce fibroblasts,the enriched myoblast population can be propagated as either mass cultures or clones. The potential advantages of mass culture growth are that it provides cells with a culture adjustment period that is not as immediately stringent as the necessity of clonal growth, and it permits an accurate counting of secondary cells following dissociation of the mass cultures and their replating at clonal densities. The potential disadvantages of an initial mass culture growth period are that subsequent clonal derivatives could have originated from the same single primary cell. The potential advantage of immediate cloning is that the progeny from each clone are known to be derived from different primary cells, and that presenescent cells go through only a few of the limited number of doublings prior to the cloning steps. The potential disadvantage of immediate cloning is due to inaccuracies in cell counts, to the presence of undissociated small cell clusters (causing the risk that supposed clonal populations did not originate from single cells), and to wide variations in clonal plating efficiency. Cell counting inaccuracy, especially in dissociated adult muscle, is due to large numbers of small muscle fiber fragments and blood cells that are difficult to distinguish from muscle cells. Undissociated cell clumps and muscle fiber debris can be partially removed by passing dissociated cells through gauze and then silk or Nitex filters, using a h a 1 exclusion size of about 20 microns, but this still permits occasional small cell clusters to pass through. Plating efficiency variability is best handled by establishing cultures at


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many different cell densities (e.g., 30,100,300,1000,3000,10,000,30,000,100,000, and 300,000cells inoculated per 100-mm dish). This procedure assures one of obtaining at least several dishes with a sufficient number of widely separated clones to permit the clean isolation of single colonies; furthermore, the highest density dishes provide the option of replating an in vim-adjusted population of secondary cells at clonal densities, and of freezing the excess cells as a backup if it is necessary to isolate new sets of clones. As a rough prediction of clonal plating efficiencies based upon cell counts obtained with a hemocytometer, it is reasonable to assume at least 1-10% for primary cultures and 5 2 0 % for secondary cultures. 3. When clonal cultures have been initiated (whether from primary or secondary cells) it is best to minimize the frequency with which the cultures are disturbed, since this decreases the probability that cells will detach from one clone and either reattach in the vicinity of another clone or initiate an identical sister clone. Unfortunately, the maintenance of undisturbed cultures is counteracted by the necessity of feeding cultures sufficiently often to prevent spontaneous differentiation. It is best to check the highest-density culture dishes (if primary cultures were established at many different cell densities, as described earlier) within 18-24 hr of plating. All cultures whose densities are greater than seems useable for subsequent clonal isolation (typically,any cultures that appear to have greater than 16viable cells per 100-mm dish at 24 hr) should then be fed according to the schedules described later for MM14 cells, with complete medium changes at least every 24 hr, and FGF additions every 12 hr. If CEE is being used in addition to FGF, an equivalent of 1%CEE, as well as 6 n g / d bFGF, should be added at each feeding. After 3-5 days growth, depending on population density and whether cell differentiation (fusion) has started, these higher-density cultures should then be passaged and used for establishing secondary clonal cultures and/ or frozen for subsequent experiments. Lower-density cultures should be left undisturbed and should not be fed until about 72 hr. At this point they should receive a complete medium change and should be started on a feeding schedule in which 2 ng/ml bFGF is added approximately every 12 hr (if cultures are also being supplemented with CEE, an equivalent of 1% should be added every 24 hr); these additions should be accompanied by complete medium changes at least every 48 hr. Supplementation with CEE can be discontinued after 4-5 days. Within 5-7 days it should be possible to detect macroscopic muscle colonies. This can be done either laboriously under an inverted phase microscope, followed by marking the clone’s location with a commercially available objective marking device, or more rapidly by naked eye. This is done by leaning back in a chair, holding the dish above one’s head, and gradually rotating and tilting the dish such that the medium covers only half the surface and such that the overhead lights reflect off the moist plate surface in a manner that accentuates individual colonies-all the while being careful not to pour the medium onto one’s face. With practice and starting out with dishes at the highest clonal densities after

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at least 7 days growth, clones will be discernible as faintly refractory dots (muscle clones are whitish because of the rounded cell morphology, and fibroblast clones will be grayer because of the flattened cell morphology). Clones are circled with a pen, after which they can be examined individually under phase optics. When cultures are handled according to the procedure just described, it is not unusual to observe a few fused cells within muscle clones within about a week of initiating the cultures. As long as the fraction of fused cells is low, this is actually beneficial since it proves that a clone is myogenic. Even when fusion has started, it is usually possible to maintain sufficient numbers of proliferating cells to attain clones containing several hundred single cells by continuing to feed the cultures on the 12-hr schedule described earlier. Although primary clones can be passaged after attaining any desired cell number, clones containing several hundred cells have several advantages: (i) certainty that the cells grow well, (ii) greater assurance that a reasonable number of secondary cells will be obtained (i.e., secondary clonal plating efficiency may be only 10-30%), and (iii) sufficient numbers of cells that multiple secondary cultures can be set up from a single subclone; these can be used for ascertaining the subclone’s cellular purity or any other attributes one wishes to screen for prior to carrying out the subsequent work of creating a permanent cell line from the particular clone. If the experimental plan does not require that the permanent myogenic cell lines each originate from a single muscle colony, then individual clones can be combined and replated as a set of pooled muscle clones, This has the advantage of increasing the number of cells and simplifies the logistics of handling many individual clonal sublines. Pooled clonal cultures obviously should not be established if the only muscle clones available for passage seem to be potentially contaminated by fibroblasts. In this case, the primary clones should be passaged, replated at clonal densities, and reisolated as clean secondary muscle clones. 4. Propagation of individual colonies is done as follows. Cloning cylinders and silicon grease are sterilized by autoclaving in glass Petri dishes (the silicon grease can be reused many times). Trypsin, growth media, and plates to receive the passaged cells are set up as described below. A sterile cloning cylinder is placed onto the grease to coat its bottom surface and twisted back and forth using sterile forceps. The cylinder is then placed, bottom down, over a premarked colony and tapped lightly on the top surface to seal the cylinder to the dish. It is preferable, especially if many clones are to be picked from the same culture dish, not to remove the growth medium from the dish. This keeps the remaining clones happier while cloning cylinders are being positioned; usually, the hydrophobic nature of the grease forces the medium out of the region directly above the clone as the cylinder is being positioned. If this occurs and if many clones are being picked from the same dish, it is preferable to add the trypsin solution immediately after placing the cloning cylinder so that the cells remain moist. Cylinders should be filled to about two-thirds of their volume with trypsin. The


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culture dish is then placed on a 37°C warming bar for about 5 min, or until inspection by phase microscopy indicates that the trypsinized cells have rounded up. Each clone is then pipetted up and down 10-20 times with a separate Pipetman, and the dissociated cells are transferred to individual tubes containing several milliliters of chilled serum-containing medium. These are centrifuged and aspirated, the invisible cell pellet is resuspended in chilled medium to the desired cell concentration (based on a previous estimate of cell number in the clone), and the cells are inoculated into preequilibrated dishes. 5. The goal from this point onward is to maintain the clonal cultures under continuous log-phase conditions until they have undergone 30-50 population doublings. After this range of doublings, most of the cells will become proliferatively senescent, while a few rare cells undergo spontaneous transformation and continue to proliferate. Since it is difficult to estimate when this will occur, it is best to expand each clonal or pooled population to about 3 X 105 cells when it is estimated that the total in v i m doublings have reached about 30. At each subsequent passage, dishes should be inoculated at a range of cell numbers (105, 104, I d , 102) and monitored for continuous exponential growth. If rapid growth persists, the higher-density cultures should be passaged and replated as above each time they complete another 4-6 doublings. The excess cells should be frozen to ensure that the line will not be lost. When proliferative senescence and spontaneous transformation occurs, dishes inoculated at 105cells will exhibit very slow growth and a few isolated colonies will continue rapid proliferation. These colonies can be cloned as described earlier. Even though such clones will be surrounded by slow-growing senescent cells, these will rapidly disappear as each transformed clone is expanded. As soon as it becomes clear that the putatively transformed clonal population is maintaining rapid growth, while other cells are senescing, the transformed cells should be expanded to high cell numbers, tested for the desired traits, and frozen for future studies. Experience with deriving hundreds of such clonal lines indicates that at least 25-50% of the original primary clones can give rise to permanent lines as long as each primary clone exhibits rapid growth at the beginning of the selection process. One can therefore be virtually assured of successfully deriving a permanent myoblast line by picking 6-8 original clones. An occasional clonal population will be encountered in which senescence does not appear to occur, even after 50-60 population doublings. This probably occurs because one or more cells within the population became converted to the transformed phenotype during an earlier passage, and gradually replaced cells that were senescing.

3. Rat Primary Muscle Cultures This procedure is from the laboratory of Steven Burden with adaptations made according to the mouse protocol in Section 1I.A. Typically, one rat hindlimb will yield 6-8 X 105cells, which is sufficient to be plated in one 100-mm tissue-culture

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dish. Many of the plated cells will not proliferate. Rat myoblasts are refed on day 2 in culture and are usually transfected on day 3. These cells may be passaged one or two times, but caution must be taken because each manipulation will increase the number of fibroblasts present with a concomitant loss of myoblasts. Dissections and isolation procedures are camed out as described in Section I1.A. for mouse primary cultures, with the following modifications:

1. Two tubes of 7 ml of enzyme solution each are used per litter of about 12 rat pups. Enzymatic incubations are camed out for three periods of 16 min. On the final trituration a Pasteur pipet flame-polished to 4 of the original diameter is used. 2. Isolated cells are preplated 15 min on uncoated Petri dishes to reduce the number of contaminating fibroblasts. 3. Rat myoblasts can be grown on normal tissue-culture plastic and should be fed every 2 days. Rat plating medium 20 ml fetal calf serum 2 ml200 mM glutamine 12.5 unitdm1 PS 78 ml DMEM

Rat differentiation medium 4 ml horse serum 2 ml200 mM glutamine 12.5 unitslml PS 94 ml DMEM

4. Chick and Quail Primary Muscle Cell Cultures Chick and quail embryonic muscle cells are perhaps among the easiest of all primary cells to prepare and culture. Avian embryos are large and give reproducibly high yields of myogenic precursor cells with relatively little experimental variation. They are quite amenable to standard transfection techniques, and introduced mammalian muscle-specific genes appear to be regulated appropriately. Cultured primary avian myotubes are considered to be the best system to mimic muscle fibers in vivo as they undergo considerable spontaneous contraction. References to alternative protocols and specialized chick and quail embryo muscle procedures are included at the end of the protocol. I 1. Incubate chick or quail eggs at 38.5"C in a humid atmosphere. The eggs should be rotated 180" daily. 2. Wash the exterior surface of an 11-day chick egg or 9-day quail egg and then sterilize with 70% ethanol. Stand egg in a small beaker, pointed side down. Sterilize all instruments immediately prior to use by dipping in ethanol, flaming and cooling off in a 50-ml tube of sterile phosphate-buffered saline (PBS). 3. Cut a window at the blunt end of the egg with a pair of sterile scissors. Enlarge window to the edge of the air sac to expose the chorioallantoicmembrane with blood vessels. Pierce the membrane with sterile sissors or forceps.

a. Protocol


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4. Remove the embryo by the neck with a pair of large sterile forceps and transfer it to a Petri dish. As the embryo is considered sterile, do not rinse it with ethanol. 5. Sacrifice and dissect away the skin overlying the breast. Remove the muscle and place it into another Petri dish, along with a few milliliters of PBS. Mince the muscle well with a pair of very fine dissecting scissors. 6. Replace the PBS with growth medium (1 ml/embryo) and transfer the muscle to 50-ml Falcon conical centrifuge tubes (4 embryodtube). 7. Vortex vigorously for 30 sec. As avian embryos are much softer than those of rodents, there is no need to resort to enzymatic disaggregation of the tissue. 8. After the large fragments settle, transfer the supernatant to a new tube. Add 4 ml medium to the original pellet and vortex again for another 30 sec. 9. Combine the two supernatants and filter through three layers of Nitex mesh or 100 pm disposable Falcon cell strainers (#2360). 10. If it is important that the myoblasts be as free of contaminating cells as possible, the suspension can be preplated briefly (30 to 60 min) on uncoated tissue-culture dishes. Fibroblasts will adhere while most myoblasts remain in suspension. The myoblasts are plated as per step 11. This step is usually not required and will necessarily result in the loss of some myoblasts. 11. Plate cells at a density of 104-106 cells/35-mm plate. 12. The following day, aspirate off medium, rinse with PBS to remove contaminating cellular debris, and add fresh medium. Avian growth medium

500 ml Eagle’s MEM with Earle’s salts 100 ml fetal calf serum 25 ml chick embryo extract (50 ml for quail muscle cells) 5 ml penicillidstreptomycin solution (5000 IU/ml and 5000 pg/rnl, respectively) 1.25 ml fungizone (250 pg/ml amphotericin B) Avian differentiation medium 500 ml Eagle’s MEM with Earle’s salts

10 ml fetal calf serum 5 ml penicillidstreptomycin solution (5000 IU/ml and 5000 pg/ml, respectively) 1.25 ml fungizone (250 p g h l amphotericin B)

b. Protocol 2 Alternative protocols for propagating high-density and clonal cultures of chick embryo myoblasts can be found in Konigsberg (1963), Hauschka and Konigsberg (1966), Bonner and Hauschka (1974), White et al. (1975), Rutz (1982), Seed and Hauschka (1981,1984), and Miller and Stockdale (1986a,b). The procedure utilized by the Stockdale lab (Feldman and Stockdale, 1992) for establishing fetal myoblast clones from E10-E20 chicken and E8-El7 quail embryos is as follows:

1. Remove and dissect specific muscles as described earlier. Store muscles in iced HBSS prior to step 2.

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2. Transfer muscle tissue to a separate Petri dish and mince the tissue into a fine slurry with iris scissors. 3. Add about 10 ml 37°C 0.2% trypsin in HBSS per 1 ml of muscle slurry and transfer to a plastic test tube. 4. Incubate at 37°C for 20 min with occasional gentle pipettings to disperse cells. 5. Add an equal volume of chilled HBSS containing 10% horse serum (HS). 6. Centrifuge 5 min at 500g. 7. Aspirate medium, and rinse and pellet the cells two additional times using 10 ml chilled growth medium (77% Ham’sF10, 15% HS,5% chicken embryo extract, 1% 0.132 mg/ml CaC12, 1%glutamine (lOOX, GIBCO #12420-014), and 1% antibiotic mix (penicillin, streptomycin, Fungizone). 8. Inoculate the cell suspension into 100-mm uncoated tissue-culture dishes containing 10 ml preequilibrated growth medium. 9. Incubate 30 min at 37°C to allow selective attachment of fibroblasts. 10. Remove the unattached cells, count, and for clonal cultures plate 250 cells into gelatin-coated 60-mm dishes containing 4 ml preequilibrated medium (1:1 growth medium and conditioned growth medium; see later section). c. PIOtOCOl 3

The procedure used by the Stockdale lab (Feldman and Stockdale, 1991,1992) for establishing avian satellite cell cultures from newly hatched to adult birds was modified from that of Yablonka-Reuveni et al. (1987). The technique is as follows: 1. Sacrifice birds by C02 asphyxiation. 2. Dissect muscles aseptically and store muscle in iced HBSS until ready for

step 3. 3. Add about 10 ml 0.2% collagenase (Worthington, CLSIII) in HBSS per 1ml volume of muscle. 4. Digest for 10-45 min at 37°C (about 10 min for 1-3 day birds and as long as 45 min for birds older than 40 days) with gentle agitation. 5. Triturate 10 times with a 10-ml serological pipet, add an equal volume of iced HBSS,centrifuge 5 min at SOOg, and discard the supernatant. 6. Resuspend the pellet in 0.1% trypsin (Difco) in HBSS, using about 10 ml per 1 ml of cell pellet, and incubate at 37°C for 10-45 min (depending on age of birds); gently agitate the mixture every few min during the digestion period. 7. Terminate digestion by addition of an equal volume of iced HBSS containing 10% HS, and centrifuge as before. 8. Resuspend the cell pellet in about 10 ml iced HBSS containing 10% HS per 1 ml cell pellet.


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9. Pass cell suspension through a stainless steel screen (or gauze), followed by passage through a 20 pm Nitex filter to remove undigested tissue and muscle fibers. 10. Centrifuge as before, resuspend the pellet in 5 ml chilled growth medium (77% Ham’s F10,15% HS, 5% chicken embryo extract, 1% 0.132 mg/ml CaC12, 1%glutamine (lOOX, GIBCO #12420-014), and 1%antibiotic mix (penicillin, streptomycin, fungizone). 11. Determine the cell number, dilute appropriately, and inoculate the desired number of cells into gelatin-coated culture dishes preequilibrated with growth medium. Additional procedures for primary quail myogenic cultures have been described in detail by Konigsberg (1979). A potentially convenient aspect of quail myoblasts is that they appear to replicate almost indefinitely without going through a recognizable senescent phase. In contrast, El2 chick myoblasts reach proliferation senescence after 40-50 doublings. In addition to primary-derived quail cells, several permanent quail myogenic cell lines (QM1-4 and 6-8), as well as a differentiation-defective variant line, have been isolated from the quail fibrosarcoma cell line QT6 (Antin and Ordahl, 1972).The myogenic lines behave appropriately in myogenic gene transfection studies and they also exhibit the capacity of muscle formation when transplanted into chick embryo limb buds (Antin ef aZ., 1991). The primary cultures can now be transfected with DNA using standard calcium phosphate protocols (see Chapter 19). The day following transfection, media is aspirated off, the plates are rinsed with PBS, and differentiation medium is added. After differentiation is allowed to proceed for 2 days, the newly formed myotubes are harvested for analysis. In some instances, it may be important to know the fraction of primary cells that are myogenic. This can be done on a parallel plate of cells that has been allowed to differentiate. The nuclei are stained with hematoxylin and the cells counterstained with eosin to visualize and quantitate the percentage of nuclei incorporated into myotubes. Alternatively, the fluorescent membrane probe merocyanine 450 (Sigma #M6390) can be used to distinguish myotubes and mononucleated myocytes with electrically excitable membranes from myoblasts and fibroblasts that lack this phenotype (Easton et aZ., 1978). In this procedure, plated cells must be thoroughly washed free of serum proteins, as they will bind the dye indiscriminately. This is done with a solution containing 5 mM Tris-HC1pH 7.4,150 mM NaCl. Expose the cells to 1.8 pM Metocyanine 540 in the preceding buffer for 10 min at room temperature under ambient lighting conditions. Rinse the cells again thoroughly to remove all unincorporated dye and examine with microscopic darkfield fluorescence at 500-550 nm. With I S 0 1600 film (FujiChrome), typical exposure times are 10-20 sec.

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d. Conditioned Medium Chick muscle cultures have been used extensively for the clonal analysis of vertebrate myogenesis, most recently for the delineation of fast and slow muscle myoblast subpopulations within the myogenic lineage (Miller and Stockdale, 1986a,b). Many of these studies have utilized conditioned medium (CM) for clonal cell growth. CM is particularly useful for myoblasts from embryos younger than E6 when virtually all muscle colony-forming cells are CM-requiring (White et al., 1975).These media are typically prepared by exposing fresh growth medium to high-density chick musclehibroblast cultures for 1-3 days, refiltering the medium, and diluting appropriately with fresh medium prior to use. The first of the CMs (Konigsberg, 1963) was shown to contain soluble collagen (Hauschka and Konigsberg, 1966) and was required for provision of a collagen substrate upon which myotube formation is potentiated. The necessity of this CM type has been eliminated by utilizing gelatin-coated Petri dishes. A second CM type accelerated the onset of muscle differentiation (Konigsberg, 1971), and was shown in both quail and mouse studies to function via the depletion of serum- and chick embryo extract-derived mitogens from the fresh medium (Linkhart et al., 1981). The necessity of this CM type has been eliminated by simply switching log-phase cultures to media containing low serum levels and zero embryo extract (or defined growth factors such as FBS). A third CM type (White and Hauschka, 1971) allows myotube formation to occur in some clones that would not undergo myotube formation in fresh medium. The percentage of muscle colonies requiring CM for myotube formation is embryonic stage specific (White et al., 1975). The active component in this CM type appears to be a glycoprotein that binds to the surface of gelatin-coated culture dishes and thereafter exerts its effect on muscle colony differentiation even in the presence of fresh medium. The latter CM type has also proven useful for propagating human muscle colony-forming cells from 5-week embryonic limb buds (Hauschka, 1974b). Conditioned medium for the growth of chick embryo myoblast clones is prepared by inoculating 5 X lo6 primary chick embryo muscle cells (typically from day 11-12 leg muscle) into each of about six 100 mm gelatin-coated Petri dishes. The cells are grown in 10 ml fresh medium (FM) consisting of 79% Ham’s FlOC (see MM14 protocol), 5% chick embryo extract prepared according to Konigsberg (1968), 15%pretested horse serum, and 1%antibiotics. These cultures are fed daily for the first 3-4 days and then allowed to differentiate without further feedings. This should result in a confluent cell layer containing many myotubes overlaying a lawn of fibroblasts. The cultures are then passaged, strained through a 20-pm silk or nylon filter to remove myotubes, and replated at 2 X lo6 cells into 20-30 100-mm gelatin-coated dishes. These cultures, which consist primarily of fibroblasts with a few interspersed myotubes, are fed daily with FM until confluent (usually by day 3). Starting on the day of confluency, the cultures are then fed on successive days with 10,15,20,20, and 20 ml FM.After each 24-hr interval the CM is removed and stored as a pooled lot at 4°C. If the conditioned cultures continue to look healthy, several additional 20-ml feedings can be done.


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The pooled CM is then passed through a 0.45-pm filter and frozen at -70°C. Under these conditions CM is stable for more than a year. For reasons that have not been ascertained, some CM batches are superior to others, and some appear to work better when diluted by as much as 1:1 with FM. When freezing the original pooled batch it is thus advisable to freeze several smaller aliquots that can be thawed and tested at full strength and at various dilutions prior to setting up more complex experiments in the medium. Tests with 5-6 day (stage 28-30) chick embryo leg muscle should exhibit 50-60% muscle colonies in a good batch of CM, whereas only about 5-10% of the clones will form muscle when cultured in FM; when tested with muscle cells from 1012-day chick embryos, 80-90% of the clones should be fused when grown in CM and 30-40% should be fused when grown in FM (see White et al., 1975, for typical clonal data from 3-12-day embryonic muscle). In clonal studies of human fetal muscle cells grown in chick CM, differentiation starts earlier than in cultures grown in FM, but the h a 1 percentage of differentiated muscle clones is about the same in both media (Hauschka, 1974a). B. Established Muscle Cell Lines Established myogenic lines do not always retain all of the morphological and biochemical characteristics of primary cultures, but they have distinct advantages. First, they are clonal and not contaminated with other nonmuscle cell types. This feature enhances uniformity between experiments and eliminates the laborintensive preparation involved in primary cultures. Second, under the right culture conditions, proliferative myoblasts can be effectively prevented from spontaneous terminal differentiation, which is inducible by a relatively short exposure to serum-deficient medium or long-term culturing. Third, the presence and the temporal expression of myogenic regulatory factors and many muscle-specific markers are well characterized in these cell lines, both for the undifferentiated and the differentiated states. In transfection studies, test DNA is introduced with high efficiency into proliferating cells, which then can be induced to differentiate at will (see Chapter 19). This feature facilitates the dissection of the myogenic differentiation process and allows for stable selection of clones bearing exogenous genes.

1. The Mouse h4h414 Line MM14 cells were derived in 1978from a single thigh muscle satellite cell clone obtained from a 60 day-old BALB/c male mouse (Hauschka et al., 1979; Linkhart et al., 1981). Their experimental advantages include a rapid population doubling time (12.5 hr), highly reproducible differentiation kinetics in response to mitogen withdrawal, and ease of handling for both transient and stable transfection assays (Amacher et al., 1993). M-Creatine kinase gene expression is activated within 5 hr of fibroblast growth factor withdrawal (Chamberlain et al., 1985), and

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greater than 90% of the population irreversibly committed to terminal differentiation within 14 hr (Clegg et aL, 1987). The potential disadvantages of MM14 cells are their labor-intensive and generally finicky cell culture requirements (necessity of frequent feeding and passaging), the difficulty of growing cultures to high population densities (>lo6 per 100-mm dish) without increasing numbers of cells undergoing spontaneous differentiation, the difficulty of growing cells on glass coverslips, and the general lack of spontaneous contractility in fully differentiated cultures. Similar to the behavior of all rodent muscle cell lines, MM14 cultures also exhibit the spontaneous appearance of differentiationdefective cells. The frequency of this occurrence varies with culture conditions, but under optimal log-phase growth it is typically in the 0.1-18 range. Experimental details as to the standard behavior of MM14 cells have been described (Linkhart et al., 1980,1981;Lim and Hauschka, 1984; Clegg et al., 1987). Persons unfamiliar with these papers should read them before trying to grow the cells. The initial derivation of MM14 cells was via the spontaneous transformation technique described earlier (Hauschka et al., 1979). The original transformed MM14 clone was diploid, and frozen foundation stocks of these cells are still available. However, during early studies with MM14 cells it was found that the diploid line gradually became tetraploid unless it was frequently subcloned. For ease of handling, these tetraploid (80-chromosome) and slightly subtetraploid (76-79-chromosome) derivativesof the original MM14 have been used for virtually all of the published experiments involving these cells. a. Reagentsfor MMI4 Cell Propagation and Dfghrentiation Culture Dishes. All standard tissue-culture dishes and flasks work well, but these should be gelatin- or collagen-coated. Sixty-millimeter dishes for clonal experiments and 100-mm dishes are commonly used for most molecular biology experiments. Exponential expansion of cell stocks is always done on 100-mm plates. Attempts to grow MM14 cells in roller bottles and on microcamer beads have been disappointing; either technique works for growing very large numbers of cells,but culture uniformity is not optimal, especiallyfor experimentsinvolving differentiation kinetics. The use of 96-well plates for MM14 cells studies is possible, but is difficult because of the frequency of feedings required for maintaining proliferation (see later discussion). Celatin-Coating Culture Dishes. Early in the history of skeletal muscle cell culture it was found that most primary myoblasts grow and differentiate better on collagen or gelatin surfaces than on glass or tissue culture plastic (Hauschka and Konigsberg, 1966; Hauschka and White, 1972). Most of the techniques used in these earlier studies have been described in a previous review (Hauschka, 1972). Although many permanent lines of rodent myoblasts do not require gelatin-coated dishes, MM14 cells exhibit a very strong preference for such surfaces. Tissue-culture plastic dishes are thus routinely coated with gelatin. Attempts to replace gelatin-coated dishes with Corning Primaria culture dishes have not proven fully successful, although MM14 cells grow and differentiate


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better on Primaria dishes than on uncoated tissue culture plastic, their behavior is not optimal. Commercially available gelatin contains soluble impurities that should be removed prior to using the gelatin for coating culture dishes. Stock gelatin solutions are prepared as follows: 1. Chill 700 ml glass-distilled water on ice. 2. Add 15 ml iced glass-distilled water to each of four 50-ml plastic screwcapped tubes and place these in an ice bucket. 3. Weigh out four 1-g quantities of commercial gelatin. 4. Gradually add each gram of gelatin to a separate tube while stirring continuously with a stainless steel spatula to prevent clumping. If clumps persist, vortex at a slow setting but avoid foaming. (Keep tubes on ice between these procedures to prevent the gelatin granules from dissolving.) 5. Centrifuge 5 min (4°C) at 2000g. 6. Pour off the slightly yellowish supernatant, and repeat the iced distilled water gelatin granule washing step nine times. (This many washings may not be required for cleaner commercial gelatin preps, but it is necessary for some. If it is yellow, wash it.) 7. Resuspend each of the h a 1 gelatin pellets in about 30 ml of room temperature distilled water, pour into a 1-liter glass bottle, and use additional distilled water rinses of the tubes to bring the final volume to 600 ml. 8. Cap the bottle loosely and autoclave 25 min using a slow exhaust setting. 9. Swirl the bottle well to mix the contents after removing from the autoclave, and cool to about 40°C. 10. Pass the warm gelatin through a 0.45-pm filter, bottle the sterile gelatin in 25-ml aliquots, and store at 4°C. Such solutions are stable for more than a year. 11. Prior to coating plates, incubate the gelatin for 30 min in a 37°C bath. 12. For coating loo-, 60-, and 35-mm plates, transfer 100,30, and 10 p1 gelatin to the center of each plate. (This can be done to 20 plates at a time.) 13. Spread the gelatin evenly over the plate surface using a sterile glass rod bent to the shape of a hockey stick. 14. Gelatin-coated plates can be used immediately, but they are typically coated in batches of hundreds at a time, allowed to dry, and stored at room temperature for use days to months later.

For coating tissue-culture flasks, put several milliliters gelatin in the flask, rock to cover the culture surface, then pipet remaining gelatin into the next flask; tip the flask so that excess gelatin collects in a corner and aspirate it. The gelatin can be transferred from flask to flask until an insufficient amount remains to coat the next flask. Nufrienr Medium. MM14 cells are routinely grown in FlOC (Ham’s F10 Antibiotics, 15% presenutrients with the calcium level adjusted to 1.26 a).

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lected horse serum, and 2 ng/ml bFGF are added to make the final growth medium (see later discussion). Other nutrient media (NM) such as Ham’s F12 (also with the calcium level adjusted to 1.2 mM, or DMEM, 1:l DMEM-F12, RPMI, and Ham’s MCDB media (Ham et al., 1988), all without calcium adjustments, work equally well. FlOC is prepared from commercial powdered Ham’s F10 medium according to the manufacturer’s instructions, except than an additional 1.41 g CaC12 2HzO is added with continuous stirring after the standard step of adding sodium bicarbonate to the medium. When gentamycin is used as the antibiotic, 12 ml of a stock 50 mg/ml gentamycin solution is added per 10 liters prior to filter sterilization of the medium. Growth Medium. Growth medium (GM) consists of 85% NM, 15%pretested horse serum, and a h a 1 basic fibroblast growth factor concentration equivalent to 2 ng/ml bFGF. As a rule of thumb, acidic FGF is about 30 times less active than bFGF (Clegg et al., 1987); thus, it should be used at higher concentrations. Typically, 150 ml pretested horse serum (see later section) is added to 850 ml NM, and this mixture is added to 100-mm and 60-mm plates in volumes of 10 and 4 ml. bFGF (typically 2 ng/ml final concentration) is then added directly to the plates (see later discussion). GM can be stored several weeks at 4°C without deterioration, but it is preferable to warm only the amount necessary for each feeding rather than repeatedly warming and chilling a larger stock bottle of GM. DiflerentiufionMedium. Differentiation medium (DM) consists of 98.5% NM, 1.5%pretested horse serum, 1pM insulin, and antibiotics. Excellent MM14 differentiation also occurs in any of the nutrient media listed above with the horse serum and insulin concentrations as stated. Although added insulin is not required for MM14 differentiation, its presence enhances the survival of differentiatedcells. The high concentration of insulin required, relative to physiological levels, may be due to its ability to mimic the binding of insulin-likegrowth factors to IGF receptors (Florini et aZ., 1993; James et aZ., 1993; Montarras et al., 1993). Pretesting Home Serum. Unlike many permanent lines of rodent myoblasts, MM14 cells do not grow well in nutrient media supplemented with various types of calf serum, unless FGF is also added to the medium. Since calf sera are typically much more expensive than horse serum, it is more cost-effective to use horse serum. Horse serum tests consist of clonal assays to check for plating efficiency and clonal growth rates. The best sera are then compared in highdensity cultures with respect to their ability to support exponential growth (12.5-hr population doubling), for their ability to prevent spontaneous differentiation during log-phase growth, and for their ability to support optimal differentiation when FGF is removed from the medium (the differentiationmedium contains 1.5% horse serum). Occasionally a serum lot may be excellent for growth and suboptimal for differentiation;it is thus sometimesadvisable to purchase separate serum lots that are optimal for each culture phenotype. In a typical 12-lot horse serum test, 2-4 lots are usually as good or slighly better than the old current lot, 2-4 are slightly poorer, and 2-4 are atrocious. High-quality pretested serum


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lots typically retain good growth and differentiation characteristics for at least 6-12 months when stored frozen according to the supplier’s recommended conditions. Horse serum dose-response studies indicate a steep linear response up to 70% of the maximal rate between 1and 5% and a more gradual increase between 5 and 15% (Linkhart el al., 1982). Serum levels up to 30% are not toxic but are not beneficial with respect to supporting higher or longer exponential growth rates. Fibroblast Growth Factor. MM14 cells require FGF for growth and G1phase cells initiate terminal differentiation in its absence (Linkhart el al., 1980, 1981; Clegg et al., 1987). During the first 37 hr of culture MM14 clones exhibit an essentially linear dose-response to concentrations of bFGF between 0.01 and 0.1 ng/ml and to aFGF between 0.3 and 3.0 n g / d (Clegg et al., 1987). For both FGFs the highest concentrations are required to maintain exponential clonal growth with a 12.5-hr population doubling time; and even at 0.1 ng/ml bFGF or 3 ng/ml aFGF, exponential growth ceases between 37 and 50 hr if additional FGF is not added at about 36 hr. The reasons for rapid FGF depletion are not fully understood, but the process appears to be primarily population and medium volume dependent; that is, it occurs more rapidly in high-density cultures grown in low medium volumes, and it occurs extremely slowly if culture dishes containing the standard GM are simply incubated in the absence of cells. In high-density cultures maximal exponential growth rates can be maintained for longer periods of time, without repetitive FGF addition, by continuously rocking the culture dishes. Optimal FGF concentrations for routine studies involving MM14 cells depend on the FGF isoform, its purity, the cell density of the cultures to be studied, and the duration of the culture period. Irrespective of their specified activity, all commercial lots should be tested carefully before large quantities are purchased and before a decision is made as to the most cost-effective concentration and FGF feeding schedule. The Hauschka lab routinely uses a homemade highly purified human recombinant bFGF produced in yeast at a concentration of 2 ng/ml. This level supports maximal exponential growth rates in clonal cultures for 50-60 hr and in newly inoculated mass cultures (l@-l@ cells per 100-mm dish) for 14-20 hr. For ease of feeding cultures, bFGF is handled as follows: 1. Purified FGF should be reconstituted following the manufacturer’s instructions. (Homemade purified recombinant bFGF is typically eluted from a heparin column at 2.5 M NaC1, adjusted to a concentration of 200 pg/ml in 2.5 M NaC1, and stored frozen in 1-ml aliquots in Nunc cryogenic vials in liquid nitrogen. bFGF remains stable and fully active for at least 8 years under these conditions. As necessary, stock tubes are thawed and dispensed in convenient smaller volumes, e.g., 15 p1 in 0.5-ml Eppendorf tubes, and refrozen at -70°C. 2. Depending on the amount of bFGF that will be required for feeding cultures during a 5- to 7-day period, an appropriate volume (e.g., 15 ml) of chilled GM containing 15% horse serum is placed in a 50 ml screw-cap plastic tube. 3. A 15-pl tube of bFGF (containing 3 pg bFGF in 15 p1 2.5 M NaC1) is thawed and transferred to the tube of GM without mixing, and the stock tube

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is rinsed out 3 times by removing 2 0 0 4 aliquots of medium from the top of the 50-ml tube, squirting it up and down several times in the Eppendorf tube, and then transferring it back to the 50-ml tube. 4. Cap the 50-ml tube and mix thoroughly by inverting the tube 10 times. 5. Store the 200 ng/ml (bFGF stock) solution at 4°C. 6. For feeding culture dishes containing 10 ml GM, add 100 pl bFGF stock and rock the dish well before reincubating. This provides a final medium bFGF level of 2 ng/ml. It is not advisable to add bFGF to media in glass bottles due to the rapid adsorption of bFGF to glass surfaces.

6. Exponential Expansion of MMi4 Cells Because of the effect of cell density on FGF depletion and the resulting spontaneous differentiation that then occurs, it is preferable to maintain logphase cultures of MM14 cells at populations below lo6 per 100-mm dish. It is also preferable to plan on a culture passaging schedule which subdivides the cultures every 4-5 cell doublings. An optimal 4-cell cycle expansion of MM14 cells over a 2-day period is camed out as follows:

1. Warm GM to 37"C, add 9 ml per 100-mm dish, add 100 pl bFGF stock per dish, rock dishes well, and preequilibrate the dishes for 1 hr in a COz incubator. 2. Dissociate log-phase MM14 cultures (see later discussion), resuspend cells to a concentration of 6-8 X 104 cells per milliliter in iced GM (minus FGF), add 1ml cells per new culture dish, and rock the dishes thoroughly so that cells are dispersed evenly over the dish surface. Since MM14 plating efficiency is typically 50-60%, this will initiate cultures with 3-4 X 104 proliferative cells per plate. 3. Feed cultures at approximately 12-hr intervals. The first feeding can be accomplished by simply adding an additional 100 pl bFGF stock, but subsequent feedings should involve a complete change of GM plus the bFGF addition. (It is possible to use the shortcut of changing the medium completely only every 24 hr and adding FGF at 12-hrintervals, but this results in a slightly slower growth rate and is likely to increase the background of differentiated cells to 2-3%.) 4. At the time of the second feeding it is preferable to place the culture dishes on a rocking platform operating at about 2 rocking cycles/min. (This is not an absolute necessity, but it will provide a slightly lower background of spontaneously differentiated cells.) 5. Because FGF depletion is cell number dependent, it is important that the interval between the last feeding (at about 36 hr) and dissociation not be more than 12-14 hr. If it must be a few hr longer, an additional 2 ng/ml FGF should be added to the old medium as soon as possible (e.g., at 48 hr). If this is not done, cells will start committing to terminal differentiation. If one knows ahead of time that the last feeding-to-dissociation interval will be more than 14 hr, the use of 3-4 ng/ml FGF for the 36-hr feeding


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will help delay the onset of spontaneous differentiation. However, although it may seem logical that the spontaneous differentiation problem could be routinely overcome by using higher FGF levels for all feedings, this practice is counterproductive because of an accompanying gradual increase in the levels of bFGF required to maintain proliferation. In actual practice exact 12-hr feeding intervals are often very inconvenient. The use of less constant daily feeding intervals such as 10 plus 14 hr is not that detrimental, especially if low levels of spontaneously differentiated cells will not adversely affect the experimental outcome. Cultures inoculated and fed as just described should contain about 5-6 X 105 cells by 48-50 hr. If the total number of undifferentiated cells needed for an experiment makes the logistics of handling the required number of plates at densities less than lo6 per 100-mm dish too difficult, cell densities can be pushed toward the 5 X lo6 level; however, the background of spontaneously differentiated cells will then approach the 5 1 0 % range. In experimental situations such as this it is preferable to inoculate cultures at 5-6 X 105 cellddish and to carry these through only about 4 population doublings rather than to inoculate at lower cell numbers and attempt to take the cultures through 8 cell cycles. In contrast to the logistical difficultiesfaced when one wishes to grow large numbers of undifferentiated MM14 cells, few problems are faced if the goal is to grow large numbers of differentiated cells, especially if it is not detrimental to have some cells differentiate prior to switching the entire culture to DM. In this case 100-mm dishes should be inoculated with 1-2 X 105 cells and the 12-hr feeding regime should simply be continued for 65-75 hr (until the cultures are slightly subconfluent). Cultures should then be switched to differentiation medium as described later. c. Passaging Cells

For optimal exponential expansion of MM14 cells it is best to passage the cultures every 4 doublings. Trypsin-EDTA (e.g., Gibco #610-5405AE) works well for dissociation. Reconstitute powdered trypsin-EDTA as recommended by the manufacturer and dilute to a final trypsin concentration of 0.025% in a physiological saline such as Ham’s saline A (10 mM glucose, 30 mM HEPES, 3 mM KC1, 130 mM NaCl, 1 mM Na2HP04, 0.0033 mM phenol red, pH 7.6). Freeze in aliquots of convenient size for typical experiments. The procedure for passaging 100-mm plates is as follows: 1. Warm 10 ml saline A to 37°C for each plate to be passaged. 2. Thaw an equivalent of 1 ml stock trypsin for each plate to be passaged; add an equal volume of saline A and warm to 37°C. (The trypsin concentration is now 0.0125%.) 3. Aspirate medium, rinse plates twice with 5 ml saline A (37°C).

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Craig Neville el al.

4. Add 2 ml trypsin (37°C) and incubate plates on a 37°C warming bar. Cells usually detach within 3 min. 5. Pipet cells 10 times through a 5-ml pipet and check under an inverted microscope to ascertain that the suspension consists of single cells. 6. Add dissociated cells to conical centrifuge tubes containing iced GM (minus FGF) equivalent to the volume of trypsin-dissociated cells to be added. 7. Serially rinse sets of 5-6 plates with 4-5 ml additional iced GM and add the rinses to the tubes of dissociated cells. 8. Centrifuge 5 min at 150g. 9. Aspirate the supernatant and resuspend cells in 1 ml iced (minus FGF) by pipetting 10 times with a 1-ml pipette tip. 10. Dilute cells such that the desired cell number for each plate is contained in 0.5-1.0 ml and distribute evenly into plates that have been preincubated with complete GM. Rock the plates well to obtain an even cell distribution. (The anticipated cell yield is calculated either by counting the resuspended cells in a hemocytometer or Coulter counter, or by predetermining the cell number per plate prior to dissociation by counting cells in 25-50 randomly selected microscopic fields encompassing a prescribed fraction of the total plate area.) d. Freezing and Thawing Cells Optimal viability of frozen cells is achieved by decreasing the temperature of cells during the freezing process at a carefully controlled rate, and then transferring the frozen cells to liquid nitrogen for long-term storage. The procedure described below is a poor-folks protocol that routinely yields MM14 cells with a 30% plating efficiency upon subsequent thawing.

1. Label the necessary number of cryogenic tubes with the appropriate coded information. 2. Resuspend dissociated cells in iced GM (minus FGF) to 2X the final cell concentration desired. (Slightly higher cell viability is achieved by freezing cells at 106/rnlor higher, but cell concentrations as low as 104/mlare at least 20%viable.) 3. Add an equal volume of iced 15% DMSO made up in GM (minus FGF) to the dissociated cells and mix gently. 4. Add the appropriate volume of diluted cells, usually 1-to 2-ml quantities, to the labeled cryogenic vials. 5. Wrap each vial in two lint-free laboratory tissues and tape sets of 2-3 vials together. 6. Place each set of vials into the depressions in a standard plastic foam egg carton. 7. Close the egg carton and place it into a -70°C freezer for at least 12 hr (not longer than 2-3 days). 8. Transfer tubes to liquid nitrogen for long-term storage.


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To thaw frozen MM14 cells: 1. Thaw cells immediately after removal of the cryovial from liquid nitrogen by immersing the lower three-fourths of the frozen vials in a 37°C water bath for about 2 min. 2. Gently resuspend the cells, dilute to the desired concentration in GM (minus FGF), and inoculate cultures. Since thawed cells tend to form small cell clusters that can lead to precocious spontaneous differentiation, it is preferable to passage and replate frozen cultures within 24-36 hr of plating. e. Dixerentiation Protocol

Exponentially growing MM14 cells differentiate rapidly in response to removing FGF from the GM. Because residual FGF disappears faster at low serum concentrations, more rapid differentiation kinetics can be achieved by simultaneously reducing the horse serum concentration to 1.5%. It is important to note that MM14 cells differentiate in the absence of FGF even when the serum level is maintained at 15%, and irrespective of whether serum is present at all. The general health of the cultures does, however, appear compromised in the total absence of serum. The standard differentiation technique for 100 mm plates is as follows: 1. Warm (37°C) 10 ml Puck’s saline G per culture dish that will be switched to differentiation conditions. (The acidic pH of saline G, approximately 5.5, is thought to potentiate the removal of bFGF such that the onset of differentiation occurs more rapidly; however, roughly equivalent differentiation kinetics occurs when cultures are rinsed with FlOC NM. A 20X stock of Saline G can be made by mixing the following chemicals, in the order listed, into 800 ml glass-distilled water, then bringing the volume to 1 liter and filter sterilizing: 160 g NaC1, 8 g KCl, 3 g KH2P04,5.8 g Na2HP04* 7H@, 22 g glucose, 20 mlO.5% phenol red. Convenient volumes of the 20X stock are then diluted with sterile distilled water prior to use.) 2. Warm (37°C) 10 ml DM (minus insulin, see earlier discussion) for each plate to be switched to differentiation conditions. 3. Aspirate GM. 4. Add 5 ml Saline G, rock plate several times, and aspirate. 5. Repeat step 4. 6. Add 10 ml DM. 7. Add stock insulin to bring final concentration to 1 pM (approximately 1 pglml). 8. Rock plates well to distribute insulin and reincubate.

f. Differentiation-Defective MM14 cells These spontaneous variants can be handled as described for their MM14 parents. However, differentiation-defective (DD) cell differentiation occurs

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much more slowly and only a small fraction of the DD cells become differentiated (Lim and Hauschka, 1984). DD cells respond mitogenically to FGF but do not require it for growth. Since they grow in horse serum alone, the serum level should be reduced to 1%to achieve more rapid differentiation. DD cell revertants that exhibit higher differentiation indices can be isolated by selecting DD clones that exhibit a more rapid onset of differentiation when deprived of growth factors (Lim and Hauschka, 1984).

2. The Mouse C2 Line The C2 muscle line was first established by Yaffe and colleagues (Yaffe and Saxel, 1977) from adult C3H mouse leg muscle that were crush-injured to induce satellite-cell proliferation. These cells maintain a fibroblast-like phenotype in media containing high concentrations of fetal calf serum, but rapidly withdraw from the cell cycle and undergo myogenic differentiation when the medium is replaced with lower concentrations of horse serum. These cells express a wide variety of adult muscle-specific markers, including the myogenic factors and the insulin-like growth factors (see Table I). A fast-fusing subclone, GC12, was derived by Blau and colleagues (Blau et aL, 1983). GCI2myoblasts are also very proliferative in culture, with a doubling time of approximately 16 hr. The fast growth and differentiation characteristics of the GCI2line are advantageous for transfection studies, but their growth and maintenance requires particular attention. Unlike certain other cell types, GCI2 myoblasts are not very motile and form colonies in situ during proliferation. Since cell-cell contact induces myoblasts to differentiate, GCI2myoblast cultures that are not passaged often soon lose their fast-fusing characteristics, as only the slower-fusing myoblasts remain. This feature also precludes the extensive passaging of a culture, since even during normal proliferation with frequent splitting, fast-fusingcells differentiate and drop out of the population. It is therefore advisable to initially prepare a large supply of identical frozen cell aliquots that can be regularly thawed to initiate new cultures.

Table I Muscle Cell Lines and Their Myogenic Factor Expression Patterns

Fusion MyoD Myogenin Myrs MRF4

c2

MM14

L6

BC3Hl

+

+ + +

+

-

+ + (+I-)

-

-

-

-

+ + (+/-)

-

+ (+I-) -


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a. C2 Culture Conditions

The procedures that follow are those routinely used in the Rosenthal laboratory. C, myoblasts are propagated in DMEM containing 20% fetal bovine serum, 2 ml freshly added L-glutamine (aliquotted and stored frozen as a 200-mMstock), and 50 unitslml penicillin-streptomycin. To induce differentiation, the medium is replaced with DMEM containing 2% horse serum in place of the fetal bovine serum. Cells start to fuse within first 24 hr in this media. The C,CI2myoblasts divide very rapidly (16 hr) and differentiate quickly. To maintain a cell stock, cells are passed at 50-70% confluence. Passing confluent cells will result in the selection of a subpopulation of slow-fusing cells. Therefore, the regular freezing of short-term cell cultures is required. Passage of any stock for more than 2-3 weeks will cause the cells to lose their ability to differentiate. Expand and freeze multiple vials from every stock vial used. Throw away cells remaining after an experiment and start a new batch for the next one. 6. C2Culture Tips To maintain stock plates, it is very important that the cells be passaged before they become confluent. Once C, myoblasts are in contact with each other, a subpopulation leaves the cell cycle and begins to differentiate,even in high serum conditions. These differentiated cells are then lost, and over time a subpopulation of the original culture is selected that will not fuse as effectively. The C,Clz subline, originally selected for its high capacity for fusion, is particularly susceptible to this problem. c. Protocol for Freezing Cells

This protocol is described for C, cells, but can be applied to most cell lines. Cells to be frozen down should have been initially dispersed evenly to avoid colonies of differentiated myotubes. Using 150-mm tissue-culture plates allows a larger number of cells to be grown while keeping the cell confluence low, thereby generating a greater number of seeding vials. 1. Rinse the plate with PBS (10 ml/l50-mm plate). 2. Add 3-5 mlO.25% trypsin or pancreatin per 150-mm plate. Leave plate at room temperature for 5 min or until all cells come off the plate. Tap the sides of the plate to loosen the cells if necessary. 3. Neutralize the enzyme with an equal volume of growth medium (the addition of serum inactivates the trypsin). 4. Transfer cells into a 15-ml tube (at this point the cells can be counted, so that a known concentration of cells can be frozen per vial) and spin at 150g for 5 min. Higher speeds will damage the cells. 5 . Aspirate the supernatant, leaving the pellet intact. 6. Resuspend the pellet in an appropriate volume of medium containing 20% FBS. Slowly add DMSO to a final concentration of 10%while mixing carefully.

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DMSO in high concentration is toxic to cells, so do not allow it to contact the pellet. We usually resuspend 1 X lo6 cells/ml and freeze l-ml aliquots. 7. Transfer the cells into cryogenic vials. 8. Cells must be frozen at -20 to -80°C overnight. An excellent method for freezing cells is to use a Nalgene Cryogenic Controlled-Rate Freezing Container (Nalge #5100-0001). The vials can be placed in the container and stored directly in a -80°C freezer overnight. Do not store these cells at -80°C for longer than 2-3 days. Cells are then transferred into liquid nitrogen where they can be stored indefinitely. d. Protocolfor Thawing Cells 1. Remove a vial of cells from liquid nitrogen and hold by hand in a 37°C water bath in order to thaw it quickly. As soon as the cells are thawed, the tube should be removed. 2. The DMSO present with cells must now be removed. Transfer the cells into a polypropylene 15-ml tube. Gradually add 10 ml PBS. 3. Resuspend cells and pellet them by centrifuging at 150g for 5 min. 4. Aspirate the supernatant and resuspend the cells in 10 ml growth medium. Plate cells in a 100-mm plate. If lo6 cells have been plated, they should be split the day after thawing. The media must be changed the next day to remove dead cells and any residual DMSO.

3. The Rat L6 Line The L6 line was derived by YafFe (1968) from a rat neonatal hindlimb preparation. Like the C, line, L6 cells remain as proliferating myoblasts in DME supplemented with 20% fetal calf serum, although they do not proliferate as rapidly as C, myoblasts. They can be induced to differentiate upon serum reduction (2% horse serum), but the rate of differentiation is much slower than that of the C2 line. This may be because L6 cells do not express their own IGFs and are dependent on the IGFs present in the medium both for stimulation of proliferation and for differentiation induction (Ewton and Florini, 1981).Subclonal derivatives of the L6 line include L6E9 (Nadal-Ginard, 1978) and L6A1 (Ewton et al., 1984), both of which reportedly differentiate faster than the parental L6 line. The L6 lines also express a more limited number of myogenic cell markers upon differentiation and are characterized by the lack of MyoD in both myoblasts and myotubes (Lathrop et aZ., 1985).

4.The Mouse BC3H1 Line The mouse muscle cell line BC3H1 was originally obtained from a nitrosoethylurea-induced brain tumor, which may account for some of its unusual characteristics (Schubert et aZ., 1974). While maintained in DME supplemented


113

with 20% fetal calf serum, BC3H1 cells exhibit a fibroblast-like morphology and like other muscle cell lines, do not express muscle-specificdifferentiation markers. Upon reduction of serum concentration (2% horse serum), the myoblasts exit the cell cycle and induce a limited array of skeletal muscle gene products, including myogenin. However, unlike other muscle cell lines, BC3Hl cells do not fuse to form multinucleate myotubes or commit to terminal differentiation, since reapplication of serum causes reentry into the cell cycle (Lathrop et al., 1985). As differentiated mononucleate myocytes, BC3H1do not express MyoD, cardiac a-actin, or fast myosin light chain 1at detectable levels (Davis et al., 1987;Strauch and Reeser, 1989; Taubman et al., 1989). Transfection of a MyoD expression vector causes the cells to fuse and restores expression of both actin and myosin light chains, suggesting that the principal defect in this cell line is the absence of MyoD.

5. Obtaining Muscle Cell Lines MM14 and MM14-DD cell stocks are available by request from Steve Hauschka, Department of Biochemistry, University of Washington, Seattle, WA 98195-7350 (telephone 206-543-1797; fax 206-685-1792). In the interest of maintaining relatively uniform MM14 and MM14-DD cell stocks among all investigators using these cells lines, it is beneficial for each new user to obtain cells from the set of foundation stocks available in this lab. Other cell lines that have been deposited with the American Tissue Culture Collection are C& (ATCC CRL 1772), L6 (ATCC CRL 1458), and BC3H1 (ATCC CRL 1443).

111. Summary and Future Prospects The recent progress in the molecular characterization of myogenic differentiation has depended largely on the successful analysis of the muscle phenotype in cell culture. As evident from the preceding descriptions, myoblasts from numerous sources can be cultured to mimic different aspects of myogenesis, from proliferation to withdrawal from the cell cycle, fusion into myotubes, and expression of contractile protein gene subsets. Certain aspects of myogenesis have been more difficult to reproduce in culture, such as the full recapitulation of contractile function and the further modulation of muscle differentiation that occurs during the generation of different fiber types. A more thorough analysis of the external environment required for the manifestation of these properties in vivo may ultimately be necessary, since it is likely that specific extracellular matrix components or heterologous cell interactions play a significant role in determining these phenotypes. In addition, the establishment of subcellular domains within the mature myotube may determine the heterogenous distribution of key factors responsible for other features of mature muscle fibers, such as articulation with

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cartilage and bone. These are future challenges for the muscle cell biologist and will broaden the spectrumof questionsto which muscle cell culture can be applied. Acknowledgments Dr. Frank Stockdale is thanked for the avian cell culture protocols, Jeanne Powell and Steve Burden for rodent protocols, and Dr. Jean Buskin and DeeGregory for their many helpful contributions to protocols provided for MM14 cells. Dr. Mary Pat Wenderoth is thanked for help with the manuscript.

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Montmas, D., Pinset, C., P'erez, M. C., Ilan, J., and Gros, F. (1993). Muscle differentiation: Insulinlike growth factors as positive modulators of myogenic regulatory genes? C. R. Acad. Sci. ZII 316,1025-1031. Nadal-Ginard, B. (1978). Commitment, fusion, and biochemical differentiation of a myogenic cell line in the absence of DNA synthesis. Cell 15,855-864. Powell, J. A., and Fambrough, D. M. (1973). Electrical properties of normal and dysgenic mouse muscle in culture. J. Cell. Physiol. 82, 21-38. Rando, T. A., and Blau, H. M. (1994). Primary mouse myoblast purification, characterization, and transplantation for cell-mediated gene therapy. J. Cell Biol. 125,1275-1287. Rhodes, S . J., and Konieczny, S. F. (1989). Identification of MRF4 A new member of the muscle regulatory factor gene family. Genes Dev. 3,2050-2061. Rutz, R., Haney, C., and Hauschka, S. D. (1982). Spatial analysis of limb buds myogenesis: A proximodistal gradient of muscle colony-forming cells in chick embryo leg buds. Dev. Biol. 90, 399-411. Schubert, D., Harris, A. J., Devine, C. E., and Heinemam, S. (1974). Characterization of a unique muscle cell line. J. Cell Biol. 6 1 , 3 9 8 4 3 . Seed, J., and Hauschka, S. D. (1981). Clonal analysis of vertebrate myogenesis VIII. Fibroblast growth factor (FGF)-dependent and FGF-independent muscle colony types during chick development. Dev. Bid. l28,40-49. Seed, J., and Hauschka, S. D. (1984). Temporal separation of the migration of distinct myogenic precursor populations into the developing chick wing bud. Dev. Biol. 106,389-393. Seed, J., and Hauschka, S. D. (1988). Clonal analysis of vertebrate myogenesis. VIII. Fibroblasts growth factor (FGF)-dependent and FGF-independent muscle colony types during chick wing development. Dev. Biol. U8,40-49. Stockdale, F. E. (1992). Myogenic cell lineages. Dev. Biol. W, 284-298. Strauch, A. R., and Reeser, J. C. (1989). Sequential expression of smooth muscle and sarcomeric a-actin isoforms during BC3H1 cell differentiation. J. Biol. Chem. 264,8345-8355. Taubman, M. B., Smith, C. W. J., Izumo, S., Grant, J. W., Endo, T., Andreados, A., and NadalGinard, B. (1989). The expression of sarcomericmuscle-specificcontractile protein genes in BC3H1 cells: BC3H1 cells resemble skeletal myoblasts that are defective for commitment to terminal differentiation.J. Cell Biol. 108, 1799-1806. Thayer, M. J., Tapscott, S. J., Davis, R. L., Wright, W. E., Lassar, A. B., and Weintraub, H. (1989). Positive autoregulation of the myogenic determination gene MyoD1. Cell 58,241-248. White N. K., and Hauschka, S. D. (1971). Muscle development in vitro. A new conditioned medium effect on colony differentiation. Exp. Cell Res. 67,479-482. White, N. K., Bomer, P. H., Nelson, D. R., and Hauschka, S. D. (1975). Clonal analysis of vertebrate myogenesis. IV.Medium-dependent classification of colony-forming cells. Dev. Biol. 44,346-361. 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. Yablonka-Reuveni, Z., Quinn, L. S., and Nameroff, M. (1987). Isolation and clonal analysis of satellite cells from chicken pectoralis muscle. Dev. Biol. 119,252-259. Yaffe, D. (1968). Retention of differentiation potentialities during prolonged cultivation of myogenic cells. Proc. Natl. Acad. Sci. USA 61,477-483. Yaffe, D., and Saxel, 0. (1977). Serial passaging and differentiation of myogenic cells isolated from dystrophic mouse muscles. Numre 270,725-727.

CHAPTER 6

Avian Cardiac Progenitors: Methods for Isolation, Culture, and Analysis of Differentiation Maureen Gannon and David Bader Department of Cell Biology Vanderbilt University Medical Center Nashville, Tennessee 37232

I. Establishment of Embryonic Axis 11. Pregastrulation Location of Cardiac Progenitors 111. Postgastdation Location of Cardiac Progenitors IV. Removal of Embryos from the Egg A. Filter-Paper Method B. The New Method V. Removal of Cardiac Progenitors VI . Removal of Endoderm A. Mechanical Removal B. Sodium Citrate C. Enzymatic Digestion VII. Culturing Whole Embryos A. Culturing on Uncoated Dishes B. Culturing on Agar-Coated Dishes VIII. Culturing Explants of Cardiogenic Mesoderm A. Fibronectin Coating B. Collagen Coating C. Culture Media IX. Culturing Primary Cardiomyocytes A. Myocyte Isolation X. Analysis of Differentiation A. Immunohistochemistry B. In Situ Hybridization with Digoxigenin-Labeled Riboprobes C. Whole-Mount in Situ Hybridization D. Explant in Situ Hybridization E. In Situ Hybridization Using '%-Labeled Probes References METHODS IN CELL BIOLOGY, VOL. 52 Copyright 0 1998 by Academic Press. All dghe of reproduction in any form reserved 0091-679X/98 125.00

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The vertebrate heart tube is a mesodermally derived structure that arises from initially paired primordia located on either side of the embryonic midline. The avian embryo is ideal for studying the early events in heart development for several reasons, including its accessibility,the availability of fate maps, the ability to culture embryos ex ovo, and low cost. To analyze and manipulate cardiac progenitors experimentally,it is essential to be able to locate this population of cells precisely. Using the avian embryo as a model system, it has been possible to localize those cells with heart-forming potential from the early stages of gastrulation throughout organogenesis. The staging series for avian development established by Hamburger and Hamilton (1951) will be used throughout this chapter. Although hours of incubation at 37°C are given as a guideline, developmental timing may vary and should be established morphologically. Generation of a functional, beating heart occurs relatively early in avian development,at stage 10 (approximately36 hr of incubation). Examination and manipulation of cardiomyocyte commitment and differentiation can therefore be completed within a short time frame. Although the population of cardiogenic precursors will be considered as a whole, it should be noted that there exist stage-dependent changes in cell-cell contacts within the cardiogenic mesoderm as well as between germ layers (Stalsberg and DeHaan, 1969;Linask, 1992). By dissecting embryos at various stages, one should be able to get a feel for these differences. In addition, throughout heart development there exists a rostral-to-caudal difference in cell-cell contact, cell commitment, and gene expression such that the more anterior population of cells develops ahead of the posterior progenitors (Stalsberg and DeHaan, 1969; Han et al., 1992;Yutzey et al., 1994).The position of cardiac precursors in the developmental pathway must be considered heterogeneous along the rostrakaudal axis. These differences will be discussed in more detail in Section X, Analysis of Differentiation.

I. Establishment of Embryonic Axes In avian development, the first embryonic axis to be determined is the dorsal/ ventral axis. The ventral surface of the developing embryo is in direct contact with the underlying yolk. The anteroposterior axis develops next with the formation of a thickened area of cells in the posterior blastoderm known as Koller’s sickle (stage 2, 7 hr). This thickening is the result of the migration of cells from the lateral regions of the posterior epiblast toward the midline. As these cells move anteriorally, they form the definitive primitive streak (stage 3-4, 13-20 hr) through which gastrulation of cells located within the epiblast will occur to yield embryonic mesoderm and endoderm. The primitive streak reaches its most anterior position at stage 4 (20 hr), at which time Henson’s node becomes apparent at the rostra1 tip. Although the primitive streak begins to form at stage 2, ingression of cells does not begin until midway through stage 3 (Vakaet, 1970).

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11. Pregastrulation Location of Cardiac Progenitors The mapping of early primitive streak stages 2-3 (Rosenquist, 1970, 1985; Garcia-Martinez and Schoenwolf, 1993) has revealed that cardiac progenitors are located in the most rostral half, with the exception of the extreme rostral tip, which will give rise to Henson’s node. Cardiac progenitors can be found in a region extending 125-750 pm from the rostral tip of the streak (Garcia-Martinez and Schoenwolf, 1993). Cardiac precursors within the primitive streak are arranged in a generally rostrocaudal sequence. Cells destined to become incorporated into the conus arteriosus and the ventricle (anterior heart tube structures) are located in more rostral regions of the streak, whereas those that will give rise to the atrium and sinus venosus (posterior heart tube structures) are localized to more caudal portions of the streak. In addition, cells destined to give rise to more anterior regions of the heart tube migrate away from the streak before those cells that will become incorporated into the caudal heart tube. Thus, it seems that there is little to no mixing of cardiac precursors along the anteroposterior axis of the primitive streak. These studies also showed that most precardiac cells have left the streak and migrated laterally to form lateral plate mesoderm by stage 4. At this stage, Henson’s node contains cells that will give rise only to the endocardium (Garcia-Martinez and Schoenwolf, 1993).

111. Postgastrulation Location of Cardiac Progenitors The position of cardiogenic mesoderm within the anterior lateral plate mesoderm following gastrulation has been well established (Rawles, 1943; DeHaan, 1963; Rosenquist and DeHaan, 1966; Stalsberg and DeHaan, 1969; GonzalezSanchez and Bader, 1990). These primordia can be visualized under light microscopy as regions of increased opacity and are located bilaterally on either side of the primitive streak with a separation of approximately 800 pm (Stalsberg and DeHaan, 1969). The caudolateral borders of the cardiac primordia remain ill defined. The single, linear heart tube results from the rostral-to-caudal fusion of the left and right heart-forming regions (LHFR and RHFR) above the anterior intestinal portal (AIP) beginning at stage 6 (25 hr) and finishing by stage 17. The generation of the coelom at stage 6-7 physically separates precardiac cells within the splanchnic lateral plate mesoderm from the more dorsally located somatic lateral plate mesoderm. Grafting of labeled progenitors into isochronic hosts and analyzing their incorporation into the heart tube has shown that there is little to no mixing of cells within the cardiogenic mesoderm (Rosenquist and DeHaan, 1966;Stalsbergand DeHaan, 1969;Gonzalez-Sanchez and Bader, 1990). Thus, the precardiac splanchnic mesoderm moves and folds without losing its integrity as a cohesive sheet and is considered an epithelium rather than a mesenchymal layer (Rosenquist and DeHaan, 1966, Linask, 1992). Therefore, it follows that any given region of the heart tube has a well-outlined precursor in

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the cardiac primordia. Stalsberg and DeHaan (1969) have shown this to be the case. The precardiac mesoderm can be subdivided approximately along its anteroposterioraxis with the most rostromedial cells giving rise to the conoventricular region, the middle portion giving rise to the ventricle, and the caudolateral cells generating the atria and sinus venosus. There is some asymmetry in the size of these general regions on the two sides, with the LHFR contributing more cells to the caudal part of the heart tube and the RHFR contributing more cells to the anterior heart tube (see Figure 8, Stalsberg and DeHaan, 1969). This has been confirmed in our laboratory using a gene expressed solely in the posterior population of cardiogenic cells throughout development (Gannon, and Bader, 1995).

IV. Removal of Embryos from the Egg As mentioned earlier, the avian embyro provides a good system for studying the early events in cardiogenesis because it develops outside its mother and is easily removed from the egg for experimental manipulations. With practice, one should be able to remove 12 embyros from their eggs and dissect out the cardiac primordia in less than 2 hr. In addition, whole embyros can be grown in culture from the earliest stages of primitive streak formation (stage 3) through the formation of a beating heart (stage 12). The following are two methods used for removing embryos up to 72 hr old from their eggs. Our laboratory prefers the filter-paper method. A. Filter-Paper Method

Embryos at early developmental stages (stages 1-20, up to 72 hr) can be easily removed from the egg using filter-paper rings. Whatman grade 3 filter circles provide a sturdy support for embryo culture. Fisherbrand P5 filter circles are thinner and less expensive and are good for removing embyros for fixation or from which the cardiogenic region will be dissected. Precut rings with an outer diameter of 2.5 cm can be made with an inner diameter of 1 cm for stage 1-5 embryos and 1.4 cm for stage 6-20. The tools needed to remove embryos from the egg include forceps, small curved scissors, filter rings, sterile PBS (150 mM NaC1, 10 mM phosphate, pH 7.4), and Petri dishes. Using the forceps, make a small crack in the upper part of the shell, cut around the top, and remove the “lid.” Be careful not to break the yolk. Surrounding the yolk are thick and thin albumin. The thick albumin must be removed. This can be done by using the forceps to scoop it out from beneath the yolk as well as to scrape it gently off the surface of the yolk. The yolk’s surface will change from a highly refractile to a more grainy appearance with the removal of the thick albumin. Following removal of thick albumin from the egg, the ring is placed around the area occupied by the embryo using forceps.


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The location of the embryo can be identified as a ring of white with a central clear area. If there is any thick albumin remaining on the surface of the embryo, the ring will not adhere to the vitelline membrane, making removal of the embyro difficult. Once the ring has adhered, cut around its circumference and lift the ring with its attached embryo off the yolk surface using forceps. To remove excess yolk, rinse the embryo gently with sterile PBS. The embryo is then placed ventral side up in PBS in a 35-mm culture dish for dissection. If the embryo is going to be cultured to a later developmental stage ex ovo, a second filter ring should be placed over the ventral surface. This ring sandwich decreases the likelihood of the embyro coming off the ring during culture. B. The New Method In the method developed by New (1955), an equatorial cut is made around the vitelline membrane and the embryo transferred, ventral side up, to a watch glass coated with albumin. The vitelline membrane is supported and held taut by a glass ring 1 inch in diameter and incubated at 37°C for 1 hr to allow the membrane to adhere to the ring. Once the membrane has adhered, the embryo and glass ring can be transferred to a culture dish with PBS for dissection.

V. Removal of Cardiac Progenitors Explants of cardiac precursors from postgastrulated embryos (stages 4-9) can be removed using Fig. 1 as a guide. From stages 7-9 only those cells posterior to the AIP can be considered undifferentiated cardiac precursors since those cardiogenic cells located over the AIP have already initiated cardiac-specific gene transcription (Han et al., 1992; Yutzey et al., 1994; Gannon and Bader, unpublished observations). Since the embyro is ventral side up, the RHFR will be on the left and the LHFR will be on the right. Prior to heart tube formation, cardiogenic mesoderm remains in close association with the anterior endoderm (Linask and Lash, 1986). It is therefore easier to remove cardiac precursors with their associated endoderm. Removal of endoderm will be discussed later. Recalling that the embryos have been placed endoderm side up, needles made of tungsten or pulled 1.0-mm glass capillary tubes can be used to cut through the endoderm and underlying mesoderm in a rectangle that includes prospective cardiac cells. These two germ layers can then be teased off the less adherent underlying ectoderm using the dissectingneedle. At earlier stages of development (stages 4-5) the mesoderm itself is not as cohesive as at later stages and may break into smaller fragments while being separated from the ectoderm. Explants of mesoderm plus endoderm can be removed more easily as an intact rectangle from stages 6-9. Explants of all stages can be transferred using a micropipet. It is sometimes easier to transfer the tissue by first applying 2 pl PBS directly

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Stage 3

Stage 4

Stage 9

Stage 10

Stage I

B

Stage 13

Fig. 1 Localization of cardiogenicprecursors and differentiated cardiomyocytes in the developing chick embryo. (A) Location of ventricular (stippled) and atrial (solid) precursors within the primitive streak and undifferentiated anterior lateral plate mesoderm based on fate mapping studies. (Stage 3: Garcia-Martinezand Schoenwolf,1993;Stage 4 and 7: OUT observations). (B) Location of differentiated ventricular (stippled) and atrial (solid) cardiomyocytes in the tubular heart based on the expression patterns of chamber-specific myosin heavy chains (Yutzey et al., 1994).

to the explant and immediately aspirating the fluid containing the explant to be transferred.

VI. Removal of Endoderm A. Mechanical Removal Endoderm can be mechanically teased away from the underlying cardiogenic mesoderm as an intact sheet of cells using either tungsten or pulled glass needles.


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For removal of endoderm, it is important to use as fine a needle as possible. The endoderm is more easily removed prior to explantingthe cardiogenic region. Beginning at one end of the cardiogenic region, a cut can be made through the first cell layer only (i.e., endoderm) and the endoderm gradually rolled back using the dissecting needle to reveal the underlying mesoderm (Fig. 2). The mesoderm can then be removed en bloc as above. This technique allows for separate or coculture of germ layers with minimal tissue fragmentation; it is the one we have used preferentially in our lab.

B. Sodium Citrate Sodium citrate may be used to dissociate endoderm from the mesoderm and ectoderm beneath it (DeHaan, 1963). Solutions of 40-50 mM in a volume of 10 pl can be added to embryos at stages 4-7+ and incubated overnight at 37°C. In the majority of cases, this results in the ventral surface being almost completely denuded of endoderm. The mesoderm remains intact and is able to continue to develop and differentiate (DeHaan, 1963). This mesoderm can then be explanted as described earlier and transferred to culture. We have not used this method extensively in our lab as it sometimes results in dissociation of all germ layers and/or incomplete removal of endoderm.

Fig. 2 Stage 4 embryo, ventral side up, showing removal of endodermal cell layer to reveal the underlying mesoderm and ectoderm. HN, Henson’s node; ps, primitive streak.


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C. Enzymatic Digestion

The use of proteases has been found to facilitate the mechanical removal of endoderm from the underlying mesoderm (Orts-Llorca and Gil, 1965; Stalsberg and DeHaan, 1969; Kokan-Moore et al., 1991; Sugi and Lough, 1994). There are two methods commonly used: (i) Incubate embyro in 0.5% trypsin in calcium/ magnesium-free saline for 10 minutes at 37°C. Enzyme action is stopped by adding fetal bovine serum and rinsing in Earle’s balanced salt solution. (ii) Incubate embryo in a mixture of collagenaseldispase (1 mg/ml) in calcium/ magnesium-free saline for 12 min at 0°C. Enzyme action is stopped by adding 0.02% EDTA in PBS at 4°C for 5-10 min. This method is the procedure of choice in several labs, although we have never used it.

VII. Culturing Whole Embryos Embryos can be cultured up to the time when they would normally begin to rotate (i.e., stage 12-13). Beyond this stage, the increased need for blood flow and the increase in tension on the extraembryonic membranes due to embryonic flexures and rotations results in arrested development. Embryos removed from the egg from stage 4 and later can be grown to stage 12 in approximately 24 hr (note that the development of embryos in culture is slower than seen in vivo). The majority of these embryos are morphologically normal and show proper spatial regulation of cardiac-specific genes (Yutzey et al., 1994; Gannon and Bader, unpublished observations). Thus, these early embryos can be experimentally manipulated in the culture system without developmental complications arising from the culture conditions themselves. The type of medium one uses depends on the experimental analysis being done. Media that are enriched with potential growth factors (for example, those containing serum or embryo extract) should not be used in studies where the effects(s) of the addition of an exogenous factor(s) is being examined. In these cases, a defined minimal media such as M199 should be used. A. Culturing on Uncoated Dishes

Eggs are removed at the desired stage as described earlier and placed in 35-mm culture dishes coated with thin egg albumin. The medium we use is one of the following: (i) a mixture of 50%thin egg albumin :50% high-glucose DMEM containing 10% fetal bovine serum (heat inactivated), 1 mM sodium pyruvate, 0.1 mM nonessential amino acids, and penicillin (100 U/ml)/streptomycin (100 pg/ml); (ii) a mixture of 50% thin egg albumin :50% M199 medium containing penicillinlstreptomycinas before; (iii) high-glucose DMEM containing 1mM sodium pyruvate, 0.1 mM nonessential amino acids, 1 mM L-glutamine, 1 X MEM nonessential vitamins, 20% fetal calf serum, 1%chick embyro extract,


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and penicillinlstreptomycin as before; and (iv) M199 medium alone containing antibiotics as before. B. Culturing on Agar-Coated Dishes Embyros are removed at the desired stage and placed on 35-mm dishes previously coated with agar-albumin medium by one of two methods:

1. Agar with NaCl (123 mM NaCl containing 0.6% Difco Bacto agar) is first boiled and allowed to cool to 49"C, then mixed with an equal volume of albumin/glucose(thin egg albumin containing 0.3% w/v glucose) and poured into 35-mm dishes at a thickness of 0.6 cm. Plates are cooled at 4°C and used within 1 hr, allowing them to equilibrate to room temperature before use (Sundin and Eichele, 1992). The embryos are then overlaid with 1 ml yolk-Tyrode solution: 1 part yolk extract (equal volumes of fresh yolk and Tyrode's solution are mixed, spun at 13,000 rpm for 10 min at room temperature, and the supernatant collected on ice) to 9 parts Tyrode's solution (Sundin and Eichele, 1992). 2. A solution of 1%Difco Bacto agar in media (high-glucose DMEM plus 7% fetal calf serum) is autoclaved, allowed to cool to 49"C, and used to coat 35-mm culture dishes. Just enough agarlmedium is used to completely cover the bottom of the dish and the remainder removed. The plates can be stored at room temperature for 2 days or at 4°C for up to 1 month. The embyros are then overlaid with 1 ml medium used above with the addition of 100 U/ml penicillin and 100 pg/ml streptomycin.

Embryos are incubated at 37°C in 5% C 0 2 with high humidity to the desired stage.

VIII. Culturing Explants of Cardiogenic Mesoderm Explants of cardiogenic mesoderm with or without associated endoderm are best cultured on an extracellular matrix component such as fibronectin or collagen. Explants can be cultured in a volume of 200 pl medium on 35-mm culture dishes or 8-mm 8-well polystyrene chamber slides (Lab-Tek chamber slides, Nunc, Inc.). We prefer using chamber slides as they minimize volume and allow for antibody detection and in situ hybridization right on the slide. As with the whole embyro culture, the choice of medium used will depend on the type of experimental analysis being done. A. Fibronectin Coating Under sterile conditions, 150-300 p1 of a 20 p g / d fibronectin solution (human; Sigma Chemical Co.) is added to each chamber and incubated for 30 min at

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37"C, followed by aspiration and rinsing with PBS. Slides must be dried well in a laminar flow hood before they are used. B. Collagen Coating Collagen Type I solution is prepared as follows: Add collagen to 0.1 M acetic acid to obtain a 0.01% w/v solution. Stir at room temperature 1-3 hours until dissolved. Add 10%(the volume of collagen solution) chloroform to the bottom of an autoclaved glass bottle without shaking. Incubate overnight at 4°C. Aseptically remove the top layer and transfer to an autoclaved glass bottle. Under sterile conditions, add 150-300 ~10.01% collagen solution to each chamber and incubate 1 hour at 37"C, followed by aspiration. Dry slides well in a laminar flow hood before using. C. Culture Media

1. Medium M199 (Gibco) plus 100U/ml penicillin and 100pglml streptomycin. This unfortified, minimal medium is sufficient to support cardiogenic differentiation and may be used for mesodermal explants at all developmental stages, although the survival rate of stage 4-6 explants may be lower (Gannon and Bader, unpublished observations). 2. M199 with 5 yglml insulin, 5 pglml transferrin, 5 nglml selenium, and penicillirdstreptornycinas before. This medium has been shown to support cardiogenic differentiation (Lough et al., 1990; Sugi et al., 1993). 3. DMEMIMcCoy's: 75 :25 mixture of DMEM and McCoy's 5A basal media (Gibco) plus 1 pg/ml linoleic acid and 250 pglml BSA as a carrier. This medium has been shown to support cardiogenic differentiation of mesodermal explants at all developmental stages (Antin et al., 1994). 4. DMEM with 10%fetal calf serum. This medium supports cardiogenicdifferentiation of mesodermal explantsfrom all developmentalstages but contains potential signaling molecules. Cultured explants of cardiogenic mesoderm develop at a slower rate than cardiogenic mesoderm in viva Explants from stage 4-6 embryos will begin to contract at approximately 48 hours of culture, whereas those from stage 7-9 may beat within 24 hr.

IX. Culturing Primary Cardiomyocytes The primitive heart tube consists of mainly two cell types: myocardial and endocardial.In contrast, older embryonichearts contain an increasing population of fibroblasts and other cell types. For primary mycoyte cultures, it is possible to enrich for cardiomyocytes using differential adherence (Kasten, 1973). Differ-


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ential adherence does not remove all nonmyocytic cell types, however, and the percentage of these cells in myocyte cultures increases with the age of the donor heart. In addition, cardiomyocytes isolated from hearts from embryonic day 4-6 (E4-6) show higher mitotic rates than those from older embryos (Clark and Fischman, 1983). We and others have observed that after 7 days in culture, spontaneous contractions of cardiomyocyte cultures decrease, and this is paralleled by a reduction in myofibrils as detected with the anti-myosin heavy chain monoclonal antibody, MF20 (DeHaan, 1967;Bader et al., 1982;Zadeh et al., 1986). A. Myocyte Isolation

Hearts should be removed from the embryo aseptically and cut into fragments of approximately 0.5-1.0 mm in sterile PBS. If desired, the atria can be separated from the ventricles for separate culture and analysis. The earliest time at which we have been able to cleanly separate atria from ventricles for primary cell culture is at E6-7. Cells are then released by enzymatic digestion. The digestion cocktail includes 0.5% trypsin, 0.1% collagenase, and 0.1% hyaluronidase Type Is, filter sterilized in PBS. Tissue fragments may be stirred gently in the enzymatic solution for 10min and then incubated at 37°C for an additional 30 min. Digestion is terminated by dilution with growth medium (M199 medium, 10% fetal calf serum, 10 microunitslml insulin, 200 pg/d cephalothin, and 1pg/ml glutathione) containing 20% fetal bovine serum. Cells are recovered by centrifugation (500 g) at 4°C. Cells are resuspended in growth medium and filtered through Nitex filters (two layers, 50- and 22-pm pore size). The myocyte population may be enriched by differential adherence for 60-90 minutes on untreated tissueculture plastic (Kasten, 1973). Cells are plated at densities of 100-200 cells/mm2 and maintained in growth medium changed every 24 hr.

X. Analysis of Differentiation In addition to the appearance of rhythmic contractions, cardiac differentiation can be analyzed by assaying for the presence of cardiac-specific proteins and/or mRNAs by using antibodies or riboprobes, respectively. This can be done in whole mount on intact embryos or on cultured mesodermal explants. Recipes for all standard stock solutions used in RNA detection may be found in Maniatis et al. (1982). A. Immunohistochemistry

Embryos are fixed and reacted with antibody while still on the filter ring. Embryos removed from the egg just prior to fixation must be rinsed free of yolk with PBS. Embryos grown in culture must have the medium removed and be rinsed free of residual medium with PBS prior to fixation. Cultured explants can

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be fixed on the chamber slides following removal of media and rinsing twice in PBS. Fixation is done for 1 hour at room temperature in 70% methanol. Wash twice in PBS and then block with 1%BSA in PBS for at least 1 hr at room temperature. Wash twice in PBS and add primary antibody at 4°C overnight (time of incubation with primary antibody may have to be adjusted according to the antibody being used). Following removal of primary antibody, the embryos or explants should be washed all day with 7-10 changes of PBS and then incubated with secondary antibody. For fluorescence-conjugated antibodies, this can be done at room temperature for 1-2 hr or at 4°C overnight (the timing and temperature of this incubation may vary depending on the type of secondary antibody being used). The embryos or explants are then rinsed several times in PBS, fixed in neutral buffered formalin for 30 min at room temperature, and then rinsed in PBS. Figures 3A and B (see Color Plates) depict a stage 11 embryo reacted with the MF20 monoclonal antibody (Bader et al., 1982).Positive reactivity is detected only in differentiated cardiomyocytes within the tubular heart. Figure 3C (Color Plates) shows an example of a cultured explant of cardiogenic mesoderm plus ectoderm removed from the embryo at stage 7 and reacted with MF20, demonstrating that differentiation has occurred in culture. B. In Situ Hybridization with Digoxigenin-Labeled Riboprobes

Digoxigenin-labeled riboprobes can be used on whole embyros or on tissue explants (Continho et al., 1992; Yutzey et al., 1994; Yutzey et al., 1995; Gannon and Bader, 1995). This procedure has the advantages of being very sensitive and nonradioactive and taking less than 1week to complete. Whole embryos can be embedded and sectioned following the in situ hybridization procedure with little to no loss in reactivity. All solutions used for fixation, storage, and in situ hybridization should be RNAse-free, and gloves should be worn throughout the entire procedure.

1. Fixation and Storage Prior to fixation, embryos must be rinsed free of yolk or culture media with PBS. Embryos (stages 4-20) can be fixed directly on the filter-paper rings in 4% paraformaldehyde in PBS for 30-60 min (depending on the developmental stage) at room temperature. Tissue explants rinsed free of medium with PBS can be fixed on chamber slides in 4% paraformaldehyde in PBS for 20 minutes at room temperature. Following removal of paraformaldehyde, embryos or explants are stored in 70% ethanol at -20°C at least overnight. For longer storage, embryos on rings should be dissected free of extraembryonic membranes. We have stored embryos or explants in 70% ethanol at -20°C for up to 2 months.

2. Making Riboprobes Digoxigenin UTP-labeled riboprobes are generated using the Boehringer Mannheim Genius 4 system according to the instructions provided in the kit.


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Sense and antisense riboprobes are synthesized from linearized vectors containing the desired cDNA cloned in frame between two different promoters (for example, SP6 and T7).Probes are resuspended in hybridization buffer and stored at -20°C. Probes can be stored for a few months and reused up to eight times. C . Whole-Mount in Situ Hybridization

We use the method developed by Continho et al. (1992, 1993) with several modifications (see Yutzey et al., 1994). Proteinase K digestion at 37°C should be adjusted according to the stage of the embryos being used. Embyros at stages 4-9 can be incubated for 3-5 min, while older embryos can be incubated for 5-10 min.For hybridization detection, we use a 1:2000 dilution of antidigoxigenin antibody (Boehringer Mannheim) at 4°C overnight. The antibody is detected using either the Genius detection system or BM purple AP-substrate precipitating (both from Boehringer Mannheim). The BM purple seems to give less background staining than the Genius system. Figure 4 shows stage 9 (A) and stage

Fig. 4 Whole-mount in situ hybridization of chick embryos using a digoxigenin-labeledantisense riboprobe to VMHC1. (A) Stage 9 embryo showing differentiatedcardiomyocytes in the unfused cardiac primordia above the anterior intestinal portal. (B) Stage 11 embryo showing differentiated cardiomyocytes throughout the linear, beating heart tube. Embryos are shown with anterior at the top.

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11 (B) embryos reacted with the antisense digoxigenin-labeled riboprobe for ventricular myosin heavy chain (VMHC1; Bisaha and Bader, 1991). At stage 9, the bilaterally located cardiac precursors have initiated myosin heavy-chain gene expression,but have not yet fused in the midline. By stage 11,the heart primordia have fused, and the tubular heart is beating. Myosin gene expression is detected in all differentiated cardiomyocytes. D. Explant in Sit# Hybridization The solutions and procedure used are essentially the same as for whole-mount in situ hybridization, with a few modifications. The volume used for all washes and incubations is 200 pl. Because the chamber slides are made of polystyrene, the xylene step is omitted. Instead, following dehydration, the explants are incubated in a second 100% ethanol wash for 30 min and then rehydrated. The proteinase K incubation is done at room temperature for 3-5 min. All 63°C incubations are done in a humid chamber with circulating air. The incubation times for washes following probe removal can be halved without an increase in background. The antidigoxigenin antibody incubation may be done overnight at 4°C or for 2 hr at room temperature, allowing for color detection the same day. Figure 5 shows a cultured explant of cardiogenic mesoderm plus endoderm removed from the embyro at stage 5 and reacted with the Vh4HCl antisense riboprobe demonstrating that the explant has differentiated in culture.

Fig. 5 I n situ hybridization of cultured explant of cardiogenic mesoderm plus endoderm removed from the embryo at stage 5. Reactivity with the digoxigenin-labeledVMHCl riboprobe demonstrates that the explant has differentiated in culture.


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E. I n Situ Hybridization Using %Labeled Probes The procedure used by Cox et al. (1984) may be used with some modifications. Whole chick embryos are fixed in 4% paraformaldehyde in PBS overnight at 4°C. Fixed embryos are dehydrated in a series of ethanol washes and embedded in paraplast. 8-pm sections are mounted on poly-L-lysine (50 pg/ml, Sigma) coated slides. Hybridization conditions and autoradiography are as described in Bisaha and Bader (1991).

References Antin, P. B., Taylor, R. G., and Yatskievych, T. (1994). Precardiac mesoderm is specified during gastrulation in quail. Dev. Dyn. u)o, 144-154. Bader, D., Masaki, To, and Fkhman, D. A. (1982).Immunochemicalanalysis of myosin heavy chain during avian myogenesis in vivo and in vitro. J. Cell BioL 95,763-770. Bisaha, J. G., and Bader, D. (1991).Identification and characterization of a ventricular-specific avian myosin heavy chain, VMHC1: Expression in differentiating cardiac and skeletal muscle. Dev. Biol. 148,355-364. Clark, W . A., and Fischman, D. A. (1983).Analysis of population cytokinetics of chick myocardial cells in tissue culture. Dev. Biol. 97, 1-9. Continho, L. L., Morris, J., and Ivarie, R. (1992).Whole mount in sim detection of low abundance transcripts of the myogenic factor qmfl and myosin heavy chain protein in quail embryos. Biotechniques l3,722-724. Continho, L. L., Moms,J., Marks, H. L., Buhr, R. J., and Ivarie, R. (1993).Delayed somite formation in a quail line exhibiting myofiber hyperplasia is accompanied by delayed expression of myogenic regulatory factors and myosin heavy chain. Development 117,563-569. Cox,K. H., Deleon, D. V., Angerer, L. M., and Angerer, R. C.(1984).Detection of mRNAs in sea urchin embryos by in situ hybridization using asymmetric RNA probes. Dev. Biol. 10% 485-502. DeHaan, R.L. (1963).Organization of the cardiogenicplate in the early chick embyro. Acta Embryol. et Morph. Exper. 626-38. DeHaan, R. L. (1%7). Regulation of spontaneous activity and growth of embryonic chick heart cells in tissue culture. Dev. Biol. 16,216-249. Gannon, M. and Bader, D. (1995). Initiation of cardiac differentiation occurs in the absence of anterior endoderm. Development 121,2439-2450. Garcia-Martinez, V., and Schoenwolf, G. C. (1993). Primitive streak origin of the cardiovascular system in avian embryos. Dev. Biol. 159,706-719. Gonzalez-Sanchez,A., and Bader, A. (1990).In vitro analysis of cardiac progenitor cell differentiation. Dev. Biol. l39,197-209. Hamburger, V., and Hamilton, H. L. (1951). A series of normal stages in the developing chick embryo. J. Morph. 88,49-92. Han, Y., Dennis, J. E., Cohen-Gould,L., Bader, D., and Fischman, D. (1992).Expression of sarcomeric myosin in the presumptive myocardium of chicken embryos occurs within six hours of myocyte commitment. Dev. Dyn. 193,257-265. Kasten, F. H. (1973).Mammalian myocardial cells. I n “Tissue Culture: Methods and Applications” (P. F. Kruse, ed.), pp. 72-81. New York Academic Press. Kokan-Moore, N. P., Bolander, D. L., and Lough, J. (1991).Secretion of inhibin BA by endoderm cultured from early embryonic chicken. Dev. Biol. 146,242-245. Linask, K. K. (1992). N-Cadherin localization in early heart development and polar expression of Na+, K+-ATPase, and integrin during pericardial coelom formation and epithelialization of the differentiating myocardium. Dev. Biol. 151,213-224.

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Linask, K. K.,and Lask, J. W. (1986).Precardiac cell migration: Fibronectin localization at mesodermendoderm interface during directional movement. Dev. Biol. 11487-101. Lough, J. W., Bolander, D. L., and Marwald, R. R. (1990). A culture model for cardiac morphogenesis. Ann. NY Acad. Sci. S,421-424. Maniatis, T., Fritsch. E. F., and Sambrook, J. (1982). “Molecular Cloning: A Laboratory Manual.” Cold Spring Harbor Laboratory, Cold Spring Harbor, NY. New, D. A. T. (1955). A new technique for the cultivation of the chick embryo in vitro. J. Embryol. EXP.Morph. 3,326-331. Orts-Llorca, F., and Gil, D. R. (1965). Influence of the endoderm on heart differentiation. Wilhelm Roux’ Archiv. Entwicklungsmechanik WS, 368-370. Rawles, M. E. (1943). The heart-forming regions of the early chick blastoderm. Physiol. Zool. 16922-42. Rosenquist, G. C. (1970). Location and movements of cardiogenic cells in the chick embryo: The heart-forming portion of the primitive streak. Dev. Biol. 22,461-475. Rosenquist, G. C. (1985). Migration of p r m d i a c cells from their origin in epiblast until they form the definitive heart in the chick embyro. Zn “Cardiac Morphogenesis” (Ferrans, Rosenquist, and Weistein, ed.). Amsterdam: Elsevier Science Publishing Co., Inc. Rosenquist, G. C., and DeHaan, R. L. (1966). Migration of precardiac cells in the chick embyro: A radioautographic study. Carnegie Znst. Wash. Publ. 625, Contrib. to Embryol. 38,111-121. Stalsberg, H., and DeHaan, R. L. (1969). The precardiac areas and formation of the tubular heart of the chick embyro. Dev. Biol. 19,128-159. Sugi, Y., and Lough, J. (1994). Anterior endoderm is a specific effector of terminal cardiac myocyte differentiation of cells from the embryonic heart forming region. Dev. Dyn. 200,155-162. Sugi, Y., Sasse, J., and Lough, J. (1993). Inhibition of precardiac mesoderm cell proliferation by antisense oligodeoxynucleotide complementary to fibroblast growth factor-2 (FGF-2). Dev. Biol. l57,28-37. Sundin, O., and Eichele, G. (1992). An early marker of axial pattern in the chick embyro and its respecification by retinoic acid. Development 114,841-852. Vakaet, L. (1970). Cinephotomicrographic investigations of gastrulation in the chick blastoderm. Arch. Biol. 81,387-426. Yutzey, K. E., Rhee, J. T., and Bader, D. (1994). Expression of the atrial-specific mysin heavy chain AMHCl and the establishment of anteroposterior polarity in the developing chicken heart. Development 120, 871-883. Yutzey, K. E., Gannon, M., and Bader, D. (1995). Diversification of cardiomyogenic cell lineages in vitro. Dev. Biol. 170,531-541. Zadeh, B. J., Gonzalez-Sanchez,A., and Bader, D. (1986). Myosin heavy chain expression in embryonic cardiac cell cultures. Dev. Bwl. 115,204-214.

CHAPTER 7

Vascular Smooth Muscle Cell Cultures Rebecca R. Pauly, Claudio Bilato, Linda Cheng, Robert Monticone, and Michael T. Crow Vascular Biology Group Laboratory of Cardiovascular Science National Institute on Aging-National Institutes of Health Baltimore, Maryland 21224

I. Introduction A. The Sources of Vascular Smooth Muscles for Culture B. Phenotypic Modulation of Vascular Smooth Muscle Cells 11. Establishment of Cell Cultures A. Commonly Used Cell Culture Solutions B. Enzymatic Isolation Methods-Medial VSMCs C. Enzymatic Isolation Methods-Neointimal Cells 111. Characterization of Cultured Vascular Smooth Muscle Cells A. Differentiation-Specific Markers B. CellMigration IV. Specialized Methods for Human Tissues A. Establishment of Cultures from Normal Human Tissues B. Establishment of Cultures fiom Human Atheromatous Tissue V. Specialized Methods to Maintain the Differentiated State A. Cocultivation of Vascular Smooth Muscle and Endothelial Cells B. The Role of the Extracellular Matrix VI. Conclusions and Perspectives References

I. Introduction A. The Sources of Vascular Smooth Muscles for Culture Vascular smooth muscle cells (VSMCs) are the predominant cell type in blood vessels of the arterial tree. They are responsible for maintaining vascular tone in response to various hormonal and hemodynamic stimuli and they are the METHODS IN CELL BIOLOGY, VOL. 52

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major cell type involved in the process of vascular repair, growth, and response to injury. Understanding how VSMCs execute these different functions is one of the keys to understanding the pathogenesis of vascular diseases and, in turn, to developing rational strategies for their treatment. The development of cell culture systems has greatly aided in this pursuit by providing a controlled environment to study the unique characteristics of these cells in isolation from other influences. A review of blood-vessel histology provides the necessary background for understanding some of the principles involved in VSMC isolation. Figure 1A shows a cross-section of an uninjured carotid artery from the rat. The vessel consists of three well-defined layers or tunica: the intima, the media, and the adventitia. The intima is defined as the layer of cells and extracellular matrix

Fig. 1 Blood vessel histology and cultured vascular smooth muscle cell morphology. (A) Crosssection of the carotid artery from a mature (6-month) rat. The lumen and vessel layers or tunica are labeled. Adv.: tunica adventitia. Arrows mark the elastic laminae that are part of the tunica media. The internal elastic lamina (IEL) and external elastic lamina (EEL)mark the boundaries of the media. The intima is not readily definable in this section, because it consists only of the endothelial cell layer lining the lumen. (B) Cross-section of balloon-injured carotid artery from a mature rat 14 days after injury. A large cellular structure (neointima) has developed in the intima. (C) Photomicrograph of cultured medial VSMCs. (D) Photomicrograph of cultured VSMCs isolated from the neointima.

7. Vascular Smooth Muscle Cell Cultures

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(ECM) that lies between the lumen and the internal elastic lamina (indicated by arrow marked “IEL”). In many vessels, including the rat carotid artery shown in Fig. lA, the intima consists only of the endothelium and the continuous basement membrane in contact with its abluminal surface (these are not visible at this magnification without specific staining). The next layer of cells and ECM is the tunica media. Its boundaries are defined by the internal (IEL) and external (EEL) elastic laminae (Figs. 1A and 1B) and contains mesenchymal cells referred to as medial VSMCs trapped within these and the other elastic laminae (indicated by the arrows). In the rat carotid artery, the tunica media consists of 4-5 elastic laminae interspersed by single layers of VSMCs. The outmost layer, the tunica adventitia (Adv in Fig. l), is composed of fibroblasts and loose connective tissue and may contain small blood vessels and nerves. In almost every medium to large artery, the elastic laminae form a mechanical barrier that separates the media from the intima and adventitia, a fact that is exploited during dissection to easily isolate the media. VSMCs are then freed from the surrounding extracellular matrix using a combination of elastase, which digests the elastic laminae, and crude collagenases, which digest the ECM surrounding each VSMC. When the blood vessel is mechanically injured and its endothelial lining removed, there is significant remodeling of the vessel. Medial VSMCs migrate across the elastic laminae and basement membrane and accumulate in the intima. There they proliferate and secrete increased amounts of interstitial matrix proteins. This new intimal structure is appropriately referred to as the neointima and the cells within it are neointimal VSMCs. Figure 1B shows a rat carotid artery 14 days after mechanical injury and the development of a substantial neointima. This is a model experimental system to study the response of VSMCs to injury, such as that occurring after balloon angioplasty. Results from these types of experiments may have ramifications for more “chronic” vascular diseases, such as atherosclerosis. What is clear from a number of experiments is that medial and neointimal VSMCs represent quite different smooth muscle cell types and that the differences that separate them persist in cell culture and throughout repeated cell passages. Morphologically,for example, cultured medial VSMCs are elongated cells (Fig. lC), whereas cultural neointimal cells are cuboidal (Fig. 1D). Methods for culturing both medial and neointimal VSMCs are described in this chapter as well as for culturing VSMCs obtained from atheromatous lesions. B. Phenotypic Modulation of Vascular Smooth Muscle Cells Since VSMCs are the only cell type present in the media of mammalian arteries, they are responsible not only for maintaining arterial wall tension but also for vascular growth, remodeling, and repair, such as that following mechanical injury (Fig. 1B). These latter functions require that these cells retain certain basic mesenchymal cell properties distinct from the differentiated properties of VSMCs. These are the abilities to synthesize extracellular matrix, to migrate,

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and to replicate. This multiplicity of functionsmay be achieved through a number of different mechanisms. Maintaining a reserve stem-cell population to respond to injury is one possible mechanism (see Majesky et al., 1992, and references therein). On the other hand, it is also clear that medial VSMCs can undergo a reversible interconversion of phenotypes that could account for the different functional roles of VSMCs. This process is referred to as phenotypic modulation. Cell-culture studies of arterial medial smooth muscle cells have been instrumental in the development of the notion that VSMCs can, in fact, modulate along a continuum of phenotypes. In this simplest conception, this notion identifies two extreme VSMC states. VSMCs in the synthetic state have a fibroblast-like appearance, proliferate readily, and synthesize increased levels of various extracellular matrix components, particularly the subcomponentsof interstitial-type ECMs such as fibronectin, collagen types I and 111, and tropoelastin. To support this phenotype, these cell contain a large amount of free ribosomes, rough endoplasmic reticulum, Golgi complexes, and mitochondria, but few myofilaments. Contractile VSMCs, on the other hand, have a muscle-like or spindle-shaped appearance and a welldeveloped contractile apparatus resulting from the expression and intracellular accumulation of thick and thin muscle filaments. These cells actively regulate blood vessel tone in response to chemical and mechanical stimuli. (A detailed review of the properties of synthetic and contractile VSMCs is in Campbell and Campbell, 1981, and Thyberg et al., 1990.) Contractile cells can readily assume a synthetic phenotype. They do so within 1-2 passages after they are placed in culture as well as when the vessel is injured. This modulation from a contractile to syntheticphenotype appears to be an important early event in the pathogenesis of many vascular diseases, such as atherosclerosis (Mosse et al., 1986) and may be a prerequisite for the migration of VSMCs that form the neointima following balloon catheter injury of the vessel (Pauly et al., 1994a).

11. Establishment of Cell Cultures There are two general methods for the preparation of primary cultures of vascular smooth muscle cells. One involves direct isolation of cells from enzymatically digested vessels (enzymatic isolation method), and the other involves isolating cells that migrate out from small tissue samples (explant method). These preparations yield smooth muscle cell cultures that at least initially have quite distinct properties (see Section 111). We will describe an enzymatic isolation procedure for VSMCs from elastic vessels such as the rat thoracic aorta. For more muscular vessels, such as the rat caudal artery, the digestion with elastase may be omitted. Section IV describes a variety of explant methods for use in the isolation of human VSMCs.


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A. Commonly Used Cell Culture Solutions

Many different media have been used for the cultivation of VSMCs without any obvious differences in the behavior of the cells. We describe below in detail the composition of the media we use for cultivation of VSMCs, which is based on Dulbecco’s modified Eagle’s medium (DMEM). 1. Supplemented growth medium. DMEM with high glucose (4 g/liter) (Gibco/ BRL No. 11995) is supplemented to a final concentration of 1mMnonessential amino acids (GibcoiBRL No. 11140 is 100X stock), 50 pg/ml penicillin, 50 pg/ml streptomycin, 100 pg/ml neomycin (PSN, GibcoiBRL No. 15640 is 100X stock), and 2 mM glutamine (GibcoiBRL No. 25030 is 100X stock). 2. Complete growth medium. Supplemented growth medium with 10% heatinactivated fetal bovine serum (FBS) (GibcolBRL No. 16140 or other

vendors). 3. Serum starvation medium. Supplemented growth medium with either 0.5% FBS or 1 pg/ml insulin, 0.67 nglml sodium selenite, 0.55 pg/ml transfemn, and 11 p g / d sodium pyruvate (GMS-G supplement, GibcolBRL No. 41400). 4. Supplemented growth medium with Hepes and PSF (DMEM/Hepes). DMEM (low glucose (1 g/liter) (GibcoiBRL #12320 or MA Bioproducts #12-708B) is supplemented to a final concentration of 2 mM glutamine (GibcoiBRL #25030 is l00X stock), 100 U/ml penicillin, 100 pg/ml streptomycin, and 0.25 pglml amphotericin B (Fungizon) (GibcoiBRL #15245 is 1OOX stock). 5. Complete growth medium with additional bicarbonate and PSF (DMEMI bicarb/FBS). DMEM (high glucose (4 @liter)) (GibcoiBRL No. 11995) is supplemented to a final concentration of 1 mM non-essential amino acids (GibcoiBRL No. 11140 is l00X stock), 2 mM glutamine (GibcoiBRL No. 25030 is 1OOX stock), 100 U/ml pencillin, 100 pg/ml streptomycin, and 0.25 pg/ml amphotericin B (Fungizone) (GibcoiBRL No. 15640 is lOOX stock), and an additional 0.75 m g / d sodium bicarbonate. 6. Cryopreservation medium. Supplemented growth medium with 20% FBS and 7.5% dimethyl sulfoxide (Sigma Chem. No. D-2650; use only unopened vial to avoid oxidation products). 7. TrypsidEDTA solution. 0.25% trypsin and 1 mM EDTA in phosphatebuffered saline (GibcoiBRL No. 25200). Note: Although this commercial trypsin solution works, we routinely use a modified solution containing 3 mM EDTA as this is more effective in removing cells for subcultivation. Also note that the concentration of trypsin used is about 5 times that used for other mesenchymal cell lines. This higher concentration is particularly needed for medial VSMCs and especially for the removal from the tissue culture container of postconfluent, serum starved VSMCs that exhibit a

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“hill and valley” appearance. Neointimal VSMCs can be passaged using either 0.25% trypsidl mMEDTA or 0.05% trypsin/0.53 mMEDTA (Gibco/ BRL No. 25300).

B. Enzymatic Isolation Method-Medial VSMCs 1. Material and reagents ‘ a. Autoclaved tools: Three sets of autoclaved scissors, forceps, and beakers, glass transfer pipets, and a stainlesssteel mesh in a 13-mmSwinexfilter (Millipore Corp., Bedford, Massachusetts). b. Sterile plasticware: 35-mm tissue-culture dishes, 15-ml plastic centrifuge tubes, and 12-cc syringes. c. Modified HBSS: Hanks balanced salt solution (HBSS)without CaCl, and MgC1, (GibcoBRL, No. 14170) containing 50 pg/ml penicillin, 50 pg/ml streptomycin, 100 pg/ml neomycin (GibcoBRL No. 15640 is l00X stock), and 0.25 pg/ml amphotericin B (Fungizone) (GibcoBRL, No. 15295 is a 50X solution). d. 2 mg/ml collagenase I (CLS I, Worthington Biomedical, Freehold, New Jersey) in modified HBSS.This is a mixture of collagenases, caseinases, clostripain, and trypsin. e. 2 mg/ml collagenase I1 (CLS 11, Worthington Products) and 0.5 mg/ml porcine elastase (Calbiochem Corp, No. 324689) in modified HBSS.

2. Procedure for cell isolation a. Wistar rats (150-200 g) are sacrificed by COz asphyxiation. The skin is wiped with 70% ethanol and cut on the left ventral side to expose the area of the thoracic aorta (left thoracotomy). b. Using another set of autoclaved scissors and forceps, remove the vessel from the origin of the left subclavian artery to its point of insertion in the diaphragm. Rinse it four times with modified HBSS. c. Remove loosely adhering adventitia from the medial layer by gross forcep dissection. (The adventitia is white and distinguished from the media, which is off-whiteltan.) d. Predigest the vessels for 30 min at 37°C in a 15-ml tube containing 2-3 ml of 2 mg/ml collagenase I solution. e. Rinse the vessels with fresh HBSS and then remove the remaining adventitia, which will appear white and fluffy, under a dissecting microscope. Transfer the vessels to a 35-mm dish and add 2-3 ml complete growth medium. Incubate overnight at 37°C. (This overnight step revives the cells and improves the viability of cells isolated from the vessel.) f. The next day, rinse the vessel twice in modified HBSS and then place it in a 15-ml tube containing 2-3 ml of the collagenase IUelastase solution.


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Incubate at 37°C for 2 hr, triturating the digest with a glass transfer pipet every 30 min. g. Stop the digestion by adding 10 ml of complete growth medium. Optional: The digest can be filtered through a sterile stainless-steelmesh fitted to a 13-mm Swinex filter (Millipore Corp., Bedford, Massachusetts) to remove large particles. h. Spin the cell suspension at lo00 rpm for 10 min in a benchtop centrifuge. Discard the supernatant, resuspend the cells in 2-5 ml of complete growth medium, and determine cell yield with a hemocytometer. i. A good preparation should yield 1-3 X lo6 cells/aorta. Because plating efficiency may vary, plate the cells from one vessel into two 35-mm tissue culture dishes in complete growth medium. Plating efficiencycan be increased by precoating the 35-mm dishes with extracellular matrix (see later procedure). This, however, may influence the behavior of the cells (see Section V). 3. Preparation of extracellular matrix coatings a. Thin layers of extracellular matrix (ECM): Collagen I (rat tail) is purchased as a concentrated (-3 mg/ml) stock solution in 0.02 M acetic acid (Upstate Biotechnology, Lake Placid, New York, No. 08-115). It is diluted in 0.02 M acetic acid to a final concentration of 100 pg/ml. Mouse type IV collagen is purchased as a concentrated solution in 50 mM Tris HC1 (pH 7.2) and 0.5 M NaCl (Gibco/ BRL No. 33018-011) and is diluted to 100 pg/ml in the same buffer. Fibronectin and laminin are purchased as concentrated stock solutions (GibcoBRL Nos. 12153-011and 23017-015, respectively) and diluted in Dulbecco’s PBS to a final concentration of 100 pg/ml. Sufficient ECM solution is added to coat the dishes at 20 pg/cm2culture surface and allowed to absorb at RT for 1-4 hr. The dishes are then rinsed twice with PBS and blocked with DMEM containing 0.1% bovine serum albumin (BSA). b. Basement membrane gel: Basement membrane from the EHS tumor is prepared inhouse as previously described (Kleinman et aZ., 1986). It is also available under the trademark Matrigel from Collaborative Research Products (Bedford, Massachusetts).‘To form a gel, the basement membrane preparation is thawed on ice, mixed with precooled pipets, and added to precooled culture dishes at 20-50 pYcm2. The gel is then placed in the 37°C incubator overnight. 4. Subcultivation of VSMCs a. Cell cultures should be examined daily, fed at 3-day intervals, and subcultured when the culture is 90% confluent or less (this usually takes 5-7 days). b. To subculture, the medium is removed, and the cells are washed with Dulbecco’s phosphate buffered saline and then incubated for 3-5 minutes with trypsiniEDTA and observed for cell detachment. c. When early signs of detachment (cell rounding, retraction) are evident, the trypsin solution is neutralized with 5 ml of complete growth medium containing serum. The solution is pipetted back and forth to assist the removal of cells from the culture plate and then subjected to centrifugation at lo00 rpm for 5 min in a benchtop centrifuge.

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d. The resulting cell pellet is then dispersed with complete growth medium and counted with the aid of a hemocytometer. Cells are routinely seeded at a density of 3 X Id cells/cm2. e. Medial VSMCs do not readily growth arrest. In fact, growth arrest (as measured by H3-thymidine uptake) does not take place for up to 5 days after serum withdrawal, even when the cells are confluent at the time of starvation. During this time, medial VSMCs exhibit a multilayered appearance known as the “hill and valley” effect. 5. Cell cryopreservation methods a. A 90% or less confluent plate is trypsinized as described earlier for subcultivation. After centrifugation, the cell pellet is suspended in 1ml of cryopreservation medium. This cell suspension is rapidly transferred to a 1.5-mlcryopreservation vial (Nunc) and rapidly transferred to a Nalgene Cryo-Freezer container placed in a -70°C freezer. (The Nalgene container is designed to reduce the temperature at a rate of 1degreelminute when placed in a -70°C freezer). The next day, the cells are transferred to a liquid nitrogen cryogenic freezer for longterm storage. b. To recover frozen cells for cultivation, rapidly thaw the vial to 37°C and transfer the thawed solution to a 100-mm plate containing 10 ml of complete growth medium. The medium is changed after 18 hr.

C. Enzymatic Isolation Methods-Neointimal Cells 1. Carotid injury (see Jenkins et al., 1997). Six-month-old male Wistar rats are routinely used. The animals are anesthetized with intraperitoneal injections of sodium pentobarbital (400 mgkg body weight (BW)), ketamine, 2 mg/kg BW, and xylazine (8 mgkg BW). Balloon catheter injury of the left common carotid artery is performed with a 2F Fogerty balloon catheter (Baxter Scientific, McGraw, Illinois) inserted through the external carotid artery and advanced the length of the common carotid artery to the aortic arch. The balloon is then inflated and passed three times along the length of the artery. The balloon is then removed and the excision site at the external carotid artery ligated. Fourteen days following injury, Evans blue dye is injected intravenously 30 min prior to sacrifice of the animal in order to determine the completeness of endothelial removal by the injury technique and to localize the neointima. (Denuded vessels stain blue with this dye.) 2. Carotid neointimal cell isolation. Prepare the artery as described earlier for the aorta. Dissect the blue staining neointima away from the nonstaining underlying media with the aid of a dissecting microscope. Place the neointima in complete growth medium in a humidified COz incubator overnight at 37°C. The next day, place the neointima in 2-3 ml of the collagenase/ elastase solution used to digest medial cells (the elastase is probably not


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necessary). Incubate at 37°C for 1 hr, triturating the sample every 1520 min. Stop the digestion by adding 10 ml of complete growth medium, centrifuge to collect the cells, resuspend in complete growth medium, and place into two or three 35-mm tissue culture dishes. When preparing parallel cultures of the medium from contralateral uninjured vessels, use the procedures described earlier for the aorta and plate the cells from a single vessel in a single 35-mm culture dish, as the yield of cells and their rate of proliferation is likely to be lower than the neointimal cells. Neointimal VSMCs exhibit a cellular morphology in culture that is distinct from that of medial VSMCs. Neointimal cells exhibit a cuboidal shape in culture, whereas medial cells are more spindle-shaped (Figs. 1D and lC, respectively). These morphologic differences, as well as differences in the expression of specific genes (Majesky et al., 1992), persist for a number of passages in cell culture. Neointimal cells also do not exhibit the “hill and valley” effect after reaching confluence. Instead, they remain as a distinct monolayer on the culture dish. Interestingly, in their morphology and gene expression patterns neointimal VSMCs resemble VSMCs isolated during early postnatal (pup) development. It has been suggested that neointimal VSMCs may arise from a residual “pup” cell population in the mature vessel (Majesky et al., 1992).

111. Characterization of Cultured Vascular Smooth Muscle Cells A. Differentiation-Specific Markers

A number of markers exists for assessing the origin of cultured cells as smooth muscle cells and assessing their state of differentiation. Following is a selected list of such markers that are expressed by VSMCs in the intact vessel and that are lost to varying extents during phenotypic modulation in culture or during vascular disease and injury. These changes for each of the markers are summarized in Table I. Although the vast majority of VSMCs in a large number of vessels exhibit a uniform pattern of reaction, significant heterogeneity in the expression of these markers can exist in normal vascular tissues (Frid et al., 1992). 1. Actin

Three different actin isoforms, designated cr, 6, and y, are detected in VSMCs by isoelectric focusing. The a-actin isoform is the product of a smooth musclespecific gene, the 6 is a product of the ubiquitous cytoplasmic (nonmuscle) actin gene, and the y-actin isoforms are products of two separate genes-nonmuscle y-actin and smooth muscle-specific y actin. During development, when VSMCs exhibit a “synthetic” phenotype, the main actin isoform expressed is 6 (Gabbiani et al., 1984;Kocher and Gabbiani, 1987). As the vessel matures, there is decreased

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Table I Expression of Smooth Muscle Cell Markers in Mature and Injured Blood Vessels and in Cell Cultures' Marker Actin

Mature vessel Injured vessel Cell culture

a

++

B

+/-

Myosin heavy chains SM1 SM2 MHCA Calponin Caldesmon h (120-150 kDa) 1 (70-80 kDa) viculin Meta-vinculin

++++ ++

+/-

++++ ++++ ++ ++

+

References

+ b.c

Gabbiani ef al. (1984) Kocher and Gabbiani (1987) Glukhova ef al. (1988) Babij et al. (1992)

++

++

+I-

+I-

Rovner et al. (1986) Glukhova ef al. (1988) Babij et al. (1992) Frid et al. (1992) Shanahan et al. (1993)

+++

++++ +I-

+ ++ ++ -

+++

++++c

+I-d

+I-

++ ++ -

Glukhova ef al. (1988) Glukhova et al. (1988)

There may be significantcellular heterogeneity with respect to the expression of these markers. Refer to the references for details. In contrast to the protein levels, the level of a-actin mRNA is actually higher in cultured cells than in the mature tissue. 'The levels of these proteins are influenced by the conditions of cell cultures (e.g., density, growth factors). Unlike other differentiation markers, the level of calponin protein does not decrease for several cell passages.

p-actin and increased smooth muscle-specifica-actin isoform expression (Kocher and Gabbiani, 1987). Smooth muscle a-actin expression persists in VSMCs even after extensive passaging in culture. Although smooth muscle a-actin has been detected in nonmuscle tissues (see reference within Frid et al., 1992),the presence of smooth muscle a-actin in cultured cells is a fairly reliable indication that the cell is of smooth muscle origin. A monoclonal antibody is commercially available for routine characterization of cultures (Sigma Chem. Co, No. A 2547). The use of smooth muscle a-actin as a marker for assessing cell differentiation, however, is subject to a number of caveats. When mRNA levels for smooth actin are compared between intact vessels and cultured VMSCs, there is, contrary to expectation, a large increase in smooth actin mRNA in the cultured VSMCs compared to the intact tissue (see Fig. 6 in Babij et al., 1992). Furthermore, proliferating (synthetic) VSMCs express higher levels of smooth a-actin mRNA than growth-arrested,confluent VSMCs. This is because total actin mRNA levels are, in general, increased in replicating cells. If instead of smooth a-actin mRNA levels, the ratio of a- to p-actin mRNA is compared, cultured VSMCs have a


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much lower ratio than the intact vessels and conditions that favor the synthetic state favor a further lowering of the ratio (Owens et al., 1986). To measure both isoforms simultaneously, a probe that encodes the translated region of either isoform should be used. This probe will react with both isoforms, because there is little divergence in the translated region of different actin isoforms, even at the nucleotide level. On a Northern blot, the a isoform runs faster than the j3 isoform.

2. Myosin-Heavy Chains Molecular cloning has established that there are two smooth muscle-specific myosin heavy chain (SMHC) cDNAs expressed in intact blood vessels and that these cDNAs are encoded by a single gene. Differential RNA processing of the single gene transcripts gives rise to a mRNA encoding a 204-kDa protein (SM1) and a 200-kDa protein ( S M 2 ) (Babij and Periasamy, 1989). Ribonuclease protection assays indicate that SM1 and S M 2 mRNA are coexpressed in intact vascular tissue at a ratio of approximately 4 :1. In both primary and passaged cultures of smooth cells, S M 2 mRNA levels decrease by over 80%, whereas those of SM1 decrease by only 30-40% compared to levels in the intact tissue. In these same cultures, nonmuscle MHCs increase substantially. Analysis of SM1 and SM2 protein using specific antibodies yields essentially the same results. Because SM1 expression persists even in passaged VSMCs, it is a useful marker for identifying cells of smooth muscle origin. SM2 expression, on the other hand, appears to be a fairly useful marker of the state of differentiation and in primary cultures appears to be sensitive to the growth state. Both isoforms are readily detected by immunoblotting using a commerciallyavailable monoclonal antibody, SMMS1 (Frid et al., 1992; Sigma Chem. Co., No. M7786). In contrast to the foregoing, quite different and potentially deceptive results are obtained when mRNA levels for SM-MHCs are analysed by Northern blotting using “smooth muscle-specific” cDNA probes. Babij and co-workers (1992) report that the MHC hybridization signal from subconfluent primary and passaged VSMCs, following incubation with a cDNA probe that does not recognize nonmuscle MHCs, is much greater than that of intact vascular tissue. Apparently, the probe reacts with an as yet unidentified MHC mRNA that is not present in nonmuscle tissues but is present in cultured VSMCs. This isoform may be that of an embryonic SM-MHC identified in developing vessels and atherosclerotic lesions (Kuro-o et al., 1991). 3. VinculidMeta-vinculin Vinculin is a 130-kDa protein that is localized at a variety of cellular adhesive structures in all muscle and non-muscle cells examined. Meta-vinculin is a 150-kDa protein that is immunologically identical to vinculin but is restricted in its expression to muscle tissue. In the human aorta, for example, as much as 40%

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of total vinculin is meta-vinculin. Meta-vinculin expression is rapidly lost in cell culture. Meta-vinculin content is readily measured by immunoblotting with a commercially available antibody (Sigma Chem. Co., No. V9131). This antibody detects vinculin and meta-vinculin, which are distinguishable by their different molecular weights. 4. Calponin

Smooth muscle-specific (basic) calponin is a troponin-like molecule that is present in most vertebrate smooth muscles. It binds to the actidtropomyosin thin filament and results in inhibition of myosin MgATPase activity. It is encoded by a 1.3-kb mRNA, which in the chicken produce two isoforms, a and p, both of which are smooth muscle specific (Takahashi and Nadal-Ginard, 1991).Smooth muscle-specific calponin expression is rapidly downregulated when VSMCs are placed in culture. cDNA probes for calponin are available from the authors and a specific antibody is commercially available (Sigma Chem. Co, No. C 2687). A non-muscle or acidic isoform of calponin has also been identified (Applegate ef al., 1994). Its expression is not smooth muscle-specific. Acidic calponin is expressed by VSMCs even into late passages.

5. Caldesmon Caldesmon is the major calmodulin- and actin-binding protein in both smooth muscle and nonmuscle cells, modulating actin-myosin interactions and contraction. Immunochemical analysis reveal heterogeneity of caldesmon expression in different tissues. Smooth muscles in vivo express caldesmons in the 120150 kDa range (h-caldesmon), whereas cultured fibroblasts and VSMCs express l-caldesmons in the 71-77 kDa range (Ueki er al., 1987). Caldesmon content is readily measured by immunoblotting with a commercially available antibody (Sigma Chem. Co.,No. C 4562). This antibody detects both l-caldesmon and hcaldesmon, which are distinguishable by their different molecular weights. As markers that are independent of the phenotypic state or growth conditions and indicative only of cell origin, smooth a-actin and smooth muscle (SM)specific MHC expression (SM1, in particular) are the most reliable. Activation of these two markers is likely to require cell type-specific transcriptional activation. The best markers for assessing differentiation status, such as meta-vinculin, caldesmon, and SM2 SM-MHC, on the other hand, appear to accumulate by differential splicing of a gene product that is expressed in both smooth muscle and nonmuscle cell types. The exception to this may be calponin, which is highly differentiation-dependentin its expression,and for which there are no nonmuscle analogues that result from differential splicing. Expression of calponin, however, generally persists longer in culture than any of the other differentiation-specific markers (Pauly, Monticone, and Crow, unpublished observations). It is tempting to speculate that the control of smooth muscle cell differentiation may be governed more by RNA splicing factors rather than by transcription factors.


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B. Cell Migration In addition to the phenotypic conversion of VSMCs from the contractile to the synthetic state that occurs either at the time of primary culture or within the first few cell passages, there is also another change in VSMC behavior that occurs with later passage. This is the ability to migrate toward chemoattractants (i.e., chemotaxis). In cultures established by enzymatic isolation methods from 3- to 6-month-old rats, this transition occurs between the 5th and 7th passages. In contrast, medial VSMCs isolated by explant techniques or neointimal cells isolated by enzymatic isolation migrate almost immediately upon their establishment in culture (Pauly et al., 1994b). This indicates that in addition to phenotypic modulation to a synthetic state, additional changes must occur in VSMCs before they are capable of migrating. The study of the factors controlling cell migration is, therefore, of fundamental importance in understanding the pathogenesis of vascular disease. A method for studying cell migration of VSMCs toward a defined chemoattractant using a modified Boyden chamber (Fig. 2A) is described next. Cells added to the upper chamber attach to a ECM-coated filter and then migrate to the underside of the filter in response to a chemoattractant placed in the bottom chamber. By interposing an extracellular matrix barrier in place of ECM-coated filters, this assay has been adapted to study the invasive characteristics of VSMCs (Pauly et al., 1994a). 1. Preparation of Filters. Nucleopore PVPF-coated, 8-pm pore size, 13-mm diameter filters (Neuro-Probe, Inc., Cabin John, Maryland) are used and positioned shiny side down in an open culture dish. They are then numbered with a black ballpoint pen to identify the experiment and orientation of the filter. Coating of the filters begins with 50 p1 of 0.5 N acetic acid applied to the filter, which is then allowed to dry in a sterile hood for 8-18 hr. The filter is then coated with 50 p1 of a 100pg/ml solution of collagen I, fibronectin, or other ECM molecules. This is then allowed to dry overnight in a sterile hood. 2. Preparation of VSMCs for Migration Assays. The cells utilized for migration assays are either in the proliferative or growth-arrested state prior to their addition to the migration chamber. For proliferating cells, cultures are harvested as described above when they are 60-80% confluent. For growtharrested VSMCs, the cultures are allowed to reach the postconfluent state (1-2 days after reaching complete confluence) and then placed in serumstarvation medium for 5 days. For medial VSMCs, this excessive period of serum starvation is required for these cells to cease incorporation of H3thymidine and achieve growth arrest. During this postconfluent period, medial VSMC cultures will assume a “hill and valley” morphology. 3. Preparation and Operation of Migration Chambers. The modified Boyden migration chamber system routinely utilized is a two-stage Plexiglas con-

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Fig. 2 VSMC migration. (A) Diagram of the modified Boyden chamber used to study VSMC migration toward a chemoattractant (chemotaxis). (B) Results of migration assays using medial VSMCs that were proliferating (cross-hatched bars) or growth arrested (open bars). Migration is expressed as the average number of cells present in a 4OOX field. The results show that migration only readily occurs in proliferating VSMCS and that this migration is dependent on the establishment of a gradient of chemoattractant (see Pauly er aL, 1995, for further details).

tainer (Neuro-Probe, Inc). The lower chamber is overfilled with supplemented growth medium (220 pl) containing a chemoattractant (typically 10 ng/ml PDGF BB) to create a dome. The ECM-coated filter (dull and numbered side up) is then carefully positioned on top of the “dome” to avoid air bubbles and then fastened to the lower chamber with a screwcap holder. The cells to be studied are added to the top of the filter (2 X 105 in 200 pl of supplemented growth medium) and the upper chambers then filled with an additional 650 pl of supplemented growth medium containing 0.1% BSA. The chambers are then placed in a humidified C 0 2

7. Vascular Smooth Muscle Cell Culturea

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incubator at 37°C for 4 hr. After 4 hr, cells that have migrated will pass through the pores and will remain attached to the lower filter surface. 4. Analysis of Migration Results. To prepare the filters for analysis, the filters are removed from the chamber and pinned numbered side down onto a surface of Paraplast embedding medium melted into a Petri dish. The filters are then fixed and counterstained with the Hema-3 system (CurtinMatheson, Houston, Texas, Cat. #122911). After a rinse in distilled water, the filters are positioned numbered side up (lower chamber side down) onto a glass side. The cells attached to the top of the filter, which represent nonmigrating cells, are then removed by gentle wiping with a moist cotton swab. Filters are then permanently mounted with a single drop of Cytoseal (Stephens Scientific) followed by a coverslip pressed firmly into position. The filters can then be analyzed with an image analysis system or manually by counting the number of cells in random 4OOX (high-power) fields. At least four fields are counted per filter and each experimental chamber is run in triplicate. Results are presented as the average number of cells per high-power field. Figure 2B shows the results of a number of experiments using passage 8-12 medial VSMCs isolated from the rat thoracic aorta. Compared to BSA, proliferating VSMCs migrated robustly toward PDGF BB. Migration was totally dependent on the establishment of a gradient. If BSA was placed in the bottom chamber or PDGF was placed in both upper and lower chambers, migration was substantially reduced. The rate of migration in the chamber is initially slow as cells first attach to the ECM-coated filter. To avoid this delay and ensure that the starting numbers of attached cells are similar, we have taken advantage of the requirement for a gradient to first allow a period of cell attachment (1-2 hr) in which PDGF is present in both chambers. Following this, PDGF is then removed from the upper chamber, re-establishingthe gradient for cell movement. Note also in Fig. 2B that growth-arrested cells do not migrate toward PDGF BB. Using this migration system, we have shown that the failure of these cells to migrate is due to alterations in their ability to activate a key step in the migration program, namely the calciudcalmodulin-dependent protein kinase I1 (Pauly et aZ., 1995).

IV. Specialized Methods for Human Tissues These are all explant methods and involve placing minced fragments of tissues into culture and waiting for cells to migrate from them. The migrating cells are then separated from the tissue fragments. A. Establishment of Cultures fiom Normal Human Tissues

Two methods are presented, one for establishment of smooth muscle cells from the saphenous vein, the other for smooth muscle cells from aortic tissue.

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1. Smooth Muscle Cell Preparation from the Human Saphenous Vein. (This method was contributed by Drs. Peter Libby and Marysia Muszynski, Vascular Medicine and Atherosclerosis Unit, Cardiovascular Division, Department of Medicine, Brigham and Women’s Hospital, Harvard Medical School, Boston.) a. Rinse the vein in a sterile Petri dish with modified DMEM/Hepes (see Section 1I.A) containing 10% heat-inactivated FBS to remove blood. Rinse the inside of the vein with a 10-cc syringe and cannula. b. Cut the vein transversely into small pieces (about 2 inches each). Do not use the section of vein containing sutures that may have been used to ligate tributaries during harvest of the vein. Cut the segments longitudinally to expose the inner surface of the vessel. c. Using forceps, strip the vessel of its intima, then ilip the vessel over and remove the adventitia using a scraping action with a scalpel (No. 10 blade). d. Cut the cleaned medial tissue into small pieces and place onto the scratched surface of a 35-mm Petri dish. Spread sections flat on the plate. e. Let the dishes sit in the hood for 30-60 min to allow the explants to attach, then add a few drops of modified DMEMiHepes + 10% FBS directly on the tissue pieces to moisten them. Transfer to a humidified 5% COzincubator at 37°C. f. On the following days, gently add a small amount of medium each day so that there is a total 3-4 ml/dish by the end of the first week. Once the explants have firmly attached, change medium twice a week. Cells should migrate from the tissue within 2 weeks. g. To harvest, remove tissue pieces and retrieve attached cells with 0.25% trypsin/EDTA. h. To subcultivate the explanted cells, passage at a 1:3 split using 0.25% trypsin/EDTA as described in Section ILB. The medium for growth and subcultivation is DMEM/Hepes 10% heat-inactivated FBS. 2. Smooth Muscle Cell Preparation from Human Aortic Tissue. (This method was contributed by Dr. Zhihe Li, Laboratory of Cardiovascular Science, NIA-NIH.) a. Segments of human thoracic or abdominal aorta are removed and rinsed several times in Hank’s balanced salt solution containing 100 Ulml penicillin, 100 &ml streptomycin, and 0.25 pg/ml amphotericin B (100X stock, Gibco/ BRL No. 15245). b. The intima is stripped off with forceps under a dissecting microscope. A 0.5-1 mm thin layer is then peeled off the vessel. This layer contains about 60% of the tunica media. c. The tissue strips are rinsed in DMEM (high glucose) (GibcoBRL No. 11995) and then cut into 1 mmz fragments with a pair of surgical scalpels. d. The explants are placed in a 60-mm Petri dish and covered with 2 ml of DMEMlbicarbPSF (see Section 1I.A) containing 20% FBS and positioned in a humidified 5% COz incubator at 37°C.

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e. To allow the explants to attach to the plastic surface, the culture dish is not disturbed for the first week. From the second week, the explants are fed twice weekly with 4 ml of culture medium/dish. f. When a large clone surrounding the explant can be seen, the culture is rinsed with Hanks solution and treated with 0.25% trypsidl mM EDTA (Gibcol BRL No. 25200) at 37°C until cells begin to detach from the plastic. After addition of a few milliliters of medium containing FBS, the cell suspension is collected and subjected to centrifugation (500g, 5 min), and the pellet is resuspended in the complete growth medium and reseeded into culture dishes. Routine passaging is performed as described in the method for the saphenous vein. B. Establishment of Cultures fiom Human Atheromatous Tissue

(Details of this method were contributed by Drs. Peter Libby and Marysia Muszynski, Vascular Medicine and Atherosclerosis Unit, Cardiovascular Division, Department of Medicine, Brigham and Women’s Hospital, Harvard Medical School, Boston; see Libby et al., 1988). The preparation of primary cultures of plaque-derived cells by ordinary explant techniques presents a unique challenge because the small, irregularly shaped, and often lipid-laden fragments of tissues tend to float and lose their contact with the substratum when covered with culture medium. To avoid this, a modification of the explant method has been developed in which a collagen gel is cast around the minced fragments of vascular tissue. This method provides a threedimensional matrix for cell attachment and growth and a favorable environment for cell egress from the tissue fragments. Although cell types other than VSMCs (e.g., macrophages) may also migrate from the tissue into the collagen gel, the cell culture medium used favors VSMC growth. 1. Materials and Reagents a. Bovine dermal collagen treated with pepsin dissolved in 12 mM HCl. This is available from Collagen Corporation, Palo Alto, California, under the trademark Vitrogen 100. b. Modified Earl’s balanced salt solution (lox). This is prepared in-house from the basic components in deionized, distilled pyrogen-free water (Gibco/ BRL Laboratories, Cat. No. 15230). Per liter, add 2 g CaC12 (anhydrous), 4 g KCl, 978 mg MgS04 (anhydrous) or 2 g MgS04*7H20,83g NaCl, 10 g D-glucose, and 10 mg phenol red. c. 1 N sodium hydroxide. d. DMEM/Hepes. See Section 1I.A for precise formulation. e. 2 m g / d collagenase solution (CLS I, Worthington Biochemical Corporation, Freehold, New Jersey) in DMEM/Hepes. 2. Procedure for Cell Isolation a. Prepare specimens for cultivation by mincing into 1-2 mm3 fragments.

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b. Suspend the minced fragments in 1.35 ml DMEM/Hepes containing 10% heat-inactivated FBS. c. In another tube placed in an ice bucket, add 1.5 ml of the collagen solution (Vitrogen 100) and mix with 150 pl of 1OX modified Earl’s balanced salt solution. d. Add 40 p1 of 1NNaOH to the tissues mince in DMEMMepes and transfer this to a 35-mm tissue culture dish. e. Immediately add the collagen solution to the tissue mince in the culture dish using a precooled pipet. Swirl the dish to mix the two components being careful to avoid introducing any bubbles during the mixing. f. Place the tissue culture dish into a 37°C COz incubator for 30-60 min. A gel should form within 10-15 min after neutralization and incubation at 37°C. g. Layer 2 ml of DMEM/Hepes containing 10% FBS on top of the gel. Feed twice weekly. Cell egress from the fragments should occur within 3 weeks. h. To subcultivate explanted cells, remove the gel from the dish and place in a 50-ml centrifuge tube. Add 5 ml of the collagenase solution and incubate for 15 min at 37°C. i. Add 10 ml of DMEM/Hepes + 10% FBS and centrifuge for 10 min at -500g. j. Decant the supernatant and resuspend the cells in 10 ml of DMEM/Hepes + FBS. The cells are then maintained and passaged as described earlier (Section ILB).

V. Specialized Methods to Maintain the Differentiated State The cell cultivation techniques just described provide a reasonable model for examining VSMCs in the synthetic state, but a poor system for studying the differentiated phenotype. This may be the result of losing important tissue interactions upon cell isolation. The individual cells within all differentiated tissues, including blood vessels, are in contact with one another, through hormonal interactions, through direct cell contact, or indirectly through specialization of the extracellular matrix. In many tissues, this contact has been shown to be essential either for inducing differentiation or for maintaining it. In the vessel, one well-recognized interaction is that between vascular smooth muscle and endothelial cells. As in other mesenchymal-epi(endo)thelial interactions, this interaction is brought about through specialization of the extracellular matrix involving the production of basal lamina or basement membranes, which is composed of collagen type IV, laminin, entactin (nidogen), and heparan sulfate proteoglycans. A continuous basement membrane separates endothelial cells from VSMCs and a basement membrane surrounds many if not all of the individual VSMCs in the media (Timpl, 1989; Heickendorff, 1989). The following methods summarize current attempts to regain the differentiated state in vitro. They rely either on trying to reestablish endotheliaWSMC interac-


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tions in culture or on providing an extracellular matrix similar in composition to that surrounding VSMCs in vitro. A. Cocultivation of Vascular Smooth Muscle and Endothelial Cells

The best-described system involves “transwell” technology in which VSMCs are grown on the bottom chamber of a six-well culture dish and endothelial cells are placed on a cell-culture insert (Falcon #3090) that is positioned over the smooth muscle cell layer. This setup allows free communication of medium between the two cell layers. This type of system, however, is limited to studying heterotypic cell interactions that may be mediated through biomolecules secreted into a common growth medium. It has been used to show that endothelial cells secrete a substance that inhibits VSMC proliferation and retards the modulation of primary cultures of VSMCs from the contractile to the synthetic state. This substance was identified as either heparin, heparan sulfate, or a related glycosaminglycan (Chamley-Campbell and Campbell, 1981). B. The Role of the Extracellular Matrix 1. Phenotypic Modulation on Fibronectin and Laminin Fibronectin and laminin are adhesive glycoproteins that have opposite effects on the phenotype of freshly isolated VSMCs (Hedin et al., 1988). Within 48 hr of being isolated and placed on a substrate of fibronectin, freshly isolated cells modulate from a contractile to a synthetic phenotype. When isolated and placed on a substrate of laminin, a component of the basement membrane that normally surrounds arterial smooth muscle cells, phenotypic modulation is considerably delayed. In fact, the delayed modulation on laminin can be delayed even longer if a synthetic peptide containing the RGD motif present in fibronectin is incubated with the cell. This peptide presumably blocks the interaction of the fibronectin receptor on the surface of the cell with fibronectin secreted by the cell. Modulation does not require DNA synthesis (Thyberg et al., 1983; Chamley-Campbell et al., 1981). It is important to note that this system does not induce differentiation but only delays the modulation from the contractile to the synthetic phenotype. According, experiments with this system are limited to primary cultures in which phenotypic modulation has not already occurred. Nonetheless, with current transfection methods that result in efficient introduction and expression of recombinant molecules into nonreplicating cells (e.g., adenoviral-mediated gene tranfer), this system will be useful for experiments targeted at identifying key components in the process of phenotypic modulation.

2. Three-Dimensional Basement Membrane Gels Three-dimensional basement membrane (BM) gels have been reported to induce and maintain 8 number of cells in the differentiated state (Friedman et

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aL, 1989, and references therein). When VSMCs are placed onto such substrates, there is an immediate inhibition of cell proliferation. This occurs whether or not serum is present and is in sharp contrast to the extended period of serum starvation (5-7 days) needed by VSMCs plated on plastic to growth arrest (Pauly et al., 1992). VSMCs also can not be stimulated to divide on BM gels, although early response genes can be activated (Crow, Pauly, and Monticone, unpublished observations). Studies indicate that there is a selective decrease in certain signaling transduction pathways, particularly those involving MAP kinases, in VSMCs plated on BM gels. VSMCs on BM gels exhibit increased fractional volumes of myofilaments-whereas cells on plastic exhibit a fractional volume of myofilaments of 4 0 % (Stadler et aL, 1989), VSMCs on BM have a fractional volume >40% (Pauly et aZ., 1992).A detailed study of differentiation-specificgene expression in this experimental system, however, has not been published. The procedure for preparing such BM gels is described in Section 11. Unlike the ECM system described earlier that requires primary cultures of VSMCs, the changes in VSMC behavior on BM have been seen with VSMCs that have been passaged up to 15 times. The only apparent difference that cell passage makes to the experiments is that the duration of the experiment is limited to 1-2 days when cells from later passage are used. This is likely to be due to the increased secretion of ECM-degrading proteases by these cells (Pauly et aL, 1994a), which eventually liquefy the gel. Even with earlier passage cells, the presence of these proteases means that VSMCs embed themselves into the BM. This poses particular problems for the isolation of RNA and preparation of protein extracts from VSMCs on BM gels. We have found that the standard guanidinium isothiocyanate (GTC) procedure in which RNA is pelleted through a CsCl bed gives very low RNA yields. This is because the excess protein from the BM gel forms a plug between the CsCl bed and GTC extract, impeding the movement of RNA to the bottom of the tube. We use a modification of this protocol in which the RNA is pelleted through cesium trifluoroacetate (CsTFA). In this procedure, RNA pellets and protein moves to the top of the tube. The reagents for this procedure and a detailed protocol are available from PharmacianKB (Piscataway, New Jersey, Cat. #27-9270-01). For the isolation of protein extracts, cultures are first rinsed with HBSS and then incubated for 30 min at 37°C in Dispase (Collaborative Research, Bedford, Massachusetts). The extract is removed and the cells washed three times with ice-cold Dulbecco’s PBS. An extract of the cells is then prepared by directly dissolving the cells in SDS-sample buffer or by adding a solution that contains 0.5% Triton X-100,2 mM MgC12, and 50 mM Tris-HC1 (pH 7.4).

VI. Conclusions and Perspectives Although it is clear that the development of techniques for culturing VSMCs has aided greatly in the understanding pathogenic mechanisms involved in vascu-


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lar disease, the challenge still exists of creating an in vitro environment in which reversible phenotypic modulation can occur. Such an environment would greatly aid current efforts by a number of laboratories trying to understand the molecular basis for phenotypic modulation. Acknowledgments We thank Drs. Peter Libby and Marysia Muszynski, Brigham and Women’s Hospital, Harvard Medical School, for detailed protocols on the isolation of smooth muscle cells from human saphenous veins and human atheromatous tissue and Dr. Zhihe Li, Laboratory of Cardiovascular Science, National Institute on Aging, for detailed protocols on the isolation of smooth muscle cells from human arterial vessels.

References Applegate, D., Feng, W., Green, R. S., and Taubman, M. B. (1994).Cloning and expression of a novel acidic calponin isoform from rat aortic vascular smooth muscle. J. Biol. Chem. 269,10683-10690. Babij, P., and Periasamy, M. (1989). Myosin heavy chain isoform diversity in smooth muscle is produced by differential RNA processing. J. Mol. Biol. 210,673-679. Babij, P., Kawamoto, S., White, S., Adelstein, R. S., and Periasamy, M. (1992).Differential expression of SM1 and S M 2 myosin i s o f o m in cultured vascular smooth muscle. Amer. J. Physiol. 262,C607-

C613. Chamley-Campbell, J. H., and Campbell, G. R. (1981). What controls smooth muscle phenotype? Atherosclerosis 40, 347. Chamley-Campbell, J. H., Campbell, G. R., and Ross, R. (1981). Phenotypic-dependent responses of cultured aortic smooth muscle to serum mitogens. J. Cell. Biol. 89,379-383. Campbell, G. R., and Campbell, J. H. (1981). Phenotypic modulation of smooth muscle cells in primary culture. In “Vascular Smooth Muscle in Culture” (J. H. Campbell and G. H. Campbell, eds.), Vol. I. Boca Raton, Florida: CRC Press. Frid, M. G., Shekhonin, B. V., KoteEansky, V. E., and Glukhova, M. A. (1992). Phenotypic changes of human smooth muscle cells during development: Late expression of heavy caldesmon and calponin. Dev. Biol. 153, 185-193. Friedman, S. L., Roll, F. J., Boyles, J., Arenson, D. M., and Bissell, D. M. (1989). Maintenance of differentiated phenotype of cultured rat hepatic lipocytes by basement membrane matrix. J. Biol. Chem. 264,10756-10762. Gabbiani, G.,Kocher, O., Bloom, W. S., Vandekerckhove, J., and Weber, K.(1984).Actin expression in smooth muscle cells of rat aortic intimal thickening, human atheromatous plaque, and rat cultured aortic media. J. Clin. Invest. 73, 148-152. Glukhova, M. A., Kabakov, A. E., Frid, M. G., Omatsky, 0. I., Bekin, A. M., Mukhin, D. N., Orekhov, A. N., Koteliansky, V. E., and Smirnov, V. N. (1988).Modulation of human aorta smooth muscle cell phenotype: A study of muscle-specific variants of vinculin, caldesmon, and actin expression. Proc. Natl. Acad. Sci. USA 85, 9542-9546. Hedin, U.,Bottger, B. A., Forsberg, E., Johansson, S., and Thyberg, J. (1988). Diverse effects of fibronectin and laminin on phenotypic properties of cultured arterial smooth muscle cells. J. Cell Biol. 107,307-319. Heickendorff, L. (1989). The basement membrane of arterial smooth muscle cells. Acta Pathol. Microbiol. Immunol. Scand 97(S9), 1-32. Jenkins, G.M., Crow, M. T., Bilato, C., Ryu, W.-S., Li, Z., Stetler-Stevenson,W., Nater, C., Froehlich, J., Lakatta, E. G., and Cheng, L. (1997).The 72 kD type IV collagenase (matrix metalloproteinase2) is activated by balloon injury in the rat and preferentially expressed in the neointima. Circulation, in prev.

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Kleinman, H. K., McGarvey, M. L., Hassell, J. R., Starr, V. L., Cannon, F. B., Laurie, G. W. and Martin, G. R. (1986). Basement membrane complexes with biological activity. Biochemistry 25,312-318. Kocher, O., and Gabbiani, G. (1987). Analysis of a-smooth muscle actin mRNA expression in rat aortic smooth muscle cells using a specific cDNA probe. Differentiation 34,201-209. Kuro-o, M., Nagai, R., Nakahara, K. Katoh, H., Tsai, R.-C., Tsuchimochi, H., Yazaki, Y., Ohkubo, A., and Takaku, F. (1991). cDNA cloning of a myosin heavy chain isoform in embryonic smooth muscle and its expression during vascular development and in atherosclerosis. J. BioL Chem. 266,3768-3773. Li, X.,Tsai, P., Wieder, E. D., Kribben, A., van Putten, V., Schrier, R. W., and Nemenoff, R. A. (1994). Vascular smooth muscle cells grown on matrigel. J. Biol. Chem. 269,19653-19658. Libby, P., Warner, S. J. C., Salomon, R. N., and Birinyi, L. K. (1988). Production of platelet-derived growth factor-like mitogen by smooth muscle cells from human atheroma. N. Engl. J. Med. 31% 1493-1498. Majesky, M. W., Giachelli, C. M., Reidy, M. A., and Schwartz, S. M. (1992). Rat carotid neointimal smooth muscle cells reexpress a developmentally regulated mRNA phenotype during repair of arterial injury. Circ. Res.7l,759-768. M o w , P. R. L., Campbell, G. R., and Campbell, J. H. (1986). Smooth muscle phenotypic expression in human carotid arteries. 11. Atherosclerotic-freediffuse intimal thickenings compared with the media. Atherosclerosis 6,664-669. Owens, G. K., Loeb, A., Gordon, D., and Thompson, M. M. (1986). Expression of smooth musclespecific a-isoactin in cultured vascular smooth muscle cells: Relationship between growth and cytodifferentiation. J. Cell BioL 102,434-352. Pauly, R. R., Passaniti, A,, Crow, M., Kinsella, J. L., Papadopoulos, N., Monticone, R., Lakatta, E. G., and Martin, G. R. (1992). Experimentalmodels that mimic the differentiationand dedifferentiation of vascular cells. Circulation 86,111-68-73. Pauly, R. R.,Passaniti, A., Bilato, C., Monticone, R.,Cheng, L., Papadopoulos, N., Gluzband, Y. A., Smith, L., Weinstein, C., Lakatta, E. G., and Crow, M. T. (1994a). Migration of cultured vascular smooth muscle cells through a basement membrane requires type IV collagenase activity and is inhibited by cellular differentiation. Circ. Res. 75,41-54. Pauly, R. R., Monticone, R., Fredman, F.,Bilato, C., Sollott, S. J., Jenkins, M., Cheng, L., Lakatta, E. G., and Crow, M. T. (1994b). Differences in the migration of vascular smooth muscle cells cultured from the neointima and media of rat carotid arteries. Circulation 90,1452. Pauly, R. R., Bilato, C., Sollott, S. J., Monticone, R., Kelly, P. T., Lakatta, E. G., and Crow, M. T. (1995). The role of calcidcalmodulin-dependentprotein kinase 11in regulating vascular smooth muscle migration. Circulation, 91,1107-1115. Rovner, A. S., Murphy, R. A., and Owens, G. K. (1986). Expression of smooth muscle and nonmuscle myosin heavy chains in cultured vascular smooth muscle cells. J. Biol. Chem. 261,147740-14745. Shanahan, C . M., Weissberg, P. L., and Metcalfe, J. C. (1993). Isolation of gene markers of differentiated and proliferating vascular smooth muscle cells. Circ. Res. 73,193-204. Takahashi, K. and Nadal-Ginard, B. (1991). Molecular cloning and sequence analysis of smooth muscle calponin. J. Biol. Chem. 266,13284-13288. Thyberg, J., Palmberg, L., Nilsson, J., Ksiazek, T., and Sjolund, M. (1983). Phenotypic modulation in primary cultures of arterial smooth muscle cells. On the role of platelet-derived growth factor. Differentiation 25,156. Thyberg, J., Hedin, U., Sjolund, M.,Palmberg, L., and Bottger, B. A. (1990). Regulation of differentiated properties and proliferation of arterial smooth muscle cells. Arteriosclerosis 10,966-990. Timpl, R. (1989). Structure and biological activity of basement membrane proteins. Eur. J. Biochem. 180,487-502. Ueki, N.,Sobue, K., Kanda, K., Hada, T., and Higashino, K. (1987). Expression of high and low molecular weight caldesmons during phenotypic modulation of smooth muscle cells. Proc. Natl. Acad. Sci. USA 84,9049-9053.

CHAPTER 8

Skeletal Muscle Satellite Cell Cultures Ronald E. Allen, Constance J. Temm-Grove, Shannon M. Sheehan, and Glenna Rice Muscle Biology Group Animal Sciences Department The University of Arizona Tucson, Arizona 85721

I. Introduction 11. Physiological or Developmental Background of Subjects

A. B. C. D.

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IV.

V.

VI.

Age Muscle Disease or Injury Endocrine Status E. Species Monolayer Mass Cultures A. Substrates B. Medium C. Cell Preparation D. Characteristics Isolated Single Fiber Cultures A. Substrates and Media B. Fiber Isolation C. Characteristics Advantages and Disadvantages of Each Culture System A. Monolayer Mass Cultures B. Isolated Single Fiber Cultures (Bischoff System) Conclusions References

I. Introduction Myogenic stem cells in postnatal muscle were initially identified as small mononucleated cells located between the sarcolemma and basal lamina of frog muscle METHODS IN CELL BIOLOGY. VOL. 52 Copyright 0 1998 by Academic Press. AU righa of repduchon in any form mmcd. 0091-679W98 $25.00

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fibers (Mauro, 1961). These cells were referred to as satellite cells and were subsequently shown to have the ability to proliferate and fuse with adjacent fibers in uninjured muscle (Moss and Leblond, 1971). This observation demonstrated the myogenic potential of satellite cells and provided a mechanism for addition of nuclei to growing fibers. The addition of new nuclei to growing fibers facilitatesfurther growth in fiber length and girth. Satellite cells were also shown to be the source of myogenic precursor cells responsible for muscle fiber repair and regeneration (reviewed by Bischoff, 1994;Yablonka-Reuveni, 1995). Therefore, the biology of the skeletal muscle satellitecell is central to our understanding of skeletal muscle growth, adaptation, and repair. Because satellite cells are sparsely distributed in muscle (approximately 1-4% of muscle nuclei in fast-twitch muscle and three or four times that frequency in slow-twitch muscle; Bischoff, 1994), in vivo studies have generally been dependent on light and electron microscopy techniques. Development of in vitro satellite cell culture techniques has complemented the in vivo studies and has permitted a more detailed examination of specific cellular characteristicsand properties of satellite cells. Two primary culture systems are commonly used (i) monolayer mass cultures of dissociated satellite cells and (ii) single muscle fiber cultures with their associated satellite cells. In both cases, cells and fibers are isolated directly from experimental subjects. The general cellular processes that have been studied in satellite cell culture systems are diagramed in Fig. 1 and include activation from the quiescent state, migration, proliferation, and differentiation or return to quiescence. The purpose of this chapter is to present two types of satellite cell culture procedures and discuss their application.We apologize in advance to those whose contributions we may have inadvertently overlooked; we have narrowed our treatment of this subject and have not been able to cover all in vitro approaches to satellite cell experimentation. For example, we have not covered the use of cultured satellite cells in gene therapy as carriers of new genes to be integrated into muscle fibers. Furthermore, by restricting our discussion to primary cell or fiber cultures, we have not broadened the presentation to include immortalized myogenic cell lines, some of which were originally derived from postnatal muscle tissue and might be considered as satellite cell lines.

Differentiation Quiescence

E

r Migration rProliferation

\ Quiescence

Fig. 1 Cellular processes that have been studied with satellite cell culture systems. The diagram describes all of the general activities of satellite cells.

8. Skeletal Muscle Satellite Cell Cultures

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11. Physiological or Developmental Background of Subjects Selection of animals and muscles to be used in the preparation of satellite cells is the first consideration in studying satellite cells in vitro. One of the important attributes of primary cell culture systems is their ability to reflect the physiological or developmental background of the animal from which cells were derived. Consequently, careful consideration must be given to the selection of subjects, with respect to age, specific muscle, disease or injury, endocrine status, species, or other physiological characteristics. All of these variables can affect certain aspects of satellite cell activity in v i m ; consequently, it is important to take these things into consideration when designing experiments. Moreover, it is also important to report the characteristics of animals used as a source of satellite cells when communicating experimental results. The following brief examples illustrate this point. A. Age

Schultz and Lipton (1982) demonstrated a marked increase in the time preceding the onset of proliferation in culture and a decrease in the ultimate size of satellite cell colonies when satellite cells from rats of increasing age were plated at clonal density. A similar observation was made by Dodson and Allen (1987) when examining the growth kinetics of mass cultures of satellite cells from young (3-4 month), adult (9-12 month), and old rats (24 months); there was a lag in the onset of proliferation in adult and old rat satellite cells compared to satellite cells from young rats. Furthermore, it has been proposed that different subpopulations can be found in young and adult animals, and that one population may be predominantly responsible for growth and the other for repair in adult animals (Schultz, 1996).

B. Muscle The incidence of satellite cells associated with slow-twitch fibers is greater than with fast-twitch fibers (Kelly, 1978), and it has been proposed that satellite cells associated with different fiber types represent distinct subpopulations.There is reasonable evidence for this in chicken and rat, in that satellite cells from predominantly fast-twitch and slow-twitch fibers express different isoforms of myosin heavy chain upon differentiation in culture (Feldman and Stockdale, 1991; Rosenblatt et aL, 1996). C. Disease or Injury

As previously mentioned, there is a lag in the onset of proliferation in satellite cells from adult and old rats when placed in culture. This lag is not observed if

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the muscle is injured a few days prior to harvesting cells. A significant percentage of satellite cells from bupivicaine-injected muscle (a myotoxic agent that has been used to study muscle regeneration) can be found in the S phase of the cell cycle immediately after culturing, in contrast to satellite cells from uninjured adult muscle (Allen and Rankin, 1990). In addition, satellite cells in mdx mice experience repeated cycles of degeneration and regeneration (McGeachie et al., 1993), and therefore, a higher percentage of satellite cells would be expected to be active than in control animals. D. Endocrine Status This area has not been examined in great detail; however, experiments conducted by Thompson et al. (1989) demonstrated that cultured satellite cells from female rats that had received anabolic steroid treatments for two weeks exhibited greater responsivity to fibroblast growth factor 2 (FGF2) and to insulin-like than cells from control rats. This observation has been growth factor I (IFG-I) verified with bovine satellite cells from anabolic steroid-treated and untreated animals (Frey et al., 1995).

E. Species Satellite cells from different species may not exhibit the same responses to certain growth factors or even show the same pattern of gene expression. For example, FGF2 is a potent inhibitor of mouse (Clegg et al., 1987) and bovine satellite cell differentiation (Greene and Allen, 1991), but it only weakly suppresses differentiation of rat satellite cells (Allen and Boxhorn, 1989). In another example, transforming growth factor beta-1 (TGF-P1) inhibits rat satellite cell differentiation and severely depresses proliferation (Allen and Boxhorn, 1989); in cultures of ovine satellite cells, TGF-P1 inhibits differentiation but stimulates proliferation (Hathaway et al., 1991). Finally, rat satellite cells express desmin early in culture, prior to proliferation or differentiation, but satellite cells from bovine muscle do not express desmin until after differentiation (Allen et aZ., 1991). Clearly, it is important to be aware of how the source of satellite cells may influence the characteristics exhibited by satellite cells in culture. Of particular importance is the issue of whether the cells would be expected to be quiescent or cycling at the time of preparation. Quiescent cells are not responsive to certain growth factors and may not express many of the genes that are expressed in cycling cells, as will be presented later.

III. Monolayer Mass Cultures Primary cultures of dissociated muscle satellite cells are derived by enzymatically and mechanically liberating satellite cells from their position between the muscle fiber plasmalemma and basal lamina. Once liberated, they can be sepa-


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rated from much of the tissue debris and plated into cell culture dishes where they will attach, divide, migrate, and differentiate. The culture environment permits direct examination of these fundamental satellite cell properties. The procedure we will present is an adaptation of the original procedure of Bischoff (1974).

A. Substrates Cell culture conditions for satellite cells do not differ greatly from those employed in culturing myogenic cells of fetal or embryonic origin. Polystyrene tissue-culture dishes are used and can be purchased from a variety of suppliers. As with other muscle cell culture techniques, optimal growth requires a coating of one of several extracellular matrix proteins or combinations of proteins. The most commonly used coating agents are listed below: 1. Collagen. Using a 0.2 mg/ml solution of soluble collagen in 1%acetic acid, coat dishes by pouring collagen solution into dish and decanting excess. Allowed to air dry or place in a warm oven at 50-60°C. 2. Polylysine. Using a solution of 0.1 mg/ml polylysine in sterile distilled water, coat dishes with 20 kuYcm2. Remove excess solution after 5 min and rinse with tissue culture grade water. Air dry for a minimun of 2 hr. (We often coat tissue culture dishes and coverslips with polylysine, followed by fibronectin coating.) 3. Fibronectin. Human or bovine fibronectin can be used as attachment factors for cultured satellite cells. Initially, we used bovine fibronectin in culture experiments employing serum-free defined medium in order to examine growth factor effects on satellite cells. The rationale behind its application was that it may provide a more defined substrate than collagen extracts or Matrigel. Apply in a 10 pg/ml solution in Dulbecco’s modified Eagle’s medium (DMEM), pour off excess solution, and air dry dishes. 4. Matrigel (Collaborative Biomedical Products). Matrigel is the soluble basement membrane extract from the Engelbreth-Holm-Swarm tumor cells, and its major components are type IV collagen, laminin, heparan sulfate proteoglycan, and entactin. Although this is an excellent substrate and promotes high plating efficiency and growth, consideration should be given to the possible presence of unidentified components and growth factors. A 1:10 dilution of Matrigel in DMEM has been found to work well for rat satellite cells. Cover the bottom of chilled tissue-culture dishes with chilled Matrigel solution. Remove excess solution after 1min and place dishes in cell culture incubator for 15 to 20 min before using. B. Medium

A variety of cell culture media have been used for the cultivation of satellite cells. Most formulations consist of basal medium plus serum. Various basal

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media have been used, such as Eagle’s minimum essential medium, DMEM, and McCoy’s medium. An alternative to serum- or extract-containing medium is serum-free “defined” medium. Both types of media have been used in satellite cell culture experimentation, and each has distinct advantages and disadvantages. 1. Serum-containing medium. Optimal growth of satellite cells in culture generally requires medium containing serum and often chick embryo extract. Furthermore, satellite cell proliferation and differentiation can be manipulated by changing serum source and concentration and by addition of insulin or other growth factors. For example, chick embyro extract and fetal calf serum have often been employed in medium to enhance proliferation. In contrast, differentiationis generallypromoted by medium that does not have high mitogenic activity. In order to promote differentiation,we routinely use DMEM with 1%antibiotic-antimycotic, 1-2% horse serum, 1 pM insulin, and 1 pg/ml linoleic acid/BSA complex (Collaborative Biomedical Products). The supraphysiologicalconcentration of insulin is used as a surrogate for insulin-like growth factors that stimulate differentiation of myogenic cells. Medium containing low serum has also been used to study the effects of various growth factors that may be found in serum and whose effects may be obscured by high serum concentrations. The medium formulation used in our laboratory to grow rat satellite cells is composed of DMEM containing 10% horse serum, 1% antibioticantimycotic (Gibco), 0.5% gentamicin (Gibco). This medium promotes proliferation and subsequent differentiation of rat satellite cells. 2. Serum-free defined medium. Serum-free defined media formulations were developed for use in studies of the influence of extracellular factors on satellite cell activities. These media permitted growth of satellite cells for various periods of time and proved to be useful in that the complex mixtures of factors found in serum or tissue extracts could be replaced with a minimal number of added reagents. The term “defined medium” is clearly a misnomer, however, because satellite cells actively secrete many growth factors and matrix molecules that can have autocrine effects on cell activities. Nonetheless, “defined” medium eliminates one of the “black boxes” in satellite cell culture experimentation by restricting the list of extracellular factors to those added in medium plus those that are produced by the cells of interest. Two defined medium formulations are presented, one for rat and one for human satellite cells. Rat satellite cell defined medium (Allen et al., 1985; Allen and Boxhorn, 1989): 75%DMEM (Gibco) 25%MCDB-104 or McCoy’s (Gibco) 1%antibiotic-antimycotic (Gibco) 0.5%geotamicin (Gibco) 1 p g / d linoleic acid/BSA complex (Collaborative Biomedical Products) lo-’ M dexamethasone (Sigma) lo-’ M Insulin (Collaborative Biomedical Products)

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500 pg/d Deutsch Fetuin (Gibco) 100 pg/d heparin (Sigma; specific lots may vary and must be screened) 50 ng/ml bovine fibronectin (Gibco) 0.5 pg/ml biotin (Sigma) 3.75 n g / d selenium (Collaborative Biomedical Products) 0.1 pg/ml vitamin E (Gibco) Notes: (1) FGF2 may be used at a concentration of 2 n g / d to enhance survival. (2) Insulin levels used in defined medium are close to physiological concentrations and not supraphysiological (micromolar range), which can be employed to mimic the effects of insulin-like growth factors, as in the differentiation promoting medium described previously. (3) Deutsch Fetuin is no longer readily available and must be prepared or specially ordered (at significant expense).

The formulationjust listed is a minimal formulation that allows cells to survive but does not promote proliferation or differentiation to any great extent. It was purposely designed this way to provide a serum-free environment in which to study the effects of growth factors and their interactions. It is important to note that satellite cells from adult rats require an initial period in serum-containing medium prior to switching to serum-free defined medium, and after 120 to 144 hr, cell survival diminishes. It is obvious that the factor(s) required for activation are not present and that there are other missing factors that are necessary for long-term growth. The just-mentioned formulation, however, provides a 3- to 4day window for studying proliferation and the onset of differentiation. Human satellite cell defined medium (Ham et nl., 1988): MCDB-120 0.5 m g / d bovine serum albumin (insulin RIA grade) M dexamethasone 10.0 n g / d epidermal growth factor 0.5 m g / d Pederson fetuin 3 X lo-’ M insulin

C . Cell Preparation

The procedure for preparing primary monolayer mass cultures of rat satellite cells was originally presented by Bischoff (1974). We have made a few minor adaptations to the procedure as it has evolved in our laboratory. Three key elements are (i) selection of the appropriate protease, one that theoretically degrades basal lamina while having minor effects on the collagen matrix associated with connective tissue fibroblasts, (ii) careful trimming and mincing of muscle tissue, which eliminates major sources of connective tissue cells, and (iii) minimizing microbial contamination. The basic steps of the procedure are as follows:

1. Instruments 1. One package of sterile instruments containing: a. 4 scissors b. 1 scalpel handle

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c. 1 hemostat d. 2forceps 2. One sterile metal strainer with 0.5-mm screen 3. One sterile 200 mesh cell dissociation sieve (Sigma) 4. One sterile glass baking dish (9-inch by 13-inch) 5. Six sterile 2 5 0 4 glass beakers 6. 150-mm sterile culture dish for cleaning tissue 7. Sterile scalpel blade (number 22) 8. Sterile 50-ml centrifuge tubes 9. One 2-liter plastic beaker 10. Sterile pipets, 1,5, and 10 ml 11. Coated tissue culture plates (fibronectin, polylysine, collagen, or Matrigel) 12. Latex gloves

2. Reagents 1. 70% ethanol 2. Sterile PBS 3. DMEM + 10% horse serum with antibiotics 4. DMEM + 20% fetal bovine serum and 10% DMSO (for freezing cells)

3. Protocol 1. Euthanize rat in carbon dioxide chamber. 2. Soak rat in 70% ethanol in 2-liter beaker for 5 min. 3. Excise only the large back and thigh muscles and place in sterile 250-ml beaker with warm PBS. 4. Rinse muscle tissue twice with warm PBS. 5. Trim away tendons, fascia, and as much connectivetissue and adipose tissue as possible. 6. Mince trimmed muscle tissue in a sterile culture dish (if large amounts of muscle are being processed, a presterilized kitchen meat grinder can be used for initial mincing step). 7. Weigh tissue in sterile preweighed culture dish and aliquot approximately 10 g150-ml tube. 8. Add 30 ml of filter-sterilized PBS containing 1.25 mg/ml protease type XVII from Streptomyces griseus (Sigma P5147). This protease is similar to pronase, originally described by Bischoff (1974). (Note: Use 1.0 mg/ml for juvenile rats, 1 month of age or less) 9. Incubate tubes in 37°C water bath for 1hr, shaking every 15 min.


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10. Centrifuge at l500g for 4 min, and discard supernatant. 11. Add warm PBS (40 ml total) to the tubes and mix on a vortex mixer for 20 sec. 12. Centrifuge at 500g for 10 min and decant supernatant into sterile 50ml tubes. 13. Repeat step 11 and centrifuge at 500g for 8 min and retain supernatant. 14. Repeat step 11 and centrifuge at 500g for 5 min and retain supernatant. 15. Pool supernatants and centrifuge at 1500g for 3 min to pellet cells. 16. Combine cell pellets into one tube using a small amount of DMEM + 10%HS and bring to a final volume equal to 1g of trimmed muscle/ml of medium. 17. Pour through sterile metal strainer and rinse with DMEM + 10% HS. 18. Pour through cell dissociation sieve and rinse with DMEM + 10% HS. 19. Centrifuge at 1500g for 3 min and discard supernatant. 20. Resuspend pellet in DMEM + 10% HS to give a final volume equivalent to 1 g of trimmed muscle per ml. 21. Pre-plate cells in 150-mm uncoated tissue-culture dish and maintain in culture incubator for 2 hr. 22. Gently agitate medium and decant cell suspension into sterile tube. 23. Plate cells in coated tissue-culture dishes at the equivalent of 1 g tissue/ 400 mm2,which will give an initial cell density of approximately 75 cells/mm2.' 23a. Satellite cells may be frozen for future use by centrifuging cells at 1500 g for 4 min and resuspending pelleted cells in freeze medium containing fetal calf serum and DMSO (3g tissue/ml freeze medium). Cells suspension is pipetted into cryotubes and placed at -70°C for 24 hr before transfemng to liquid nitrogen. 24. Incubate cell cultures in a humidified atmosphere at 37°C in 5% C02, and replace medium daily. D. Characteristics 1. Culture Homogeneity Several factors may affect the percentage of satellite cells in primary cultures. The first critical point in the process is in the selection of protease. Bischoff (1974)

* All fiber fragments and debris cannot be removed with this procedure; therefore, it is difficult to get accurate cell counts immediately following cell preparation. We have found it easier and more reproducible to plate cells based on the initial amount of starting material. Furthermore, there is a limit to the amount of cell suspension that can be plated per unit area; beyond a certain point, plating efficiency decreases dramatically, possibly because of the amount of debris that settles onto the surface of the tissue culture plate and blocks the attachment of cells. If Percoll (Pharamacia, Piscataway, New Jersey) gradients are used to remove debris and some additional nonmuscle cells (Yablonka-Reuveni et al., 1987), cleaner cell suspensions are obtained, and cells can be counted directly before plating. However, if large numbers of primary cultures of satellite cells are needed, the use of Percoll gradients may require too much time and labor.

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originally demonstrated this point when comparing the effects of collagenase, pronase, and trypsin on the preparations of satellite cells from minced rat muscle, He showed a correlation between digestion of the basal lamina and liberation of satellite cells; collagenase was least effective of the three, and pronase most effective. It is apparent that satellite cells and fibroblasts can be selectively liberated depending on the type of protease used. The second important aspect of the cell preparation process is the selection of muscle and the amount of associated connective that is subsequently carried through the proteolytic digestion step. This point is demonstrated in Figs. 2 through 4. Satellite cell cultures were prepared from the major muscle of the back after the fascia and other connective tissue had been trimmed away or from a combination of muscles from the lower leg that were not trimmed free of associated connective tissue (these muscles included the gastrocnemius, tibialis, soleus, and extensor digitorum longus). The percentages of satellite cells in these preparations were assessed by determining the presence of MyoD, desmin, and c-met using immunofluorescence (Figs. 2 and 3). These three proteins are only found in the myogenic cells in these cultures and are not expressed by muscle fibroblasts. MyoD is expressed very rapidly following satellite cell activation (Grounds et al., 1992; Smith et aL, 1994; Yablonka-Reuveni and Rivera, 1994); desmin is expressed in proliferating satellite cells prior to 24 hr in these cultures (Allen et al., 1991); and c-met message and protein, the receptor for HGF/SF, has been shown to be present as soon as satellite cells can be isolated (Allen et al., 1995; A. Cornelison and B. Weld, personal communication). Satellite cell percentages were greater when muscle preparations with minimal connective tissue were used as a source and when extra care was taken to trim away fascia and connective tissue (Fig. 4). Pre-plating satellite cell suspension in uncoated culture dishes for 2 hr to allow preferential attachment of fibroblasts further increases the purity of the cell preparation. Unfortunately, many of the larger muscles that may be easier to trim connective tissue from are predominantly composed of fast-twitch fibers and consequently have a lower frequency of satellite cells. As a result, fewer cells are harvested per gram of muscle. The cultures in this demonstration were prepared from adult (9-12 month old) male rats and were evaluated at 30 hr in culture. This represents a time prior to proliferation and therefore reflects the satellite cell composition of the original cell preparation. Even though the percentage of nonmuscle cells is relatively small at this time, when cultures are allowed to grow in high-serum medium for an extended period, the percentage of nonmuscle cells will increase. This results in part from the differentiation and withdrawal of myogenic cells from the population of proliferating cells. Some experimentsmay require that specificmuscles be used, and those muscles may be relatively high in connective tissue content. Other experiments may require a higher degree of satellite cell purity. In these cases, additional purification steps may be taken. The most effective step may be centrifugation through Percoll gradients (Pharamacia; Yablonka-Reuveni et al., 1987). Following the


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Fig. 2 Immunofluorescence detection of proteins that are useful markers for cultured satellite cells. Desmin (a and a’) and c-met (b and b‘) are present only in myogenic cells as shown in cultures from untrimmed muscles from the lower leg of male rats; cultures were fixed at 30 hr. Panels c and c’ show MyoD immunofluorescence and phase images of a group of myogenic cells from 24-hr cultures. Desmin antibody D3 was obtained from the Developmental Studies Hybridoma Bank; polyclonal rabbit anti-MyoD antibody was a gift from Dr. Steve Konieczny; rabbit polyclonal antic-met antibody was purchased from Santa Cruz Biotechnology (Cat. No. sc-162). Rhodamhe-labeled secondary antibodies were used for all three procedures. Magnification bar = 25 pm.

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Fig. 3 Desmin and c-met double immunofluorescence. Examples of rat satellite cells cultured for 30 hr showing the presence of c-met (a' and b') in myogenic cells containing desmin (a and b). In these cultures cells either had both proteins present, or they had neither. This indicates that c-met may be a useful marker for satellite cells in culture. Goat anti-mouse rhodamine-labeled second antibody was used for desmin and goat anti-rabbit Bodipy FL second antibody was used for c-met. Magnification bar = 15 pm.

previously described procedure, the final cell pellet is resuspended in medium and layered over a discontinuous gradient of 11.5 ml of 20% Percoll and 1.5 ml of 60% Percoll in a Corex tube that has been pretreated for 2 hr with horse serum. Cells are centrifuged for 5 min at 15,OOOg with the brake off. Satellite cells are collected at the 20/60% interface; fibroblast and fiber debris are found in the upper fractions. Bischoff and Heintz (1994) describe a different set of conditions for separation of satellite cells with Percoll; cells are layered on a step gradient of 35, 50, and 70% Percoll and centrifuged at 1,250g for 20 min with the brake off. Satellite cells are collected at the 35/50% and the 50/70% interfaces; fibroblasts are found from 0 to 35% and red blood cells are found in the 70% pellet. A high degree of purity can be achieved with these techniques, although total yield is compromised and a great deal of labor and time are required to process large amounts of tissue. Satellite cell separation using a fluorescence activated cell sorter has been used with human satellite cells labeled with a monoclonal antibody to cell surface protein (Webster et d., 1988). Not all species bind this antibody, however, and other satellite cell-specific antibodies are not available, although antibodies to c-met may be new candidates.

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T

~Mti M p D Dg

Untrimmed

c - MM ~ p D Da

Trimmed

C - MM ~ p D Da

Trimmedmreplnted

Fig. 4 The influence of muscle connective tissue on the percentage of satellite cells in monolayer mass cultures. Cultures were prepared from two sources: (1) A combination of muscles from the lower leg (gastrocnemius, tibialis, soleus and extensor digitorum longus) that were not trimmed free of associated connective tissue prior to digestion and plating, and (2) the major muscle of the back with the fascia and other connective tissue carefully trimmed away prior to enzymatic digestion. Cells were plated directly into coated tissue culture dishes or were pre-plated on uncoated culture dishes for 2 hr. Three satellite cell markers were used to assess culture heterogeneity by immunofluorescence; desmin, c-met, and MyoD. Bars represent the means and standard errors for percentages of cells containing each of the three marker proteins in 30-hr cultures from untrimmed, trimmed, and trimmed and pre-plated muscle preparation.

2. Activation/Proliferation As previously mentioned, the kinetics of growth, including cell cycle kinetics and gene expression, depend on the background of the cells in viva The vast majority of satellite cells harvested from adult rat are quiescent. In recent years, it has become increasingly apparent that differences in gene expression exist between quiescent and proliferating satellite cells. The distinction between the two states is only beginning to be appreciated. All of the differences noted thus far are in the absence of expression of certain genes in quiescent cells and the induction of gene expression as satellite cells become activated and enter the cell cycle. Table I lists genes or cellular responses that our laboratory has observed in quiescent andor activated rat satellite cells. As is apparent in Table I, most of the activities or genes noted thus far center around growth control or control of differentiation. It is important to recognize that if cultures are established from muscle in which satellite cells are active, the higher ratio of active to quiescent satellite cells would result in detection of the previously indicated genes or responses from the outset in cultures. For example, satellite cells from 3-week-old rats specifically bound and responded to FGF2 by 18 hr in culture, whereas cells from adult rats did not specifically bind or respond until after 36 hr in culture (Johnson and Allen, 1995).

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Table I Genes and Responses Expressed during Activation of Quiescent Satellite Cells in Monolayer Cultures GeneResponse c-Met Respond to HGFlSF HGF/SF FGF2 specific binding FGF Receptor 1 (FGFRl) FGFR2 FGFR3 FGFR4 Respond to FGFl & 2 Respond to IGF-I IGF-I IFG-I1 Respond to PDGF-BB Respond to EGF Cyclin D PCNA MyoD MRF4 Myfs Myogenin

Quiescenta

Activated

+ +

+

- (traces)b

- (traces) - (traces) - (traces) - (traces) - (traces)

+ -

-

+ +

+ + +

- (traces) - (low)

+ +

+ + + + + + + +

+ +

mRNA and protein mRNA and protein mRNA mRNA mRNA mRNA mRNA and protein mRNA protein protein mRNA and protein mRNA and protein mRNA and protein mRNA and protein

Freshly harvested satellite cells from adult rat were used as quiescent cells; activated cells were cells that had been in culture for 36 hr or more. By RT-PCR very small amounts of mRNA were detected; in some cases, this may be due to the small fraction of satellite cells that are already active in vivo at the time of harvest.

3. Differentiation In rat satellite cell cultures prepared from adult animals, the first evidence of terminal differentiation can be found between 72 and 96 hr. Sarcomeric myosin heavy chain can be localized in a few mononucleated cells as early as 72 hr and in small myotubes by 96 hr. Myogenin is also detected in the nuclei of mononucleated and fused cells during the same time frame. The typical array of muscle specific proteins are subsequently expressed in the differentiated myotubes, and as previously mentioned,some fiber-typespecificgene expressionhas been detected that reflects the muscle of origin. Large myotubes developin these cultures and spontaneous contractions are seen if cultures are allowed to grow for extended times. As myotubes mature and contractionsbecome stronger and more frequent,they often detach from the substrate. Although a high degree of differentiation can be attained in culture, it is possible to find myogenic precursor cells that persist through several passages and differentiation cycles. This observation has not been studied in detail, but it may be a reflection of the “stem cell” nature of satellite cells.


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Figure 5 illustrates the time course of some of the key events occurring in cultured adult rat muscle satellite cells as they become activated, start to proliferate, and begin to differentiate. As previously mentioned, activation occurs in a relatively synchronous manner in cultures of adult rat satellite cells. Most cells complete their first division at approximately 48 hr. Because this is a primary culture system, however, perfect synchrony is not possible; a minor fraction of the cells will be cycling at the time of harvest and can therefore be found in the S phase of the cell cycle at earlier times. Cyclin D and PCNA are expressed in G1 of the cell cycle, and the percentage of cells expressing these proteins can be seen to increase dramatically prior to 48 hr. DNA synthesis, as demonstrated by pulse labeling with BrdU, follows the increased expression of cyclin D and PCNA. With regard to the basic helix-loop-helix myogenic regulatory genes, MyoD is expressed very early and persists throughout the time period examined. Expression of MRF4 and myf5, however, peaks at about the time cells undergo their first division, and then the percentage of cells expressing these factors decreases. The transient nature of MRF4 and myf5 expressionmay be responsible

A

100

180 0

%"* a 20 0

Fig. 5 Summary of the time course of DNA synthesis and the expression of key proteins during the activation, proliferation, and differentiation of cultured adult satellite cells; data were compiled from several experiments conducted with this culture system. Panel (A) shows the temporal relationship among two cell cycle regulated proteins, cyclin D and PCNA, DNA systhesis (BrdU incorporation), and cell proliferation during the activation phase in satellite cell cultures from adult male rats. The time course of expression of four myogenic regulatory genes in relationship to the onset of cell proliferation in these cultures is show in panel (B). Labeling indices were determined by irnmunofluorescence using the following antibodies; anti-MyoD, -MyfS, and -MRF4 were provided by Dr. Steve Konieczny; anti-myogeninwas provided by Dr. Woodring Wright; anti-cych D was purchased from Santa Cruz Biotechnology; anti-PCNA was purchased from Boehringer Mannheim; antibromodeoxyuridinewas acquired from Developmental Studies Hybridoma Bank.

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for the observation that the percentage of cells expressing either of these two factors never approaches the level of MyoD expression. Myogenin expression increases as cells begin to differentiate; sarcomericmyosin heavy chain expression followsmyogenin expression closely (data not shown). In experimentsthat extend to 120 or 144 hr, the percentage of cells expressing myogenin approaches the percentage of MyoD expressing cells. In general, the pattern of myogenic regulatory factor protein expression is consistent with the previously reported pattern of message expression in similar rat satellite cell cultures (Smith et al., 1994). Figure 5 only describes the expression patterns of a few important regulatory proteins. As others are added, clues to the control of satellite cell activities may emerge.

IV.Isolated Single Fiber Cultures The isolated single fiber culture system is an in vitro model for studying satellite cells that is one step closer to the microenvironment found in living muscle. Bekoff and Betz (1977) described a single fiber tissue culture procedure that was subsequently adapted by Bischoff for studying satellite cells (1986a). This procedure entails proteolytic digestion of muscle tissue and the liberation of intact muscle fibers with their associated basal lamina and encased satellite cells. The obvious advantage of this system is the authenticity of the satellite cell environment. The associationswith sarcolemma and muscle fiber basement membrane are obviously not possible to duplicate in monolayer cultures. Extracellular matrix certainly affects satellite cell activity, and evidence exists that communication between fiber and satellite cell can influence satellite cell activity (Bischoff, 199Ob). Consequently, this culture system has been useful in providing insight into the regulation of satellite cells.

A. Substrates and Media Some of the substrates used for monolayer mass cultures of satellite cells have also been used for culturing myofibers. The size of fibers and their attachment properties, however, make substrate attachment more critical. The three substrates that have been used most extensively are clotted plasma or collagen (Bischoff, 1986a) and Matrigel (Rosenblatt et al., 1995). Serum-containing medium is used in these culture systems and is composed of Eagle’s minimal essentialmedium or Dulbecco’smodified Eagle’s medium, 1% antibiotic-antimycotic mixture (Gibco, Grand Island, New York) and variable concentrations and sources of serum. Chick embryo extract may also be employed, depending on the desired growth response. For example, high concentrations of fetal calf serum, chick embryo extract, and horse serum have been used to promote extensive migration and proliferation of satellite cells (Rosenblatt et al., 1995), whereas 10% horse serum alone maintained satellite cells in a


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quiescent state in association with myofibers (Bischoff, 1986a). The addition of chick embryo extract promoted activation of fiber-associated quiescent satellite cells, as did the addition of a soluble extract from lightly crushed muscle (Bischoff, 1986b).Culture medium can also be altered to facilitate differentiation of satellite cells that have proliferated within the basal lamina tube or have migrated out and proliferated in association with the substrate. In general, differentiation medium has low mitogenic potential, often contains low concentrationsof horse serum and may not contain embryo extract or other mitogenic agents, just as with monolayer mass cultures.

B. Fiber Isolation The basic procedure that follows was originally outlined by Bischoff (1986a); specific adaptations of this procedure that have been described by Rosenblatt et al. (1995) will be presented following the basic procedure. 1. Carefully remove flexor digitorum brevis (FDB), taking care to handle only by the tendons. This is a small muscle between the toes of the rat; it was selected because of its short, uniform fibers that could be readily liberated and isolated in large numbers. 2. Remove epimysium and treat with collagenase in PBS at 37°C for 1.5 to 2 hr with gentle agitation. Bischoff fractionated crude collagenase (Sigma, type I) on SP-Sephadex to separate proteolytic activity into three fractions, neutral protease, collagenase, and colostripain. Collagenase (0.6 mg/ml) and neutral protease (0.4 mg/ml) are used for digestion. The addition of colostripain causes damage to the basal lamina, which results in activation and liberation of satellite cells; therefore, it is omitted. 3. Wash muscle with PBS and place in Eagle’s medium plus 10%horse serum and 1%antibiotic.Separate muscle fibers by pulling apart the three distal tendons. Fibers are liberated by gentle trituration using a wide-mouth pipet. Up to 100 trituration cycles may be required to disperse fibers; progress is checked microscopically. According to Bischoff (1986b) fiber hypercontraction or uptake of vital dye indicate that fibers have been killed by triturating too vigorously or by toxicity of the enzyme preparation. 4. Muscle debris and free mononucleated cells (many of which may be fibroblasts) are removed from the fibers by sedimentingfibers through culture medium at lg. Three cycles of sedimentation are used. 5. Attach fibers to culture dishes in a fibrin clot or collagen gel. 6. Maintain fibers in Eagle’s medium plus 10% HS and 1%antibiotic.

The following are some of the variations on this procedure that have been reported by Rosenblatt et al. (1995).

la. Other muscles with longer fibers are used, such as the extensor digitorum longus, soleus, and tibialis anterior. These muscle are frequently used in other

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types of physiological studies, and therefore it is possible to integrate in vivo and in vitro experimentation. 2a. Collagenase is not fractionated to remove colostripain. This may not be necessary because the goal of this procedure is not to maintain quiescent satellite cells on living fibers; consequently, the integrity of the basal lamina is not as important. 4a. and 5a. Isolated fibers are not separated from debris or mononucleated cells by sedimentation at unit gravity because the longer fibers used in this procedure do not survive the sedimentation cycles. Individual fibers are removed from suspension and plated directly into tissue culture dishes that have been coated with 10% Matrigel, instead of collagen or fibrin clot. 6a. Fibers are initially grown in 10% horse serum, 5% chick embryo extract in DMEM. After 3 to 4 days, many spontaneously activated satellite cells have migrated away from the fiber and have proliferated. At this stage the fiber is removed and the monolayer culture of satellite cells is grown in proliferationpromoting medium that consists of 20%fetal bovine serum, 10%horse serum, and 1%embryo extract in DMEM. When cell differentiation is desired, differentiation medium containing 2% fetal calf serum, 10% horse serum, and 0.5% embyro extract in DMEM is used. C. Characteristics

Single fiber cultures have been used to study two basic aspects of satellite cell activity. Bischoff (1986a, 1986b, 1990a, 199Ob) used the system to study the activation of quiescent satellite cells; this required a culture system that did not result in significant spontaneous activation. Consequently, the integrity of the basal lamina was critical, as was the absence of significant numbers of dying fibers that might release activation signals. The selection of protease used in digestion and the selection of the small FDB muscle from the foot of rats were important in minimizing fiber injury and damage to the basal lamina that would result in satellite cell activation. In the procedure described by Rosenblatt et al. (1995), the primary application of this culture approach was to generate monolayer cultures of pure populations of satellite cells from specilk fibers or groups of fibers of the same fiber type for the subsequent study of gene expession following differentiation (Rosenblatt et al., 1996). In this case, having quiescent satellite cells was of little importance, and therefore spontaneous activation was not a problem. This permitted the use of other muscles with longer myofibers, such as the EDL, soleus, and tibialis anterior,that have been extensivelyused in physiological studies and have specific fiber type compositions.The important elements in this adaptation of the isolated myofiber system were the absence of connective tissue cells and the ability to identify the specific fiber from which a group of satellite cells was derived.


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1. Culture Homogeneity As previously mentioned, if care is taken in the fiber isolation procedures, connective tissue cells can be eliminated and pure myogenic cultures can be obtained. 2. Activation/Proliferation The most important attribute of the Bischoff preparations is the ability to maintain satellite cells in a quiescent state in association with a living fiber. This has permitted the examination of the activation process. Once activated, however, the kinetics of entry into the cell cycle and proliferation are similar to that found in monolayer cultures of satellite cells from adult animals. In addition, the temporal expression patterns of the myogenic regulatory factors MyoD and myogenin and the expression of certain cell cycle regulated proteins such as PCNA are very similar to that observed in monolayer cultures (YablonkaReuveni and Rivera, 1994). 3. Differentiation Differentiation of the progeny of satellite cells occurs within the basal lamina tube if it is undamaged, and satellite cells fuse together to form new myotubes. This occurs if the fiber is first killed or even in the presence of a living fiber if satellite cells are activated by chick embryo extract or crushed muscle extract (Bischoff 1986a, 1986b). It is noteworthy that Bischoff reported fusion of satellite cell progeny with each other but not with the adjacent living myofiber as occurs in living muscle. When myofiber cultures are used to generate satellite cells that are subsequently studied as monolayer cultures, differentiation occurs as in monolayer cultures generated from minced muscle.

V. Advantages and Disadvantages of Each Culture System There are advantages and disadvantages to each of the culture systems discussed, and one single culture system may not be adequate for addressing all experimental questions. The following lists some of the advantages and disadvantages of each culture system. A. Monolayer Mass Cultures

1. Advantages 1. Large numbers of satellite cells can be generated from any muscle or group of muscles. This may be essential for providing adequate amounts of material

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for biochemical or molecular analyses, especially when studying the activation of quiescent satellite cells. 2. Important properties of satellite cells that are affected by the physiological state of the donor animal may be studied, whereas the “physiologicalimprinting” may be lost if extensive passaging and expansion of a smaller number of cells is required for analysis. 3. Procedures may be less difficult and timeconsuming than single fiber culture procedures. 2. Disadvantages 1. Although a very high percentage of myogenic cells can be obtained in the primary cultures, it is very difficult to completely eliminate all nonmuscle cell contamination. Monolayer cultures derived from cultured single fibers can, however, provide pure myogenic cultures. 2. Satellite cells often come from muscles of mixed fiber type, and for some experiments this may present a problem, If monolayer cultures are derived from isolated muscle fibers of known fiber type (Rosenblatt et d.,1995), this problem is eliminated, but only small numbers of satellite cells are obtained, and the population must be expanded significantly if large amounts of protein or RNA are required. The importance of fiber type for various aspects of satellite cell activity has not been explored to any significant degree. B. Isolated Single Fiber Cultures (Bischoff System)

1. Advantages 1. Satellite cells can be maintained in a quiescent state for variable periods of time. 2. Nonmuscle cell contamination can be eliminated.

2. Disadvantages 1. Some biochemical and molecular analyses may not be possible because of limitations on amounts of protein and RNA that can be isolated, and the isolated protein and RNA may come from myofibers as well as from satellite cells. 2. Fiber isolation procedures may be more difficult and time-consuming than other techniques.

VI. Conclusions Skeletal muscle satellite cell culture techniques have provided a means for coupling muscle structural and physiological observations to cellular and molecu-


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lar explanations. Intrinsic properties of these myogenic cells have been studied in vitro, and it has become apparent that the satellite cell has properties that are distinctly different from those of embryonic or fetal myogenic cells (reviewed in Bischoff, 1994; Yablonka-Reuveni, 1995). Extrinsic factors that may be involved in regulating the activity of satellite cells in muscle tissue have also been identified by application of satellite cell culture techniques. A variety of growth factors have been shown to have specific effects on activation, proliferation, and differentiation of satellite cells. Several of these have been shown to be present in muscle tissue during growth or repair, and therefore represent candidates for important physiological regulatory roles. By integrating in vivo and in vitro approaches, greater understanding of the cellular and physiological mechanisms of muscle repair and growth are being achieved.

References Allen, R. E., and Boxhorn, L. K. (1989). Regulation of skeletal muscle satellite cell proliferation and differentiation by transforming growth factor-beta, insulin-like growth factor I and fibroblast growth factor. J. Cell. Physiol. l38,311-314. Allen, R. E., and Rankin, L. L. (1990). Regulation of satellite cells during skeletal muscle growth and development. P.S.E.B.M. 19481-86. Allen, R. E., Dodson, h4. V., Luiten, L. S., and Boxhorn, L. K. (1985). A serum-free medium that supports the growth of cultured skeletal muscle satellite cells. I n Vitro Cell. Dev. Biol. 21,636-640. Allen, R. E., Rankin, L. L., Greene, E. A., Boxhorn, L. K., Pierce, P.R., and Johnson, S. E. (1991). Desmin is present in proliferating rat muscle satellite cells but not in bovine muscle satellite cells. J. Cell. Physiol. 149, 525-535. M e n , R. E., Sheehan, S. M., Taylor, R. G., Kendall, T. L., and Rice, G. M. (1995). Hepatocyte growth factor activates quiescent skeletal muscle satellite cells in vitro. J. Cell. Physiol. 165,307-312. Bekoff, A., and Betz, W.(1977). Properties of isolated adult rat muscle fibers maintained in tissue culture. J. PhysioL 27& 537-547. Bischoff, R. (1974). Enzymatic liberation of myogenic cells from adult rat muscle. Anat. Rec. 180, 645-662. Bischoff, R. (1986a). Proliferation of muscle satellite c e b on intact myofibers in culture. Dev. Biol. 115,129-139. Bischoff, R. (1986b). A satellite cell mitogen from crushed adult muscle. Dev. Biol. 115,140-147. Bischoff, R. (199Oa). Cell cycle commitment of rat muscle satellite cells. J. Cell BioL Ill, 201-207. Bischoff, R. (199Ob). Interaction between satellite cells and skeletal muscle fibers. Development 109,943-952. Bischoff, R. (1994). The satellite cell and muscle regeneration. In “Myology,” Vol. I, 2nd edition (A. G. Engel and C. Franzini-Armstrong, eds.), pp. 97-118. New York McGraw-Hill. Bischoff,R., and Heintz, C.(1994). Enhancement of skeletal muscle regeneration: Develop. Dynamics 201,41-54. Clegg, C.H., Linkhart, T. A., Olwin, B. B., and Hauschka, S. D. (1987). Growth factor control of skeletal muscle differentiation: Commitment to terminal differentiation occurs in G1 phase and is repressed by fibroblast growth factor. J. Cell Biol. 105, 949-956. Dodson, M.V., and Allen, R. E. (1987). Interaction of multiplication stimulating activitykat insulinlike growth factor I1 with skeletal muscle satellite cells during aging. Mech. Ageing and Develop. 39,121-128. Feldman, J. L., and Stockdale, F. E. (1991). Skeletal muscle satellite cell diversity: Satellite cells from fibers of different types in cell culture. Dev. Biol. 143,320-334.

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Ronald E. Allen ct al. Frey, R. S.,Johnson, B. J., Hathaway, M. R., White, M. E., and Dayton, W. R. (1995). Growth factor responsiveness of primary satellite cell cultures from steers implanted with trenbolone acetate and estradiol-1%. BAM 5, 71-80. Greene, E. A., and Allen, R. E. (1991). Growth factor regulation of bovine satellite cell growth in vitro. J. Anim. Sci. 69, 146-152. Grounds, M. D., Garrett, K. L., Lai, M. C., Wright, W. E., and Beilharz, M. W. (1992). Identification of skeletal muscle precursor cells in vivo by use of MyoDl and myogenin probes. Cell Tissue Res. 267999-104. Ham, R. G., St. Clair, J. A., Webster, C.. and Blau, H. M. (1988). Improved media for normal human muscle satellite cells: Serum-free clonal growth and enhanced growth with low serum. I n Vitro Cell. Dev. Biol. 24,833-844. Hathaway, M. R., Hembree, J. R., Pampusch, M. S.,and Dayton, W. R. (1991). Effect of transforming growth factor beta-1 on ovine satellite cell proliferation and fusion. J. Cell. Physiol. 146,435-441. Johnson, S. E., and Allen, R. E. (1995). Activation of skeletal muscle satellite cells and the role of fibroblast growth factor receptors. Exp. Cell Res. 219,449-453. Kelly, A. M. (1978). Satellite cells and myofiber growth in the rat soleus and extensor digitorum longus muscles. Dev. Biol. 65, 1-10. Mauro, A. J. (1961). Satellite cell of skeletal muscle fibers. J. Biophys. Biochem. Cyrol. 9,493-495. McGeachie, J. K., Grounds, M. D., Partridge, T. A., and Morgan, J. E. (1993). Age-related changes in replication of myogenic cells in mdx mice: Quantitative autoradiographic studies. J. Neurol. Sci. 119,169-179. Moss, F. P., and Leblond, C. P. (1971). Satellite cells as the source of nuclei in muscles of growing rats. Anat. Rec. 170,421-436. 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. D. V. Biol.-Animal31,773-779. Rosenblatt, J. D., Parry, D. J., and Partridge, T. A. (1996). Phenotype of adult mouse muscle myoblasts reflects their fiber type of origin. Diferentiation 60,39-45. Schultz, E. (19%). Satellite cell proliferative compartments in growing skeletal muscles. Dev. Biol. 175,84-94. Schultz,E., and Lipton, B. H. (1982). Skeletal muscle satellite cells: Changes in proliferative potential as a function of age. Mech. Aging Dev. 20,377-383. Smith, C. K., Janney, M. J., and Allen, R. E. (1994). Temporal expression of myogenic regulatory genes during activation, proliferation, and differentiation of rat skeletal muscle satellite cells. J. Cell. Physiol. 159,379-385. Thompson,S. H., Boxhorn, L. K., Kong, W., and Allen, R. E. (1989). Trenbolone alters the responsiveness of skeletal muscle satellite cells to fibroblast growth factor and insulin-like growth factor I. Endocrinology W, 2110-2117. Webster, C., Pavlath, G. K., Parks, D. R., Walsh, F. S.,and Blau, H. M. (1988). Isolation of human myoblasts with the fluorescence-activated cell sorter. Exp Cell Res. 174,252-265. 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. Yablonka-Reuveni, Z., Quinn, L. S., and Nameroff, M. (1987). Isolation and clonal analysis of satellite cells from chicken pectoralis muscle. Dev. Biol. 119,252-259. Yablonka-Reuveni, Z. (1995). Development and postnatal regulation of adult myoblasts. Microscop Res Tech 30,366-380.

PART I11

Viral and Cellular Gene Delivery Systems in Muscle


CHAPTER 9

Retroviral Gene Delivery Mark J. Federspiel* and Stephen H. Hughest ' Molecular Medicine Program Mayo Clinic and Mayo Foundation Rochester, Minnesota 55905 + ABL-Basic

Research Program NCI-Frederick Cancer Research and Development Center Frederick, Maryland 21702

I. Introduction A. Properties of Retroviruses B. Retroviral Vector Systems C . Targeting Gene Expression in Vivo D. Gene Transfer Efficiency E. The ALV-Based Retroviral Vector System 11. Vector Construction and Virus Propagation A. ALV-Based Retroviral Vectors B. Adaptor Plasmids C. Virus Stocks 111. Avian Model A. In Vitro B. In Vivo IV. ALV/Receptor Mouse Model A. I n Vitro B. In Vivo: Targeting Delivery to Muscle C. In Vivo: Experimental Animals Generated by Infecting Early Embryos D. Amphotropic ALV Vectors V. Summary/Reprise References

I. Introduction The major advantage of retroviral vectors over other gene transfer techniques is the ability of retroviral vectors to deliver genes efficiently and stably to eukaryoMETHODS IN CELL BIOLOGY. VOL 52 Copynghr 0 1998 by Academic Press. AU ngha of reproducaon in any form mewed. 0091-67YX/Y8 $25.00

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tic cells. In animal systems, not only is it important that gene delivery be extremely efficient but, in most cases, stringent control of gene expression in the target tissue and/or at a particular stage of development is also required. In this chapter, we describe the properties of retroviruses and how these properties affect the properties of retrovirus-basedvectors. We compare the advantages and disadvantages of retroviral vectors to those of several other gene transfer techniques. The chapter will also describe the retroviral vector system based on the avian leukosis virus (ALV) retrovirus family we have developed.

A. Properties of Retroviruses 1. The Retroviral Life Cycle We will describe the retrovirallife cycle only briefly (Fig. 1). For more extensive reviews, see Varmus and Swanstrom (1982, 1985), Varmus and Brown (1989), and Coffin (1990a). One of the primary determinants of host range is the ability of a protein on the surface of the virus, the viral envelope glycoprotein (the product of the env gene), to specifically interact with a host cell surface protein, the host receptor. There appear to be at least two distinguishable steps in viral entry; the first involves recognition and binding of the viral envelope glycoprotein to the host receptor, and the second involves steps that lead to the fusion of the

Fig. 1 A diagrammatic representation of the retroviral life cycle.

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host and viral membranes. Following membrane fusion, the viral core is released into the cytoplasm where the single-stranded RNA genome found in the virion is copied into double-stranded DNA by the viral enzyme reverse transcriptase (RT). This reverse transcription process produces a linear dsDNA that is longer than the RNA genome from which it derives. The ends of the linear DNA contain terminally redundant sequences known as long terminal repeats (LTRs). The core particle, which now contains the linear double-strand DNA version of the viral genome, enters the nucleus. Oncoretroviruses and the vectors derived from them appear to require the breakdown of the nuclear membrane that occurs during mitosis for the viral DNA to gain access to the cellular genome. Once this occurs, the linear viral DNA is inserted into the host genome by the virally encoded enzyme integrase. The integrated viral DNA is called a provirus. Efficient production of progeny viruses requires the integration of the viral genome into host DNA. The viral DNA can be inserted into a large number of different sites in the host genome; however, sites where the viral genome are joined to host DNA are precise: the provirus is exactly two bases shorter at each end than the linear DNA from which it is derived. The viral LTRs, which are found at the ends of the double-stranded DNA version of the viral genome and the integrated proviruses, contain all the sequences necessary to direct the synthesis of viral RNA. The structural ( g a g ) proteins of the virion core and the enzymatic (gag-pol)components are translated from genomic length RNA transcripts (Fig. 2). Other viral genes, including the env gene, which codes for the envelope glycoproteins, are translated from spliced RNA that is derived from a genomic-sized RNA precursor. Simple oncoretroviruses make only two RNAs, full-length and the spliced env mRNA; complex retroviruses (such as HIV-1) make additional spliced mRNAs that encode regulatory proteins. Gag and gug-pol are synthesized as polyproteins that subsequently self-assemble. In the majority of retroviruses this process takes place on the inner aspect of the outer membrane of the host cell. The assembly process also incorporates into the virion core two copies of the viral RNA, and a number of small host RNAs that include the tRNAs that serve as the primer for the reverse transcription process. The virus is then budded from the cell and, in so doing, acquires an outer membrane that is studded with the viral envelope glycoprotein. The gag and gag-pol polyproteins are cleaved into the smaller proteins found in the mature virion by a virally encoded protease. Maturation of the virus, which is a result of the proteolytic cleavage of gag and gug-pol, takes place either while, or shortly after, the core is budded through the membrane of the host cell. 2. env Glycoprotein and Receptor Interactions The interaction between the retroviral envelope glycoproteins and a specific cell surface receptor is the first step in determining the susceptibility of a cell to a particular retrovirus (Fig. 3A) (Weiss, 1982). Retroviruses efficiently penetrate the membrane of only those cells that express a specific receptor. Resistance to

A

B Provlrus GAG

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Genomlc RNAI full-length message

Gag polyproteln Gag-pol polyproteln

Spliced message

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AAA,

Envelope glycoprotein

Spliced message

src protein

Fig. 2 RSV virion structure and expression. Panel A shows a diagrammatic representation of a mature Rous sarcoma virus (RSV)virion. The legend at the right identifies individual proteins found in the mature virion. The outer membrane of the virion contains the transmembrane protein (TM), which is associated with the surface protein (SU).The matrix protein (MA) lies just under this outer membrane. The core of the virion is structurally delimited by the capsid protein (CA). Inside the


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retroviral infection can occur at the cell surface in two ways: (i) The cell is genetically resistant, that is, a functional version of the specific receptor is not present on the surface of the cell; or (ii) the receptors are saturated with viral envelope glycoproteins that physically block the receptor, a phenomenon known as receptor interference. The envelope glycoproteinsproduced by an exogenously acquired provirus or from an endogenous provirus can bind to and block the corresponding host cell receptor. Detailed studies of host susceptibility and resistance have been done with both avian and murine retroviruses. The members of the Rous sarcoma virudavian leukemia virus (RSV/ALV) family of avian retroviruses have been classified into five major envelope subgroups (A-E). The susceptibility of avian cells to viruses that express these envelope subgroups is determined by differences at three genetic loci, rv-a, rvb, and rv-c. Tv-a controls susceptibility to subgroup A, tv-c to subgroup C, and rv-b to subgroups B, D, and E. The susceptibilities of a number of strains of chickens have been characterized using RSVlALVs with different envelope subtypes. There may also be variation in the level of susceptibility of various tissues in the animal and at different stages in development. The host range of murine retroviruses has been classified into three groups based on the viral envelope glycoprotein and host receptor interactions: ecotropic, xenotropic, and amphotropic. In general, ecotropic strains of murine leukemia virus (MLV) replicate efficiently only in mouse and rat cells. Xenotropic strains of MLV can replicate efficiently in a wide variety of species including rat, mink, human, and avian cells, but do not replicate efficiently in mouse cells. The amphotropic murine retroviruses can replicate efficiently in both murine and nonmurine cells.

capsid are two viral RNA genomes, shown partially covered with nucleocapsid protein (NC). The two genomic RNAs are hydrogen bonded near their 5’ ends. The core also contains reverse transcriptase (RT), integration protein (IN), and protease (PR). Panel B shows the relationship of the proviral DNA, the open reading kames, viral RNAs, and proteins of RSV. The LTRs of the provirus are shown as a series of three boxes (U3, R, and U5). The viral genome is divided into gag, pol, enu, and src genes. Adjacent host DNA is shown as a wavy double line. In the case of RSV, the gag and env genes are in the same reading frame; pol and src are in a different reading frame. Host DNA-dependent R N A polymerase transcribes the provirus, yielding a full-length RNA that serves both as the viral genome and as the message for the gag and gag-pol polyproteins. This RNA is capped (indicatedby pppG) and polyadenylated (AAAJ. In RSV, approximately 5% of the time that ribosomes reach the end of the gag open reading kame, a ribosomal hrneshifi event occurs that allows the translation of the gagpol polyprotein. The gag and gag-pol polyproteins are processed by the viral protease to yield the proteins found in the mature virion. The relative positions of these components in the polyproteins are shown. The RSV polyproteins contain a p10 protein whose hnction is not well understood. A portion of the full-length viral R N A i s processed by the host splicing machinery, which gives rise to the enu and src mRNAs. The env mRNA is translated into the envelope glycoprotein, which is cleaved into SU and TM by a host cell protease. The src mRNA is translated into the src oncoprotein.

Fig. 3 Biological properties of retroviruses. Panel A depicts the interaction of retroviral env glycoprotein and cellular receptors. (1) A cell that expresses the appropriate receptor proteins can be efficiently infected by both subgroup A and B RSV. An infected cell expresses viral proteins, and produces new infectious virus. If viral envelope glycoproteins are expressed, they bind to and block the specific cell receptor. If the transforming gene carried by RSV, src, is expressed, it alters the cell morphology. (2) A cell that does not express the appropriate receptor for subgroup A RSV cannot be infected by RSV(A) and remains untransformed. A cell that expresses the subgroup B receptor is susceptible to infection by RSV(B) and is transformed. (3) A cell that has been infected by a virus of a particular subgroup is resistant to reinfection by virus of the same subgroup, a phenomenon called receptor interference. For example, if a cell is infected with a subgroup


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3. Viral Gene Expression As we have already discussed, a cell is susceptible to infection by a retrovirus only if the appropriate receptor protein is expressed on the surface of the target cell. This specific recognition allows the viral core to enter the cytoplasm of both replicating and quiescent cells, but does not guarantee that viral infection will be successful. To complete the infection process, the viral dsDNA must integrate into the cellular genome (Fig. 3B). Oncoretroviruses require the infected cell to replicate, with the breakdown of the nuclear membrane, for this to occur (Varmus et aL, 1977; Fritsch and Temin, 1977; Humphries et af., 1981). The requirement that the target cell must replicate limits the delivery of retroviral vectors in animals. Although most or all of the cells in a target tissue express the receptor and will bind and internalize the retrovirus, viral DNA will be integrated in the host genome in only the subset of cells that divide. For example, at birth, the skeletal muscles of mice and rats contain two cell types, the actively dividing myoblasts and the nondividing myotubes (Kelly and Zacks, 1969; Kelly, 1978; Hughes and Blau, 1990). Although virus will bind to and enter both types of cells, only the myoblasts will be infected. Thus, “effective” delivery will be limited by the ratio of replicating to quiescent cells. Moreover, virus that enters nondividing cells will be lost. If only the dividing myoblasts express the retroviral receptor, the efficiency of delivery should increase since the nontarget cells would not “remove” virus by internalization. The skeletal muscles of mice and rats that are more than 15-20 days old contain primarily myotubes, quiescent myoblasts (satellite cells), and very few dividing myoblasts. As a consequence, infecting the skeletal muscles of older animals with a retrovirus is inefficient. Protocols (injury, for example) that induce cell division in adult muscle can be used to generate cellular targets that can be successfully infected with retroviral vectors. B. Retroviral Vector Systems There are two types of vectors derived from retroviruses: replicationcompetent and replication-defective. A retroviral vector that contains in its ge-

(A) avian leukosis virus [ALV(A)], which is related to RSV but does not contain the src gene, the cell is resistant to RSV(A) infection and transformation because the subgroup A receptors are blocked. Receptor interference is specific. Infection by ALV(A) will not prevent the cell from being infected by RSV(B). Viral infection by the simple retroviruses requires that the host cell undergo mitosis (Panel B). The retrovirus gains entry into the cytoplasm of the cell by membrane fusion following the specific interaction of the viral envelope glycoprotein and the host cell receptor. Retroviral entry can occur in both replicating (1) and quiescent (2) cells. Efficient expression of viral proteins only occurs after integration of the viral DNA into the cellular genome in the nucleus. RSV appears to require the breakdown of the nuclear membrane during mitosis to gain access to the cell DNA (1). Redrawn, in part, with permission from Weiss (1982).

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nome all the information needed to complete the viral life cycle will be replication-competent. We have developed a replication-competent retroviral vector system that was modeled after the naturally occurring avian retroviral vector, RSV. ALVs are replication-competentviruses whose genomes contain the three required viral genes gag, pol, and env. RSV is, in essence, an ALV that acquired a cellular oncogene src, which is not required for virus replication. We deleted the src gene from a cloned DNA copy of an RSV genome and inserted in its place the recognition site for a restriction enzyme. This site can be used to insert genes of experimental interest. The advantage of a replication-competent vector is the ease with which high-titer virus stocks can be produced. Specialized helper cell lines are not required. Since the vector replicates, high titers are achieved rapidly by virus spread. Replication-competentvectors are not usually well suited for experiments where widespread dissemination of virus is undesirable, such as cell lineage studies. A replication-defective retroviral vector cannot replicate because it is missing the genetic information for one or more of the viral proteins. Infectious virus can be produced if the missing proteins are supplied in trans in a helper cell designed to complement the defect in vectors. If a replication-defective vector is introduced into a normal cell, there should be no spread of the virus subsequent to the initial infection. Defective vectors can be used to study cell lineage and in experiments where the widespread expression would be toxic. In general, the upper limit on the size of a retroviral genome is about 10 kb. Since portions of the viral genome are deleted in replication-defective vectors, the size of an inserted experimental gene (or genes) can range up to 8 kb depending on the design of the particular vector. In contrast, only about 2.5 kb can be inserted into a replication-competent vector. However, the replication-defective vectors suffer from two major problems: (i) Generally, the infectious titer produced by the vectorhelper cell system is lower, and in some cases, substantially lower, than those of the replication-competent viruses from which they were derived. The efficiency of delivery and gene expression is determined by the initial viral titer. (ii) Recombination among vector, helper, and related endogenous sequences can occur, which results in virus stocks that contain both the defective vector and a contaminating replication-competent virus. Recombination can occur in the helper cell during the production of viral vector stocks or in the target cells (Purcell et al., 1996). Careful monitoring for contaminating viruses must be done to minimize the risk of unintended spread of the vector. Moreover, contaminating replication-competentviruses may grow to detectable levels only after the vectors have been introduced into the target animal.

C.Targeting Gene Expression in Vim For many experimental applications it is necessary to restrict the expression of a transferred gene to a particular tissue or cell type, or to a defined stage of development. Several different approaches have been used successfully. The


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most common approach involves the use of a vector that will deliver the gene of interest to most or all of the cells in a tissue or animal, but in which the expression of the experimental gene is controlled (and restricted) by an internal tissue-specific promoter. The success of this approach is determined by the specificity of the promoter sequences. Internal promoters have been used both in replication-competent and replication-defective vectors (Emerman and Temin, 1984; Palmer et a!., 1987; Overall et al., 1988; Hantzopoulos et aL, 1989; Petropoulos and Hughes, 1991; Petropoulos et al., 1992). An alternative approach is to express the experimental gene constitutively but restrict the delivery of the vector and the gene it carries to a particular target tissue. Retroviral vectors can be restricted to specific cells or tissues if only those cells or tissues express the appropriate receptor. One approach is to prepare chimeras that contain susceptible tissue grafted into an otherwise resistant embryo. This has been done using embryos from appropriately chosen strains of chickens (Fekete and Cepko, 1993; Morgan and Fekete, 1995). Genes can be delivered by retroviral vector into embryos at a variety of developmental stages either before or after the chimera has been surgically constructed. Replicationcompetent vectors can be used and they will spread efficientlywithin the susceptible tissue. The procedure can be used to prevent the gene of interest from causing adverse effects to the rest of the animal. Although the production of surgical chimeras from avian embryos is fairly routine, the technique is labor intensive and requires special skills. An alternative to using surgical chimeras to control which cells or tissues are susceptible to a retroviral vector would be to generate animals that genetically restrict receptor expression to only the target tissue. We have developed two transgenic mouse systems that contain the receptor for ALV-based retroviral vectors: one system in which the receptor is specifically expressed only in striated muscle (Federspiel et aL, 1994), and a second system in which the receptor is under the control of a constitutive promoter and is expressed in most or all tissues (Federspiel et aZ., 1996). These transgenic mouse systems will be described in Section IV. D. Gene Transfer Efficiency

Retroviral vectors remain the delivery vehicle of choice for the stable expression of genes because they can be used to transfer genes efficiently and because the viral genome (including any inserted marker) is stably integrated into the cellular genome. The initial titers of retrovird vector systems vary from 107-108 infectious units (ifu) per milliliter for the replication-competent ALV-based vectors to 16-106 ifu per ml for replication-defectivevectors. To try to address the limitations of retroviral vectors, alternative gene delivery methods have been developed (Partridge and Davies, 1995). The advantages and disadvantages of four alternative systems are outlined next. Adenovirus-based vectors have several advantages (Rosenfeld et aL, 1992; Bett et al., 1993; Ragot et aL, 1993; Tripathy et al., 1994; Ghadge et al., 1995;

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Mitani et al., 1995).These vectors can theoretically carry much larger segments of DNA, grow to very high titers (-loll) and can infect nondividing cells efficiently. However, two major disadvantages appear to limit the potential applications of adenovirus-based vectors in vivo. The viral genome is not stably integrated into the cellular genome, so that, with time, gene expression will be lost. Repeated applications of the vector to restore gene expression may well be limited by the fact that these vectors express adenovirus genes and are likely to stimulate a potent immune response. A recent study reported that, in some cases, firstgeneration adenovirusvectors (those with the Ela and E l b genes deleted) elicited a strong cellular immune response upon the initial vector infection, which led to development of pathology including damage from inflammation and destruction of the genetically modified cells (Yang et al., 1994;McCray et al., 1995; Yang et al., 1995).Further improvements in the vectors that would reduce or eliminate the expression of the adenovirus genes by the vectors will be needed to moderate the cellular immune response. In addition, it is not known whether it is possible to limit the expression of an experimental gene to particular cells or tissues (with internal tissue-specific promoters, for example) by adenovirus vector delivery. Adenoviral vector gene delivery will be discussed in another chapter. Gene delivery vectors have been developed based on the adenoassociated virus (AAV), a replication-defective parvovirus (for a review, see Flotte and Carter, 1995). AAV replication requires co-infection with a helper virus, an adenovirus,herpes virus, or vaccinia. In the absence of a helper virus, the AAV genome integrates into the host genome and may be rescued by helper virus infection. AAV vectors have been shown to infect and express genes in both dividing and quiescent cells. However, it is unclear whether AAV integrates in nondividing cells and gene expression is less robust. The ability of AAV vectors to integrate, persist, and be expressed in vivo is still unclear and will require additional studies. A potential problem that has not been tested is the fate of an integrated AAV vector upon a subsequent helper virus infection. Will recombinant AAV virus be generated? The size limit of AAV vectors appears to be 4.7 kb. The titers of AAV vectors ( e l @ ) has been limited by technical problems in preparing stable helper lines and the purification of helper-free stocks. It is not clear whether AAV vector infection and expression can be made specific for particular cells or tissues. Several methods of physically introducing naked DNA into cells have been tested. Plasmid DNA, plasmid DNA/liposome complexes, and plasmid DNA in complexes designed to target specific cellular receptors have all been shown to deliver DNA into cells and result in the expression of the encoded protein. All naked DNA delivery techniques have one major problem: the efficiency of delivery is so low that it is impractical for many applications. However, this technique does show promise for immunization, especially if the DNA is injected intramuscularly or intradermally. Enough cells take up the DNA and express the encoded protein to initiate an immune response (Wolff et aL, 1990; Acsadi et al., 1991;Ulmer et al., 1993;Wang et al., 1993;Raz et al., 1994;Wang et al., 1995).


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Genetic engineering of somatic cells ex vivo, and the subsequent transplantation of the altered cells into the animal, has been moderately successful. Both retroviral vectors and adenovirus vectors, as well as DNA transfection, have been used to transfer genes into cells in culture. Engineered skeletal muscle myoblasts have produced several proteins, including human growth hormone and a minidystrophin protein in mice (Barr and Leiden, 1991; Shawan et al., 1991; Ragot et al., 1993; Rando et al., 1995). The highest levels of expression in animals were achieved using engineered cells from transformed muscle cell lines (e.g., C2C12). Clearly there are physical limits on the number of engineered cells that can be delivered into a particular tissue. This is obviously a problem in the skeletal musculature. It is not yet clear whether normal cells, for example, myoblasts, will express proteins to as high a level as the transformed cells. If special cell lines are required, this raises important issues of histocompatibility and host rejection of the transplanted cells. Myoblast transfer into muscle is the subject of another chapter in this book. E. The ALV-Based Retroviral Vector System

The rest of this chapter describes the ALV-based retroviral vector system that we have developed (Fig. 4) (Hughes et al., 1987; Greenhouse et al., 1988; Petropoulos and Hughes, 1991; Petropoulos et al., 1992; Boerkoel et al., 1993; Federspiel and Hughes, 1994). This system can be used in both avian and murine animal models. The ALV retroviral vector system is the only replicationcompetent retroviral vector that can deliver experimental sequences of reasonable size (up to 2.5 kb). These vectors can be used to transfer genes efficiently into avian cells. The avian embryo is uniquely accessible for experimental manipulation and an extensive repertoire of in vitro and in vivo techniques has been developed to take advantage of this accessibility. The ALV retroviral vectors can be used to express experimental genes under the control of the constitutive viral LTR promoter, or from an internal tissue-specific promoter. The gene for the cellular receptor for the subgroup A ALVs has been cloned (Bates et al., 1993; Young et al., 1993). This cloned receptor has been used to generate lines of transgenic mice that are susceptible to the ALV-based retroviral vector system (Federspiel et al., 1994,1996). Since mammalian cells lack receptors for efficient infection by ALV (Weiss, 1992), targeting viral infection, and as a consequence, gene expression, can be accomplished by generating transgenic mouse lines that express the receptor in a specific tissue. There are several advantages to using the ALV-based retroviral vector system rather than a murine retroviral vector in mammalian cells. ALV vectors can be quickly and easily grown to high titers (107-108ifu per ml) as replicationcompetent viruses on avian cells but are inherently replication-defective upon infection of mammalian cells (Weiss, 1982; Wills et al., 1989; Berberich et al., 1990; Nasioulas et al., 1995). It is a simple matter to produce the ALV vector stocks on avian cells that lack related endogenous sequences with which the


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A Replication Competent GAG

I

RCOSIRCON

I

t

R

K II [ID

RCOSBPIRCONBP

RCAWRCAN

RCASBPlRCANBP I

I

I

I

I

I

I

I

0

1

2

3

4

5

6

7

1

8 kb

B Replication Defective Cia I BBAN

&

penvA

0

I

I

I

I

I

I

I

1

2

3

4

5

6

7

I

8 kb

Fig. 4 Schematic representation of the ALV-based retroviral vectors. Panel A shows the replication-competentvectors that contain three genes gag, pol, and env and a unique CluI cloning site, flanked by long-terminal repeats (LTRs). The RCOS and RCON vectors contain LTRs derived from Rous-associated virus 0 (open). The RCAS and RCAN vectors contain Schmidt Ruppin-A (SR-A) RSV LTRs (closed). The BP (Bryan polymerase) vectors contain the pol region from the Bryan strain of RSV (BH-RSV) in place of the prototypic SR-A RSV sequences. The pol regions were moved on EcoRI (R) to KpnI (K) fragments. Some of the vectors contain a splice acceptor (SA) required for the expression of an experimental gene under the control of the viral LTR (e.g., RCOS). Other vectors lack the SA and require an internal promoter to express an inserted gene (e.g.. RCON). Panel B shows the replication-defectiveretroviral vector system. The vector BBAN was created by deleting the env region of the RCASBP vector. The envelope glycoproteins are supplied in trans by the QenvA helper cell line that expresses the subgroup A SR-A RSV env under the control of the murine leukemia virus LTR promoter. The env gene was linked to the SV40 polyadenylation signal.

vectors could recombine (Astrin et al., 1979). Recombination of the ALV vectors with either exogenous or endogenous virus is unlikely to occur in mammalian cells since mammalian cells lack related viral sequences and the avian viruses do not replicate in these cells. The replication-defective murine retroviral vectors usually grow to titers of 1@-106 and are produced on cells that contain homologous viruses that can recombine with the vector and result in contaminating replication-competent helper virus and loss of experimental sequences.

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11. Vector Construction and Virus Propagation A. ALV-Based Retroviral Vectors

We have developed a series of replication-competent retroviral vectors that derive from the Schmidt-Ruppin A (SR-A) strain of Rous sarcoma virus (RSV). The genome of the viral vectors is present in proviral form on a pBR-based plasmid (Fig. 4A) (Hughes et af., 1987;Greenhouse er uf., 1988).The characteristics of these vectors are summarized in Table I. The RSV src gene, which is not required for viral replication, was replaced by a unique CfuI recognition site that can be used for the insertion of a wide variety of DNA sequences. The insert size is limited to 2.5 kb. Larger inserts, and sequences of any size that are toxic to the cell, will be deleted from the virus during propagation. There are four types of vectors that differ in the transcriptionalelements that are used to control the expression of the inserted gene: RCAS, RCAN, RCOS, and RCON [RCAS, Replication-Competent, ALV LTR, Splice acceptor; RCAN, ReplicationCompetent, A L V LTR, No splice acceptor; RCOS Replication-Competent, Rous-associated virus type 0 (RAV-0) LTR, Spxce acceptor; RCON, Replication-Competent, RAV-5 - LTR, No splice acceptor]. Two of these groups Table I ALV-Based Retroviral Vectors Vector

Subgroups Viral genes available in vector

LTR

RCASBP A,B,D,E

gag, pol, env ALV (enh+)

A,B,D,E

gag,pol, env ALV (ehn+)

RCAS

RCANBP A,B,D

gag,pol, env ALV (enh+)

RCAN

gag, pol, env ALV (enh+)

A,B,D

RCOSBP A,B,D

gag, pol, env RAV-0 (enh-)

RCOS

A,B,D

gag,pol env RAV-0 (enh-)

RCONBP A,B,D

gag, pol, env RAV-0 (enh-)

RCON

A,B,D

gag,pol, env RAV-0 (enh-)

BBAN

A

gag,Pol

ALV (enh+)

None

ALV (enh+)

TFANEO None

Purpose

Titer

Comments

Replication-competent; High level expression from Bryan-RSV pol LTR via a spliced RNA Replication-competent High level expression from LTR via a spliced RNA Expression of genes from Replication-competent; Brym-RSV POI an internal promoter Replication-competent Expression of genes from an internal promoter Replication-competent; Moderate level expression from LTR via a spliced Bryan-RSV pol RNA Low level expression from -16 Replication-competent LTR via a spliced RNA Expression of genes from >Id Replication-competent; Bryan-RSV pol an internal promoter Expression of genes from -16 Replication-competent an internal promoter Expression from an internal 104-106 Replication-defective; promoter or add a Bryan-RSV pol splice acceptor Transfection vector Direct expression from with neo selection LTR

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of vectors use the promoter region contained in the viral LTR to control the expression of an experimental gene via a spliced RNA transcript (RCAS and RCOS). In the RCAN and RCON vectors, which lack a splice acceptor, an internal promoter can be used to control the expression of an experimental gene (Petropoulos and Hughes, 1991; Petropoulos et al., 1992). The origin of the viral vector LTRs affects both the viral expression titer and the level of expression of an internal gene. The SR-A LTRs that are present in the RCAS/RCAN vectors contain a powerful enhancer that enables the virus to replicate efficiently and grow to high titers. However, these viruses have oncogenic potential in vivo that results in tumors and mortality in birds older than 4-6 months. RCOS/ RCON viruses contain the Rous-associated virus 0 (RAV-0) LTRs, which do not contain a powerful enhancer, and consequently these vectors grow to lower titers. The RCOSLRCON viruses have little or no oncogenic potential in vivo. The viral titers and level at which experimental genes are expressed by all four of these vector viruses can be increased by substitutingthe pol region of the Bryan strain of RSV (BH-RSV) for the SR-A RSV pol region (RCASBP, RCANBP, RCOSBP, and RCONBP) (Nemeth et al., 1989; Petropoulos and Hughes, 1991; Petropoulos et al., 1992; Federspiel and Hughes, 1994). Any one of the five ALV envelope subgroups (A-E) can be used in these vectors. The specific envelope gene in a vector is shown in parentheses after the vector name [e.g., RCASBP(A), the RCASBP vector with the subgroup A envelope gene]. We have also developed a replication-defectiveALV vector system based on the Bryan strain of RSV (Fig. 4B). The Bryan virus contains the gag, pol, and src genes, but lacks most of the env gene (Lerner and Hanafusa, 1984; Sudol et al., 1986). The env glycoproteins can be supplied in trans from either an endogenous or an exogenous source. In the system we developed (Boerkoel et al., 1993) the env glycoproteins are produced in the QT6 cell line (QenvA) from an expression cassette that contains the U3 promoter region of the murine leukemia virus LTR, the ALV subgroup A env gene, and a polyadenylation site from SV40. The plasmid that carries the defective retroviral vector BBAN (BryanRSV defective vector, Bryan-RSV pol gene, ALV LTR, No splice acceptor) contains two cassettes: aproviral cassette thatencodes the%BAN vector, and a selection cassette that consists of the neomycin resistance gene under the control of the chicken P-actin promoter. In an effort to reduce recombination events that would result in the production of replication-competent viruses, the vector and helper sequences were designed to minimize the amount of shared sequence. Experimental sequences of 4-4.5 kb can be accommodated in this vector. BBAN does not contain a splice acceptor for the expression of an experimental gene from a spliced RNA. BBAN also contains a unique ClaI site for the insertion of experimental sequences. Inserts can be subcloned as described later in either the SACLA12NCO adaptor plasmid for expression under the control of the viral LTR, or into the CLA12 plasmid if an internal promoter is used. Virus-producing cell lines are generated by transfecting the BBAN/

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experimental gene plasmid into the QenvA cell line and selecting for neomycinresistant cells. B. Adaptor Plasmids

Experimental sequences must be modified to produce CluI ends before being inserted into the CluI site of the ALV vectors. A series of adaptor plasmids have been designed that contain multiple cloning sites flanked by CluI sites (Hughes et ul., 1987). The simplest adaptor plasmid contains only an array of cloning sites designed to avoid any sequences that interfere with transcription or translation. Adaptor plasmids have been designed to supply an efficient initiator ATG and a eukaryotic leader suitable for use in the RCAS/RCON vectors. The characteristics of the three most widely used adaptor plasmids are summarized in Table 11: CLA12 contains a restriction site polylinker flanked by CluI sites; CLA12NCO contains a eukaryotic transcription leader and ATG start codon (contained in an NcoI site) followed by the CLA12 polylinker; SACLA12NCO contains a splice acceptor site in addition to the ATG and leader sequences present in Clal2NCO. These three plasmids were all derived from a deleted version of pBR327 and are relatively low-copy plasmids in E. coli. A high-copy version of CLA12NCO has been constructed (pUCCLA12NCO) that produces significantly higher amounts of DNA. However, the adaptor plasmids are only used as intermediates in vector construction and if there is a concern that the inserted sequence might be somewhat toxic in E. coli, the higher copynumber adaptor plasmids may cause problems. CLA12 was designed so that the polylinker sequence can be used in either orientation without interfering with the transcription or translation. The CLA12 adaptor plasmid can be used to add CluI sites conveniently to the ends of promoterlgene cassettes. In this case, the transcriptional control elements for the experimental gene are contained on the internal promoter.

Table I1 Adaptor Plasmids Adaptor

Polylinker

CLA12

pUC12

CLAl2NCO

pUC12N

SACLA12NCO pUC12N

Purpose

Comments

Does not interfere with transcription or translation in either orientation Prototype NcoIlATG adaptor is To provide the insert with a used to ensure efficient eukaryotic leader sequence and expression of inserts in RCASI an initiator ATG RCOS vectors To provide insert with a eukaryotic Used with the defective vector BBAN to express genes from leader and initiator ATG and a the LTR promoter splice acceptor To convert a variety of DNA sequences into CIaI fragments


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To achieve the highest level of gene expression, we recommend using CLA12NCO in combination with the RCASBP vector. This adaptor both contains a transcriptional leader sequence and has a consensus ATG start codon contained in an NcoI site (CCATGG). These sequences work very efficiently with the promoterlenhancer elements of our retroviral vectors and can be used to express genes at high levels. Synthetic oligonucleotidescan be used to replace a region of the inserted sequence between the ATG and a convenient downstream restriction site, or the 5 ‘ region of the sequence to be inserted can be modified using PCR to produce appropriate restriction sites. If the nucleotide coding sequence does not begin ATGG . . . , two versions of the vector can be constructed one that contains the wild-type protein coding sequence and one that generates an NcoI site at the ATG (which will change the second amino acid of the protein). In some cases we have found that using the consensus translational start site CXATGG . . . significantly increases the levels of protein translated from viral RNA. If both types of sequences are used, the effects of these changes, both on the levels of protein that are produced and on the ability of the protein to function appropriately, can be tested directly. The polylinker downstream of the NcoI site in CLA12NCO derives from the E. coli expression plasmid PUC12N. This means that exactly the same DNA sequence engineered into CLA12NCO can be introduced into both the E. coli expression vector and a retroviral vector and that the same protein will be produced in both prokaryotic and eukaryotic cells. Adaptor plasmids that contain different sets of cloning sites as well as an alternate initiator ATG contained in the restriction site of NdeI (CATATG) have also been generated. An adaptor plasmid can easily be designed for a specific application, but it is important to keep in mind that the sequences contained in the polylinker should not interfere with transcription or translation. To optimize viral titer and gene expression we recommend that the gene under study be modified for insertion into the vector. The inserted DNA fragment should not contain introns or a polyadenylation site. Introns will be removed during the viral replication cycle, and the presence of an internal polyadenylation site will interfere with the production of full-length viral RNA. In general, it is advisable to delete essentially all of the 5’ and 3’ untranslated sequences. This should remove many of the elements that could interfere with viral gene expression. These manipulations can be easily carried out using the adaptor plasmids.

C. Virus Stocks Virus stocks are produced on chicken embryo fibroblasts (CEF) cultures established from individual 10-11-day embryos. The embryo is taken from the egg and the head, legs, and internal organs are removed. The remaining body wall is passed through a 5-ml syringe and is then mixed with 5 ml of a solution containing 0.5 mg per ml collagenase and 0.125 mg per ml dispase in PBS


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(phosphate buffered saline, no Ca2+or Mg2+),and digested for 30 min at 4°C. The mixture is then vortexed gently but thoroughly, and the resulting suspension is allowed to stand for 5 min to let the large debris settle. Cells are collected from the supernatant by centrifugation at 2000 rpm for 10 min at 4°C. The cell pellet is resuspended in 30 ml CEF medium and plated onto three 100-mm Falcon tissue culture plates. CEF medium consists of Dulbecco's modified Eagle's medium (Gibco-BRL, Bethesda, Maryland), 10% tryptose phosphate broth (Gibco-BRL), 5% fetal bovine serum (Hyclone Inc., Logan, Utah), 5% newborn bovine serum (Gibco-BRL), 100 units penicillin per ml, and 100 pg streptomycin per ml (Advanced Biotechnologies Inc., Silver Spring, Maryland). It is important to test individual lots of serum to determine which lots support efficient growth of CEFs. CEFs are grown at 39°C at 5% C 0 2 and split 1:3 when confluent using Trypsin de Larco (Advanced Biotechnologies Inc.). Some embryos do not produce CEFs that grow well in culture. If cells from a particular embryo grow well, these cells should be passaged until approximately 20 plates are obtained and the bulk of the cells frozen for future use. CEFs are stored in liquid nitrogen by mixing 1 part cells to 1 part freezing medium (10 ml CEF medium, 2 ml dimethylsulfoxide, 1 ml fetal bovine serum), frozen at -7O"C, and transferred to liquid nitrogen. Virus propagation is initiated by transfection of the plasmid DNA containing the proviral form of the retroviral vector into low-passage CEF (Fig. 5). For transfecting CEF, we prefer the calcium phosphate precipitation method (Kingston et al., 1989) followed by a brief glycerol shock. Plasmid DNA (5-10 pg) is mixed with sterile water to a volume of 437.5 pl, and 62.5 p12 M CaClz is added. While vortexing, add 500 pl 2X Hepes solution dropwise and let the mixture stand at room temperature for 5 min. The 2X Hepes solution consists of 270 mM NaCl, 10 mM KC1,l mM HaHP04, and 42 mM Hepes adjusted to pH 7.08 (the pH is critical), filter sterilized, and stored at -20°C. The DNA, CaC12, and Hepes mixture should form a fine, diffuse, bluish-white precipitate. The precipitate is added to 10 ml of CEF medium on a 100-mm plate of CEF at approximately 30% confluence and incubated at 39°C for 4 hr. After incubation, the medium should be removed, and 5 ml of a CEF medium containing 15% glycerol solution is then added to the transfected cells. Incubate at 39°C for 35 min, wash once with 10 ml PBS, and add fresh CEF medium. Depending on the retroviral vector employed, maximum viral titer is achieved after 4-10 cell passages. Infected cells can be stored in liquid nitrogen for future virus harvest. Virus stocks can also be stored by removing supernatants from infected cultures. Floating cells and debris are removed by centrifugation at 2000 rpm at 4°C for 10 min, and the stocks are transferred to a fresh tube and stored frozen at -70°C indefinitely or at -20°C for short periods of time. Viral titer will decrease from 2- to 10-fold with each freezekhaw cycle. Since retroviruses are inherently unstable, it is good practice to initiate the preparation of fresh viral stocks by transfection. Do not passage the virus from cell to cell unless it is absolutely necessary for the experimental protocol.

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Fig. 5 General procedure for generating and applying ALV-based retroviral vector virus stocks. High-titer retroviral vector stocks and/or virus infected chicken embryo fibroblasts (CEFs) can be used to deliver experimental genes to both avian and mammalian cells. Examples of in vitro and in vivo applications of the retroviral vector system are outlined.

Viral spread can be monitored by assaying cell-culture supernatants for viral GAG antigens by Western transfer or by assaying for reverse transcriptase activity (Petropoulos and Hughes, 1991). Virions are collected by ultracentrifugation of cell culture supernatants in an SW40 rotor at 35K for 1hr at 4°C. Genomic DNA can be prepared from infected cultures and analyzed by Southern transfer to verify the structure of the proviruses. Protein expression from the experimental gene can be analyzed by Western transfer with appropriate antisera and/or by a functional assay (if one is available) to demonstrate that the expressed protein has the expected biological activity.

III. Avian Model A. In Vitro The avian experimental system has relatively few permanent cell lines, and most of these are not suitable for the generation of ALV vector stocks. The

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viruses do grow on QT6 cells, a chemically transformed quail fibroblast cell line (Moscovici et al., 1977); unfortunately, the viral titers are much lower on QT6 cells than what can be achieved by growing the virus in CEF (Petropoulos and Hughes, 1991). However, careful attention to reagents and culture conditions makes it possible to passage CEF about 25 times. ALV virus stocks are produced on Line 0 CEF whenever possible, since Line 0 chickens do not contain endogenous sequences that are closely related to the ALV vectors. This eliminates recombination between the vector and endogenous viruses and makes it significantly easier to monitor viral DNA, RNA, and protein structure and expression. Line 0 cells are susceptible to ALV env subgroups A-D but are resistant to subgroup E (CE). Line 0 eggs can be obtained from the USDA-ARS, Avian Disease and Oncology Laboratory (3606 East Mount Hope Road, East Lansing, Michigan 48823), for a nominal charge. Currently, ALV retroviral vectors are available that have envelope subgroups A, B, D, and E, and it would be a simple matter to produce subgroup C versions if they are needed. An experimental gene can be expressed at a variety of levels simply by using an appropriately chosen vector to deliver the gene. Table I11 summarizes the range of chloramphenicol acetyl transferase (CAT) activity expressed by RCOS, RCOSBP, RCAS, and RCASBP in CEF (Federspiel and Hughes, 1994). Since vectors with different subgroups are available, this system can also be used to deliver multiple genes (Fig. 5 ) (Givol et af., 1994). CEF can be infected first with a replication-competent ALV vector with, for example, a subgroup A envelope, followed by a second infection with a vector with another envelope, subgroup B. This results in a culture in which two genes have been delivered and expressed virtually in every cell. This type of “double-hit” experiment is quite convenient for studying how genes cooperate in cultured cells. The replication-competent ALV vector system can also be used to infect primary cell cultures to study the effect of genes involved in, for example, muscle differentiation. RSV has been shown to infect and transform a wide variety of cells in vitro, including myoblasts (Kaign et aZ., 1966; Fiszman and Fuchs, 1975). Myoblasts transformed by RSV fail to differentiate and fuse into myotubes Table III CAT Expression in Injected CEFs“ ~~

Passage no.

1

3

6

9

RCOSlCAT (A) RCOSBPlCAT (A) RCASlCAT (A) RCASBPKAT (A)

3 13 1,300 25,000

2 28 7,000 21,000

4 63 8,000 30,000

10 350 12,000 33,000

The CAT activities given in the table derive from the percentage of chloramphenicol acetylated in a standard reaction and quantitated using radioanalyticimaging. High levels of CAT activity were determined using serial dilutions of the protein extract to keep the assay in the linear range. Reprinted with permission from Federspiel and Hughes (1994).

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(Fiszman and Fuchs, 1975; Holzer et al., 1975) and show altered muscle-specific protein expression (Miskin et aL, 1978). We have used the replication-competent ALV vectors to deliver and express a variety of proteins in primary cultures of myoblasts and in myotubes derived from these myoblast cultures (C. Gruber, S. Forry-Schaudies, and S. H. Hughes, unpublished observations). B. In Vivo

1. Gene Delivery by Somatic Infection Standard DNA microinjection techniques developed for generating transgenic mammals are not practical in birds. The avian egg is fertilized in the oviduct before the shell has been added and is relatively inaccessible at this stage. By the time the egg is laid, the embryo consists of approximately 50,000 cells. Consequently, the efficient delivery of experimental genes in chickens has been limited to infection of somatic and germ cells by retroviral vectors. Two groups have generated germ-line insertions by infecting embryos of newly laid but unincubated eggs with retroviral vectors (Fig. 5 ) (Salter et al., 1987; Bosselman et al., 1989a). We routinely inject 50-100 p1 of either a virus stock or virusproducing CEF (lo6cells) near the blastodisc of unincubated Line 0 eggs. Virusinfected chicks are identified at hatch using an ELISA assay for the viral GAG protein. The efficiency of the resulting somatic infection varies from one bird to another. In some of the birds the germ line is also infected. It should be noted that the susceptibility of particular cell types to infection by different ALV envelope subgroups has not been extensively characterized. It is likely that the distribution of host receptors for ALV varies between cell types and during development. If it is important to infect a particular cell or tissue type, we recommend that the susceptibility of that target cell be tested with vectors of different env subtype. The percentage of viremic chicks produced and the degree of somatic infection will vary depending on which of the vectors is used. The variability is due to the growth characteristics of the vectors. About 20-30% of the chicks injected with vectors that grow to relatively low titers (i.e., RCOS) are viremic. If high-titer vectors (i.e., RCASBP) are used, 50-8096 of the chickens are viremic. If the high-titer vectors are used, the experimental gene will be present in a majority of cells and tissues. The low-titer viruses are less efficient. The amount of virus that can be injected is limited. A simple method to increase the efficiency of infection is to inject infected CEFs rather than a virus stock. The injected CEFs produce an amount of virus that far exceeds the amount of virus that can be delivered in 50-100 pl of virus stock. The result is an increase in the frequency of viremic chicks and in higher levels of infected cells and tissues. For long-term experiments (where the birds must survive for more than 4 months), for example, for the generation of transgenic lines, it is important to use vectors with low oncogenic potential and to accept the lower efficiency of gene delivery.We have generated 23 transgenicchicken lines using vectors similar


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to the prototype replication-competent RCOS and RCOSBP vectors (Salter etal., 1987;Crittenden et al., 1989). The somatically infected male chickens transmitted ALV retroviral vector DNA to their offspring at frequencies ranging from 1 to 11%.The proviral inserts expressed both RNA and protein products (Salter and Crittenden, 1989; Federspiel et al., 1991). One drawback of the chicken model is that the reproductive cycle of chickens is longer than that of mice; it takes 2-3 years to produce an experimental stock of transgenic chickens.

2. Tissue-Specific Expression The utility of the somatic infection approach was tested in experiments in which a muscle-specific promoter, the chicken aSk-actinpromoter, was linked to the reporter gene chloramphenicol acetyl transferase (CAT). We had previously shown that the cYsk-actinlCATgene cassette that was used in the RCANBP vector was muscle specific in transgenic mice (Petropoulos et al., 1989) (Fig. 6A). To test whether an ALV retroviral vector system carrying this same muscle-specific gene cassette would deliver the gene to all of the tissues but limit the expression of the experimental gene to the skeletal muscles, CEFs infected with the RCANBP vector carrying the ask-actin/CAT gene cassette were used to infect Line 0 embryos. Viremic chicks were identified and expression of viral GAG protein and CAT activity determined in various tissues (Petropoulos et al., 1992). As shown in Fig. 6B, essentially all of the tissues in the infected birds expressed high levels of the viral capsid protein (p27) showing that the vector had infected a wide variety of tissues. However, CAT activity was limited to skeletal muscle and heart (Fig. 6B; CAT), showing that the expression of the internal a,k-actin promoter was independent of viral gene expression. This in vivo experimental system can be used to produce a number of experimental animals in a few weeks. The somatic infection model may be especially useful for the in vivo characterization of promoters and the study of genes involved in development. An advantage of the avian system is that development occurs primarily in ovo, and the embryo is easily accessible for vector infection and for physical manipulations and observation.

3. Replication-Defective Retroviral Vectors In an effort to limit viral spread after initial gene delivery, we and others have constructed replication-defective avian and murine retroviral vector systems (Watanabe and Temin, 1983; Cepko et al., 1984; Miller et al., 1995; Hock and Miller, 1986; McIvor et al., 1987; Price et al., 1987; Danos and Mulligan, 1988; Galileo et al., 1990; Lynch and Miller, 1991; Palmer et al., 1991; Scharfmann et al., 1991; Fields-Berry et al., 1992; Snyder et al., 1992; Boerkoel et al., 1993; Riviere et al., 1995; Sadelain et al., 1995). Bosselman and co-workers have generated transgenic chicken lines by infecting day 0 embryos (as described earlier) with a replication-defective reticuloendotheliosis virus (REV)-based vector de-


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A

6 CAT

P27

Fig. 6 Tissue-specific expression of a gene inserted into an ALV-based retroviral vector. Panel A shows a representative chloramphenicol acetyl transferase (CAT) assay performed on tissue homogenates prepared f'rom transgenic mice carrying a chicken aSk-actinpromoter-CAT construct. The acetylated forms of chloramphenicol are marked with arrows. Panel B shows tissue-specific CAT expression in infected chickens. Tissue homogenates were prepared from a chicken somatically infected with RCANBP/q&AT/F, a replication-competentRCANBP vector that contains the identical Gk-actin promoter-CAT cassette that was tested in transgenic mice (Panel A). CAT activities (CAT) and the levels of the retroviral capsid protein (p27) were quantitated from the same tissue homogenates. Proteins were separated by polyacrylamide gel electrophoresis and transferred to nitrocellulose. The viral capsid protein p27 was detected using a polyclonal rabbit anti-p27 antibody (see Petropoulos er al., 1992). The position of the p27 capsid protein is marked with an arrow.


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veloped in Howard Temin’s lab (Bosselman et al., 1989a). They observed a relatively high incidence of viremic chicks as a result of infection by recombinant replication-competent REV (Bosselman et al., 1989b; Briskin et al., 1991). This illustrates the major problem of using defective vector systems. However, in these experiments the viremic birds could be eliminated and experimental birds were obtained that contained stable replication-defectiveproviruses. The ALV-based replication-defective QenvAIBBAN vector system has been tested in vim and in vivo in an effort to detect recombinant replication-competent viruses (Boerkoel et al., 1993). Sanes and co-workers (Galileo et al., 1990) have reported that the vector system they developed (62QT-6) generated little or no recombinant replication-competent virus. We compared the two systems both in vitro and in vivo. The P-galactosidase gene @gal) was used as a phenotypic marker gene in both vector systems to quantitate the titers of the viral vectors. No replication-competentvirus was detected with the QenvAIBBANfigalvector system either in the helper cell line used to prepare the virus stocks or in the subsequent infection of Line 0 embryos. However, the 62QT-6 vector system produced recombinant virus that was detectable in vitro and upon infection of Line 0 embryos. These results should serve as a warning that great caution is needed if defective retroviral vector systems are used for an experimental application that requires absolutely no virus spread, for example, cell lineage analyses. As described earlier, replication-defective retroviral vector systems can produce significant levels of recombinant replication-competentvirus. Sensitive techniques are available for the detection of recombinant viruses. For example, long-term assays carried out on endogenous virus-free cells have been used to detect as few as 10 replication-competent avian retroviruses. Assays are now being developed that increase the level of detection of replication-competent murine retroviruses in replication-defectivevirus stocks. A number of laboratories have shown that retroviruses are very adept at recombination and that vectors derived from retroviruses recombine at high rates to yield recombinant viruses (Dougherty and Temin, 1986; Dougherty et al., 1989; Coffin, 1990b; Katz and Skalka, 1990; Pulsinelli and Temin, 1991; Olson et al., 1992; Varela-Echavarria et al., 1993). This makes it extremely difficult to develop a replication-defective retroviral vector system that will not generate replication-competent recombinant viruses. This is in part the rationale behind the development of the ALV receptor mouse system. The ALV vectors are replication-competent in avian cells but are naturally replication-defective in mammalian cells. We do not believe the virus can overcome the inherent replication defect it has in mammalian cells.

IV. ALV/Receptor Mouse Model A. In Vitro Because they lack appropriate receptors, mammalian cells are not susceptible to efficient infection by ALVs. In addition, if an ALV provirus is introduced

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into a mammalian cell, new infectious viruses are not produced. This appears to be primarily due to differences in viral RNA splicing and protein transport in avian and mammalian cells. The genetically defined locus for the subgroup A ALV receptor, fv-a, has been cloned from quail-cell DNA (Bates ef al., 1993; Young et aL, 1993). Mammalian cells that express tv-a are susceptible to efficient infection by ALV(A); however, these infected mammalian cells do not produce virus. The ALV(A) receptor can be introduced into cell lines by transfection to produce lines that are susceptible to ALV(A) retroviral vectors (Fig. 5). Recombination and instability will be rare because neither the avian cells used to produce the vector stock nor the mammalian target cells contain sequences that are closely related to the vector. The viral titers (107-108 ifu per ml) are high enough to infect the majority of the mammalian cells in a culture so that efficient gene delivery can be achieved without drug selection. Since ALV does not produce high levels of envelope glycoprotein in mammalian cells, the infected cells can be infected with a second vector containing a second gene.

B. h UVO: Targeting Delivery to Muscle

The cloning of the gene for the ALV(A) receptor made it possible to develop a mouse model system in which the delivery of the ALV vector and the consequent expression of the experimental gene would be restricted to particular cells or tissues (Fig. 5). To restrict ALV susceptibilityto muscle, several transgenic mouse lines were generated that express a fv-a cDNA under the control of the musclespecific chicken ask-actin promoter (Federspiel et al., 1994). Previous work showed that the particular promoter segment caused the expression of a linked CAT gene primarily in the skeletal muscles and heart of transgenic mice (Fig. 6A) (Petropoulos ef al., 1989).

1. Gene Delivery to Muscle: Virus Stock The susceptibility of the ask-actin/receptortransgenic lines to infection by ALV(A) was initially characterized by injecting thigh muscles of 5-, lo-, and 15day old mice with retroviral vector stocks (0.02-0.05 ml at day 5; 0.05-0.07 ml at day 10; 0.07 ml at day 15) using a tuberculin syringe. Two viruses were used: RCASBP/AP(A) that contains the human placental alkaline phosphatase gene (AP),and RCASBP/CAT(A).Expression of the AP gene can be easily visualized by standard histochemical procedures. The RCASBP/CAT(A) virus was used to quantify viral delivery and gene expression. One week after infection, animals were sacrificed and their tissues analyzed. All four transgenic mouse lines were susceptible to infection by ALV(A) virus stocks. The highest level of susceptibility was found in line 1494 (Fig. 7A, see Color Plates), which was used for all of experiments discussed next.

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9. Renoviral Gene Delivery

2. Mitotic Requirements The efficiency with which the leg muscles of line 1494 mice can be infected declines rapidly after 5 days of age (Table IV). All simple retroviruses (including ALV) appear to require cell division to complete their life cycle. If we extrapolate from data on skeletal muscles in rats (Kelly and Zack, 1969; Kelly, 1978; Hughes and Blau, 1990), the ability to infect the skeletal muscles of line 1494 mice correlates with the mitotic potential of the myoblasts. Since the expression of experimental genes delivered by any of the available retroviral vector systems is restricted to dividingcells,the design of the experiment must take this limitation into account. Certain complex retroviruses (Le., HIV-1) can successfully infect nondividing cells, so it may eventually be possible to overcome this limitation. 3. Long-Term Expression The ability of RCASBP/CAT(A)-infected muscle to maintain the expression of CAT was determined at 1, 5, 12, 20, and 35 weeks after infection of 5-dayold line 1494 mice (Table V). The total CAT activity in the leg increases as the animal ages in parallel with the increase in the mass of the leg muscle and the increase in muscle-specificproteins (e.g., myosin, actin). There is an increase in the specific activity of CAT between weeks 1 and 5; however, after that time, the level of CAT specific activity remains relatively constant through at least 35 weeks of age. In all experiments, the infection was confined to the injected leg and did not spread to the contralateral leg. As expected, nontransgenic siblings that were injected in parallel were not infected.

4. Gene Delivery to Muscle: Infected CEFs In an effort to increase the efficiency of vector delivery into muscle, an alternative virus delivery procedure was tested. CEFs producing the retroviral vectors Table IV Susceptibility of (UAKE Line 1494 Leg Muscle to ALV (A) Infection as a Function of Age" Injected

Animals tested

Average CAT activity (range)

Relative activity

6 4 4

2500 (1400 to 4330) 200 (100 to 240) 85% confluent. (a) To split the cells, aspirate off the media and add 1-1.5 ml0.05% trypsinl 0.53 mM EDTA. Incubate the culture dish at 37°C for 2-5 min. (b) Once the cells begin to lift off the plate, add 5 ml DMEM + 10% FBS + P/S. Trypsinized cells are then transferred into 50-ml tubes and briefly centrifuged at loo0 rpm in a Beckman GPR centrifuge. (c) To have cells at 80% confluency in 2 days, one dish of 90-100% confluent cells should be split 1:2 or 1:3. To have cells about 80% confluent in 3-4 days, split 1:5 or 1:6. If the cells are plated too sparsely they will not proliferate. 2. Construction of Recombinant Adenovirus: Cotransfection of HEK 293 Cells (a) Cotransfection is carried out using HEK 293 cells at 80-90% confluency. Cells are maintained in 60-mmdishes with DMEM + 10%FBS P/S and fresh media is added =24 hr prior to the transfection. (b) The adenovirus DNA (e.g., pJM17) and the shuttle plasmid containing the expression cassette are cotransfected into 293 cells using calcium phosphate

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precipitation. This reaction is started by adding 10 pg of the shuttle vector and 10 p g of pJM17 to 500 pl of 2X HBS and then adding H 2 0 to a volume of 937.5 pl in a sterile 1.5-ml microfuge tube. Invert the tube several times to mix, add 62.5 pl of 2M CaC12, and mix the reaction again. Allow the precipitate to form at room temperature for 45-60 min. (c) For transfection, add 500 p1 of the Capo4 precipitate dropwise to each 60-mm dish of HEK 293 cells and incubate at 37°C for 4-5 hr in a 95% 0 2 : 5 % C 0 2 incubator. (d) Remove the cloudy and turbid medium by aspiration, wash the cells with PBS, and then add 4 ml DMEM + 10% FBS + P/S medium. (e) Exchange the medium with 3 ml of fresh medium the next day to remove dead cells. (f) Add medium (2 ml) every 3-4 days without removing any medium. (g) Plaques begin appearing 7-10 days following transfection and the full cytopathic effect (CPE) will be seen within 3-5 days of plaque appearance. (h) Collect cells after full CPE is evident into 15-ml polypropylene tubes. (i) Expose the cell suspension to three cycles of freezing and thawing to lyse any intact cells and release the virus. 6) Centrifuge at 2500 rpm for 10 min to separate cellular debris from the virus. The supernatant, which contains the viral stock, is aliquoted and stored at -20°C. C. The Plaque Assay

The plaque assay is used to obtain clonal recombinant adenovirus, and also for titering viral stocks. The basic protocol, which is adapted from earlier protocols (Graham and Prevac, 1991), is described next, along with the steps that are unique for each application of this assay. Southern blots are used to confirm the structure of each recombinant adenovirus clone. (a) One day prior to the plaque assay, HEK 293 cells should be split 1:3 into 60-mm dishes. (b) To start the plaque assay, serial dilutions of viral stock ranging from to are prepared in DMEM + 2% FBS + P/S. (c) Aspirate the medium from the HEK 293 cells, add a 1-ml aliquot of each virus dilution to individual dishes of HEK 293 cells, and allow adsorption for 1 hr at 37°C. (d) Remove the medium and overlay the dishes with 8 ml MEM/Agar per dish. (e) Allow the agar to solidify at least 30 min at room temperature before returning the dishes to the incubator. (f) Plaques begin appearing in 7-10 days and can then be counted for titering purposes.

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(g) To plaque purify recombinant adenovirus, collect individual plaques by transfering a plug of agar containing a well-isolated plaque to a cryotube containing 1 ml DMEM + 10% FBS + P/S. Freeze at -20" until ready to expand. (h) Plaques are expanded by adding the entire 1 ml of media containing the plug to a 60-mm culture dish containing nearly confluent 293 cells. (i) After 1hr, add 3 ml DMEM + 10% FBS + P/S and incubate at 37°C until CPE is visible. 6 ) Plaque expansions are collected using three cycles of freezing and thawing followed by centrifugation as described earlier. (k) The supernatant containing the viral lysate is collected and stored in cryotubes at -20°C. CsClz purification of crude virus is performed using established protocols (Graham and Prevec, 1991).

IV.Analysis of Myotilament Protein Expression and Incorporation in Adenovirus-Infected Ventricular Myocytes Myofilament protein isoform remodeling can be characterized using both Western blot analysis and immunohistochemicalstaining of the cardiac myocytes. The actual ratio of the myofilament protein isoforms is determined using Western blot analysis. In addition, the stoichiometry of the myofilament protein under study can also be determined in relation to that of other cellular proteins. Immunofluorescence is used to determine whether the uniform striation pattern is maintained at the level of the single myocyte and can be used for direct evidence of the incorporation of a delivered myofilament gene product into the sarcomere. Together, these techniques, which are described next, determine the extent of isoform remodeling taking place within the myofilaments of the ventricular myocytes (Westfall et al., 1997).

A. Western Blot Analysis (a) Samples are separated by SDS-PAGE at constant current (20 mA) for 5 hr, transblotted onto PVDF membrane (0.45 pm) for 2000 volt-hours, and then fixed in PBS containing 0.25% glutaraldehyde as described by Westfall et al. (1996). (b) Immunodetection is carried out by initially blocking nonspecific binding sites with Tris-buffered saline (TBS; Tris = 100 mM, pH 7.5, NaCl = 150 mM) containing 5 % nonfat dry milk overnight. Milk can be increased up to 10% w/v to remove background unless biotinylated antibodies are desired, in which case Tween, gelatin, or albumin should be used as blocking agents. 0.5% milk containing the primary (c) Blots are then incubated in TBS monoclonal Ab (mAb) for 1.5 hr and then washed three times in TBS with 0.5% milk for 5 min each.

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(d) Incubate blot for 1 hr in TBS + 5% milk containing a horseradish peroxidase-conjugated goat anti-mouse Ab. (e) Antibody binding is detected by enhanced chemiluminescence. (f) Background is minimized if conjugated Abs are affinity purified and preadsorbed against serum proteins from other species. Both primary and secondary Abs should be titered to optimize protein detection. B. Indirect Immunofluorescence with Dual Monoclonal Antibodies

The strategy used for dual immunofluorescence detection depends largely on the ectopic protein expressed in the cell under study. Expression of an exogenous protein not found in a particular cell type is readily detectable, and in this case, only a single primary Ab is needed for indirect immunofluorescence detection. However, studies of contractile protein structure and function will necessitate detection of diminished endogenous protein andlor appearance of exogenous protein within the contractile apparatus. In the example shown in Fig. 3, immunofluorescence is used to follow the loss of the endogenous myofilament protein isoform and thus gives an indication of the number of cells that are completely remodeled. Antibodies that can detect the appearance of an isoform would give the first indication of when the protein is incorporated into the myofilament. Although the number of commercially available antibodies that are specific for individual contractile protein isoforms continues to expand, mAbs for isoforms of each contractile protein are not generally available. In addition, few mAbs are capable of recognizing mutant proteins. One option to overcome primary mAb availability is to use tag sequences on the protein under study, which can then be identified with a mAb. The primary concern with this option is that incorporation of the exogenous protein into a complex system such as the myofilament, as well as the function under study, may be directly affected by the tag sequence itself. The protocol for indirect immunostaining with two monoclonal Abs (mAbs) is shown in Fig. 3 and involves incubations with a primary mAb that,recognizes only one isoform of the protein under study (either the new isoform or, as shown in Fig. 3, step 1, the disappearing isoform) and a Texas Red (TR)-conjugated secondary Ab (step 2), followed by neutralization of unreacted sites on these Abs (step 3). A second set of incubations is subsequently carried out with a second primary Ab that recognizes an alternative isoform of the protein or recognizes both isoforms (step 4) and a fluorescein isothiocyanate (F1TC), conjugated secondary Ab (step 5). For example, this staining procedure can be used to distinguish the cardiac isoform of troponin I (TnI) from other TnI isoforms (Westfall et d.,1996,1997)within adenovirus-infected ventricular myocytes in primary culture. (a) Fix cells on coverslips in 3% paraformaldehyde in phosphate-buffered saline (PBS, pH 7.40; 2.3 m M NaH2P04, 8.0 mM Na2HPO4, 150 mM NaC1,

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2

4

5

Fig. 3 Dual monoclonal antibody labeling strategy. In this protocol, one mAb recognizes the endogenous contractile protein isofom (detected with Texas Red-conjugated secondary Ab), while the second mAb recognizes all isofoms of this protein (detected with FITC secondary Ab). Using this strategy, the loss of the endogenous isofom is then followed at various time points after gene transfer.

2.5 mM KCl) for 30 min. Note; Do not allow the paraformaldehyde solution to be heated above 70°C during preparation of the fixative. (b) Wash samples three times in PBS for 5 min each. (c) To minimize background from excess aldehydes, incubate coverslips in 50 mM W C l (in PBS) for 30 min, followed by three 5-min washes in PBS. (d) Prepare humidity chamber for blocking and antibody incubation steps. This chamber typically consists of a shallow container with a lid, H20-saturated gauze to line the container, and wood sticks to elevate coverslips or slides above the gauze. (e) For the initial blocking step, add -200 pl PBS containing 0.5% Triton X100 (PBS-TX) and 20% normal goat serum (NGS) to each coverslip. Incubate at room temperature for 30 min. The PBS-TX + NGS solution should be passed through a 0.2-pm filter to remove debris.

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(f) Drain off blocking solution and add primary mAb diluted in PBS-TX containing 2% NGS. Incubate at room temperature for 1.5 hr. The concentration of NGS can be adjusted in all blocking and mAb incubation steps to optimize mAb binding and protein detection. (g) Wash samples three times in PBS-TX for 5 min each. (h) Nonspecific secondary Ab binding is blocked by incubating samples in PBS-TX containing 20% NGS for 30 min. (i) Incubate samples with TR-conjugated goat anti-mouse Ab diluted in PBSTX plus 2% NGS for 1 hr. Wash each sample three times in PBS-TX for 5 min each. (j) Neutralize unreacted sites on the first primary mAb and TR-conjugated secondary Ab by incubating samples in goat anti-mouse IgG (Sigma) overnight at 4°C. (k) Incubate samples with goat anti-mouse Fab fragments (Jackson Immunochemicals) for 1.5 hr at room temperature. Both the IgG and Fab fragments are diluted 1:20 in PBS-TX with 2% NGS. These steps eliminate any crossrecognition by the second set of primary and secondary Abs. (1) Block samples with 20% NGS in PBS-TX for 30 min at room temperature. (m) Incubate samples in a second primary mAb diluted in PBS-TX plus 2% NGS for 1 hr at room temperature. Wash each sample three times in PBS-TX for 5 min each. (n) Block sample in PBS-TX containing 20% NGS for 30 min at room temperature. ( 0 ) Binding of the second primary mAb is detected with a fluorescein isothiocyanate-conjugated goat anti-mouse Ab diluted in PBS-TX plus 2% NGS. Samples are incubated with this secondary Ab for 1 hr at room temperature. (p) Rinse samples in PBS, and mount as described by Kleyman et al. (1991). Samples are now ready for microscopic evaluation.

V. Summary and Future Directions In summary, recombinant adenovirus vectors can be used to accomplish highly efficient myofilament gene transfer, expression, and incorporation into adult ventricular myocytes in primary culture. This approach is made possible, in part, by a primary culture system that allows for the maintenance of the both the cellular and subcellular differentiated state of adult cardiac myocytes in primary culture. This new approach will be an important adjunct to transgenic technology, as both share the goal of molecular dissection of the cardiac contractile apparatus. The primary strengths of the viral delivery system are the rapid and efficient transfer of myofilament genes into adult cardiac myocytes. These features have the potential to facilitate greatly the timely and comprehensive analysis of myo-

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filament protein structure-function relationships in the context of the intact cardiac myocyte. However, whether this system can be adapted for functional studies of other myofilament proteins besides troponin I has not yet been demonstrated. The system outlined here depends critically on the “window of time” over which the differentiated state of the adult myocyte is retained in culture, and also depends on the normal protein turnover rate of the myofilament protein under study. In this regard, it is known that there is significant variation in the half-lives of key myofilament proteins in the heart. For example, the half-life of troponin I is 3.2 days, whereas for the myosin light chains the half-life approaches 9 days (Martin, 1981). Thus, structure-function analysis of myosin light chains using this system will likely require a further increase in the time window over which the differentiated state of the adult cardiac myocyte is maintained in primary culture. Finally, the demonstration of myofilament gene transfer into cardiac myocytes in vitro raises the exciting prospect of using recombinant viral vectors to modify myofilament gene expression, and thus the function, of the working myocardium in vivo. Results using reporter gene constructs generally demonstrate the modest and transient expression of delivered genes in the heart (Kass-Eisler et aL, 1993). Whether this can be improved via secondthird generation viral vectors or, in fact, whether myofilament genes will be expressed and incorporated into the contractile apparatus in myocytes in vivo must await further experimentation. Acknowledgments We thank Mr. Nathaniel Lee for technical assistance. This work was supported in part by grants from the National Institutes of Health and the American Heart Association, National Center and Michigan affiliate. J. M. Metzger is an Established Investigator of the American Heart Association. M. Westfall is a trainee in the Molecular and Cellular Cardiology program at the University of Michigan Cardiovascular Research Center.

References Becker, T. C., Noel, R. J., Coats, W. S., Gomez-Foix,A. M., Alam, T., Gerard, R. D., and Newgard, C. B. (1994).Use of recombinant adenovirus for metabolic engineering of mammalian cells. In “Methods in Cell Biology,” Vol. 43. (M. G. Roth, ed.) New York Academic Press. Geisterfer-Lowrance, A. A. T., Christe, M., Conner, D. A., Ingwall, J. S., Schoen, F. J., Seidman, C. E., and Seidman, J. G. (1996).A mouse model of familial hypertrophic cardiomyopathy.Science

272,731-734. Graham, F. L., and Prevec, L. (1991).Manipulation of adenovirus vectors. In “Gene Transfer and Expression Protocols”(E. J. Murray, ed.). Clifton, New Jersey: Humana. Haworth, R. A., Hunter, D. R., and Berkoff, H. A. (1980).The isolation of Caz+-resistantmyocytes from the adult rat. J. Mol. Cell. Cardiol. 12,715-723. Kass-Eisler A., Falck-Pedersen, E., AIvira, A., Rivera, J., Buttrick, P. M., Wittenberg, B., Cipriana, L., and Leinwand, L. (1993). Quantitative determination of adenovirus-mediated gene delivery to rat cardiac myocytes in vitro and in vivo. Proc. Natl., Acad Sci USA. 90,11498-11502. Kirshenbaum, L. A., MacLeUan, W. R., Mazur, W., French, B. A., and Schneider, M. D. (1993). Highly efficient gene transfer to adult rat ventricular myocytes by recombinant adenovirus.J. Clin. Invest. 92,381-387.

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Kleyman,T.R., Kraehenbuhl, J.-P., and Emst, S.A. (1991). Characterization and cellular localization of the epithelial Nazt channel. J. Biol. Chem. 266,3907-3915, Martin, A. F. (1981). Turnover of cardiac troponin subunits. J. Biol. Chem. 236,964-968. Metzger, J. M., Parmacek, M. S., Barr, E., Pasyk, K., Lin, W., Cochrane, K. L., Field, L. J., and Leiden, J. M. (1993). Skeletal troponin C reduces contractile sensitivity to acidosis in cardiac myocytes from transgenic mice. Proc. Natl. Acad Sci USA 90,9036-9040. Muthuchamy, M., Grupp, I. L., Grupp, G., O’Toole, B. A., Kier, A. B.,Boivin, G. P., Neumann, J., and Wieaorek, D. F. (1995). Molecular and physiologicaleffects of overexpressingstriated muscle p-tropomyosin in the adult heart. J. Biol. Chem. 270,30593-30603. Palermo, J., Gulick, J., Colbert, M., Fewell, J., and Robbins, J. (19%). Transgenic remodeling of the contractile apparatus in the mammalian heart. Circ. Res. 78,504-509. Rust, E. M., Westfall, M. V.,Johnson, R., Lee, N., and Metzger, J. M. (19%). Efficient gene transfer into primary ventricular myocytes mediated by recombinant adenovirus. Biophys. J. 70, A170. Rust, E. M., Michele, D., Westfall, M. V., and Metzger, J. M. (1997). Adenovirus-mediated troponin T gene transfer and expression in adult cardiac myocytes. Biophys. J. (abstract), 72, A175. Westfall, M. V., Samuelson, L. C., and Metzger, J. M. (1996). Troponin I isoform switching in embryonicstem cell-derived cardiac myocytesfollowsthe vertebrate cardiac development program. Devel. Dynamics 206,2638. Westfall, M. V., Rust, E. M., and Metzger, J. M. (1997). Slow skeletal troponin I gene transfer, expression, and myoflament incorporation enhances adult cardiac myocyte contractile function. Proc. Natl. Acad. Sci. 94,5444-5449.

CHAPTER 16

In Vivo Approaches to Neuromuscular Structure and Function Rita J. Balice-Gordon Department of Neuroscience University of Pennsylvania School of Medicine Philadelphia, Pennsylvania 19104-6074

I. Introduction 11. In Vivo Analysis of Skeletal Muscle and Neuromuscular Innervation A. Equipment B. Preparation of the Animal and Surgical Exposure of Muscle C . In Vivo Visualization of Synaptic Components of Neuromuscular Junctions D. Digital Imaging of Skeletal Muscle and Neuromuscular Innervation 111. Manipulation of Motor Neurons, Muscle Fibers, and Synapses in Vivo IV. In Vivo Observations and Manipulations of Muscle Fibers and Neuromuscular Innervation: Practical Considerations V. In Vivo Approaches: New Insights into Neuromuscular Plasticity A. Development of Neuromuscular Innervation B. Maturation of Synaptic Maintenance C. Age-related Alterations in Synaptic Maintenance VI. Summary References

I. Introduction The nervous system communicates with skeletal muscle via connections between motor neurons and muscle fibers called neuromuscular junctions. Compared to synapses in the central nervous system, neuromuscular junctions are large, simply organized, and relatively accessible for structural and functional analyses in vivo and in vitro. For these reasons, neuromuscular junctions have METHODS IN CELL BIOLOGY, VOL. 52 Copyright 0 1998 by Acadenuc Prm. All nghu of rcproducoon in any form mewed 0091-h79X/98 $25.00

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been used for several decades as a model synapse for studying how neural input affects muscle function. Neural modulation of muscle properties involves communication between motor neurons and muscle fibers in both the antero- and retrograde directions. This has been appreciated from a large number of experimental studies as well as analyses of motor neuron and muscle fiber diseases in which neural input to muscle is compromised. Even with as advantageous a preparation as nerve and muscle, most studies involve making observations at single time points, meaning that the dynamic aspects of neuromuscularinteractions have been inferred rather than observed directly. Given that the interdependence among motor neurons and muscle fibers is essential for the maintenance of normal function, and that these interactions are ongoing processes and are difficult to recapitulate in vitro, significant insights can be derived from studies of neuromuscular plasticity in living animals. The inability to observe neuromuscular innervation directly over time under different circumstances has hampered our understanding of neural regulation of muscle properties and of the dynamic balance between plasticity and stability of neuromuscular junctions. For example, damage to muscle fibers induces degeneration, followed under normal conditions by muscle fiber regeneration. Motor neurons respond rapidly to muscle fiber damage, within several dozen hours extending long axonal and nerve terminal sprouts that regress over many days as muscle-fiber integrity is restored. The sorts of interactions between muscle fibers and motor neurons that are altered as a result of muscle fiber damage and how these change over time can be studied to best advantage in situ. A number of technical advances have permitted longitudinal studies of neuromuscular innervation over hours to months in vivo. Lichtman and his colleagues (Lichtman et al., 1987; Purves and Voyvodic, 1987), among several other groups (Hill and Robbins, 1991;Hill etal., 1991;Herrera etal., 1990,Herrera and Banner, 1990, Chen et al., 1991; Chen and KO,1994; Wernig et al., 1991; LangenfeldOster et al., 1993; Wigston, 1989, 1990), have developed staining and imaging techniques that have allowed the same neuromuscular junctions and muscle fibers to be monitored over time in living mice. This approach has provided new insights into the mechanisms underlying neuromuscular development and maturation in normal, diseased, and manipulated animals. This chapter describes the practical aspects of these techniques and discusses their some of applications, strengths, and limitations.

II. In Vivo Analysis of Skeletal Muscle and Neuromuscular Innvervation Skeletal muscle and its innervation are ideal for in vivo analysis as they are accessible in living animals without extensive invasive surgery,muscle innervation is relatively simply organized, and motor neurons and muscle fibers can be easily

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and separately manipulated. Studying dynamic alterations in neuromuscular innervation in living animals over time requires several complementary experimental approaches.Modifications to commonly used microscopy equipment, repeated surgical exposure of muscles of interest, vital fluorescent staining of selected cells, digital imaging of neuromuscular junctions and muscle fibers of interest, and manipulation of innervation or of muscle fibers are discussed next.

A. Equipment Visualization of fluorescently labeled structures of interest can be performed with a modified upright epifluorescence microscope. Modifications to the microscope body and to the stage facilitate surgical exposure of the muscle of interest and optical access to neuromuscular junctions and skeletal-muscle fibers.

1. Microscope The optical train of the microscope can be spatially dissociated from the stage in one of several ways. In the configuration shown in Fig. 1, we separated the stage from the body of an infinity-corrected epifluorescence microscope (Leica, DMR) mounted on linear bearings (Newport). Both were mounted to a vibration isolation table (Technical Manufacturing Corp., Micro-G). When pulled forward on the linear bearings, the optical train of the microscope engages a magnetic stop, positioning the microscope objectives in their usual position over the stage. A planetary gear focus mechanism was added beneath the stage and mounted to a coarse positioning jack (Newport Inst.) mounted to the table. When pushed back from the stage on the linear bearings, the surface of the stage becomes accessible to accommodate an intact animal. A stereomicroscope (Leica MZ8 on a boom stand; Diagnostics, Inc.) can then be pulled into place over the stage and used to perform the surgical exposure of the muscle(s) of interest, position the animal and the muscle, and, at the termination of the experiment, allow wound closure and animal recovery. With this configuration, the micromanipulators and other devices used to position the tissue can remain in position as the microscope is pushed back from the stage. This is especially useful if the animal needs to be repositioned, re-anesthetized, or otherwise accessed during the experiment. Moreover, micromanipulators can be mounted on the stage and positioned for intracellular injection or physiological analysis, and these do not have to be repositioned if the microscope is pushed away from the stage. A second configuration used by Lichtman and his colleagues contains a pivot in the microscope body so that the optical train can be swung away to the left or right over the stage (see Fig. 1 in van Mier et al., 1994; Fig. 1 in Purves and Voyvodic, 1987). Limited clearance between the objectives and the stage surface makes it necessary to reposition micromanipulators and other devices each time the pivot is engaged. Although an upright epifluorescence microscope is quite useful, similar longitudinal studies of neuromuscular junctions have been performed on modified inverted microscope configurations as well (Robbins and Polak, 1988; Hill and Robbins, 1991; Hill et al., 1991).

Fig. 1 Video miscroscopy setup for in vivo imaging. (A) Photograph of an infinity-corrected epifluorescence microscopy set up to facilitate imaging of living animals. The optical train of the microscope (Leica DMR, focusable stage) has been mounted on linear bearings (Newport) below the base. The linear bearings are in turn mounted to the surface of an optical bench vibration isolation table (Technical Manufacturing Corp., Inc.). The stage and focusing mechanism have been separated from the body of the microscope and are mounted directly to the table. The microscope can be pushed away from the stage, allowing access to the surface of the stage to position a living animal. A magnetic stop allows the original position to be accurately restored by pulling forward. Incident light is provided via 100-W Hg or 75-WXe lamps mounted to the rear of the microscope body (not visible in photograph) and shuttered under software control (Lambda 10, Sutter Inst.). A silicon intensified target camera (SIT)camera (Dage-MTI, Inc.) is mounted in the camera port on top of the microscope. The output of the camera is sent to a high-resolution imaging board (Matrox, Inc.) and images are digitized, acquired, displayed, analyzed, and processed using a PCbased image-processing system (MetaMorph, Universal Imaging, Inc.) (B)Close-up view of setup shown in A. Magnetic stop is in lower right corner and linear bearings are visible adjacent

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2. Stage and Animal Plate Variable-focusstages or fixed stages can be adapted to this sort of design, and each has strengths and limitations depending on the application. We have two set ups in the lab, one with a variable focus stage (fixed objectives) as shown in Fig. 1, which we find useful for in vivo imaging, and a second setup in which the stage is fixed with variable-focus objectives. This is useful for intracellular injection and physiological recordings, as electrodes, micromanipulators, and so forth are more easily positioned beneath water immersion lenses (not shown). Because of their small size, mice are easily accommodated on a modified stage, although any animal is feasible with the configuration shown in Fig. 1 with an appropriately designed animal plate and suitably large stage (large stages, up to 18 X 18 inch, designed for industrialapplications,are readily available for biological use). The animal plate consists of an acrylic base approximately 4 inch in thickness with a ca. 3 X 4 X 1/3inch well positioned in the middle (Fig. lB, C). The edges of the plate are covered with magnetized stainless steel.

3. Other Equipment Several additional pieces of equipment are useful. Compact three-axis hydraulic micromanipulators(Narashige,Inc., Newport, Inc.) for injection or physiologi-

to stop. The microscope has been pushed back on the linear bearings so that the surface of the stage is exposed. A stereomicroscope has been swung into place over the stage and is used for animal preparation and exposure of muscles of interest. An adult mouse has been placed supine on a Plexiglas animal plate surfaced with magnetized stainless steel attached to the microscope stage. (C)Close-up view of stage plate and adult mouse shown in B. The animal is anesthetized and mechanically respirated via a small polyethylene tube inserted into the trachea (tubing anchored with a magnet in bottom of photograph). The animal’s teeth are anchored with a rubber band held by two magnets. The skin of the neck has been opened to expose the left sternomastoid muscle. Beneath the skin, the salivary glands overlying the sternomastoid muscle are held out of the way with miniature retractors (small magnets, top of photograph). Following staining, muscles are stabilized, if necessary, with a small firepolished glass bar slipped underneath the muscle away from the muscle nerve, and a small glass coverslip is lowered over the endplate band (three-axis micromanipulators in lower left and middle right of photograph). The muscle is supehsed with saline throughout the experiment, and vital dyes are applied directly over the muscle as necessary. (D)Following staining, the stereomicroscope is swung away from the stage, and the microscope objectives pulled back into place over the animal. Thus, excellent optical access to superficial structures in skeletal muscle can be achieved. A similar approach can be used to incorporate intracellular injection, drug or DNA vector delivery, or physiological analysis of skeletal-muscle fibers or of neuromuscular junctions, by mounting a small three-axis hydraulic micromanipulator to a magnet directly on the stage (not shown). Use of long-working-distance water-immersion objectives (e.g., Zeiss 40X 2.2-mm working distance ceramic-coated 0.75 n.a. lens, not shown) allows the electrode to be positioned between the lens and structures of interest. The electrode and muscle surface can easily be visualized with a cross-polarizing cube and epi-illumination. Photography by Lee Wojnar, Wojnar Photography, Inc., Philadelphia, Pennsylvania.

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cal analyses can be mounted on magnetic bases and positioned directly on top of the stage. A small animal respirator (Model 683; Harvard Apparatus, Cambridge, Massachusetts;Model SAR-830, C W , Pennsylvania) is useful to control breathing during image acquisition, although depending on the muscle(s) selected for study this may not be necessary. Miniaturized retractors and three axes micropositioners mounted onto magnets (Herbach and Rademan, Pennsylvania; Fig. 1C; made by E. Horn) are useful for animal positioning and exposure of muscles of interest. B. Preparation of the Animal and Surgical Exposure of Muscle

1. Adult Mice To prevent pain and discomfort to the animal during the surgical exposure of the muscle and to prevent movement, mice are deeply anesthetized. Although sodium pentobarbital (Nembutal) injected intraperitoneally or inhalant anesthetics are adequate for this purpose, respiratory depression often lengthens recovery time. A mixture of ketamine and xylazine (0.17 mg/ml ketamine (KetaSet, Fort Dodge Lab., Iowa); 1.7 mg/ml xylazine (Anased, Lloyd Lab., Iowa) in 0.9% NaCl; 5.0 d k g injected intraperitoneally) allows a deep level of anesthesia to be rapidly attained within minutes. Animals are anesthetized for 20-30 min and recover quite rapidly (10-15 min) when warmed. Following anesthesia, animals are positioned with rubber bands attached to small magnets so that the muscle of interest can be exposed (Fig. lB, C). To control respiration during image acquisition, mice are intubated with a small polyethylene tube inserted into the trachea and attached to a small animal ventilator (see Section I.A.3). Care should be taken not to overinflate the lungs inadvertently, especially in young animals. Many muscles are easily accessible with small incisions in the skin and retraction of the underlying fascia. These include the sternomastoid muscle in the neck, the cutaneous pectoris over the rib cage, the gluteus in the hindquarters, and the tibialis anterior and gastrocnemius muscles of the hindlimb. The skin over the muscle of interest is prepared by shaving followed by treatment with a depilatory (Nair, Carter-Wallace, Inc.) to remove remaining hair. The skin can then be swabbed with 70% EtOH or betadine. For most studies the sternomastoid or gluteus muscles are used because of their easily accessible location, ease of exposure, minimal dissection, and easily located endplate band, The sternomastoid muscle is exposed by placing the animal on its back, making a small (3-5 mm) incision in the ventral skin of the neck between the chin and the sternum, and reflecting the skin and underlying salivary glands laterally with small retractors (Fig. 1C). Following the skin incision, tissues are superfused with sterile mammalian Ringers solution (e.g., Travenol, Abbott Labs). Muscle fibers or neuromuscular junctions can then be stained and visualized as described later (Section 1.C).

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Following staining, using the stereomicroscope, a ca. 1-mm-wide fire-polished stabilizing bar mounted to a micropositioner is placed beneath the belly of the muscle, away from the nerve, and is raised slightly. A small fire-polished circular or oval glass coverslip (thickness 1 or 1.5; lens coverslip correction is typically 0.17 mm; ca. 4-5 mm in diameter) mounted to a micropositioner is lowered over the muscle. These damp movements caused by arterial pulsations and allow optical access to superficial neuromuscular junctions and muscle fibers. The gluteus muscle, a large, flat, and simply organized muscle in the hindquarters, is exposed with a 5- to 6-mm incision in the overlying skin, which is then deflected laterally, and the underlying connective tissue is freed by blunt dissection with care taken not to damage underlying muscle fibers. The bulbocavernosus muscle in the perianal region of male mice is also easily accessible, with a small incision in the scrotum and reflection of the gonads (Balice-Gordon et al., 1990). No stabilizing bar is necessary, and a larger coverslip (10 mm wide) can be used. Other groups have used superficial muscles such as the cutaneous pectoris (Hill and Robbins, 1991; Hill et al., 1991), the gastrocnemius complex (Wigston, 1990) and the deeper soleus (Wigston, 1989). A number of other muscles are also readily accessible. At the end of the experiment, tissues are returned to their original position and the skin sutured with 6-0 silk (American Cyanamid Corp., Connecticut). Mice are allowed to recover under a heating lamp or a thermoregulated blanket (Harvard Apparatus).

2. Neonatal Mice The procedures described above have been modified to study neonatal animals (Balice-Gordon and Lichtman, 1993). In pups less than 1 week old, lowering body temperature is a safe, fast, and effective way to achieve anesthesia. Pups can be placed on ice for 5-6 min, and cryoanesthesia can be maintained with a small cold brass block placed in the well in the animal plate on the stage. Respirations and heartbeat are slowed so that artificial respiration is often unnecessary. Pups older than 1week can be safely anesthetized with ketamine/xylazine, although longer times are necessary for recovery than in adult animals. 8-0 silk can be used to suture the skin, and the ends of interrupted sutures should be kept short so that the wound is not reopened by the mother. The skin is also painted with a liquid bandage preparation (e.g., NexaBand, Veterinary Prod. Lab., Phoenix, Arizona) so that the suture line is protected and the wound completely sealed. Pups can be warned under a heating lamp or on a thermoregulated blanket and then returned to their mother. More than 80% of pups survive the surgical, staining, and imaging procedures. Some care in handling pups is necessary, however, so that manipulated pups survive and are cared for after being returned to their mothers. It is at this step that the highest mortality is observed. Wearing gloves when handling pups, removing only a few pups at a time, sham operating on remaining pups in a

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litter, removing all blood from the skin before returning fully recovered pups to their mother, using experienced mothers, fostering pups with experienced mothers, and/or anesthetizing the mother until all of the pups can be returned seem to improve the survival rate.

3. Embryonic Mice Embryonic ages are also accessible to in vivo approaches. We have begun to observe axon outgrowth and synapse formation in embryos over several hours from about the time a well-formed limb bud is present (ca. E l 3 in rodents). Pregnant females are anesthetized as described earlier, the abdomen prepared by shaving, depilatory treatment, and swabbing with alcohol, and a midline incision made to expose the abdominal cavity. One uterine horn is removed and placed on wet gauze adjacent to the incision. Tissues are superfused with sterile mammalian Ringers solution containing0.01 m g / d terbuteline sulfate (Brethine; Ciba-GeigyLtd.) to block uterine contractions.An incision is made in the uterine wall, with care taken to avoid the placenta, and the embryo partially removed. The embryo is placed onto gauze soaked with ice-cold Ringers solution, is manipulated into position, and a small skin incision is made either in the limb bud or in the hindquarters (to expose the area of the gluteus muscle). Embryos are often not adequately anesthetized when the mother is, and thus cryoanesthesia is necessary. Embryos can be stably maintained in this fashion for several hours. In older embryos (El6 and older), we recently have begun to remove the embryo completely, cannulating the umbilical artery with a fine glass needle attached to polyethylene tubing and maintaining oxygenation by slowly perfusing ice-cold oxygenated DMEM through the cannula with a syringe pump. Preliminaryresults suggest that embryos maintain their heart rate and will resume spontaneous movements following warming after ca. 4 hours of perfusion. At the end of the experiment, embryos are placed on ice and are euthanized by decapitation. Pregnant females are euthanized with Nembutal. C. In Viuo Visualization of Synaptic Components of NeuromuscularJunctions

1. Vital Staining Following exposure of the skeletal muscle of interest, muscle fibers and neuromuscular junctions can be stained with a wide variety of fluorescent dyes. Although unfortunately there is no such thing as a truly “vital” dye, many compounds can be used to stain living tissue in an essentially nontoxic fashion (Table I). Each compound should be carefully evaluated and appropriate controls performed to evaluate toxicity and the possibility of photodamage over both the short and long term. Many compounds can be washed out of the tissue at the end of the experiment. Styryl pyridinium derivatives such as 4-Di-2-ASP (Magrassi et al., 1987; Molecular Probes, Eugene, Oregon), a dye that selectively stains

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mitochondria (Lichtman et al., 1989), can be used to visualize motor nerve terminal branches and myofibrillar sarcomeres (Lichtman et al., 1987). Although 4-Di-2-ASP has been found to transiently block neuromuscular transmission when used at high concentrations (> mM; Magrassi et al., 1987), at ca. 10 p M it can be used repeatedly without apparent short- or long-term structural or functional effects. Compounds such as RH 795, RH 414, and their relatives (Grinvald, 1985; Grinvald et al., 1986; Molecular Probes) can be used to stain nerve terminal and axon membranes, and to a lesser extent muscle-fiber membranes, muscle spindles, and blood vessels (Balice-Gordon and Lichtman, 1993). These compounds have also been used to label motor nerve terminals in an activity-dependent fashion, based on internalization of the compound during membrane recycling (Betz and Bewick, 1992;Betz et al., 1992).Terminal Schwann cells, which are believed to play a role in nerve terminal maintenance and reinnervation (Son and Thompson, 1994a,b), can be stained with calcein blue AM ester (Huitink et al., 1974;Balice-Gordon, 1994;Molecular Probes). Staining is the most robust if the animal is maintained at 37"C, probably because intracellular nonspecific esterases (which cleave the ester group, generating a fluorescent product that is cell impermeant) are more active. The concentrations of dyes and staining conditions for these compounds summarized in Table I have been extensively evaluated by a number of investigators. However, the possibility of photodamage when labeled structures are illuminated with high-intensity light must be carefully evaluated. Postsynaptic specializationscan also be simultaneously visualized, for example, with a highly selective and essentially irreversible marker for acetylcholine receptors (AChRs), alpha bungarotoxin (available from Biotoxins, Inc.; Sigma), conjugated to a fluorescent molecule such as rhodamine using the protocol of Ravdin and Axelrod (1977; a number of different fluorophores conjugated to alpha bungarotoxin also available through Molecular Probes; Sigma; see Table I and Fig. 2). By keeping the concentration of toxin low (2-10 pg/d of Ringers for 1-3 min), ca. 10-20% of AChRs are labeled; thus, synaptic transmission is unaffected (Lingle and Steinbach, 1988).If higher concentrations are used, synaptic transmission can be blocked for many hours. 2. Nonvital Staining A number of nonvital fluorescent dyes can also be used to stain synaptic components. The synaptic basal lamina interposed between motor nerve terminals and skeletal muscle fibers can be visualized using fluorescently conjugated lectins such as peanut agglutin (PNA; KO,1987), Vicia villosa B4 agglutinin (VVA; Scott et al., 1988), and Dolichos biflorus agglutinin (Sanes and Cheney, 1982). VVA appears to mark sites rich in acetylcholinesterase (Scott etal., 1988). PNA has been used to monitor the basal lamina of frog neuromuscular junctions over several weeks to months without apparent damage (Chen et al., 1991; Chen and KO, 1994). Other lectins appear to induce a robust inflammatory response,

Table I Fluorescent Dyes for visualization of Skeletal-Muscle Fibers, Supporting Cells, atld Neuromuscular Dye Nerve terminals. 4-Di-2-ASP

Chemical name (sourcea)

4-(4DiethyIaminostyryl)-Nmethylpyridinium iodide (MP)

Fluorescenceb FLTC

Staining properties Mitochondria; motor

nerve terminal

Concentration

References

5-10 pA4 in Ringers for Magrassi er d.(1987) 1-5 min Lichtman et uL (1987)

branches, preterminal axons; diffuse staining of muscle fikrs;spotty staining of Schwann cells, fibroblasts,

satellite and other

R H 414

RH 795

l7-C

N-(3-(Triethylammonium)propyl)- FITC, FUTC 4-(4-( pdiethylanimopheny1)butadieny1)pyridiniu, dibromide

Tetanus, toxin, fragment C Texas Red (CalBioChem, Inc.), Texas Redconjugated

cells. Membranes; motor 20-50 pA4 in Ringers for nerve terminat 5-10 min branches, myelin; preterminal axons; satellite cells; diffuse staining of musclefiber membrane and connective tissue Motor nerve terminal 16 p M €or 10 min membranes

Grinvdd (1985) Grinvald et al. (1986) Betz and Bewick (1992) Betz er aL (1992) Balice-Gordonand Lichtman (1993) Robbins and Pol& (198) Hill and Robbins (1991) Hill et af. (1991)

Postsynaptic AChRs:

aBTX

RITC,FITC Alpha-bungarotoxin (BioToxins, near W Inc.; S;MP), rhodamine- (MP, S), fluorescein- (MP,S), Lucifer Yellow-(MP), amca- (MP), and Cascade Blue- (MP)conjugated

a-subunit of AChR

2-10 pg/d in Ringers for 5-10 min; longer to saturate

Ravdin and Axelrod (1977)

Synaptic basal lamina

50 pg/d for 30 min

KO (1987); Chen et 01,

Basal lamina:

PNA

WAC

Peanut agglutinin (E-Y Lab., Vector Lab.)

RITC, FITC

Vick villosa 84 agglutinin (E-Y

FITC

Labs,S) DBA' khwann cells:

Dolichos biflorus agglutinin (S)

CB

Calcein blue AM ester (MF)

Nuclei: Hoechst 33258,33342 Bis-beazamide (MP, S , K)

near UV

Synaptic basal lamina; 20-50 &ml for 10-30 diffuse connective min tissue, blood vessels

(1991); Chen and KO (1994) Scott et aL (1988) Sanes and Cheney

(1982)

Cytoplasmic; terminal 50-100 p M in Ringed Huitiilk ef oL (1974) Schwatm cells, 1%DMSO 20-30 min Balice-Gordon (1%) nonmyelinating at 37°C Schwana cells; m n e diffuse staining of fibroblasts and satellite cells

near W, FITC Nuclei by intercalation 0.01-0.059b in Ringers into DNA for M. 1 min

Amdt-Jovin and Jovin

(1977) Sprick (1991)

~~

a

~~

~~

MP,Molecular Probes; S, Sigma; K,Kodak. Other sources may be available. Emission and excitation wavelengths can be obtained from manufacturer. Long-term taxicity has not been evaluated; appear to induoz local ineammation and/or damage in viva

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Fig. 2 Vital fluorescent dyes selectively stain pre- or postsynaptic components of neuromuscular junctions. Neuromuscular junction from a 3-month-old mouse gluteus muscle stained with four different vital dyes. CB, calcein blue AM ester staining of terminal Schwann cells; RH 795, staining of presynapticnerve terminal and axon membranes;4-Di-2-ASP. stainingof presynapticmitochondria within presynaptic motor nerve terminal branches; and RaBTX, rhodamine alpha-bungarotoxin staining of the postsynaptic distribution of acetylcholine receptors. Terminal Schwann cells ensheathe presynaptic nerve terminal and preterminal axon branches, and extend processes following nerve damage and reinnervation (Reynolds and Woolf, 1992; Son and Thompson, 1994a,b).The selectivity of this staining has been evaluated by comparing vital staining with CB with S-100 immunostaining of the same neuromuscularjunctions (Bake-Gordon et al., unpublished observations). Motor nerve terminal membranes stained with RH 795 encompass the cytoplasmic organelles stained with 4-Di2-ASP; these two dyes reveal complementaryaspects of presynaptic motor innervation.The alignment between presynaptic motor nerve terminals and postsynaptic AChRs is precise following the period of developmental synapse elimination until near the end of the first year of life, when age-related changes in synaptic maintenance appear to be manifested by mismatches between synaptic components (see Fig. 5). Scale bar = 20 pm.

leading to muscle damage. However, when used at the terminal experiment, these markers are useful because they mark moieties that remain at long times following loss of motor-nerve terminals or postsynaptic AChR-rich areas (Rich and Lichtman, 1989a). This is likely to be because esterase and other molecules in the basal lamina have half-lives on the order of several weeks, while AChRs, for example, have half-lives on the order of several days (Lingle and Steinbach, 1988). Thus, the configuration of the synaptic basal lamina can serve as a “road map” of the previous location of synaptic sites during periods of plasticity.

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In addition, there are a number of antibodies that selectively stain different proteins localized at neuromuscular junctions or different components of muscle fibers. Although a complete description is beyond the scope of this chapter, these can be used in terminal experiments to verify structural components labeled with vital dyes and to allow identified neuromuscular junctions and muscle fibers to be permanently stored on slides. Some skeletal-muscle and neuromuscularjunction components that can be labeled with antibodies include motor nerve terminals (e.g., neurofilament; PGP 9.5; synaptic vesicle proteins such as synaptophysin), terminal Schwann cells (e.g., S-100 antigen and numerous other antibodies; c.f. Son and Thompson, 1994a),basal lamina components (e.g., acetylcholinesterase), and postsynaptic components (e.g., AChR subunits; cytoskeletal proteins such as dystrophin and myofibrillar components). D. Digital Imaging of Skeletal Muscle and Neuromuscular Innervation

Following staining, the coverslip and stabilizingbar are positioned as described earlier (Section 3.B.1). Although stabilizing bar is optional and depends on whether the particular muscle of interest needs to be lifted out of position slightly for optimal access, a coverslip is often necessary because most water-immersion lenses are coverslip corrected. The coverslip is carefully adjusted so that the surface of the muscle is slightly depressed without damaging muscle fibers or compromisingblood flow. Thus, movements caused by respiration and heartbeat are attenuated. A low-magnification, long-working-distance, dry lens (Leica PL Fluotar l o x , 0.30 n.a.) is used to locate the surface of the muscle and the endplate band, which can be visualized using either the appropriate fluorescence filter set or a cross-polarizer filter set (Leica XPOL cube), which allows myofibrillar striations and axon bundles to be clearly seen. At low magnification (Leica PL Fluotar 16X, 0.50 n.a.), superficial junctions are mapped out in areas of interest. This map is used to relocate junctions at later views (see Fig. 8 in Balice-Gordon and Lichtman, 1993;Fig. 5 in van Mier et aL, 1994). Although individual neuromuscular junctions may undergo changes in structural features, the relationship of one junction to another and to landmarks such as blood vessels, nerve branches, or muscle edges is unchanging. It is relatively straightforward to use these cues to relocate individual muscle fibers and junctions successfully over periods of more than 1 year (Fig. 3). Higher-magnificationwater-immersion objectives are used to image individual muscle fibers and neuromuscular junctions of interest (Leica PL Fluotar 16X, 0.50 n.a.; 50X O.TY1.0 n.a.; lOOX N Plan 1.20 n.a.). Oil-immersion objectives (Leica PL Fluotar lOOX, 1.4 n.a. w/ correction collar) can also be used. In this case, the lever arm with the coverslip is removed, a small coverslip is attached directly to the outside lens of the objective with a small drop of oil, and the entire assembly is lowered into the preparation directly into Ringers, which will not displace the oil or the coverslip. Finite, 160-mm tube-length lenses from

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Fig. 3 Sternomastoid neuromuscular junction viewed nine times over the first year of life. Motor nerve terminal arborization stained with 4-Di-2-ASP for the first time in a 2-week-old mouse and imaged; the same junction was restained and imaged on eight subsequent occasions. The preterminal axon can be seen at the top of each panel, where it branches to give rise to the motor terminal arbor. Comparison of the pattern of branches in each view shows that while the branches elongate and the space between them enlarges, the pattern of branches remains largely the same over the 14-month interval during which.the junction was studied. The muscle-fiber nuclei, shown as dark ovals in and around the junction, seem to alter position from view to view. Arrows indicate a spot of staining that was added at 4 months; this new branch was subsequently stably maintained for several months. Scale bar = 20 pm. After Balice-Gordon and Lichtman (1990).

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other manufacturers can also be used with the appropriate adapter lens assembly without appreciable loss of image quality. For example, we use a ceramic-coated, long-working-distance(> 2 mm) Zeiss Achroplan 40X 0.75n.a. water-immersion lens designed to be used without a coverslip for performing intracellular injection or physiological measurements of synaptic function, as a microelectrode can easily be accommodated between the lens and the preparation. Incident light is provided by a 100-W Hg or a 75-WXe lamp, and the light is typically attenuated 50-70% using neutral-density filters to avoid photodamage. A low-light-level video camera (SIT, VElOOO, Dage/MTI, Inc.) or an intensified CCD camera (Videoscope, Inc.) is used to obtain images of structures of interest. Images are digitized with a PC-based image-processing system using a Matrox board and MetaMorph software (Universal Imaging, Inc.). For most applications, 30 frames (1 sec) are averaged to produce an image. During image capture, the ventilator is turned off so that breathing movements do not cause blur in the image. After the experiment is completed, images can be displayed, analyzed, and enhanced using interactive software. Images are stored onto rewritable magneto-optical disks (Pinnacle Sierra 1.3 GB optical drive). Hard copies of images are obtained using a 600-dpi printer (Hewlett-Packard) for notebookquality prints and a dye sublimation printer (Tektronix Phase I1SDX) for publication-quality prints.

111. Manipulation of Motor Neurons, Muscle Fibers, and Synapses in Kuo Methods have been developed by a number of groups for manipulation of motor neurons, muscle fibers, and synapses that are compatible with the basic in vivo imaging approaches described here. For example, crushing or cutting a muscle nerve allows the opportunity to study the events that make up muscle fiber de- and reinnervation in vivo. Other groups have developed clever ways to chronically paralyze or stimulate muscle nerves over long times (Thompson, 1983; Ridge and Betz, 1984; Callaway et al., 1987), and these have extended our understanding of the effects of motor-neuron activity or paralysis on the development and maturation of skeletal muscle and its innervation. We have developed methods to manipulate small synaptic regions within individual neuromuscular junctions while leaving most of the synaptic sites unaffected (described in Balice-Gordon and Lichtman, 1994). This sort of approach circumvents the wholesale disruption of motor neuron-muscle fiber interactions and permits the use of pharmacological agents and toxins that selectively affect pre- or postsynaptic activity. Muscle fibers can also be manipulated in a number of ways in vivo, for example, by inducing focal damage with a glass micropipet or by laser ablation (van Mier and Lichtman, 1994; van Mier ef al., 1994). Individual muscle fibers can be microinjected with pharmacological agents, blocking antibodies, or DNA in situ

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using glass microelectrodes and a pressure delivery device (a picospritzer; WPI, Inc.). If care is taken during delivery to avoid damage to myofibrils, such agents can be readily introduced into the muscle-fiber cytoplasm, or subsynaptic regions can be targeted. Fluorescent tracers can be used to monitor the progress of the injection, and muscle fibers readily survive injection and can then be followed in vivo over appropriate lengths of time (Balice-Gordonet aL, unpublished results). In vivo imaging approaches can also be used to monitor the effects of molecular perturbations of skeletal muscle and its innervation. A number of groups are altering the molecular environment of groups of muscle fibers in living animals using adenovirus (see Ragot et al., Chapter 11of this volume) or retrovirus (see Federspiel and Hughes, Chapter 9 of this vo1me) based gene transfer. We have used glass microelectrodes and pressure delivery of adenovirus into the arterial blood supply of individual muscles such as the sternomastoid and have observed a high degree of marker gene expression (more than 90% muscle fibers positive for /3-galactosidaseexpression; unpublished results). We have also used pressure injections into the femoral artery to achieve a high degree of marker gene expression throughout the musculature of the hindlimb in newborn mouse pups, although capillary permeability to adenovirus appears to decrease after the first week after birth and is low in normal adult mice and rats (H. Stedman et al., unpublished results). In collaboration with S. Hughes and colleagues, we have begun to deliver retroviruses encoding genes of interest into embryonic and neonatal mice of a transgenicline that express a retroviral receptor under control of a skeletal muscle-specific promoter (see Federspiel and Hughes, Chapter 9 of this volume). Thus, in combination with the approaches described earlier, these tools allow the molecular environment around or within individual muscle fibers and synapses to be manipulated and the effects in the animal to be assessed over time.

IV.In Viuo Observations and Manipulations of Muscle Fibers and Neuromuscular Innervation: Practical Considerations Muscle and other tissue have most often been studied either in devitalized (fixed, sectioned, stained) or in vitro preparations. Because plasticity in muscle innervation or in fibers themselves cannot be observed directly in these circumstances, dynamism has been inferred by analyzing a number of single time points and inferring the direction and magnitude of alterations that have occured between observations. Inferring alterations in this way is subject to a number of interpretative limitations, and thus evidence for either structural or functional dynamism is at best indirect. For this reason, in vivo observations have added and will likely continue to add to our understanding of skeletal muscle plasticity, neural influences on muscle, and the plasticity of neuromuscular innervation. However, in vivo observationsraise their own interpretative and technical limitations (see also Balice-Gordon and Lichtman, 1991, for detailed discussion). For

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example, in vivo manipulations are invasive and the very act of observing may either induce alterations or inhibit normally occurring processes. Thus, it is necessary to design control experiments that address this possibility. The staining pattern produced by vital dyes is often selective for a particular organelle (i.e., 4-Di-2-ASP for mitochondria,RH 795 for membranes) and different conclusions may be reached if one or a small number of dyes are used to the exclusion of other methods. Observations of different muscles in different species of animals may yield different results. Moreover, it is difficult to know a priori how many views spaced over what interval are sufficient for revealing the dynamics of a particular process, especially if those dynamics occur out of synch with the interval between views. These are among the issues that need to be considered when in vivo approaches are employed.

V. In Viuo Approaches: New Insights into Neuromuscular Plasticity A. Development of Neuromuscular Innervation Neuromuscular innervation undergoes a dramatic rearrangement during late embryonic and early postnatal life, during which individual neuromuscular junctions undergo a transition from multiple innervation by several motor neurons to the mature pattern of single innervation (Redfern, 1970; Brown et al., 1976; reviewed in Purves and Lichtman, 1980, and Jansen and Fladby, 1990). The transition from multiple to single innervation occurs during the fist few postnatal weeks in rodents by retraction of motor axonal collateral branches well after the period of cell death; thus, there is no change in the number of innervating motor neurons (Bfown et aL, 1976).Thus, initially each motor axon may branch to contact severalfold more target muscle fibers than each will ultimately innervate. Over the past several years we have used the approaches described earlier to study this developmental reorganization in skeletal muscle, by watching literally as it occurred. The same neuromuscular junctions were followed during the transition from multiple to single innervation in living neonatal mice (BaliceGordon and Lichtman, 1993). Vital staining of AChRs and motor nerve terminals showed that although junctions were multiply innervated by two different motor axons, several small AChR areas were lost in succession. By comparing the pattern of alpha-bungarotoxin staining at each time point before and after the addition of additional bungarotoxin, we found that postsynaptic areas were lost because existing AChRs disappeared and because no new receptors were inserted in those areas. Motor nerve terminals were observed to be eliminated from the same areas that lost postsynaptic AChRs (Fig. 4). Moreover, the density of postsynaptic AChRs was downregulated gradually over 1-2 days, and this downregulation appeared to begin well in advance of the elimination of the overlying motor nerve terminal. Similar observations were made during synapse elimina-

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Fig. 4 Transition from multiple to single innervation during neuromuscular development involves loss of presynaptic motor nerve terminals and underlying AChRs. Sternomastoid neuromuscular junction stained with 4-Di-2-ASP (upper row) and rhodamine conjugated alpha bungarotoxin (lower row) in a postnatal day 6 mouse (left column) and again at day 8 (right column). At P6, this junction is multiply innervated by two axons, entering the junction from the upper left. By P8, one axon has been eliminated and one axon remains to singly innervate the junction (arrows). In addition, several motor nerve terminal areas are lost from the upper right region of the junction (arrow). During the transition from multiple to single innervation, AChRs are eliminated from beneath the terminals that were lost (bottom row, arrows). Scale bar = 20 gm. After Balice-Gordon and Lichtman (1993).

tion induced following nerve crush and muscle reinnervation (Rich and Lichtman, 1989a). The precocious loss of AChRs suggests that changes in the postsynaptic muscle fiber membrane beneath one axon and its terminals (but not beneath the remaining axon) may lead to the loss of those terminals and the withdrawal of that axon.

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We are presently directly observing the sequence of structural and functional interactions between pre- and postsynaptic neurons that lead to synapse formation. Preliminary work has suggested that the initial events in synapse formation, such as formation of terminal sites and clustering/declustering of acetylcholine receptors beneath newly arrived motor axon growth cones, occur rapidly enough to be studied over hours in intact embryos visualized in utero. Several technical issues remain to be worked out, however. Many of these involve practical aspects of the experiments, such as how long embryos can be manipulated yet still remain viable; accomplishing adequate vital labeling of poorly differentiated structures; and obtaining fine enough temporal and spatial resolution to visualize small and rapidly changing processes. We have begun to use extracellular recordings of endplate currents to study the onset of synaptic function. Although performing such studies in vivo may seem difficult considering the option of tissue culture, it is important to note that many aspects of neuromuscular maturation have not been successfully recapitulated in vitro. B. Maturation of Neuromuscular Innervation

When the mature pattern of single innervation is established at a neuromuscular junction, we found that loss of synaptic sites was only rarely observed. Neuromuscular junctions do undergo a striking alteration, however, with respect to size, increasing ca. fourfold in area from 2 weeks to 3 months of age, and more slowly thereafter. By following junctions from 2 weeks to more than 1 year of age in two different-muscles, the sternomastoid and the bulbocavernosus, we found that this growth occurs by the intercalary expansion of the synaptic regions left after synapse elimination is complete, largely without the ongoing addition, or loss, of synaptic areas (Figs. 3 and 5; Balice-Gordon and Lichtman, 1990, 1991). The growth of presynaptic motor nerve terminals was matched by the growth of postsynaptic AChR-rich areas. These observations, combined with observations of changes in junction size following reversible manipulation of bulbocavernosus muscle fiber size with androgens (Balice-Gordon et al., 1990), suggested that motor nerve terminal growth is a physical consequence of its adhesive attachment to postsynaptic regions, and that the enlargement of postsynaptic regions is itself a consequence of the expansion of the membrane of a growing muscle fiber. Experiments in which labeled AChRs were studied over time supported this view, because previously labeled receptors were observed to spread apart as junctions grew, and new receptors were intercalated throughout this region rather than selectively at the ends or edges of the junction. Thus, neuromuscular junction growth seemed to resemble closely the expansion of a drawing on a balloon when the balloon is inflated. Although greater degrees of synaptic addition and loss have been reported for other muscles (Wigston, 1989, 1990; Hill and Robbins, 1991; Hill et aL, 1991; Langenfeld-Oster et al., 1993) and in other species (Herrera et al., 1990; Chen and KO, 1994; Chen et al., 1991), we concluded that for the most part, neuromuscular junctions grow the way a hand

Fig. 5 Maturation of neuromuscularjunctions involves expansion of pre- and postsynapticspecializations. Neuromuscular junction from a mouse sternomastoid muscle studied at 1 month (upper row), 4 months (middle row), and 6 months (lower row) of age. Shown are 4-Di-2-ASP staining of mitochondria within motor nerve terminal branches (left column), RH 795 staining of motor nerve terminal and axon membranes and myelin sheaths (middle column), and rhodamine-conjugated alpha bungarotoxin (RaBTX) staining of the postsynaptic distribution of AChRs. Presynaptic motor nerve terminal branches align precisely with postsynapticAChR-rich areas. Over 5 months of observation, two smallpostsynapticAChR-rich areas disappear as the overlyingpresynaptic element becomes myelinated (arrowsin bottom row, middle panel). A small gap appears in the postsynapticdistribution of AChRs (arrows). It is likely that synaptic areas break apart by passive expansion as the muscle fiber membrane expands to accommodate the growing myofiber (Balice-Gordon and Lichtman, 1990, Balice-Gordon et al., 1990). The loss of synaptic regions following the period of synapse elimination is relatively rare. Scale bar = 20 pm. After Balice-Gordon and Licthman (1993).

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does, by overall enlargement of existing elements, rather than the way a plant does, by continually adding branches in some areas and losing branches in others. C. Age-Related Alterations in Synaptic Maintenance

After approximately 1 year of age, preliminary work has shown that the neuromuscular synaptic stability characteristic of much of the first year of the animal’s lifetime is altered. We have observed progressive loss of motor nerve terminal and AChR sites, in some cases leading to the deterioration of the entire synapse (Fig. 6). These observations are consistent with a large body of single time-point observations that suggests that neuromuscular junctions are highly plastic, losing synaptic sites, producing motor nerve terminal sprouts, with re-

Fig. 6 Age-related alterations in the stability of neuromuscular junctions. Three sternomastoid neuromuscular junctions viewed at 10.14.18, and 20 months of age. Motor nerve terminals stained with 4-Di-2-ASP (upper row) and postsynaptic AChRs stained with rhodamine-conjugated alpha bungarotoxin (lower row). Between 10and 20 months of age, each junction shows dramatic alterations in the pattern of pre- and postsynaptic sites. For example, in the middle of the three junctions, some of these changes include loss of existing synaptic areas (compare 10-month and 14-month panels), compensatory addition of new synaptic sites (adjacent to the original junction; 14-month panel), further loss and addition in these areas (18-month panel) followed by almost complete disintegration of the junction (20-month panel). Between 14 and 18 months of age, autofluorescent granules, probably lipofuscin, accumulate in and around junctions (lower row, 18-month m panel). Scale bar = 20 pm.

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maining sites becoming broken apart (reviewed in Wernig and Herrera, 1986; Smith, 1988; Robbins, 1992). Changes in the presynaptic population of motor neurons, in the postsynaptic population of muscle fibers, or both may contribute to synaptic instability as animals age. On the one hand, deterioration and loss of presynaptic motor neurons has been suggested to lead to transient denervation of muscle fibers in aging animals with subsequent reinnervation by sprouts from remaining motor axons, although there are conflicting reports about motor-neuron death in aging animals (see Wernig and Herrera, 1986,for review). Ongoing rounds of denervatiodreinnervation have been suggested to be the cause of age-related changes in neuromuscular junctions, although this has not been tested directly (reviewed in Wernig and Herrera, 1986). Denervation and subsequent reinnervation does not appear to alter the stability of pre- and postsynaptic sites in young adult muscles, however, unless transient multiple innervation is being eliminated (Rich and Lichtman, 1989a). Moreover, as motor neurons die, one might expect that neuromuscular junctions would disintegrate abruptly rather than gradually. Preliminary observations suggest that denervated junctions are rarely if ever observed in aging animals, and that changes in synaptic maintenance occur with a protracted time course, over weeks to months (Fig. 6). Thus, changes in the ability of presynaptic motor neurons to maintain all of their terminals have been proposed to be the cause of synaptic instability observed in aging animals, although this possibility has never been tested directly. By progressively reducing the number of motor neurons innervating a muscle, the remaining motor neurons can be induced to maintain abnormally large terminal arbors (Thompson and Jansen, 1977). The effect of stressing motor neurons in this way could be studied by following synapses in partially denervated muscles over long times, that is, months. On the other hand, a number of observations argue that postsynaptic muscle fibers must also play an important role in synaptic maintenance. Muscle fibers in aged animals may be more susceptible to damage during normal use than muscle fibers from young adult animals (reviewed in Faulkner et aZ., 1990). Ongoing muscle fiber damage may lead to synaptic instability. In support of this possibility, targeted ablation of muscle fibers has been shown to lead to the loss of synaptic sites on residual basal lamina ghosts in young adult mice (Rich and Lichtman, 1989b) and regenerating muscle fibers have been shown to induce motor nerve terminal sprouts (van Mier and Lichtman, 1994). These results, among others, suggest that healthy postsynaptic muscle fibers provide factors essential for normal synaptic maintenance. It is less clear whether regeneration of muscle fibers following damage restores synaptic maintenance over long times in either young adulthood or in aging (see Rich and Lichtman, 1989a; BaliceGordon and Lichtman, 1991). Muscle fiber regeneration seems to be less robust in aged than in young-adult animals (reviewed in Faulkner etaL, 1990),suggesting that the processes that reestablish synaptic maintenance after damage may become compromised in aged animals. Thus, changes in the viability of postsynaptic

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muscle fibers may be the underlying cause of synaptic instability in aging animals. This possibility could be tested directly by damaging individual muscle fibers and studying the maintenance of synaptic sites on that fiber as de- and regeneration occur at short (Rich and Lichtman, 1989b; van Mier and Licthman, 1994) and long times. It is more probable that some interplay of pre- and postsynaptic factors is responsible for the decreased synaptic maintenance observed in aged animals. One obvious candidate for such interplay is ongoing neural activity. Thus, the decline in synaptic maintenance observed in aging animals may be due to the effects of inactivity rather than of aging per se. Modifications of postsynaptic electrical or mechanical activity during development alter synaptic maintenance, even when the innervating axons are still normally active (reviewed in Thompson, 1985). Several studies have documented decreased locomotor activity in several rodent species with advancing age (cf. Robbins and Fahim, 1985). Moreover, experimentally induced muscle disuse has been reported to lead to changes in motor nerve terminals such as increased turnover and sprouting (reviewed in Wernig and Herrera, 1986). Whether such changes in muscle activity are due to decreases in motor neuron firing rate, the ability of motor nerve terminals to activate postsynaptic muscle fibers, or the ability of muscle fibers to respond to activation is at present unclear. In large part, this is because conventional methods to manipulate synaptic activity affect both presynaptic and postsynaptic activity at the same time. For example, direct stimulation of muscle activates motor axons as well as muscle fibers, and pharmacological manipulations such as TTX administration block pre- as well as postsynaptic action potentials. Removing presynaptic innervation by denervation is effective in eliminating patterned motor activity, but also disrupts trophic interactions between nerve and muscle that themselves have pronounced effects on muscle maintenance. Moreover, the assays used to document the effects of activity manipulation, for example, in uitro measurements of muscle tension, motor unit size, etc., are difficult to interpret, because of the inherent variability present within an animal and between animals (see Callaway et al., 1987; Ridge and Betz, 1984). Thus, activitydependent mechanisms have proven difficult to unravel, but longitudinal observations in uivo may clarify some of these mechanisms. Thus, for unknown reasons and by as yet unknown mechanisms, neuromuscular junctions undergo a transition from marked stability to marked instability as animals age. Similar age-related changes in synaptic maintenance are also believed to occur in the far less accessible synapses in the central nervous system, as well as elsewhere in the periphery. Moreover, several neurodegenerative disorders, for example, Alzheimer’s disease, are characterized by a progressive and irreversible decline in the viability of neurons and their synapses. The complexity and relative inaccessibility of synaptic circuitry in the central nervous system where many of these disorders manifest themselves has made it difficult to ask how specific sets of neurons, their targets, and the synapses between them are compromised as a result of the disease, or for that matter as the result of

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normal aging. Thus, studies of the cell biology underlying changes in synaptic maintenance in skeletal muscle in aging animals will provide valuable insight into age- and disease-related processes.

VI. Summary Approaches that permit direct observation and manipulation of skeletal muscle and its innervation in living animals will continue to contribute to our understanding of neural influences on muscle function in developing and mature animals. Understanding how motor neurons interact with each other, with supporting cells such as Schwann cells, and with their target muscle fibers are fundamental issues in neuroscience, as similar mechanisms are likely to underlie the formation and plasticity of synaptic connections in the less easily accessible central nervous system. Acknowledgments I thank Drs. H.L. Sweeney, L. D. Peachey, M. Pinter. and M. M. Rich for helpful discussions and advice; Geoff Daniels of Leica, Inc., and William Penney and Michael Carman in the Johnson Foundation Biomedical Engineering group for discussions about and construction of modifications to the upright epifluorescence microscope and stage; Edward Horn for constructing the miniaturized three-way micromanipulators, the stage plate, and adaptor lenses to accommodate 160-mm tubelength finite objectives in an infinity-corrected optical train; and Lee Wojnar for photographs of the microscopy setup. This work is supported by a McKnight Neuroscience Scholar Award, a Sloan Research Fellowship, and grants from the NM.

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Betz, W. J., Mao, F., and Bewick, G. S.(1992).Activity-dependentfluorescent staining and destaining of living vertebrate nerve terminals. J. Neurosci. 12,363-375. Brown, M. C., Jansen, J. K. S., and Van Essen, D. (1976). Polyneuronal innervation of skeletal muscle in new-born rats and its elimination during maturation. J. Physiol. (Lond) 261, 387-422. Callaway, E. M., Soha, J. M., and van Essen, D. C. (1987). Competition favouring inactive over active motor neurons during synapse elimination. Nature 328,145-159. Chen, L., and KO,C.-P. (1994). Extension of synaptic extracellular matrix during nerve terminal sprouting in living frog neuromuscular junctions. J. Neurosci. 1 4 796-808. Chen, L., Folsom, D. B., and KO, C.-P. (1991). The remodeling of synaptic extracellular matrix and its dynamic relationship with nerve terminals at living frog neuromuscular junctions. J. Neurosci.

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26,288-301. Magrassi, L., Purves, D., and Lichtman, J. W. (1987). Fluorescent probes that stain living nerve terminals. J. Neurosci. 7, 1207-1214. Purves, D., and Lichtman, J. W. (1980). Elimination of synapses in the developing nervous system. Science 210, 153-157. Purves, D., and Voyvodic, J. T. (1987). Imaging mammalian nerve cells and their connections over time in living animals. Trends Neurosci. 10, 398-404. Ravdin, P., and Axelrod, D. (1977).Fluorescent tetramethyl rhodamine derivatives of a-bungarotoxin: Preparation, separation, and characterization. Anal. Biochern. 80,585-592. Redfern, P. A. (1970).Neuromuscular transmission in new-born rats. J. Physiol. (Lond) 209,701-709. Reynolds, M. L., and Woolf, C. J. (1992). Terminal Schwann cells elaborate extensive processes following denervation of the motor endplate. J. Neurocyrofogy 2l,50-66.

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CHAPTER 17

Molecular Diversity of Myofibnllar Proteins: Isoforms Analysis at the Protein and mRNA Level Stefan0 S c h i a n o and Giovanni Salviati Department of Biomedical Sciences and CNR Center of Muscle Biology and Physiopathology University of Padua 35121 Padua, Italy

I. Myofibrillar Protein Isoforms in Muscle Tissues 11. Multiple Approaches to Isoform Analysis A. Integrating SDS-PAGE and Immunoblotting with Immunohistochemistry B. Integrating SDS-PAGE and Immunoblotting with Microsequencing and Mass Profile Fingerprinting C. Integrating Studies at the Protein and mRNA Level D. Multiple Analyses on the Same Muscle Sample 111. Electrophoretic Techniques and Immunoblotting A. Myosin Isoforms B. Other Myofibrillar Proteins IV. Immunohistochemistry and in Situ Hybridization V. Other Techniques for mRNA Analysis VI . Isoform Analysis and Physiological Studies in Single Muscle Fibers VII. Summary and Perspectives References

.

I. Myofibrillar Protein Isoforms in Muscle Tissues Actin, myosin, troponin, and other myofibrillar proteins exist in multiple isoforms that are differentially distributed among the various fiber types present in skeletal and cardiac muscle. Most of these isoforms are also differentially expressed during development and can thus be used as developmental stagespecific as well as fiber type-specific markers of muscle differentiation. HeterogeMETHODS IN CELL BIOLOGY,VOL. 52 Copyright 0 1998 by Academc Press AU nghrr of reproducnon m any form reserved

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neity of contractile proteins is but one aspect of the diversity of muscle fibers, which is also reflected in other fiber type-specific features such as the variable metabolic profile and the presence of multiple forms of sarcotubular system proteins. The complexity of myofibrillar protein isoforms is apparently much greater than that of other cell components. The identification of these isoforms thus provides a powerful tool for investigating the process of muscle differentiation, as well as the adaptive changes of muscle cells in response to neural, mechanical, and hormonal stimuli. The study of myofibrillar protein isoforms is also essential to an understanding of the molecular basis of muscle physiology. Myosin isoforms are responsible for the variable maximum shortening velocity of muscle fibers (see Schiaffino and Reggiani, 1996, and Pette and Staron, 1997), whereas troponin isoforms are major determinantsof the variable Ca2+sensitivity of the myofibrils (see Schachat et aL, 1987). Molecular diversity among myofibrillar proteins has been the object of active investigation since the difference in ATPase activity between myosins from fast and slow skeletal muscles was first reported 30 years ago (Barany et al., 1965). Subsequent studies (reviewed by Pette and Staron, 1990) revealed differences in electrophoretic mobility, immunochemical reactivity, and peptide cleavage pattern among isoforms of myosin light chain (MLC), myosin heavy chain (MHC), and troponin subunits. Direct amino acid sequence analyses provided definitive evidence for the diversity of a-cardiac and a-skeletal actins and of various MLC and troponin isoforms. Additional and more extensive sequence data were recently obtained by gene cloning techniques. Molecular biology studies showed that some isoforms, such as actin and MHC isoforms, are the product of distinct genes, whereas other isoforms, such as troponin T and tropomyosin isoforms, may derive by alternative splicing from a single gene. A list of the major myosin and troponin isoforms identifled in mammalian striated muscle, some of which will be discussed in different sections of this chapter, is shown in Table I. The list is by no means complete and in particular does not include myosin isoforms present in specializedmuscles, such as extraocular muscles. In addition, striated muscles contain isoforms of other myofibrillar proteins, such as tropomyosin, a-actinin, C-protein, and M-protein. The purpose of this chapter is to describe and discuss critically the techniques used to identify the various isoforms. The approaches presented here, although mainly focused on contractileproteins, may also apply to isoforms of sarcotubular system proteins and of other muscle-cell components.

II. Multiple Approaches to Isoform Analysis Several methods are available for isoform analysis in muscle tissues. The point we want to make in this section is that any single method has intrinsic limitations; therefore, a combination of multiple approaches is required to avoid uncertainties and misinterpretationsand thus obtain a correct and complete characterization of

Table I Myofibrillar Protein Isoforms and Their Expression Pattern in Mammalian Striated Gene

Isoforms

MHC-2A MHC-2B MHC-2x MHC-embryonic

MHC-2A MHC-2B MHC-2X MHC-emb

MHC-neonatal

MHC-neo

MHC-extraocular MHC-mandibular MLC-1/3 fast MLC-1 slow-a MLC-1 slowlventricular MLC-1 embryonic/atrial MLC-2 fast MLC-2 slow/ventricular MLC-2 mandibular MLC-2 atrial actin-a-cardiac actin-a-skeletal TnC-fast TnC-slow/cardiac TnT-fast

MHC-eo MHC-m MLC-lf, MLC-3f MLC-lsa MLC-lS/V (-lsb) MLC-le/a MLC-2f MLc-2s/v MLC-2m MLC-2a actin-a-cardiac actin-a-skeletal TnC-f TnC-s/c TnT-lf, -2f, -3f, -4f and developmental isoforms TnT-ls, -2s TnT-lc, -2c, -3c, -4c TnI-f TnI-s TnI-c TM-af

TnT-slow TnT-cardiac TnI-fast TnI-slow TnI-cardiac TM-afast TM-aslow TM-P a-actinin-slowlcardiac a-actinin-fast MBP-C-fast MBP-C-slow MBP-C-cardiac MBP-H M-protein myomesin Titin Nebulin

TM-(YS

TM-P a-actinin-dc a-actinin-f MBP-C-fast MBP-c-slow MBP-C-cardiac MyBP-I1 M-protein myomesin Titin (several isoforms) Nebulin (several isoforms)

Pattern of expression slow skeletal m., heart (ventricles) mandibular m., extraocular m., spindles, heart fast skeletal m. fast skeletal m. fast skeletal m. dev. skeletal m., mandibular m., extraocular m.,spindles dev. skeletal m., mandibular m., extraocular m., spindles extraocular m. mandibular m. (carnivores) fast skeletal m. slow skeletal m. slow skeletal m., heart (ventricles) dev. skeletal m., heart (atria) fast skeletal m. slow skeletal m.,heart (ventricles) mandibular m. (carnivores) heart (atria) skeletal m., heart skeletal m., heart fast skeletal m. slow skeletal m., heart fast skeletal m., dev. skeletal m. slow skeletal m. dev. skeletal m., heart fast skeletal m. slow skeletal m., dev. heart heart fast skeletal m., heart slow skeletal m. skeletal m., heart slow skeletal m. and fast 2A fibers, heart fast skeletal m. fast skeletal m. slow skeletal m. heart skeletal m. skeletal m., heart skeletal m., heart skeletal m., heart skeletal m.

Modified from Schiaffino and Reggiani (1996). Note. The list includes only those isoforms whose existence has been established by analysis at both the protein and mRNA level. The pattern of expression may vary between species. Abbreviations: MyBP-C, myosin binding protein-C (C-protein); MyBP-H: myosin binding protein-H (H-protein); MHC, myosin heavy chain; MLC, myosin light chain; TnC, troponin C; TnT, troponin T; TnI, troponin I; TM, tropomyosin.

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the isoform composition of a muscle sample. Common methodological problems encountered in isoform analysis concern the interpretation of multiple electrophoretic bands (multiple isoforms or degradation products?), staining reactions with monoclonal antibodies (specific reactions or cross-reactionswith unrelated components?), and in situ hybridization signals (can they be taken as evidence for the presence of a given isoform?). These problems cannot be solved within the context of any single technique, though they are often easily solved when a second, different technique is applied to the same muscle sample. The importance of an integrated approach will be discussed in this section focusing mainly on (i) integration of biochemical analyses and immunohistochemistry, and (ii) integration of studies at the protein and mRNA level. A. Integrating SDS-PAGE and Immunoblotting with Immunohistochemistry

Sincemost muscle tissues are heterogeneous with respect to fiber type composition, it is important to define the cellular localization of the different isoforms. Biochemical and immunochemical procedures, such as sodium dodecyl sulfate polyacrylamide gel electrophoresis (SDS-PAGE) and immunoblotting (see Section 111),can be used to distinguish muscle isoforms and determine their relative abundance within a muscle sample, but do not allow mapping of the distribution of the isoforms among the various fiber types. Morphological procedures, such as immunohistochemistry (see Section IV),must be used for this purpose. Let us consider, for example, the case of a muscle sample that displays a mixed MHC isoform composition by SDS-PAGE and immunoblotting, say, 85% type 2X MHC and 15%Plslow MHC. To evaluate the functional implications of this MHC isoform composition, as well as the mechanism of gene regulation responsible for generatingthis particular isoform mixture, it is imperative to establish the cellular localization of the two MHC types. Anti-MHC staining with isoform-specific antibodies can easily clarify whether the sample consists of two fiber populations containing either 2X MHC or fllslow MHC, or a single fiber population with a mixed MHC composition, or both. The second alternative was found to be true in the case of the rat soleus muscle after denervation and chronic stimulation with a high-frequencyimpulse pattern (Schiaffino et al., 1988a;1989). Coexistence of different isoforms within the same fiber has been documented in both normal and experimental muscles, and indeed with certain isoforms this is the rule rather than the exception: MLCl fast and MLC3 fast coexist in variable proportions in most fast-type muscle fibers of mammalian skeletal muscles (Wada and Pette, 1993; Bottinelli et al., 1994), and almost one-third of the fibers present in the rat extensor digitorum longus muscle have a mixed MHC composition, mostly containing either 2X with 2B MHC or 2X with 2A MHC (DeNardi et al., 1993). An alternative approach for isoform analysis at the single fiber level is the biochemical study of isolated fibers (see Section VI). However, this approach is not easily applied to minor skeletal muscle fiber populations or to myocardial myocytes, including the cardiac conduction tissue myocytes: with these cells,

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immunohistochemistryremains the only practical method to map the distribution of contractile protein isoforms. There are two major limitations of immunohistochemical techniques: The first is that specific antibodies against certain isoforms are not available; the second is that these techniques provide only qualitative, not quantitative, information. Other methodological problems with immunohistochemistry, such as the risk of cross-reactions, are discussed in Section 1I.B and in Section 111. B. Integrating SDS-PAGE and Immunoblotting with Microsequencing and Mass Profile Fingerprinting A recent study on myofibrillar changes in the ischemic heart provides a good example of this combined analysis. Ischemia is known to induce depression of cardiac contractile function and altered Ca2+sensitivity of the myofibrils; thus, it is reasonable to postulate changes in myofibrillar proteins, and specifically changes in troponin, in the ischemic heart. Indeed, myofibrillar preparations from the acutely ischemic rat heart were found to display an altered pattern of reactivity with an anti-troponin T monoclonal antibody. Immunoblotting analysis revealed that ischemic heart preparations contain, in addition to the major 41kDa troponin T reactive band, another higher mobility (about 38-kDa) reactive band, which is not detected in myofibrillar preparations from control heart (Westfall and Solaro, 1992). The new band was interpreted as a degradation product of cardiac troponin T, and it was therefore suggested that the altered Ca2+ sensitivity of the myofibrils and the depressed contractility of the ischemic heart may be related to troponin degradation. We have reinvestigated this problem and found that two different monoclonal anti-troponin T antibodies prepared in our laboratory react in immunoblotting with cardiac troponin T, but not with the 38-kDa band present in myofibrillar preparations from ischemic heart (Barbato et al., 1994). The discrepancy could be explained by assuming that, whereas the antibody used in the previous study reacts with an epitope localized in the major (38-kDa) postulated degradation product of troponin T, our antibodies recognize an epitope localized in the minor (about 3-kDa) peptide, a peptide too small to be seen in the gel. However, we were not persuaded by this explanation and set out to determine by an independent approach the nature of the 38-kDa band. The band was cut out of the gel and subjected both to microsequencing, to determine the N-terminal amino acid sequence, and to tryptic digestion and subsequent mass profile fingerprinting. In the latter technique the molecular masses of the peptides produced by proteolytic digestion are analyzed by mass spectrometry, resulting in a mass profile fingerprint that uniquely defines a particular protein (James et al., 1993; Henzel et al., 1993). Both N-terminal sequence analysis and mass profile fingerprinting unambiguously showed that the 38-kDa band corresponds to the glycolytic enzyme, glyceraldehyde phosphate dehydrogenase (GAPDH). The identity of this

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electrophoretic band as GAPDH was confirmed by immunoblottingexperiments with anti-GAPDH antibodies (Barbato et al., 1996). This story shows that the interpretation of the protein composition of myofibrillar preparations, as determined by SDS-PAGE and immunoblotting,may be complicated by (i) cross-reactions of antibodies-in this case one anti-troponin T antibody apparently cross-reacts with GAPDH (see also Section 111)-and (ii) translocation of proteins from the cytosol to the myofibrils that may occur under certain experimental conditions, in this case ischemia. Erroneous interpretations can be avoided by applying additional techniques such as microsequencing and peptide mass fingerprinting. As regards the potential application of these techniques to isoform analysis in muscle tissues, it should be pointed out that although mass profile fingerprinting is likely to find useful applications, Nterminal microsequencing is impossible with many contractile proteins that are N-terminally blocked by posttranslational modification. C. Integrating Studies at the Protein and mRNA Level

It is very useful to integrate protein analysis of muscle isoforms with a study of the corresponding mRNAs by in situ hybridization (see Section IV) or by other techniques for mRNA analysis (see Section V). This is often the only way to obtain definitive evidence for the existence of new isoforms. For example, the existence of 2X MHC, a novel major MHC isoform present in mammalian skeletal muscles, was postulated on the basis of the reactivity pattern of several anti-MHC antibodies (Schiaffino et aZ., 1986, 1989). This notion was supported by the identification of a new MHC band, designated IID MHC, by SDS-PAGE (Biir and Pette, 1988; Termin et al., 1989) and the demonstration that this band corresponds to the 2X MHC (LaFramboise et al., 1990). However, results from immunochemical and electrophoretic studies cannot provide conclusive evidence for the existence of a new MHC, since the postulated new isoform could result from posttranslational modifications (see Bandman et aZ., 1982). Conclusive evidence can only derive from sequence data obtained from direct protein sequencing or deduced from gene cloning. Isolation of cDNA clones coding a novel MHC specifically expressed in type 2X fibers provided definitive support to the existence of the 2X MHC isoform (DeNardi et al., 1993). Analysis at the mRNA level is especially useful whenever protein analysis is difficult or incomplete. For example, biochemical procedures cannot be applied to the study of muscle tissues during early embryonic development because specific organs and tissues cannot be dissected. Therefore in situ hybridization has become a major technique for isoform analysis in the embryo in combination with immunohistochemistry. When isoform-specific antibodies are not available-for example, to distinguish a-cardiac from a-skeletal actin-in situ hybridization is the only possible approach to isoform analysis in the embryo (see Sassoon et al., 1988).

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A unique advantage of a combined analysis at the protein and mRNA level is the possibility of obtaining information on the mechanism of regulation of gene expression. Changes in muscle protein isoforms are often closely correlated with changes in the corresponding mRNAs, suggesting that control of isoform expression likely occurs at the transcriptional level (for example, see Lompri et al., 1984). However, it has occasionally been reported that transcripts coding for a certain isoform are detected in tissues where the protein isoform itself is not detectable. For example, cardiac troponin I transcripts but not cardiac troponin I protein are detected in the rat ventricular myocardium during early stages of embryonic development (Gorza et al., 1993). The appearance of the cardiac troponin I protein at subsequent developmental stages cannot therefore be interpreted as due to the activation of the corresponding gene, but rather reflects changes in posttranscriptional, presumably translational, control mechanisms (Andersen and Schiaffino, 1997). D. Multiple Analyses on the Same Muscle Sample Given the importance of combined isoform analyses and the fact that only one muscle sample is often available for study, as in human skeletal or cardiacmuscle biopsies, it is useful to devise procedures that allow multiple analyses on the same muscle sample. A simple method for this purpose, which is illustrated in Fig. 1, includes the following steps:

1. Muscle sample is frozen in liquid nitrogen and stored at -80°C until it is further processed (most analyses can be carried out on samples stored for years). 2. The frozen sample is transferred to a cryostat and mounted with the long axis of the muscle fibers oriented perpendicular to the blade. 3. Serial transverse sections, approximately 10 pm thick, are alternatively collected onto gelatin-coated slides or transferred to Eppendorf tubes (sections can be stored for several weeks at -20°C). SDS-PAGE, immunoblotting RT-PCR

c

\

immunohis tochemistry /in

situ hybridization

Fig. 1 Serial cryosectionsof muscle samples eithercan be collected on slides (right) for immunohistochemistry, enzyme histcchemistry, and in situ hybridization, or can be transferred to Eppendorf tubes (left) for SDS-PAGE and immunoblotting or RT-PCR.

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4. Sections on slides can be processed for immunohistochemistry, for enzyme

histochemistry (e.g., myosin ATPase staining for fiber typing or P-galactosidase staining in transfected muscles or muscles from transgenic animals), or for in situ hybridization (see Section IV). 5. Sections transferred to Eppendorf tubes can be processed for SDS-PAGE and immunoblotting (see Section 111) or for RT-PCR (see Section V).

III. Electrophoretic Techniques and Immunoblotting Contractile protein isoforms can be distinguished by electrophoretic mobility in one- or two-dimensional gels. This is a classical method for isoform analysis that is especially powerful when it is integrated with immunoblotting to identify the various proteins and with densitometry or image analysis to obtain a quantitative evaluation of their relative proportions. The basic methods for one-dimensional SDS-PAGE, two-dimensional electrophoresis, and immunoblotting are those originally described by Laemmli (1970), O’Farrell (1975), and Towbin et al. (1979), respectively. However, critical variations in the electrophoretic procedures are necessary to separate certain isoforms, for example, high amounts of glycerol must be added to the gel to separate MHC isoforms (see later discussion). In addition, appropriate transfer procedures should be applied for correct immunoblotting analysis of high- or low-molecular-masspeptides. In this section we first consider the type of preparations that are more appropriate for electrophoretic/immunoblotting analyses and discuss some general methodological problems with these techniques; we then discuss the protocols for the separation and identification of myosin isoforms and other myofibrillar proteins. Tissue homogenates, crude myofibrillar preparations, purified myofibrillar preparations, or purified proteins, such as myosin or troponin, can be used for SDS-PAGE and immunoblotting analyses. Crude myofibrillar preparations are essentially homogenates depleted of soluble proteins and microsomes and can be rapidly obtained by homogenizing the tissue in a salt medium (e.g., 0.1 A4 KC1, 30 mM Tris/HCl, and 5 mM EGTA), centrifuging for 5 min at 10,0oOg, and dissolving the pellet in SDS sample buffer. Triton X-100,0.5 to 1%,can be added to the salt medium to extract membrane proteins. Crude myofibrillar preparations represent a convenient choice for most purposes: The procedure is simpler and faster, with reduced risk of proteolysis, and allows a complete recovery of each isoform. Homogenates and crude preparations are the only choice when working with cryosections (see Section 1I.C) or with single fibers (see Section VI). One major problem with one-dimensional SDS-PAGE concerns the possibility that different peptides comigrate in the same electrophoretic band. Twodimensional electrophoresis can be used to distinguish isoforms with the same apparent molecular mass. In this procedure peptides are first separated by charge


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in a gradient of pH using isoelectric focusing, then by size in a second dimension using SDS electrophoresis. One complication of two-dimensional gels is the frequent occurrence of multiple spots that result from different phosphorylation states of a single peptide rather than from the presence of different peptides. For example, several troponin T spots can be identified in immunoblots of two-dimensional electrophoreses; some of these spots correspond to different isoforms, while others are phosphorylated variants of the same isoform. Phosphorylated variants can be eliminated by alkaline phosphatase treatment of the myofibrillar or troponin preparations, using either soluble or insolubilized alkaline phosphatase (Briggs et aZ., 1984; Hartner et aZ., 1989). Another problem frequently encountered in the interpretation of SDS-PAGE experiments is that protein degradation may occur during the preparative procedures. Various mixtures of protease inhibitors have been used, often uncritically, to avoid this risk; however, rapid preparative procedures may obviate the problem without the need for inhibitors. One major problem with immunologicaltechniques, including immunoblotting and immunohistochemistry,concerns the specificity of antibodies. Antibodies to a given protein may cross-react with unrelated peptides sharing a common epitope; therefore, one must use caution in interpreting results based exclusively on a single antibody. For example, as previously discussed in Section II.D, some antitroponin T antibodiescross-reactwith the glycolyticenzyme GAPDH, and similar amino acid sequences were identified in the two proteins (Sanders et al., 1987). A monoclonal antibody raised against the 200-kDa component of neurofilaments was found to cross-react with slow skeletal troponin T (Takagi et al., 1989). Antisera directed against dystrophin showed cross-reaction with a fast skeletal isoform of a-actinin, presumably as a result of similarities in the central rod domains of the two proteins (Hoffman et al., 1989).

A. Myosin Isoforms The myosin molecule consist of two heavy chains, two alkali light chains, and two regulatory light chains; both MHCs and MLCs comprise several isoforms that can associate in various combinations, giving rise to a wide variety of molecules (see Pette and Staron, 1990, and Schiaffino and Reggiani, 1996). Native myosin isoforms can be distinguished by electrophoresis in the presence of pyrophosphate and in the absence of SDS and other denaturing agents, as originally described by d’Albis and Gratzer (1973) and Hoh (1975). A detailed protocol for the separation of myosin isoforms under nondenaturing conditions has been reported (d’Albis and Janmot, 1993). Both MHCs and MLCs can influence the electrophoretic mobility of the molecule; therefore, when multiple MHC and MLC isoforms coexist in the muscle sample, which is usually the case with skeletal muscle, the result is a complex electrophoreticprofile with partially overlapping bands. On the other hand, a simple profile is observed in the ventricular myocardium, which contains only one type of alkali and regulatory MLC isoform: The

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three bands present in the rat ventricle, called V1, V2 and V3, correspond to the a-MHC homodiier (Vl), the a-Ip-MHCheterodimer (V2), and the P-MHC homodimer (V3). Densitometric measurements of these bands can be used to determine the relative concentration of a- and P-MHC, which are not easily separated by SDS-PAGE. MHC isoforms can be distinguished by one-dimensional electrophoresis and by immunoblotting. A critical factor for the separation of MHCs by SDS-PAGE is the addition to glycerol to the gel, as first proposed by Carraro and Catani (1983). Subsequent refinements of the procedure, including an increased amount of glycerol, allowed three MHC bands to be distinguished (Danieli-Betto et aL, 1986), and finally four MHC bands in adult rat skeletal muscle (B& and Pette, 1988). Other protocols have subsequently been published that also allow the different MHC isoforms to be distinguished (LaFramboise et al., 1990; Sugiura et al., 1990; Talmadge and Roy, 1993; Rossini et al,, 1994). A common feature of the various protocols is the high glycerol content (30 to 40%) in the gel, while there are differences with regard to acrylamide concentration, use of gradient vs nongradient gel, and other parameters. It is not yet clear what factors, beside the addition of high amounts of glycerol to the gel, are crucial for a successful separation of MHCs. Figure 2 illustrates the electrophoretic separation of the four major MHC isoforms present in adult rat skeletal muscles, as well as the analysis of the same myosin preparationsby a double immunoblottingprocedure. Standard immunoblotting protocols (e.g., Otto, 1993) can be used for the separation of MHC isoforms. An important point is that blotting conditionsmust be appropriate for complete transfer of these high-molecular-weight peptides.

Pig. 2 Identification by SDS-PAGE and double immunoblotting analysis of the four major MHC

isoforms (type 2A, 2X, 2B, and @/slowMHCs)present in rat skeletal muscle. Myosin preparations from rat diaphragm (DIA) and tibialis anterior (TA) muscles were processed for SDS-PAGEusing the procedure described by LaFramboiie et al. (1990). One gel was silver stained (A) and a parallel gel was blotted onto nitrocellulose and processed for indirect immunoperoxidase with monoclonal antibody BF-32, which reacts with 2A and @/slowMHCs (B). The paper was photographed and then stained with monoclonal antibody RT-D9, which reacts with 2X and 2B MHCs (C). (From LaFramboise et al., 1990.)


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We use overnight transfer at 250 mA and costant voltage at room temperature. On the other hand, 2 hours at 400 mA are sufficient to obtain good transfer of smaller peptides such as troponins: Longer transfer time can lead in this case to loss of the peptides that cross the sheet of paper. Efficiency of transfer can be checked by staining the blot with Ponceau Red and comparing the band pattern with that seen in a Coomassie Blue-stained gel loaded in the same way. The Ponceau Red stain is then removed by washing in distilled water and the paper is incubated overnight at 4°C in the blocking solution prior to incubation with the appropriate antibody. After incubation with anti-mouse immunoglobulins conjugated with peroxidase the paper is treated with diaminobenzidineor chloronaphthol to reveal bound antibody. The same blot can then be reacted with a different anti-MHC antibody (see Fig. 2). MLC isoforms can be separated by one-dimensional SDS-PAGE, using 15% or gradient gels. Two-dimensional electrophoresis as described by O'Farrell (1975) allows a better separation of the various MLC isoforms, especially those with similar apparent molecular weight, such as MLClsa and MLClsb, the two alkali light chains present in slow muscles of different mammalian species.

B. Other Myofibrillar Proteins The electrophoretic characterization of other myofibrillar proteins raises at least two other types of problems. One problem is related to the molecular size of the proteins. Skeletal muscle contains titin, a filamentous, elastic protein with a molecular weight of about 2500 kDa, and nebulin, with a molecular weight of over 800 kDa. These proteins can be separated on large-pore (4%) acrylamide gels or 3-15% linear gradient acrylamide gels ( G r a d e r and Wang, 1993). It is also convenient not to use the stacking gel, since titin in particular may be trapped in it during the usual electrophoretic run. The second type of problem is represented by those myofibrillar proteins, such as troponin I and T, that are basic proteins. Good separation of troponin isoforms by SDS-PAGE is obtained by using 30% acrylamide and 1.1%bis-acrylamide in 8%gels, as suggested by Anderson and Oakeley (1989). However, the resolution of the several isoforms derived by alternative splicing from each troponin T gene requires the use of two-dimensional gel electrophoresis. Under the usual electrofocusing procedure, these basic proteins are not resolved, and a nonequilibrium gradient electrophoresis (NEPHGE), as first described by 0'Farrell et al., (1977), must be used in the first dimension. A large number of fast and slow skeletal troponin T isoforms and their phosphorylated variants can be identified by this procedure in skeletal muscle (Moore et al., 1987; Hartner et al., 1989).

IV. Immunohistochemistry and in Situ Hybridization A unique feature of skeletal muscle is the large size of the muscle fibers. This allows processing of serial transverse cryosections of the same fibers with specific

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antibodies and nucleic acid probes to localize different contractile protein isoforms and the corresponding transcripts in any single fiber. The rationale of isoform analysis at the protein and mRNA level has been discussed in Section 11, and procedures combining immunohistochemistry and in situ hybridization with multiple antibodies and probes have been applied to rat skeletal muscle (DeNardi et aL, 1993) and human skeletal muscle (Smerdu et aZ., 1994). Figure 3 illustrates this type of analysis in human skeletal muscle. The investigator using this approach needs to be aware of the possibility that the isoform profile may vary along the length of the muscle fibers. Nonuniform distribution of MHC isoforms in different segments of the same fiber was observed in normal frog skeletal muscles (Edman et aZ., 1988), in chronically stimulated rabbit muscles (Staron and Pette, 1987), and in denervated rat muscles (Schiaffino et aZ., 1988b). Segmental variation in isoform distribution is also

Fig. 3 Correlation of in situ hybridization analysis with MHC-specific probes and immunohistochemistry with anti-MHC antibodies. Serial cryosections of human skeletal muscle were hybridized with 3sS-labeled cRNA probes complementary to the 3’ untranslated region of fllslow (a), 2A (b), and2X (c) MHC transcripts,then processed for autoradiography and viewed by dark-field microscopy. Most fibers contain only either fllslow (l),or 2A (A) or 2X (X) MHC mRNA, a fiber coexpressing 2A and 2X MHC transcripts is indicated by a triangle. Serial sections were processed for immunoperoxidase with monoclonal antibodies specific for fllslow MHC (d), for all type 2 MHCs (e), or for all MHC except 2X MHC (f). In this sample fibers reactive for 2X M H C transcripts correspond to fibers unstained by BF-35.(From Smerdu et al., 1994.)


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normally found in intrafusal muscle fibers (see Kucera and Walro, 1989). It is therefore advisable to examine sections at different levels of the muscle sample with the same antibodies and probes. In addition, analysis of longitudinal sections may be useful with muscles showing nonuniform isoform distribution. We have discussed in the previous section the specificity of antibodies and the risk of cross-reactions with unrelated peptides. The possibility of cross-reactions should also be considered with the in situ hybridization assays, since the transcripts coding for different isoforms may display a high degree of homology. Probes specific for unique regions of each isoform-specific mRNA must be used. The sequences corresponding to the 5’ and 3‘ untranslated regions of the transcripts are usually highly specific for each isoform even when the coding regions are highly homologous. Specific probes can therefore be prepared by subcloning untranslated regions into appropriate vectors. Indirect immunofluorescence or immunoperoxidase assays can be carried out on unfixed cryosections using standard protocols. For in situ hybridization the sections are fixed in 4% paraformaldehyde in phosphate-buffered saline (PBS) for 30 min, then washed in PBS. The following steps, including proteinase pretreatment and acetylation of the sections, hybridization with 35S-labeledprobes, washing, and visualization of the probes, are essentially similar to the protocol described by Sassoon and Rosenthal (1993). However, the alkaline hydrolysis step to reduce the probe length may be omitted when the length of the probe is less than 0.3 kb. For example, the isoform-specific probes for rat and human MHC that we have used in our studies are complementary to the 3’-untranslated region of the corresponding mRNAs and are between about 70 and 200 nt long; with these probes we found that alkaline hydrolysis even of short duration (1015 min) leads to markedly reduced hybridization signal. When multiple antibodies and probes are applied to serial sections of the same tissue it becomes unnecessary to use controls, such as preimmune serum as control for immunohistochemical reactions and sense probes as control for in situ hybridization reactions, since the different antibodies and probes provide better internal controls (see Sassoon and Rosenthal, 1993).

V. Other Techniques for mRNA Analysis Isoform-specific mRNAs can be identified by a variety of techniques, such as Northern blotting, S1 nuclease protection, RNase protection, primer extension, and reverse transcription followed by polymerase chain reaction (RT-PCR). Detailed protocols of these methods can be found in standard gene-cloning manuals (e.g., Sambrook et al., 1989). Here we will briefly discuss their pros and cons with particular reference to their application to isoform analysis. It should be stressed that, at variance with in situ hybridization, the other techniques cannot determine the cellular localization of the transcripts, except perhaps the RT-PCR technique, which is extremely sensitive and could therefore be used

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Fig. 4 Four different methods for identifying isoform-specifictranscripts. The scheme shows how these assays can be used to determine the presence and relative abundance of two mRNAs, A and B, coding for two similar yet distinct isoforms, in three muscle samples: a, which contains only A transcripts; b, which contains only B transcripts; and c, which contains both A and B transcripts. In Northem blotting, RNA isolated from the three muscle samples is separated by electrophoresis, blotted onto nitrocellulose, and hybridized to one or the other labeled probe (asterisks indicate radioactive labels). Specific antisense probes must correspond to regions showing no significant sequence homology with other transcripts, such as 3' or 5' untranslated regions. In the RNme protection assay, FWA isolated from the three muscle samples is hybridized with a labeled antisense


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for mRNA analysis in single skeletal muscle fibers. A scheme of these techniques is shown in Fig. 4. Northern blotting is a simple technique; however, it has a relatively lower sensitivity than other methods. Furthermore, comparative analyses by Northern blotting are not completely accurate because the signals produced by different isoform-specific probes cannot be precisely compared even when probes of similar size are used. S l nuclease and RNase protection assays are more sensitive and can provide precise quantitative information on the relative proportion of different transcripts. These techniques are based on the use of enzymes, S1 nuclease or RNase A, that digest single-stranded but not double-stranded nucleic acid sequences. With both techniques a single antisense probe is used, an RNA probe in the RNase protection assay or a DNA probe in the S1 nuclease assay. The probe is hybridized to total RNA and generates a longer double-stranded fragment by hybridizing with the corresponding isoform-specific transcript and shorter partially protected fragments with other similar but not identical isoform transcripts. An example of the application of the S1 nuclease protection technique in isoform analysis is a quantitative study of a-and @-MHCtranscript levels in the developing rat heart (Lompri et al., 1984). The primer extension assay has been used to quantitate the relative abundance of a-cardiac and a-skeletal actin transcripts in skeletal and cardiac muscle samples (Ordahl, 1986; Winegrad et aZ., 1990). The two transcripts have an identical sequence in the 5' coding region, so that an antisense oligonucleotide primer can anneal to both mRNAs, but differ in the length of the 5'-untranslated region, so that cDNAs of different length are generated by primer extension with reverse transcriptase. RT-PCR is a very sensitive assay that allows amplification of selected regions of multiple transcripts in a single reaction using appropriate primers. Precise quantitative evaluation of the amplification products is difficult because variation may occur both in the reverse transcriptase and polymerase chain reaction.

R N A probe corresponding to transcript A but containing a region, marked AB, with a sequence identical in transcript B. RNases that digest single-stranded but not double-stranded R N A produce a longer fully protected fiagment with transcript A, which can be separated on a sequencing gel fiom the shorter partially protected hgment obtained with transcript B. The primer extension assay can be applied to transcripts having an identical sequence in the 5' coding region but a different length of the 5' untranslated region. A labeled oligonucleotide primer complementary to the common sequence can anneal to both transcript A and B and reverse transcriptase generates extension products of different lengths that can be distinguished on a sequencing gel. In the RT-PCR assay, complementary DNAs are generated fiom the mRNAs using reverse transcriptase and oligo-dT primers, then selected regions of the cDNAs are amplified using oligonucleotideprimers specific for unique sequences ofthe two transcripts. Amplification products of different length can be directly visualized after electrophoresis in ethidium bromide-stained gels.

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However, studies indicate that it is possible to quantify levels of specific mRNAs by RT-PCR using synthetic RNAs as internal standards. For example, in one study the levels of P-MHC mRNA and other cardiac transcripts have been quantified in endomyocardial biopsies from human heart (Feldman et al., 1991). RT-PCR can also be performed on crude tissue extracts from small tissue samples, whereas the other techniques require isolation of RNA, either total RNA or poly(A)+ RNA, from larger muscle samples.

VI. Isoform Analysis and Physiological Studies in Single Muscle Fibers The distribution of contractile protein isoforms in the various types of muscle fibers can be studied using biochemical and immunochemical micromethods applied to isolated fibers (see Pette and Staron, 1990).This approach is especially interesting when combined with physiological studies, since correlated analyses may provide important indications as to the functional significance of the different isoforms. More direct evidence for isoform function can be obtained by isoform exchange experimentsin skinned fibers: these experiments consist of the selective extraction of one isoform, such as fast skeletal troponin C, and its substitution with another isoform, such as slow/cardiac troponin C (see Moss, 1992). In this section we briefly review the various single-fiber preparations most commonly used for correlated studies and discuss protocols for isoform analysis by SDSPAGE and immunohistochemistry in single fibers. Intact muscle fibers can be isolated from frog but not from mammalian skeletal muscle. On the other hand, it is possible to obtain skinned fiber segments from mammalian muscle that allow measurements of different physiological parameters such as maximum velocity of shortening, force generating capacity, and sensitivity of the contractile apparatus to Ca2+,as well as sarcoplasmic reticulum functions. Chemically skinned preparations are most commonly used in these studies: A common procedure involves chemical skinning with EGTA, which makes the plasma membrane permeable to Ca2+without disrupting the sarcoplasmic reticulum. Other protocols involve the use of glycerol or Triton X-100, which disrupt the sarcoplasmic reticulum but do not alter the contractile apparatus. These preparations can be stored for months at -20°C. Alternatively, single fibers can be isolated from freeze-dried muscle samples, including human muscle samples obtained by percutaneous biopsies (Larsson and Salviati, 1992); these preparations can be stored at -80°C for many months and are suitable for mechanical measurements in combination with isoform analysis, but not for sarcoplasmic reticulum studies since all membranes are destroyed. Electrophoretic analysis can be performed on segments of single fibers after dissection or at the end of the physiological experiments. Skinned fibers are transferred with the help of a needle under a stereomiscroscope to a small capillary tube that is iilled with 20 p1 of SDS solubilizing solution (Salviati


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et al., 1982). About 5 p1 are used for MHC electrophoresis and the remaining 15 p1 for MLC electrophoresis. Fibers are left overnight in this solution at room temperature, then processed for SDS-PAGE to identify myosin isoforms or other contractile proteins, as described in Section 1II.A. After electrophoresis the gels are first stained with Coomassie Blue and photographed, then destained with several changes of destaining solution (40% methanol-10% acetic acid), and finally stained with silver, which increases the sensitivity severalfold (Memll et al., 1981). However, we find that silver is a poor stain for some myofibrillar proteins such as tropomyosin; therefore, it is useful to stain gels first with Coomassie Blue. Zmmunohistochemical analysis of isolated single muscle fibers can be performed using the following procedure, applied by Edman et al. (1988) to frog fibers and by Bottinelli etal. (1991,1994) to rat fibers. At the end of the mechanical experiment a single fiber or fiber segment is placed in a Petri dish filled with Ringer solution in the case of intact frog fibers or of relaxing solution in the case of skinned mammalian fibers. The bottom of the dish is covered with Silgard, so that the fiber can be kept slightly stretched by fixing it by means of two pins inserted in the clips at the ends of the fiber. The solution contained in the dish is slowly replaced with a solution with the same concentration but containing gelatin 10-15 dl00 ml at 37-38°C. The dish is then transferred to a refrigerator where the gelatin is allowed to set at a temperature of 4°C. A small block of gelatin containing the fiber in the center is cut out, wrapped in aluminum foil, and frozen in liquid nitrogen. The gelatin block is mounted in a cryostat and serial transverse sections are collected on gelatin-coated slides. It is useful also to collect on each slide a section of the whole muscle or part of the muscle from which the fiber had been isolated. The sections are then processed for immunohistochemistry and the reactivity of the single fiber with each specific antibody is assessed by comparison with the staining in the whole muscle. When adjacent segments of the same rat skeletal muscle fiber are either stained with anti-MHC antibodies using the procedure described here or analyzed for MHC isoforms by SDS-PAGE, as described in Section VI.B, the concordance between the two techniques is about 98%.

VII. Summary and Perspectives Several techniques are available for identifying the multiple isoforms of contractile proteins present in skeletal and cardiac muscle. Isoforms can be distinguished and their relative abundance can be evaluated by SDS-PAGE or two-dimensionalelectrophoresis; these techniques must be integrated with immunoblotting analyses and/or with microsequencing and mass profile fingerprinting to establish the identity of electrophoretic bands and spots. The differential distribution of the isoforms among the various fiber types or the coexistence of two or more isoforms within the same fiber can be analyzed in tissue sections

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by immunohistochemical methods or in isolated single fibers by biochemical micromethods.The functionalrole of contractile protein isoforms can be explored by correlated biochemical and physiological studies of skinned single fibers. Protein isoform analysis requires the application of multiple techniques, such as SDS-PAGE combined with immunohistochemistry, to the same muscle sample; serial cryosection of skeletal muscle can be used for this purpose. In addition, studies at the protein level can be integrated with studies at the mRNA level using a variety of techniques, such as Northern blotting, RNase protection, RTPCR, and in situ hybridization. Isoform analysis at the mRNA level is essential for identlfying new isoforms and for defining the mechanisms of gene regulation implicated in differential isoform expression. The accumulation of new molecular biology data, and in particular the rapid progress in genomic studies, should soon lead to the identification of all contractile protein genes and provide the nucleotide and amino acid sequences of all muscle isoforms. Complete analyses at the mRNA level should thus be possible in the near future: In particular, one may predict the development of rapid RTPCR assays for the detection of multiple isoform transcripts in a single reaction. As a fallout of these studies one may also predict further advances in protein isoform analysis; for example, isoform-specific antibodies can be prepared using amino acid sequence information. Expansion of protein sequence databases should also increase the applications of peptide mass fingerprinting and thus contribute to isoform analysis at the protein level. A major goal for future studies will be to define the functional role of the various isoforms. In addition to the traditional comparative studies, based on the correlation between isoform composition and physiological properties of different fiber types, and the more recent isoform exchange experiments in skinned single fibers, two other powerful approaches should further progress in this area: (1) in vitro studies with reconstituted systems of purified muscle proteins, such as in vitro motility assays with specific myosin isoforms (see Lowey et al., 1993); and (ii) transgenic animals, in which one can determine the effect of overexpression of certain isoforms or their elimination by gene “knockout” or the effect of ectopically expressed isoforms, such as expression of fast skeletal troponin C in cardiac muscle (Metzger et al., 1993). Each method has its own limitations: Reconstituted protein systems in vitro do not reproduce the complexity of the sarcomere, even less that of the fiber, and on the other hand, interpretation of physiological studies on transgenic mice may be complicated by adaptive changes in other muscle isoforms induced by the presence of the transgene. Multiple approaches will thus be required for understanding the functional significance of the contractile protein isoforms. Acknowledgments Original work reviewed in this chapter was supported by grants from BIOMED Contract PL 950174, Agenzia Spaziale Italiana and from Minister0 dell’Universit8e della Ricerca Scientifica e


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Tecnologica of Italy, Telethon-Italy (grants A.58 and 692), and Progetto Finalizzato Ingegneria Genetica.

References Andersen, J. L., and Schiaffino,S. (1997). Mismatch between myosin heavy chain mRNA and protein in human skeletal muscle fibers. Am. J. Physiol, in press. Andersen, P. A. W., and Oakeley, A. E. (1989). Immunological identification of five troponin T isoforms reveals an elaborate maturational troponin T profile in rabbit myocardium. Circ. Res.

65,1087-1093. Bandman, E., Matsuda, R., and Strohman, R. C. (1982). Myosin heavy chains from two different adult fast-twitch muscles have different peptide maps but identical mRNAs. Cell 29,645-650. Bar, A., and Pette, D. (1988).Three fast myosin heavy chain in adult rat skeletal muscle. FEBS Lett. 235, 153-155. Barany, M., Barany, K., Reckard, T., and Volpe, A. (1965). Myosin of fast and slow muscles of the rabbit. Arch. Biochem Biophys. 109, 185-191. Barbato, R. Menabb, R., Dainese, P., Carafoli, E., Schiaffino, S., and Di Lisa, F. (1996). Binding of cytosolic proteins to myofibrils in ischemic rat hearts. Circ. Res. 78, 82-87. Bottinelli, R., Schiaffino,S., and Reggiani, C. (1991).Force-velocity relation and myosin heavy chain isoform composition in skinned fibres of rat skeletal muscle. J. Physiol. (London) 437,655-672. Bottinelli, R., Betto, R.,Schiaffino, S., and Reggiani, C. (1994). Unloaded shortening velocity and myosin heavy chain and alkali light chain isoform composition in rat skeletal muscle fibers. J. Physiol. (London) 478,341-349. Briggs, M. M., Klevit, R. E., and Schachat, F. H. (1984). Heterogeneity of contractile proteins. Purification and characterization of two species of troponin T from rabbit fast skeletal muscle. J. Biol. Chem. 259,10369-10375. Carraro, U., and Catani, C. (1983). A sensitive SDS-PAGE method separating myosin heavy chain isoforms of rat skeletal muscles reveals the heterogeneous nature of the embryonic myosin. Biochem. Biophys. Res. Commun. 116,793-802. d’Albis, A., and Gratzer, W. B. (1973). Electrophoretic examination of native myosin. FEBS Left.

299292-296. d’Albis, A., and Janmot, C. (1993).Separation by gel electrophoresis under nondissociatingconditions (PPi PAGE) of the native isoforms of myosin-A technical note. Basic Appl. Myol. 3,235-238. Danieli-Betto, D., Zerbato, E., and Betto, R. (1986). Type 1, 2A and 2B myosin heavy chain electrophoretic analysis of rat muscle fibers. Biochem Biophys. Res. Commun. l38,981-987. DeNardi, C., Ausoni, S., Moretti, P., Gorza, L., Velleca, M., Buckingham, M., and Schiaffino, S. (1993).Type 2X myosin heavy chain is coded by a muscle fiber type-specific and developmentally regulated gene. J. Cell BioL 123,823-835. Edman, K. A. P., te Kronnie, G., Reggiani, C., and Schiaffino, S . (1988). Maximum velocity of shortening related to myosin isoform compositionin frog skeletal muscle fibres. J. Physiol. (London)

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Feldman, A. M., Ray, P. E., Silan, C. M., Mercer, J. A., Minobe, W., and Bristow, M. R. (1991). Selective gene expression in failing human heart. Quantificationof steady-state levels of messenger RNA in endomyocardial biopsies using the polymerase chain reaction. Circulation 83,1866-1872. Gorza, L.,Ausoni, S., Merciai, N., Hastings, K. E. M., Shiaffino, S. (1993). Regional differences in troponin I isoform switching during rat heart development. Dev. Biol. 156,253-264. Granzier, H. L. M., and Wang, K. (1993). Gel electrophoresis of giant proteins. Solubilization and silver-staining of titin and nebulin from single muscle fiber segments. Electrophoresis 14,56-64. Hartner, K.-T., Kirschbaum, B. J., and Pette, D. (1989). The multiplicity of troponin T isoforms. Distribution in normal rabbit muscles and effects of chronic stimulation. Eur. J. Biochem. 179,

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Henzel, W. J., Billeci, T. M., Stultz, J. T., Wong, S. C., Grimley, C., and Watanabe, C. (1993). Identifying proteins from two-dimensional gels by molecular mass searching of peptide fragments in protein sequence databases. Proc. Natl. Acad. Sci. USA 90,5011-5015. Hoffman, E. P., Watkins, S. C., Slayter, H. S., and Kunkel, L. M. (1989). Detection of a specific isoform of alpha-actinin with antisera directed against dystrophin. J. Cell Biol. 108,503-510. Hoh, J. F. Y.(1975).Neural regulation of mammalian fast and slow muscle myosins: An electrophoretic analysis. Biochemistry 14,742-747. James, P., Quadroni, M., Carafoli, E., and Gonnet, G. (1993).Protein identification by mass profile fingerprinting. Biochem Biophys. Res. Commun. l95,58-64. Kucera, J., and Walro, J. M. (1989).Nonuniform expression of myosin heavy chain isoforms along the length of cat intrafusal muscle fibers. Histochemistry 92,291-299. Laemmli, U. K. (1970).Cleavage of structural proteins during the assembly of the head of bacteriophage T4.Nature (London) 227,680-685. LaFramboise,W. A., D a d , M. J., Guthrie, R. D., Moretti, P., SchiafEno,S., and Ontell, M. (1990). Electrophoretic separation and immunological identification of type 2X myosin heavy chain in rat skeletal muscle. Biochem. Biophys. Acta 1035, 109-112. Larsson, L., and Salviati, G. (1992). A technique for studies of the contractile apparatus in single human muscle fibre segmentsobtained by percutaneous biopsy. Acta Physiol. Scand. 146,485-495. Lomprk, A. M.,Nadal-Ginard, B., and Mahdavi, V.(1984). Expression of the cardiac ventricular aand @-myosinheavy chain genes is developmentally and hormonally regulated. J. Biol. Chem. 259,6437-6446. Lowey, S., Waller, G. S., and Trybus, K. M. (1993).Function of skeletal muscle myosin heavy and light chains isoforms by an in vitro motility assay. J. Biol. Chem. 268,20414-20418. Merrill, C. R., Goldman, D., Sedman, S. A., and Ebert, M. H. (1981). Ultrasensitive stain for proteins in polyacrylarnide gels shows regional variation in cerebrospinal fluid proteins. Science 2% 1437-1438. Metzger, J. M., Parmacek, M. S., Barr, E., Pasyk, K., Lin, W. I., Cochrane, K. L., Field, L. J., and Leiden, J. M. (1993). Skeletal troponin C reduces contractile sensitivity to acidosis in cardiac myocytes from transgenic mice. Proc. Natl. Acad. Sci. USA 90,9036-9040. Moore, G.E., Briggs,M. M., and Schachat,F. H. (1987).Patterns of troponin expressionin mammalian fast, slow and promiscuous fibres. J. Muscle Res. Cell Motil. 8, 13-22. Moss, R.L. (1992).Caz+regulation of mechanical properties of striated muscle. Mechanistic studies using extraction and replacement of regulatory proteins. Circ. Res. 70,865-884. OFarrell, P. H. (1975).High resolution two-dimensional electrophoresis of proteins. J. Biol. Chem. 250,4007-4021. O’Farrell, P. Z., Goodman, H. H., and O’Farrell, P. H. (1977). High resolution two-dimensional electrophoresis of basic as well as acidic proteins. Cell U,1133-1142. Ordahl, C. P. (1986). The skeletal and cardiac a-actin genes are coexpressed in early embryonic striated muscle. Dev. Biol. 117,488-492. Otto, J. J. (1993). Immunoblotting. Meth. Cell Biol. 37, 105-117. Pette, D., and Staron, R. S. (1990).Cellular and molecular diversities of mammalian skeletal muscle fibers. Rev. Physiol. Biochem. Pharmacol. 116,l-76. Pette, D., and Staron, R. S. (1987).Mammalian skeletal muscle fiber type transitions. Int. Rev. Cytol. 170,143. Rossini, K.,Rizzi, C., Sandri,M., Bruson, A., and Carraro, U. (1995).High-resolution SDS-PAGE and immunochemical identification of the 2X and embryonic myosin heavy chains in complex mixtures of isomyosins. Electrophoresis, 16,101-104. Salviati, G., Betto, R., and Danieli-Betto, D. (1982).Polymorphism of myofibrillar proteins of rabbit skeletal-muscle fibres. An electrophoretic study of single fibres. Biochem. J. 207,261-272. Sambrook, J., Fritsch, E. F., and Maniatis, T. (1989). “Molecular Cloning: A Laboratory Manual,” 2nd ed. Cold Spring Harbor, New York Cold Spring Harbor Laboratory. Sanders, C., Stewart, D. I. H., and Smillie, L. B. (1987).Troponin-T and glyceraldehyde-3-phosphate dehydrogenase share a common antigenic determinant. J. Muscle Res. Cell Motil. 8,118-124.


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Sassoon, D., and Rosenthal, N. (1993). Detection of messenger RNA by in situ hybridization. Meth. Enzymol. 225,384-404. Sassoon, D., Gamer, I., and Buckingham, M. (1988). Transcripts of a-cardiac and a-skeletal actins are early markers for myogenesis in the mouse embryo. Development 104,155-164. Schachat,F. H., Diamond, M. S., and Brandt, P. W. (1987). Effect of different troponin T-tropomyosin combinations on thin filament activation. J. Mol. Biol. 198, 551-554. Schiaffino, S., and Reggiani, C. (1996). Molecular diversity of myofibrillar proteins: Gene regulation and functional significance. Physiol. Rev. 76, 371-423. Schiaffino, S., Saggin, L., Viel, A., Ausoni, S., Sartore, S., and Gorza, L. (1986). Muscle fiber types identified by monoclonal antibodies to myosin heavy chains. I n “Biochemical Aspects of Physical Exercise” (G. Benzi, L. Packer, and N. Siliprandi, eds.), pp. 27-34. Amsterdam: Elsevier. Schiaffino, S., Ausoni, S., Gorza, L., Saggin, L., Gundersen, K., and Mmo, T. (1988a). Myosin heavy chain isoforms and velocity of shortening of type 2 skeletal muscle fibres. Acta Physiol. Scand. 1% 565-566. Schiaffino, S., Gorza, L., Pitton, G., Saggin, L., Ausoni, S., Sartore, S., and Lprmo, T. (1988b). Embryonic and neonatal myosin heavy chain in denervated and paralyzed rat skeletal muscle. Dev. Biof. U7, 1-11. Schiaffino, S., Gorza, L., Sartore, S., Saggin, L., Vianello, M., Gundersen, K., and Lprmo, T. (1989). Three myosin heavy chain isoforms in type 2 skeletal muscle fibres. J. Muscle Res. Cell Motil. 10, 197-205. Smerdu, V . ,Karsch-Mizrachi,I., Campione, M., Leinwand, L., Schiaffino,S. (1994). Type 2X myosin heavy chain transcripts are expressed in type 2B fibers of human skeletal muscle. Am. J. Physiol. 267,1723-1728. Staron, R. S., and Pette, D. (1987). Nonuniform myosin expression along single fibers of chronically stimulated and contralateral rabbit tibialis anterior muscles. PfIugers Arch. 409,67-73. Sigiura, T., and Murakami, N. (1990). Separation of myosin heavy chain isoforms in rat skeletal muscles by gradient sodium dodecyl sulphate-polyacrylamide gel electrophoresis. Biomed. Res. 11987-91. Takagi, M., Takano-Ohmuro, H., Nakamura, T., Kawahara, H., Shimizu,T., Obinata, T., and Kohama, K. (1989). Cross-reactivity of an antineurofilament antibody with a troponin-T isoform. Muscle Nerve 12,827-832. Talmadge, R. J., and Roy, R. R. (1993). Electrophoretic separation of rat skeletal muscle myosin heavy-chain isoforms. J. Appl. Physiol. 75,2337-2340. Termin, A., Staron, R. S. and Pette, D. (1989). Myosin heavy chain isoforms in histochemically defined fiber types of rat muscle. Histochemistry 92,453-457. Towbin, H., Staehelin,T., and Gordon, J. (1979). Electrophoretic transfer of proteins from polyacrylamide gels to nitrocellulose sheets: Procedure and some applications. Proc. Narl. Acad. Sci. USA 76,4350-4354. Wada, M., and Pette, D. (1993). Relationship between alkali light-chain complement and myosin heavy-chain isoforms in single fast-twitch fibers of rat and rabbit. Eur. J. Biochem. 214,157-161. Westfall, M. V., and Solaro, R. J. (1992). Alterations in myofibrillar function and protein profiles after complete global ischemia in rat hearts. Circ. Res. 70,302-313. Winegrad, S., Wisnewsky, C., and Schwartz, K. (1990). Effect of thyroid hormone on the accumulation of mRNA for skeletal and cardiac a-actin in hearts from normal and hypophysectomized rats. Proc. Natl. Acad. Sci. USA 87, 2456-2460.


Molecular Genetic Analysis of Muscle Gene Regulation


CHAPTER 18

Transgenic Mice: Production and Analysis of Expression Alexander Faerman and Moshe Shani Institute of Animal Science Agricultural Research Organization The Volcani Center Bet Dagan 50250, Israel

I. Introduction 11. Production of Stable Transgenic Mice A. Choice of Expression Vectors and DNA Preparation for Microinjection B. Superovulation C. Media Preparations D. Embryo Recovery E. Preparation of Holding and Microinjection Pipets F. Microinjection: Equipment and Setup G. Vasectomy and Embryo Transfer H. Identification of Transgenic Mice 111. Production of Transient Transgenics A. Advantages IV. Analysis of Expression at the R N A Level A. RNA Isolation B. Northern Blots C. RNase Protection Assay D. RT-PCR E. In Situ Hybridization to Tissue Sections F. Whole-Mount in Situ Hybridization of Mouse Embryos V. Analysis of Expression at the Protein Level A. CAT Assay B. LacZ Staining VI. Future Directions References

METHODS IN CELL BIOLOGY. VOL. 52 Copyright 0 1998 by Academic Prrrr. All ngho i w ~ i - h 7 9 ~ / 9snx o n

of

reproduction in any form reserved

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I. Introduction Production of transgenic mice become one of the most powerful tools to dissect complex developmentalprocesses such as genetic determination, pattern formation, cell lineages, cis-acting elements controlling gene expression, etc. In the study of myogenesis, transgenesis was exploited to address several important topics:

1. The cis-acting DNA sequences involved in the developmental regulation and tissue specificity of structural and regulatory genes, Elements controlling the expression of the skeletal muscle actin (Shani, 1986; Petropoulos et al., 1989; Asante et al., 1994), troponin (Hallauer et al., 1988; Banerjee-Basu and Buonanno, 1993), myosin light chain 2 (Shani, 1985), M creatine phosphokinase (Johnson et al., 1989), aldolase (Salminen et al., 1994), acetylcholine receptor (Gundersen et al., 1993), myogenin (Cheng et al., 1992, 1993; Yee and Rigby, 1993; Fujisawa-Sehara et al., 1993), and MyoD (Goldhamer et al., 1992, 1995) were characterized. 2. The mechanisms confemng fiber-type specificity. Transgenic mice carrying the rat troponin I slow gene (Banejee-Basu and Bounanno, 1993),the quail fast skeletal muscle troponin I (Hallauer et al., 1993), the rat fast skeletal muscle myosin light chain 1 (Donoghue et al., 1991), and the human aldolase A genes (Salminen et al., 1994) revealed a complex transcriptional requirements for the different muscle fibers. 3. The consequences of overexpressing muscle specific genes in skeletal muscle. To this end transgenic mice overexpressing the B subunit of creatine b a s e (Brosnan et al., 1993), the c-ski (Sutrave et al., 1990), or the rat myosin light chain 2 (Shani et al., 1988) genes were studied. 4. The consequences of ectopic expression of the myogenic regulatory genes MyoDl (Faerman et al., 1993) or myf5 genes (Miner et al., 1992; Santerre et al., 1993). 5. Delineating the cis-acting elements responsive to regulation by the nerve (Banerjee-Basu and Buonanno, 1993; Buonanno et al., 1993). 6. The possible existence of regional specification along the rostrocaudal axis (Donoghue et al., 1992, Grieshammer et al., 1992). 7. The efficiency of correcting muscular dystrophy in mdu mice (Matsumura et af., 1993). The scope of the present chapter is to describe in detail the methodology of transgenic mouse production, mainly focusing on the most commonly used technique of microinjecting into fertilized eggs, based on the experience of our laboratory. This chapter also describes methods for the analysis of transgenic mice with an emphasis on the in situ technologies.

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11. Production of Stable Transgenic Mice A. Choice of Expression Vectors and DNA Preparation for Microinjection

An expression vector should contain the appropriate promoter/enhancer sequences, the entire protein-codingregion, and a polyadenylationsignal. Although cDNAs have been used successfully in a number of studies, they are in general poorly expressed. Therefore, including genomic sequences containing introns within the expression vector is highly recommended. If the entire gene is not yet available, minigenes carrying combinations of genomic and cDNA can also be considered. There appears to be no upper limit to the size of microinjected DNA. DNA cloned into cosmids or yeast artificial chromosomes is efficiently incorporated into the mouse genome (Schedl et al., 1993). The DNA to be microinjected should be free of vector DNA sequences. The presence of prokaryotic vector sequencesmay affect expression of the introduced gene. The purity of the DNA is of utmost importance. Care must be taken to remove organic solvents and salt, which are highly toxic to fertilized eggs. We routinely isolate genomic fragments by electroelution followed by pwification of the eluted fragment by elutip-d columns (Schleicher & Schuell, Inc., Keene, New Hampshire), described in detail as follows:

1. Digest plasmid DNA (20-30 pg) with the appropriate restriction enzyme(s). 2. Electrophorese the digested DNA on 0.8-1% agarose gels at 25 volts. 3. Cut the gel fragment under UV illumination and place it in a dialysis tube. 4. Electroelute the DNA at 80 V for 1-2 hr. 5. Reverse the current for 30-60 sec. 6. Adjust to the low salt solution (0.2 M NaC1, 20 mM Tris-HC1 pH 7.5, 1 mM EDTA) and slowly pass it through an elutip-d column, preequilibrated with the same buffer. 7. Wash the column with 2-3 ml of the low-salt buffer, and dry it completely. 8. Elute the DNA with 0.4ml of the high-salt solution into a sterile Eppendorf tube and add 2 volumes of ethanol. Store overnight at -20°C. 9. Dissolve the pellet in TE (10 mM Tris-HC1, pH 7.5, 1 mM EDTA). 10. Determine the DNA concentration either by fluorometer or by running an alliquot in an analytical agarose gel with known amounts of DNA molecular markers. 11. Dilute the DNA to the desired concentration (for most purposes 1-3 pg/ ml of a 5-kb DNA fragment) with the injection buffer (10 mM Tris-HC1pH 7.5, 0.1 mM EDTA). It can be stored at -20°C. Most importantly, each DNA preparation should be checked for toxicity to fertilized eggs. We test the ability of each DNA preparation to allow development of microinjected eggs to the two-cell stage. If a large proportion of microinjected

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eggs are blocked at the one-cell stage, or show signs of morphological deterioration, the DNA solution is diluted twofold or more until about 80% of the eggs reach the two-cell stage. B. Superovulation 1. Mouse Strains Most mouse inbred strains reproduce poorly and the efficiency of gene integration is considerably lower than F1 or F2 hybrid strains. F2 hybrid zygotes generated by breeding C57BI/6J X SJL F1 mice have been most extensively studied. Other working combinations include C57BI/6J X C3H, C57BU6J X CBA, or BalblC X C57BU6J. We use the inbred mouse strain FVB/N, established at the NIH, as a donor strain for fertilized eggs (Takeo et aL, 1991). This mouse strain is characterized by vigorous reproductive performance and consistently large litters. In addition, fertilized FVBNeggs contain large and prominent pronuclei, which facilitate microinjection.

2. Mouse Colony

An access to an adequate animal care facilities is essential for the production and maintenance of transgenic mice. Mice should be housed under controlled conditions of temperature, humidity, and light cycle. The health of the colony must be monitored routinely, and serum samples should be tested for the presence of antigens of infectious viruses. Maintaining mice in pathogen-free conditions is advantageous. However, it is also extremely expensive. 3. Hormone Treatment Our mouse colony is under a 12-hr lighddark cycle. The end of the light cycle is at 6 P.M. and the end of the dark cycle is at 6 A.M. Superovulation is initiated by injecting 5 i.u. pregnant mare serum (PMS) (Sigma, St. Louis, Missouri, or Gestyl from Organon, or Folligon from Intervet Laboratories) between 2 and 3 P.M. followed by injecting 5 i.u. human chorionic gonadotropin (hCG) (Sigma) two days later between 12 and 1 P.M. Females injected with hCG are placed individually with stud males. Vaginal plugs are checked on the following morning. C. Media Preparation

We use CZB (Chatot et aL, 1989)and HEPES-CZB media for flushing, microinjection, and culturing of one-cell mouse embryos to the two-cell stage (Tables I and 11). Glutamine is added to the medium from a fresh 100-mM glutamine stock immediately before use. Media are prepared in disposable polystyrene tissue-culture tubes, using endotoxin-free, tissue-culture grade water (Sigma

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Table I Recipe for 100 ml 1OX Stock Solutions Stock A

Stock B Stock C Stock D Stock E

4.110 g NaCl 0.356 g KCI 0.293 g MgS04 . 7H20 0.161 g KH2PO4 6.180 g 60% sodium lactate 0.040 g EDTA 0.060 g penicillin 0.050 g streptomycin 2.100 g NaHCOs 0.010 g phenol red 0.360 sodium pyruvate 2.520 g CaC12 . 3H20 5.958 g Hepes 0.010 g phenol red

Chemical Co. No. W3500).Media are filter-sterilized through 0.22-pm Millipore filters in disposable filter holders, gassed with 5% COz, and stored at 4°C. We prepare fresh media every 2 weeks. The osmolarity of all media are tested by freezing-point depression and ranged from 274 to 295 mosmol. D. Embryo Recovery

Fertilized mouse embryos are recovered from the oviducts of plugged females as follows: 1. Sacrifice mice by cervical dislocation. Dissect the oviduct free of the ovary and uterus and place in a small Petri dish containing warm CZB medium. Bring an oviduct into a depression glass slide containing HEPES-CZB and 0.3 mg/ml hyaluronic acid under the dissecting microscope.

Table I1 Preparation of 10 ml CZB and HEPES-CZB Stock A

B C D E Glutamine (200 mM) HZO BSA (Sigma A4378)

CZB

Hepes-CZB

1 ml 1 ml 0.1 ml 0.1 ml 0.05 ml 1.15 ml

lml 0.1 ml 0.1 ml 0.1 ml 0.9 ml 0.05 ml 1.15 ml 5.0 g

5.0 g

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2. The fertilized eggs are located at the ampule, which because of superovulation is expanded and transparent. Through its thin wall, masses of fertilized eggs surrounded by cumulus cells can easily be observed. These masses are released from the ampule to the medium by tearing the thin wall with watchmaker’s forceps. 3. The cumuluscells are dispersed in 3-5 min because of the enzymatic activity, and the eggs are taken in a small volume and washed twice in 2 ml CZB medium. Fertilized eggs can be distinguished from unfertilized eggs by their prominent second polar body. E. Preparation of Holding and Microinjection Pipets

1 . Holding Pipet Holding pipettes are made of 5 - ~ Drummond 1 microdispenser tubes, Cat No. 105G, outside diameter 1.0 mm, as follows: a. Tubes are drawn by hand over a small Bunsen flame to an outside diameter of about 100 pm. b. The drawn capillary is clamped to the micoforge (e.g., Narashige, MF-79) and a clean break is made by positioning the filament next to the pipet, heating the filament until the glass starts fusing to the filament, cooling the filament for a few seconds, and pulling the tube away from the filament. c. Orient the capillary so that the tip is facing down and is brought close to the filament. The two should be in the same plane and in a sharp focus. The filament is turned on and the glass begins to melt. Stop heating the filament when the internal diameter of the tube is reduced to about 20 pm.

2. Microinjection Pipets We use the thin-wall glass capillaries with internal filament (TWlOOF-4 of World Precision Instruments, Inc.) with an outside diameter of 1.0 mm. There is no need to clean or pretreat these capillary tubes in any way. In fact, such pretreatments should be avoided. a. Mount a pipet in the Sutter PC-84 puller. This puller is designed for maximum control and flexibility. The temperature, the force, the distance, and the time for a pull are all programmable with a defined sequence of up to 16 steps. b. Choose an appropriate program following the detailed instruction of the manual. For DNA microinjection, a tip size of 0.3-0.5 pm gives an ideal flow. c. Determine the tip size of the pulled capillaries using the Sutter LW-87 tip calibration device. d. Store pulled pipets for several hours in a dust-free box. e. Load DNA into the microinjection pipet by placing the base of the pipet into the DNA solution. The solution is drawn by capillary action to the tip.

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F. Microinjection: Equipment and Setup

1 . Equipment

We use the fixed-stageNikon Diaphot inverted microscope with long-workingdistance Nomarski objectives and Narashige NT micromanipulators. The micromanipulators are arranged so that the syringe micrometer on the right side, controlling the holding pipet, is connected via polyethylene tubing to the instrument collar on the left micromanipulator. The polyethylene tubing connected to the holding pipet is filled up with Fluorinet (Sigma) and all air bubbles expelled from the system. The micromanipulator on the right side, controlling the microinjection needle, is connected via polyethylene tubing to the picoinjector PLI-100 (Medical Systems Corp.). The pico-injector is a single-channel, pneumatic, digital injector that delivers small liquid volumes precisely by applying a regulated pressure for a set time.

2. Microinjection Chamber Homemade 75 X 38 m m glass chamber slide is used. Several stripes of HepesCZB medium are placed in the middle of the slide and covered with silicone oil. Fertilized eggs (20-30) are positioned at one end of the microdrop under the dissecting microscope. 3. Microinjection a. Place the chamber on the microscope stage. The elongated microdrops should be parallel to the Y axis of the stage and the embryos are at the front of the stage. Bring the embryos into focus with the lower-power objective (X10). b. Insert the holding pipet into the instrument collar, making sure that all air bubbles are expelled. With the micrometer, push the Fluorinet into the pipet until a drop is hanging at the tip. Move the chamber to the rear of the stage and carefully lower the pipet into the microdrop. Make sure that the angle is such that only the tip of the pipette will touch the glass slide. Move the chamber into the front of the stage and suck up an embryo with the micrometer. c. Insert the microinjection pipet into the instrument collar. Set up the picoinjector to deliver a constant positive pressure that prevents back flow and clogging of the pipet. Align the pipet parallel to the holding pipet and slowly lower it into the microdrop. Do not allow the pipet to reach the bottom of the drop, as it might break. Also, do not allow the two pipets to collide. Switch from the X10 to the X 2 5 objective. d. With an embryo firmly held by the holding pipet, bring the male (usually bigger) pronucleus into focus. Bring the tip of the microinjection pipette into a focus, with the fine vertical drive of the manipulator, next to the zona pellucida. Move the pipette into the pronucleus, avoiding the nucleoli, and microinject by

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pressing the foot switch of the pico-injector. If the pronucleus swells, injection was successful (Fig. 1).If no swelling is visible, then either the tip is blocked or the pipette was not inserted into the nucleus. In the former case the pressure can be increased, whereas in the latter, the pipet should be withdrawn and

Fig. 1 Injection into the male pronucleus of mouse fertilizedegg. (A) Orienting the microinjection pipet to the optical plane of the male (larger) pronucleus. (B) Insertion of the pipet and injection of the DNA solution. (C) Withdrawal of the pipet.

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realigned at the optical plane of the pronucleus prior to the second attempt. Mouse embryos are usually resistant to multiple penetrations if the plasma membrane is not ruptured. e. After a successful injection, withdraw the pipet and move the chamber to the back of the stage. Expel the embryo from the holding pipet by applying pressure to the micrometer and move the chamber to the front of the stage again to suck up the next embryo. f. When all the embryos in that drop have been injected, take the injection chamber to the dissecting microscope, transfer the healthy-looking eggs into a drop of CZB medium under silicone oil, and place them in the incubator. g. Take 20-30 fresh eggs into the next microdrop and repeat the microinjection steps just described.

4. Troubleshooting Microinjection into the pronuclei of fertilized eggs is still endowed with numerous technical problems. The rule of the thumb is that trying to overcome flow problems of a micropipet is better than replacing it. Basically, problems arise in the action of either the injection needle or the holding pipet. a. The needle is blocked. This is the most common problem and is likely due to proteins sticking to the tip during penetration and withdrawal. Apply momentary high pressure with the pico-injector to clear the clogged pipets. Alternatively, the tip of the pipet can be glanced against the holding pipet to increase the size of the opening. If these remedies do not clear the pipet, replace it. b. Aged pipets become very sticky and upon withdrawal, nuclear material is removed as well. Even if such an embryo does not lyse immediately, it will not survive. The pipet should be discarded. c. The pressure of injection can push the nucleoli from the nucleus to the cytoplasm. Such embryos do not survive. d. Avoid the nucleoli! These are very sticky organelles and when the pipette is withdrawn nucleolar material will be dragged along. This will kill the egg and clog the pipet. Pipets clogged with nucleolar material must be replaced. e. The holding pipet, Oil droplets may clog the pipet. In addition, if the tip size is too big, embryos may be sucked up and clog the pipet. G. Vasectomy and Embryo Transfer

The most common practice is to transfer embryos several hours after microinjection into the oviducts of 1-day pseudopregnant mice. However, transferring can be performed after overnight culturing of injected embryos at the two-cell stage to 1-day pseudopregnant females.

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1. Vasectomy Anesthetize an adult male mouse by intraperitonealinjection of 0.4 ml Avertin. Place the mouse on its back and after gently pushing the testes into the abdominal cavity, make a small (about 1-cm) transverse incision just above the preputial gland. With blunt forceps, pull out the testicular fat pad until the testes are exposed. Cut as much of the vas deferens as possible and repeat the procedure with the other testes. Replace the testes into the abdominal cavity and apply two wound clips to the skin. We do not suture the body wall. Vasectomized mice are allowed to recover for about 2 weeks before they are caged individually and mated with CD1 females that serve as foster mothers for microinjected embryos. Avertin is prepared by dissolving 2.5 g 2,2,2-tribromoethanol (Aldrich) in 5 ml ten-amyl alcohol in warm tap water. When it is completely dissolved, add doubledistilled water to a final volume of 200 d.Store at 4°C in the dark.

2. Embryo Transfer Pipet Introduce air bubbles into a finely drawn Pasteur pipet and up to 15 embryos in a minimum volume of CZB medium.

3. Embryo Transfer Anesthetize a pseudopregnant CD1 mouse by intraperitoneal injection of Avertin. Place the mouse on the stage of a dissecting binocular (e.g., Zeiss SV8) and make a longitudinal incision at back midline at the level of the uterus. With blunt forceps pull out the ovarian fat pad along with the ovary, oviduct, and some of the uterus. Clip a serafine to the fat pad and spread it on the skin so that the ovary and oviduct remain outside the body wall. The opening of the infundibulum always faces the tail. Apply a few drops of 0.1% adrenaline and tear the ovarian bursa with two watchmaker forceps. Adrenaline treatment prevents bleeding from the injured vascularized bursa for several minutes. Push the bursa under the ovary and identify the infundibulum in the crevice between the ovary and oviduct. Insert the embryo transfer pipette into the infundibulum, being careful not to tear the delicate tissue. Expel the embryos and air bubbles and slowly remove the pipette. Return the ovary to the abdominal cavity. Repeat the procedure on the other side. Close the skin with a wound clip and keep the mice warm until they recover from the anesthesia. Embryo transfer into the oviduct is considered to be the most difficult step in the production of transgenic mice. Therefore, one should allow some time to practice with nonmanipulated eggs. H. Identification of Transgenic Mice

1. Tyrosinase Gene Marker Screening for positive transgenic mice is labor and time consuming, requiring DNA analyses of tail biopsies (see later discussion) by Southern blot hybridiza-

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tion or PCR. Therefore, the idea that transgenic mice can be identified at birth is very appealing. It was demonstrated that introducing a functional tyrosinase gene into albino mice led to pigmentation of the eye and skin with high penetrance (Beermann et al., 1991). It is also established that coinjection of two independent genes into fertilized mouse eggs results in most cases in cointegration of both genes at the same chromosomal site. Therefore, coinjecting the gene of interest with the tyrosinase gene would lead to cointegration and facilitate identification of transgenic animals over the albino genotype. However, caution should be excercised in adopting this approach, since we and others could not reproduce these results.

2. Tail DNA Analyses This is the traditional method for screening transgenic animals. a. Anesthetize 3-week-old mice with 0.3-0.4 ml of Avertin and cut about 0.5 cm of the tail into a microcentrifuge tubes. b. Digest the tail biopsy for several hours in 0.5 ml solution containing 100 mM EDTA, 10 mM NaCl, 10 mM Tris-HC1, pH 8.0,1% SDS, and 200 pg/ ml proteinase K at 55°C in a shaking water bath. c. Extract the digest with phenolkhloroforndisoamyl alcohol followed by extraction with chloroform/isoamyl alcohol. d. Adjust to 2.5M ammonium acetate and precipitate the DNA with 1volume of cold isopropanol by spooling on a glass rod. e. Wash the DNA with 70% ethanol and dissolve in 50 pl of 10 mM TrisHCl, pH 8.0,l m M EDTA (or 0.1 mM EDTA if PCR is used for the screening). This method yields DNA that is usually clean enough for dot blots, restriction digests, and Southern blots, as well as for PCR. Although isolating DNA from a large number of mice is laborious and expensive, it is highly reliable. There are several protocols for quicker DNA isolation that avoid extractions in organic solutions. However, such protocols are less reliable and might even be misleading. If one must employ these procedures, appropriate positive internal-control primers should be included.

111. Production of Transient Transgenics A. Advantages

Production of stable transgenic strains is time and labor consuming. Usually the analysis of these mice awaits the F1 or F2 generation. In addition,maintaining a large number of strains requires investing substantial resources in adequate animal facilities. However, in many cases, such as for the identification of cisacting control elements conferring a particular pattern of gene expression or for

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the analysis of the consequences of dominant gene expression on embryonic development, a quicker means would be to produce transgenic embryos and analyze them directly instead of establishing stable strains for each integration event. We have used this approach to dissect the regulatory elements of the human and quail myoD genes (Goldhamer et al., 1992,1995; Pinney et al., 1995), as well as to study the consequences of ectopic myoD expression on mouse embryos (Faerman et al., 1993).

IV. Analysis of Expression at the R N A Level A. RNA Isolation

Total tissue RNA is isolated by the urea-LiC1method of Auffray and Rougeon (1980). All solutions should be sterilized and prepared in DEPC-treated water.

Care must be taken at all steps to prevent RNase activity. Place a tissue in 5 ml of 6 M ured3 M LiCl in a sterile plastic tube on ice. Add few drops of 1-octanol to prevent foaming. Homogenize for 30 to 60 sec in a Polytron homogenizer at full speed. Leave at 4°C overnight and spin for 30 min at 10,000 rpm in the cold. 4. Dissolve the pellet in 2 ml solution containing 7 M urea, 2% SDS, 0.35 M NaC1,l mM EDTA, 10 mM Tris-HC1, pH 8.0, and 50 pglml heparin. 5. Extract the RNA with phenolkhlorofodisoamyl alcohol and then with chloroform/isoamyl alcohol. 6. Adjust to 2.5 M ammonium acetate and add 2.5 volumes cold ethanol. 7. Pellet the RNA at 10,OOO rpm for 15 min in the cold and dissolve in 100200 pl sterile water. RNA concentration is determined by reading the optical density at 260 nm. Store at -70°C until used. 1. 2. 3. 3.

B. Northern Blots 1. Prepare a 1.5% agarose gel of 10 X 15 X 6 cm dimension by dissolving 1.5 g agarose in 74 ml water. Cool to 60-65°C and add 10 ml of 1OX MOPS buffer (0.2 M MOPS, 50 m M sodium acetate, 10 mM EDTA, pH 7.0) and 16 ml of 37% formaldehyde solution in fume hood. 2. Dissolve a pellet of 20 pg total RNA in 5 pl DEPC-treated water and 19 p1 dye buffer (100 p1 1OX MOPS, 160 pl formaldehyde, 500 p1 formamide, 100 pl glycerol, and 0.1% bromphenol blue). 3. Heat the sample at 65°C for 5 min and immediately quench in ice-water

bath. Add 0.5 pglml ethidum bromide. 4. Run the gel at 10 Vlcmfor about 2-3 hr, until the bromphenol blue migrates about 10 cm.

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5. Soak the gel in 1OX SSC twice for 20 min each with gentle shaking. This removes most of the formaldehyde. 6. Photograph the gel on a short-wave transilluminator. 7. Transfer the RNA in 1OX SSC by capillary action onto Genescreen membrane. 8. Fix the RNA to the membrane by placing the nylon membrane with the RNA facing down on a short-wave transilluminator for 5 min. 9. Prepare 32P-labeledprobes (RNA or DNA) using standard procedures. 10. Prehybridize in 50% formamide, 5X SCC, 5X Denhardt, 50 mM sodium phosphate buffer, pH 6.8, 250 pglml sheared denatured salmon sperm DNA, 100 pg/ml yeast tRNA, and 1%SDS. Prehybridization temperature is determined by the type of probe being used. For RNA probe use 60-65"C, for DNA probe use 50-55°C. 11. Hybridize in 50% formamide, 5X SSC, I X Denhardt, 20 mM sodium phosphate buffer, pH 6.8, 100 pg/ml sheared denatured salmon sperm DNA, 100 pg/ml tRNA, 1%SDS, and 10% dextran sulfate.

C. RNase Protection Assay

The RNase protection assay is a highly sensitive assay to detect and quantitate mRNA. 1. Linearize the plasmid DNA on the 3' of the insert with the appropriate restriction enzyme. 2. Synthesize RNA probe using SP6, T3, or T7 RNA polymerase in a reaction mixture containing 32P-UTP(specific activity 800 CUmmol), 0.5 pg linearized plasmid, 500 p M each ATP, GTP, and CTP, 12.2 p M UTP, 30 pCi 32P-UTP, 16 units RNasin, 10 mM NaCl, 40 mM Tris-HC1, pH 7.5, 10 mM DTT, and 10 units of the RNA polymerase. Incubate for 1 hr at 37°C. 3. Remove the template DNA with 20 pg/ml RNase-free DNase in the presence of 10 pg carrier tRNA and 20-40 units RNasin. Incubate for 15 min at 37°C. Stop the reaction with 2% SDS. 4. Add ammonium acetate to 2.5 M and ethanol-precipitate. Dissolve the pellet in 200 pl hybridization buffer (80% deionized formamide, 0.4 M NaCl, Pipes, pH 6.4, and 1 mM EDTA). 5. Dissolve a pellet containing 10 pg total RNA and 10 pg tRNA in hybridization buffer containing about 200,000 cpm of the RNA probe. 6. Heat the sample at 85°C for 5 min and incubate overnight at the appropriate temperature under water. 7. Stop the hybridization by adding 180 pl of a solution containing 0.3 M NaCl, 10 mM Tris-HC1, pH 7.5,5 mM EDTA, 8 pg RNase A, and 0.4 pg RNase T1. Incubate for 1hr at 37°C.

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8. Add 10~ 1 2 0 % SDS and 50 pg proteinase K and incubate at 37°C for 15 min. 9. Add 10 pg carrier tRNA, ammonium acetate to 2.5 M,and 2.5 volumes of ethanol to precipitate the hybrids. 10. Dissolve the pellet in 5 p1 running buffer and electrophorese on 5% sequencing gel. D. RT-PCR

The greatest advantage of RT-PCR for detection and quantitation of specific RNAs is its speed. However, because of its sensitivity, false-positive results caused by contaminationwith minute amounts of DNA are the major restriction. 1. cDNA synthesis with random primers. In a total volume of 40 pl, add 1OX Taq polymerase buffer (500 mM KC1,100 mM Tris-Ha, pH 8.3,15 mM MgClz, 0.1 mg/ml gelatin), 2 pg total RNA, 100 pmol hexamers, 1.25 mM each dNTP, 40 units RNasin, 200 units Moloney murine leukemia virus reverse transcriptase. Incubate for 1 hr at 42°C. 2. PCR in a total volume of 50 pl containing 1OX taq polymerase buffer, 10 pl cDNA, 100 pmol oligonucleotide primers, 5 pl DMSO. Heat at 95°C for 2 min. Add 2 units Taq DNA polymerase. Annealing and elongation temperature and time should be tested for each set of primers. It is recommended to include internal positive control primers for each reaction. Suitable primers could be as follows: L-pyruvate b a s e 3’ S16 ribosomal protein

5’ GGGTCAGTTGAGCCACACTCG3’

5’ AAGCAACGTAGCAGCATGGAA 5‘ AGGAGCGATITGCTGGTGTGGA 3’ 5’ GCTACCAGGCCTITGAGATGGA

The number of amplification cycles depends on the method of detection. If ethidium bromide staining is the ultimate goal, then 30-40 cycles should be performed. If RNA quantitation is required, the reaction should be stopped while still linear and the amount synthesized detected by Southernblot hybridization using an internal oligonucleotide as a probe.

E. I n Sit# Hybridization to Tissue Sections In situ hybridization (ISH) refers to the set of methods combining cytological and histological techniques with methods of nucleic acid hybridization. This combination allows detection of specific nucleotide sequences within cells and tissues, thus providing visualization of genes and/or gene products. The si@cance of ISH cannot be overestimated because of its high sensitivity and resolution, permitting detection of single-copy genes on chromosome preparations or several hundred mRNA molecules in single cells on tissue sections. ISH became

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the method of choice to visualize the spatio-temporal pattern of gene expression during embryogenesis. There are several categories of ISH according to the probe used (DNA, RNA, or oligonucleotide),the method of labeling its detection (radioactivevs. nonradioactive), and the target (DNA vs RNA). Important theoretical as well as practical aspects of ISH technique are comprehensively discussed in several reviews (Wilkinson and Green, 1990; Polak and McGee, 1990; Wilkinson, 1992; Wilkinson and Nieto, 1993; Sassoon and Rosenthal, 1993). Protocols described in this chapter are used routinely in our laboratory to study the expression of muscle regulatory and structural genes in normal and transgenic embryos and can be adopted for other tissue systems. We describe procedures for detecting mRNA on tissue paraffin sections, using 35S- or digoxigenin-labeled riboprobes, and the visualization of gene expression using the whole-mount ISH on postimplantation (8.5-12.5 dpc) mouse embryos.

1. Radioactive in Sit# Hybridization to Tissue Sections a. Tissue Preparation

All steps (tissuepreparation, sectioning, and prehybridization treatments) must be carried out in RNAse-free conditions with sterile solutions and equipment. All reagents used for ISH are kept separately. Fixution. Freshly prepared fixative is made by mixing 4 g paraformaldehyde with 70-80 ml water. Ten microliters of 10 N NaOH is added and the mixture is heated in a fume hood to 70"C, until the powder is dissolved. After cooling, 10 ml of 1OX PBS is added and the volume is adjusted to 100 ml with water. The pH should be 7.4. Fix excised tissues in ice-cold fixative for 16-24 hr. The thickness of tissue blocks must not be more than 3-5 mm. The volume of fixative must be 20-30 times the volume of the tissue sample. Purufin Embeding. Wash fixed tissues in PBS at +4"C (2 X 15 min). Dehydrate through a series of ethanols: 50%, 70%, 95% and three changes of absolute ethanol. Then incubate in chloroform (twice). Impregnate in three changes of molten paraffin wax (e.g., Paraplast) at 60°C. Duration of incubations depends on the size of tissue blocks. For mouse embryos at developmental stages of 9.5-11.5 days, 30 min incubation at each step is sufficient. For later embryonic stages or large pieces of adult tissue, incubations must be prolonged. Processing can be interrupted and the material be stored in 70-100% EtOH at 4°C. Following impregnation, the sample is oriented in the molten paraffin with heated needles and left at room temperature until it solidifies. Paraffin blocks can be stored indefinitely at +4"C. Sectioning. High-quality sections are prerequisite for unequivocal ISH results. Therefore, it is a good practice to select only the best and most informative sections in terms of morphology. This can be done by collecting some sections for histological evaluation and choosing neighboring sections according to the presence and quality of the desirable structures for ISH. Sections are collected

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on slides subbed with 3-aminopropyltriethoxysilane(TESPA) to prevent detachment of sections during the harsh treatments of the ISH procedure. The slides are prepared by washing in hot 10% laboratory detergent in water, rinsing in running hot water and twice in distilled water, baking in an oven at 200-250°C for 2-3 hr, dipping in 2% TESPA in acetone for 10 sec, rinsing twice in acetone and once in DEPC-treated water, and drying at room temperature or at 37°C. Subbed slides can be stored indefinitely in dust-free conditions. Float separate sections or ribbons of serial sections (4-6 pm) on a drop of 2% EtOH in DEPCtreated water on a subbed slide. Place the slide on a slide-warming table heated at 40-45°C. Let them dry overnight at 40-50°C and store in a refrigerator. Under these conditions slides can be maintained over a year. b. Riboprobe Preparation Riboprobes are synthesized by in vitro transcription reaction with SP6, T3, or 'I7 RNA polymerases as described in Section 3 IV.D, using [CZ-~~SIUTP and 1 p10.75 M dithiothreitol (DTT). Importantly, there is no need to hydrolyze the probe. The amount of probe applied is critical. More than 105 cprdpl results in a significant increase in the background, whereas the intensity of the hybridization signal remains essentially the same. For most purposes we use probes at 2 X lo4cprdpl, which results in exposure time that varies from 3 days (for highly abundant RNAs) to 3 weeks (for low-abundance RNAs).

c. Prehybridiration The aim of the prehybridization step is to make the target RNA accessible to the probe. This is done by treating the biological samples with proteinase K. The treatment described next appears to be suitable for different tissues and for various probes. The most important parameter is the temperature of hybridization, which must be determined empirically for each probe. All washing steps can be performed in Coplin jars (glass or plastic) for a small number of slides or in a stainless steel slide rack, for 30-50 slides. Solutions must be preheated prior to treatment. If not stated otherwise, incubations are at room temperature.

1. Bring the slides to room temperature and deparaffinize by 2 X 10-min changes in xylene. 2. Rehydrate sections by passing sequentially through absolute ethanol (2 X 5 min), 95%, 70%, and 50% ethanols (2 min each). 3. Wash in DEPC-treated water (5 min). 4. Incubate in 2X SSC for 30 min at 70°C. 5. Wash in DEPC-treated water (5 min). 6. Treat with proteinase K solution (2 p g / d in 0.2 M Tris-HC1, pH 7.6, 0.1 M EDTA) for 20 min at 37°C. 7. Refm in 4% paraformaldehyde in PBS for 20 min. 8. Treat slides in freshly prepared 0.2% glycine in PBS for 5 min.

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9. Wash in DEPC-treated water for 5 min. 10. Transfer slides into freshly prepared triethanolamine solution (5.53 ml triethanolamine per 0.5 liter DEPC-treated water) and add acetic anhydride (1.25 ml per 0.5 liter). Incubate for 5 min with continuous mixing over a magnetic stirrer. Add the same amount of acetic anhydride and incubate for another 5 min. Wash the slides in DEPC-treated water for 5 min. The purpose of the acetylation step is to decrease the nonspecific binding of the probe to the section. However, in our hands this step did not lead to noticeable improvement of the results. 11. Wash in DEPC-treated water for 5 min. 12. Dehydrate in graded ethanols (2 min each) 50%, 70%, 95%, 100%. 13. Air-dry the slides and proceed to the hybridization step. d. Hybridization 1. Apply the hybridization solution (Table 111) containing the probe onto pretreated sections (approximately 20 p1 per cm2). Do not allow the drop to dry.

2. Cover the drops with precut pieces of Parafilm and displace air bubbles. 3. Arrange slides horizontally (with sections facing up) in a slide box containing filter paper soaked 50% formamide 5x SSC, and seal the box with an electric tape. For a large number of slides we use a stainless steel slide rack placed vertically in a heat-sealable bag. Incubate overnight at the hybridization temperature (50-65°C). e. Posthybridization Washings

To prevent the oxidation of the 35S-labeledprobe, leading to high background, all solutions used prior to RNAse digestion must contain 10 mM D l T or 1%pmercaptoethanol. 1. Place slides in an appropriate container with 5x SSC, 1% P-mercaptoethanol at 65°C. Incubate for 30-60 min until the Parafilm pieces disengage. 2. Wash slides in 2x SSC, 50% formamide, 1%mercaptoethanol at 65°C for 30 min followed by washing in 2x SSC at 37°C 3 X 5 min. Table III Composition of In Situ Hybridization Buffers 50% formamide 4 X SSC (pH 8.0) 1 X Denhardt’s 0.5 mg/ml herring sperm DNA 0.25 m g / d yeast RNA 10 mM DTT 10%dextran sulfate

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3. Treat the slides with RNAse A (10 pg/ml) in 0.4 M NaCl, 0.01 M Tris-HC1 (pH 7.5),5mM EDTA for 30 min at 37°C. 4. Repeat the high-stringency washing (step 2) followed by 2x SSC washing. 5. Wash in 3 liters of 0 . 1 ~SSC at 37°C for 15 min. 6. Dehydrate rapidly through 50,70,90% ethanols containing 0.3 M N€€,AC and finally by two changes of absolute ethanol. Air dry.

f. Autoradiography For a rapid assessment of the hybridization,expose slides to P-max film (Amersham) for 12-24 hr. The hybridization signal on the film can give some indication about the abundance of the transcripts and the localization of the expressing cells. To localize the signal more precisely, high-resolution autoradiography using nuclear track emulsion is required. Important theoretical and practical aspects of histoautoradiography are discussed in Rogers (1979). 1. In the darkroom under safelight conditions (e.g., 25-watt lamp with Ilford 920 filter), melt Kodak NTB-2 emulsion gel in a 50-ml graduated plastic tube containing double-distilled water (1 :1) for 1-2 hr at 43°C. 2. Dip a clean blank slide into the emulsion and remove slowly. Repeat this step with blank slides until no air bubbles appear at the surface of the slide. 3. Dip the hybridized slides into the emulsion. The volume of emulsion needed to cover a standard histological slide is 0.3-0.5ml. 4. Withdraw the slides from the emulsion vertically and blot excess emulsion. 5. Dry slides vertically in the dark for 2-3 hr at room temperature. Using a slide dryer significantly accelerates drying. 6. Store dipped slides overnight at room temperature in a light-tight box. 7. Add into each box a bag with silica gel, seal the box with electric tape, and store at +4"C.

To determine the optimal exposure time (several days to several weeks), test developments should be performed. g. Development and Staining

Warm the box to room temperature for 1-2 hr to avoid condensation. Solutions used for developmentmust be freshly prepared and brought to room temperature. The volumes of developer and fixer should be 5-10 ml per slide. 1. Develop slides in Kodak D-19 developer for 3 min in the dark. 2. Rinse briefly in distilled water. 3. Fix in 30% sodium thiosulfate for 5 min. 4. Wash in at least three changes of distilled water (20 min each) and dry.

For cytologicalobservation the sections must be stained. Usually, light nuclear

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staining allows visualization of both cells and silver grains. We routinely use hematoxylin (Mayer’s or Erlich’s) for counterstaining: 1. The slides are incubated in hematoxylin for 10-30 sec, then washed in 2-3 changes of distilled water followed by washing under tap water. 2. When the color of the sections turn from purple to blue, wash again in distilled water and air-dry. 3. To prepare permanent preparations, put dried slides into xylene and then mount with covedlips using DPX (BDH) or other mountant.

The hybridization signal can be visualized and photographed using either bright- or dark-field illumination. With bright-field illumination, silver grains appear as black dots above the section. If the signal is strong, bright-field photography is adequate. However, when the signal is moderate or weak, dark-field illumination should be used. Under these conditions the signal appears as bright shining dots over a black background and no tissue details can be discerned. Therefore, the standard presentation of radioactive in situ hybridization includes a combined black-and-white photograph of the same section taken with brightand dark-field optics (Fig. 2A,B). Some hematoxylin stained tissues shine brightly under dark-field illumination. To overcome this problem, photographs of unstained sections can be taken with phase-contrast optics or the dark-field photographs can be taken after destaining in ethanol. Another way of presenting results is to make a double exposure of bright- and dark-field images of the same frame on a color film.This results in a combined color picture displaying both silver grains and tissue section details.

2. Nonradioactive in Situ Hybridization with Digoxigenin-Labeled Probes Nonradioactive ISH (nrISH) is based on probes labeled with haptens such as biotin, digoxigenin (dig), or fluorescein. Antisense riboprobes are prepared by in vitro transcription using the appropriate modified nucleotides. The protocols for nonradioactive and radioactive ISH are essentially the same. The most significant difference is the detection of the bound probe. With digoxigenin-labeled probes, anti-digoxigenin antibodies conjugated with marker enzyme (horseradish peroxidase or intestinal alkaline phosphatase) are employed for the immunohistochemical detection. There are several advantages of nrISH, the most important of which is the high resolution not achievable with 35S-labeledprobes and autoradiography. In addition, it avoids the problem of handling, storage, and wasting of radioactive materials, and the histochemicaldetection is quicker, simpler, and more controllable than autoradiographic detection of rISH signal. a. Synthesis of Dig-Lubefed Probes Dig-labeled antisense riboprobes are synthesized using a Dig RNA labeling kit (Boehinger-Mannheim) according to the manufacturer’s protocol. Alternatively,

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Fig. 2 In situ hybridizations.Bright-field (A) and dark-field (B) photomicrographsof a transverse section through a somite of 9.0-dpc mouse embryo hybridized to "S-labeled myogenin riboprobe. (C) Photomicrographof aparasagittal section through the cervical somites of 11.5-dpcmouse embryo hybridized to dig-labeled myoD riboprobe. (D) Whole-mount in situ hybridization showing myogenin expression in 9.5-dpc mouse embryo.

one can use the components of the in v i m transcription reaction purchased from other brands in combination with Boehringer's nucleotide mixture containing dig-labeled UTP. The probe can be stored at -70°C for months. The amount of probe synthesized can be quantitated using dig-labeled control RNA (Boehringer-Mannheim) as a standard. Serial dilutions of the probe and

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control RNA are spotted onto a nylon membrane and revealed with anti-dig antibody conjugate according to the manufacturer’s protocol. For most purposes a concentration of 0.2-1.0 pg/ml of probe is needed. However, it is recommended to determine the optimal concentration by running trial hybridizations with serial dilutions of the probe. The composition of the hybridization buffer is the same as for rISH but without DTT. b. Prehybridization and Hybridization Tissue section treatments and hybridization conditions for nrISH do not differ from these described for rISH. Notably, in contrast to rISH, acetylation decreases the background of hybridizations with dig-labeled probes. Perform stages 1-10 of the prehybridization treatment protocol for rISH and hybridize at 50-65°C as described earlier. c. Posthybridization Washings and Immunodetection The slides are washed according to steps 1-6 of the rISH posthybridization protocol. There is no need to add D l T or @-mercaptoethanol to the washing solutions. The imrnunohistochemicaldetection step is performed at room temperature. Avoid drying of the sections at all steps. 1. Wash slides 3 X 5 min each in TBS (0.15 M NaC1,50 mM Tris-HC1, pH 7.4). 2. Apply a blocking solution (50-100 pl TBS with 3% normal goat or sheep serum). Incubate in a humid chamber for 30-60 min. 3. Tip off excess blocking solution and do not wash. 4. Apply alkaline phosphatase-conjugated anti-digoxigenin antibodies (antidigoxigenin-AP, Fabfragments, Boehringer-Mannheim) diluted 1:500 in TBS with 1%normal goat or sheep serum and incubate for 2 hr. If background staining (due to nonspecific ineraction of the conjugate with the section) appears, the conjugate can be absorbed with heat-inactivated acetone powder prepared from liver of the species used as the source of material for hybridization (Table IV). 5. Wash 3 X 10 min in TBS. 6. Wash 2 X 5 min each in alkaline phosphatase buffer (APB - 100 mM TrisHC1, pH 9.5, 150 mM NaCl, 25 mM mgC12). 7. Reveal alkaline phosphatase activity by incubating sections in the dark with a freshly prepared staining solution. To prepare staining solution, dissolve 5 mg levamisole in 10 ml APB and add 50 pl stock solution of 5-bromo-4-chloro-3indolyl phosphate, 4-toluidine salt (BICP), and 37.5 pl stock solution of nitro blue tetrasolium chloride (NBT).

BCIP stock solution: 50 m g h l in dimethylformamide. NBT stock solution: 100 mg/ml in 70% dimethylformamide. Stock solutions are stable for several months at -20°C in the dark. Ready-for-use stock solutions can be purchased from Boehringer-Mannheim.

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Table IV Preparation of Liver Acetone Powder ~~~

~~

1. Homogenize liver in a minimal volume of PBS on ice. 2. Add 4 volumes of cold acetone and mix well. Keep on ice for 30 min with occasional mixing. Centrifuge at l0,oaOg for 10 min and discard the supernatant. 3. Resuspend the pellet with cold acetone and mix vigorously. Keep on ice for 10 min. Repeat centrifugation. 4. Air-dry the pellet at room temperature on a clean piece of filter paper and store at 4°C in an airtight container. To prepare 1 ml of absorbed antibodies (1:500): 1. To an Eppendorf tube containing 2 mg acetone liver powder add 0.5 ml TBS and heat at 70°C for 30 min. Cool on ice. 2. Add 2 pl anti-digoxigenin antibodies alkaline phosphatase conjugate and incubate with shaking on ice for 1hr. 3. Centrifuge at 10,OOOg for 15 min at +4"C. To the supernatant add 10 p1 normal goat or sheep serum and 0.5 ml TBS.

The appearance of staining varies significantly depending on the abundance of the mRNA. With abundant mRNAs, staining appears after 15-30 min (and up to several hours) of incubation at room temperature. Longer incubations (from 12-16 hr to several days) must be performed at +4"C because of higher stability of the substrate solution at low temperature. Staining is monitored by periodic microscopic observations. The reaction is stopped by washing in TE (50 mM Tris-HC1,l mM EDTA, pH 8.0). The slides are rinsed in distilled water and counterstained with methyl green. Cover the sections with water soluble mountant (e.g., glycerol-gelatin or GVA mountant, Zymed). An example of an nrISH result is presented in Fig. 2C. As an alternative to alkaline phosphatase, horseradish peroxidase conjugates can be used. However, this alternative is recommended only for abundant mRNAs, because of the instability of the peroxidase substrate solutions and the inhibition of enzyme activity by the color product. Nevertheless, for double ISH, employing both 35S- and dig-labeled probes, peroxidase detection is preferable because the product of the alkaline phosphatase reaction interfereswith autoradiography. Another advantage of the peroxidase reaction product is its insolubility in xylene, allowing the use of xylene-based mountants (e.g., DPX, Permount) with superior optical properties. For the peroxidase detection protocol a slight modification must be introduced. After deparaffinization in xylene, sections are incubated for 30 min in absolute methanol with 0.3% H202to inhibit endogenous peroxidase activity. The sections are then transferred into absolute ethanol and processed according to the nrISH protocol as described earlier. The immunodetection is performed as follows: 1. Wash slides in 0 . 1 ~ SSC followed by 3 X 5 min rinses in PBS. 2. Incubate for 45 min in PBS with 3% of normal goat or sheep serum.

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3. Tip off excess fluid and apply anti-digoxigenin antibodies conjugated to peroxidase (anti-digoxigenin-POD, Fabfragments, Boehringer-Mannheim) diluted 1:100 and preabsorbed (if needed) with the appropriate acetone liver powder, and incubate in a humid chamber for 1 hr. 4. Wash in PBS (3 X 10 min). 5. Incubate with freshly prepared substrate solution: 0.15 M NaC1,O.l M TrisHCl, 0.01 M imidazole, pH 7.6,0.5 mg/ml3.3-diaminobenzidinetetrahydrochloride, and 0.01% Hz02. Staining develops within several minutes and incubation longer than 30 min does not result in the intensification of staining.

F. Whole-Mount in Situ Hybridization of Mouse Embryos The major advantage of whole-mount ISH ( W I S H ) is the ability to study the spatial pattern of gene expression directly in the embryo, instead of the timeconsuming sectioning, hybridization, and three-dimensional reconstructions. We tested several protocols of W I S H and found the one developed by Barth and Ivarie (1994), for quail embryos, to be relatively simple and highly reproducible for 8.5- to 12.0-dpcstage mouse embryos. W I S H also employs dig-labeled riboprobes. 1. Fixation of Embryos

Dissect embryos free of extraembryonic membranes in PBS and fix overnight in 4% paraformaldehyde in PBS. Wash fixed embryos in 2-3 changes of 70% EtOH and proceed for hybridization or store in 70% EtOH at -20°C (for several months). Using watchmaker’s forceps and tungsten microneedles, open body cavities to allow free flow of solutions; otherwise, entrapped reagents may cause unacceptable high background. This can be performed while embryos are still in paraformaldehyde or in alcohol. With unturned embryos (8.0-8.5 dpc stage), just remove the extraembryonic membranes and tear the remains of the amnion, to stretch the embryo. With 9.0-10.0 dpc embryos, we make a sagittal incision in the frontal surface of the forebrain to open the brain cavity. From this embryonic stage, the heart must also be teased. With embryos of 10.5-12.0 dpc stages, additional incisions are made in the forebrain vesicles and the roof of the fourth ventricle must be ruptured. During pre- and posthybridization washings, embryos are kept in a roundbottom polypropylene 10-15 ml tube (e.g., Corning 17 X 100 mrn tube, Cat. No. N-25225). Incubations with probe and antibody solutions are done in 2-ml polypropylene test tubes with screw caps (e.g., Sarstedt Cat. No. 72.693). Use polyethylene Pasteur pipets (e.g., Sterilin, Cat. No. PP89) to remove solutions from tubes and to transfer embryos from tube to tube (for bigger embryos, cut the tip of the pipet). When changing solutions, always leave some liquid in the tube to avoid flattening of embryos.


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For washings, the tubes are placed horizontally on a rocking platform. Washings are performed at room temperature, unless stated otherwise. Hightemperature incubations are performed in a hybridization oven.

2. Hybridization Protocol a. Take embryos from 70% EtOH, transfer to PBT.3 (PBS with 0.3% Triton X-100), and wash, 3 X 5 min. b. Treat with Proteinase K (30 pg/ml in PBT.3) at 37%C. With 8.5-9.0 dpc embryos incubate for 10 min, with 9.5 dpc embryos incubate for 15 min, and with 10.5-12.0 dpc embryos incubate for 30 min. c. Rinse in freshly prepared 0.2% glycine in PBT.3 for 5 min. d. Rinse in PBT.3 3 X 5 min. e. Transfer embryos into 2-ml polyethylene tubes with screw caps and fill each tube with a mixture (1 :1)of PBT with hybridization buffer. Incubate for 10 min. f. Replace with hybridization buffer and prehybridize by rotating the tubes in a hybridization oven for 2-3 hr at 65°C. g. Replace with preheated (70"C, 15 min) hybridization buffer containing the probe (250 ng/ml) and hybridize in a hybridization oven at 65°C for 24 hr. h. Remove the hybridization solution (do not discard the hybridization solution; it can be stored at -20°C and reused at least five additional times) and rinse with hybridization buffer without dextran sulfate: once briefly, once for 10 min, once for 20 min at 65°C in a hybridization oven, and once overnight at 65°C in a hybridization oven. i. Transfer embryos into 15-ml tubes, rinse with shaking in PBT.3 3 X 30 min at room temperature, and proceed to the immunodetection step.

3. Immunodetection All steps of the immunodetection are performed at room temperature on a rocking platform. a. Rinse with PBT.3 containing 2% blocking reagent (Boehringer-Mannheim) for 1hr. The 10% stock solution of the blocking reagent is prepared according to manufacturer's protocol, treated with DEPC, autoclaved,and stored at -20°C. b. Absorb the anti-dig antibodies with heat-inactivated mouse-liver acetone powder. To prepare 4 ml of absorbed conjugate (1:2OOO in PBT.3 with 2% blocking reagent), weigh into an Eppendorf tube 4 mg acetone liver powder, add 0.5 ml PBT.3, and heat at 70°C for 30 min. Cool on ice, add 2 p1 of antidigoxigenin antibodies alkaline phosphatase conjugate (Boehringer-Mannheim), and incubate with shaking on ice for 1 hr. Centrifuge at 10,OOOg for 15 min at +4"C. To the supernatant add 800 p 10% blocking reagent and bring to 4 ml with PBT.3.

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c. Incubate embryos with preabsorbed antibodies for 2.5 hr in 2-ml tubes. d. Rinse briefly with PBT.3 and transfer embryos into 15-ml tubes. Rinse 2 X 5 min with PBT.3. e. Rinse with PBT.5 (PBS with 0.5% Triton X-100) for 3 X 20 min. f. Rinse APB.3 (APB with 0.3% Triton X-100) for 15 min. g. Incubate overnight with APB.3. h. Place embryos in 75 X 10 mm glass test tubes, fill the tubes with APB containing 10% poly(viny1alcohol) (APBPVA), and incubate for 30 min. [APBI PVA is prepared by dissolving poly(viny1 alcohol) (MW 30,000-7O,OOO, Sigma P-8136) in APB in a boiling water bath. i. Replace the APBPVA solution with alkaline phosphatase substrate solution, prepared by adding 4.5 pl NBT and 3.5 p BCIP per ml of APBPVA. Incubate in the dark with periodic inspection under the stereomicroscope until the desired intensity of staining is achieved. The rate of color development varies from 1-2 to 24 hr. To avoid overstaining, overnight incubations can be performed at +4"C. To stop the reaction, wash twice in TE. Hybridized embryos can be stored in TE at +4"C in the dark. Photographs of early embryos (up to 9.5 dpc) are taken with a photomicroscope equipped with low-power (2X-4X) objectives (Fig. 2D). If an embryo is too transparent, the staining on the other side of the body is revealed as well and reduces the quality of the pictures. To make embryos less transparent, dehydrate partially in 70% EtOH. Alternatively, embryos can be fixed in 10% formalin.

V. Analysis of Expression at the Protein Level A. CAT Assay

One of the most popular reporter genes for the characterization of control elements in transgenic mice is the bacterial chloramphenicol acetyl transferase (CAT) gene. The qualitative CAT assay is performed as follows: 1. Homogenize tissue samples in 1-5 m10.25 M Tris-HC1, pH 7.8, using Polytron homogenizer. 2. Heat the homogenate for 10 min at 65°C. Centrifuge, collect the supernatant, and store at -70°C. 3. Determine protein concentration and adjust to 20-100 ug, in a final volume of 60 pl. 4. Prepare a mixture containing 20 p1 20mM acetyl coenzyme A, 0.2 pCi 14C-chloramphenicol,and 5 p1 20 mM Tris-HC1, pH 7.8. Adjust the volume to 40 p1 with H 2 0 and add it to the protein sample.


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5. Incubate at 37°C for 1 hr. Stop the reaction with 1 ml ethyl acetate and vortex vigorously. Centrifuge, collect the supernatant, and dry in a Speedvac. 6. Dissolve in 20 pl ethyl acetate, load on a thin-layer chromatography (TLC) plate, and chromatograph with chlorofomdmethanol (955). Dry the plate and autoradiograph.

The quantitative CAT assay is done according to Sleigh (1986): 1. To the protein extract (54 pl) add 20 pl 8 mM chloramphenicol, 5.7 pl 2.0 M Tris, pH 7.8, and 20 p10.1 pCi 14C-acetylcoenzyme A in 0.5 mM nonradioactive acetyl coenzyme A. 2. Incubate for 1 hr at 37°C. 3. Extract with 200 pl ethyl acetate and remove 160 p1 of the supernatant to a minivial containing 2 ml xylene/xylofluor (9 :1). 4. Reextract the sample with 200 pl ethyl acetate and add 200 p1 of the supernatant to the minivial. 5. Read radioactivity in a scintillation counter, 6. A standard curve is made with a commercial CAT enzyme (P. L. Biochem) at a range of 0.002-0.08 unitslreaction.

B. LacZ Stab

‘g

The bacterial LacZ gene, encoding bacterial /3-galactosidase, provides an ideal in situ visual marker for the analysis of regulatory elements in the developing mouse embryoand to follow the spatiotemporalpattern of lacZ in whole embryos. It is also extensively used as a vector for promoter or enhancer traps. The main advantage of lacZ reporter genes is the high sensitivity of detection at the singlecell level (Fig. 3). 1. Dissect embryos in PBS and transfer them into ice-cold freshly prepared 4% paraformaldehyde in PBS. The volume of fixative must exceed 20-30 times the total volume of fixed material. For better penetration of fixative and staining solutions into internal parts of the body, embryos older than 12-13 dpc must be “opened”: make a longitudinal cut in the midline of the abdomen or cut the embryo into two halves along the body axis. In embryos older than 16 dpc the skin should be removed. 2. Incubate for 2-24 hrs at 4°C. 3. Wash embryos in PBS at room temperature (at least X 3) for 30-60 min. The volume of PBS must exceed 20-30 times the total volume of fixed material. 4. Incubate while rocking in staining solution at 37°C for the first 2-3 hr and then at 30°C overnight. Staining solution contains 0.1% X-gal (a 4% stock solution is prepared in dimethylformamide),2 mM MgC12,0.01% (w/v) sodium desoxycholate, 0.02% (vlv) NP-40, 5 mM K3Fe(CN)6, 5 m M We(CN),, and 10% X 10 PBS. The solution is passed through a 0.22-pm filter prior to staining.

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Fig. 3 Expression of the bacterial lacZ reporter gene. Histochemical detection of lacZ activity in transgenic mouse embryo (10.5 dpc) carrying the reporter gene driven by the regulatory elements of the quail myoD gene. (A) Whole-mount stained embryo. The staining is concentrated mainly in the somites. (B) A sagittal section through the cervical somites of the same embryo showing staining in myotomal cells.

5. Wash embryos in several changes of tap water at room temperature, until the yellow staining is removed (10 min to several hours, depending on the size of the embryos). 6. Place the embryos in 50% EtOH (15 min to 1hr, depending on the size of the embryo) and finally in 70%EtOH. Stained embryos can be store in 70%EtOH. To make embryos more transparent, further dehydrate by passing them through the following reagents: 70%, 80%,95%, 3 X loo%,1:1mixture of 100% EtOH with isobutyl alcohol, pure isobutyl alcohol, 1:1mixture of isobutyl alcohol with mineral oil, and pure mineral oil. The duration of each step vanes according to the size of the embryo: from 15 min (embryos prior to 9.5 dpc) to 1 hr (at later stages). The procedure can be stopped at any EtOH stage. However, too long contact of stained embryos with isobutyl alcohol should be avoided.

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7. For embedding into paraffin, transfer embryos into melted paraplast at 56-60°C and impregnate in three changes of melted paraplast during 1.5-3 hr. Orient the embryos in the embedding tray, remove the tray from the oven, and let it cool down. 8. Paraffin sections can be counterstained with nuclear fast red and mounted with any permanent mountant. However, to avoid stain dissolution care must be taken to minimize the time of exposure to xylene.

VI. Future Directions It appears that transgenic technology by microinjection into mouse fertilized eggs contributed to a number of questions related to myogenesis. Some of these topics are indicated in the Introduction. However, one direction for improvement is to enlarge the size of introduced DNA into the mouse genome. For example, the control elements of the myogenic regulatory gene myf5, the first of the myoD gene family to be expressed during somite formation and differentiation, are not yet known. This gene is closely linked to the MRF4 on human chromosome 12. Attempts to express genomic myf5 gene in transgenic mice have failed to recapitulate the embryonic pattern of expression (Patapoutian et af., 1993). Perhaps control elements of this gene are located further upstream or downstream. The ability to introduce very large DNA fragments, either cosmids or yeast artificial chromosomes, into the mouse genome would permit addressing this question. Another question of interest is whether each of the myogenic regulatory genes are expressed in distinct cell populations during myotome formation. This can be addressed by studying the consequences of expression of toxigens driven by specificelements of these genes or by specificactivation of toxigenes in specific myogenic cells types using the cre-lox recombination approach (Sauer, 1993). It is assumed that members of the MyoD family are functionally identical. The multiplicity of genes serves to fine-tune their expression during embryonic and adult stages. To address this question, MyoD can be activated by myogenin regulatory elements and vice versa in wild-type or in null mutants. Finally, gene trap is one of the most exciting approaches to identify novel genes involved in the establishmentof pattern formation and specificcell lineages. Although it is a blind test, many interesting patterns leading to the identification of novel genes have been reported (for a review, see Skarnes, 1993). We have recently identified a transgenic mouse strain in which the lacZ reporter gene is driven by putative regulatory DNA sequences of an endogenous gene expressed specifically in myogenic cells that migrate from the ventrolateral edge of the somite to populate the limb buds. The spatio-temporal pattern of expression of this gene is distinct from that of another gene (Pax3) expressed in these cells (Williams and Ordahl, 1994). Other$genesimportant in the specification of the myogenic cell lineages and in patterning skeletal muscles would likely be identified using this approach.

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References Asante, E. A., Bosswell, J. M., Burt, D. W., and Bulfield, G. (1994). Tissue-specific expression of an a-skeletal actin-lacZ fusion gene during development in transgenic mice. Transgenic Res. 3,59-66. Auffray, C., and Rougeon, F. (1980). Purification of immunoglobulin heavy chain mRNA from total myeloma tumor RNA. Eur. J. Boichem. 107,303-313. Barth, J., and Ivarie, R. (1994). Polyvinyl alcohol enhances detection of low abundance transcripts in early stage quail embryos in a nonradioactive whole mount in situ hybridization technique. Biotechniques 17, 324-327. Banejee-Basu, S., and Buonanno, A. (1993). cis-Acting sequence of the rat troponin I slow gene confer tissue- and development-specifictranscription in cultured muscle cells as well as fiber type specificity in transgenic mice. Mol. Cell Biol. 13, 7019-7028. Beermann, F., Ruppert, S., and Schutz, G. (1991). Tyrosinase as a marker for transgenic mice. Nucl. Acids Res. 19, 958. Buonanno, A., Edmondson, D. G., and Hayes, W. P. (1993). Upstream sequences of the myogenin gene convey responsiveness to skeletal muscle denervation in transgenic mice. NUC.Acids Res. 21,5684-5693. Brosnan, M. J., Raman, S. P., Chen, L., and Koretsky, A. P. (1993). Altering creatine isoenzymes in transgenic mouse muscle by overexpression of the B subunit. Am. J. Physiol. 363, c151-cl60. Chatot, C. L., Ziomek, C. A., Bavister, B. D., Lewis, J. L., and Rorres, I. (1989). An improved medium support development of random-bred 1-cell mouse embryos in vitro. J. Reprod. Fert. 86,679-688. Cheng, T.-C., Hanley, T. A., Mudd, J., Merlie, J. P., and Olson, E. N. (1992). Mapping of myogenin transcription during embryogenesis using transgenes linked to myogenin control region. J. Cell Biol. 119,1649-1656. Cheng, T.-C., Wallace, M. C., Merlie, J. P., and Olson, E. N. (1993). Separable regulatory elements governing myogenin transcription in mouse embryogenesis. Science 261,215-218. Donoghue, M. J., Alvarez, J. D., Merlie, J. P., and Sanes, J. (1991). Fiber type- and position-dependent expression of a myosin light chain-chloramphenicol acetyltransferase (CAT) transgene detected with a novel histochemical stain for CAT. J. Cell Biol. 115,423-434. Donoghue, M. J., Morris-Valero,R., Johnson, Y.R., Merlie, J. P., and Sanes, J. R. (1992). Mammalian muscle cells bear a cell-autonomousheritable memory of their rostrocaudal position. Cell 69,67-77. Faerman, A., Pearson-White, S., Emerson, C., and Shani, M. (1993). Ectopic expression of myoD in mice causes prenatal lethalities. Devel. Dynamics l96,165-173. Fujisawa-Sehara,A., Hanaoka, K., Hayasaka, M., Hiromasa-Yagami,T., and Nabeshima,Y-I. (1993). Upstream region of the myogenin gene confers transcriptional activation in muscle cell lineages during mouse embryogenesis. Biochem. Biophys. Res. Comm. 191,351-356. Goldhamer, D. J., Faerman, A., Shani, M., and Eemerson, C. P. (1992). Developmental regulation of the myogenic regulatory gene myoD by a distal enhancer. Science 256,538-542. Goldhamer, D. J., Brunk, B. P., Faerman, A., Shani, M., and Emerson, C. P. (1995). Embryonic activation of myoD by a highly conserved distal control element. Development 121,637-649. Greishammer, U., Sassoon, D., and Rosenthal, N. (1992). A transgene target for positional regulators marks early rostral specification of myogenic lineages. Cell 69,79-93. Gundersen, K., Sanes, J. R., and Merlie, J. P. (1993). Neural regulation of muscle acetylcholine receptor E- and a-subunit gene promoters in transgenic mice. J. Cell Biol. U3,1535-1544. Hallauer, P. L., Hastings, K. E., and Peterson, A. C. (1988). Fast skeletal muscle-specificexpression of a quail troponin I gene in transgenic mice. Moll. Cell Biol. 8,5072-5079. Hallauer, P. L., Bradshaw, H. L., and Hastings, K. E. (1993). Complex fiber-type specific expression of fast skeletal muscle tropanin I gene construct in transgenic mice. Development 119, 691-701. Johnson, J. E., Wold, B., and Hauschka, S. (1989). Muscle creatine kinase sequence elements regulating skeletal and cardiac muscle expression in transgenic mice. Mol. Cell Biol. 9,3393-3399.

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Matsumura, K., Lee, C. C., Caskey, T., and Campbell, K. P. (1993). Restoration of dystrophinassociated proteins in skeletal muscle of mdx mice transgenic for dystrophin gene. FEES Let 3209276-280. Miner, J. H., Jeffrey, B. M., and Wold, B. J. (1992). Skeletal muscle phenotypes initiated by ectopic myoD in transgenic mouse heart. Development 114,853-860. Patapoutian, A., Miner, J. H., Lyons, G. E., and Wold, B. (1993). Isolated sequences from the linked myf5 and MRF4 genes drive distinct pattern of muscle-specific expression in transgenic mice. Development 1%61-69. Petropoulos, C. J., Rosenberg, M. P., Jenkins, N. A., Copeland, N. G., and Hughes, S. H. (1989). The chicken skeletal muscle a-actin promoter is tissue specific in transgenic mice. MoL Cell Biol. 9,3785-3792. Pinney, D. F., Charles de al Browse, F.,Faerman, A., Shani, M., Maruyama, K., and Emerson, C. P. (1995). Quail MyoD is regulated by a complex array of cis-acting control sequences. Devel. Biol. 170,21-38. Polak, J. M.,and McGee, J. O’D. (1990). “In Situ Hybridization. Principles and Practice.” Oxford University Press. Rogers, A. W. (1979). “Techniques of Autoradiography,” 3d ed. ElsevierBIorth Holland Biomedical Press. Salminen, M., Maire, P., Concordet, J-P., Moch, C., Forteu, A., Kahn,A., and Dagelen, D. (1994). Fast-muscle specific expression of aldolase A transgenes. Mol. Cell Biol. 1%6797-6808. Santerre, R. F., Bales, K R., Janney, M. J., Hannon, K., Fisher, L. F.,Bailey, C. S.,Moms, J., Ivarie, R., and Smith, C. K. (1993). Expression of bovine myf5 induces ectopic skeletal muscle formation in transgenic mice. Mol. Cell BioL l3,6044-6051. Sassoon, D., and Rosenthal N. (1993). Detection of messenger RNA by in situ hybridization. In “Methods in Enzymology: Guide to Techniques in Mouse Development” (P. M. Wassarman and M. L. DePamphilis, eds.), Vol. 225. New York Academic Press. Sauer, B. (1993). Manipulation of transgenes by site-specificrecombination: Use of Cre recombination. I n “Methods in Enzymology: Guide to Techniquesin Mouse Development” (P. M. Wassarman and M. L. DePamphilis, eds.), Vol. 225, pp. 890-900. New York Academic Press. Schedl, A., Montoliu, G., Kelsey, G., and Shutz, G. (1993). A yeast artificial chromosome covering the tyrosinase gene confers copy number-dependent expression in transgenic mice. Nature 362, 258-261. Shani, M. (1985). Tissue-specific expression of rat myosin-light chain 2 gene in transgenic mice. Nature 314, 283-286. Shani, M. (1986). Tissue-specificand developmentallyregulated expression of a chimeric actin-globin gene in transgenic mice. MoL Cell BWL 6,2624-2631. Shani, M., Dekel, I., and Yoffe, 0. (1988). Expression of the rat myosin-light chain 2 gene in transgenic mice: Stage specificity, developmental regulation and interrelation with the endogenous gene. Mol. Cell Biol. 8,1006-1009. Skarnes, W. C. (1993). The identification of new genes: Gene trapping in transgenic mice. Curr. Op. Biotechnol. 4,684-689. Sleigh,M. J. (1986). A nonchromatographicassay for expressionof the chloramphenicolacetyltranferase gene in eucaryotic cells. Anal. Biochem. 156,251-256. Sutrave, P., Kelly, A. M., and Hughes, S. H. (1990). ski can cause selective growth of skeletal muscle in transgenic mice. Genes & DeveL 4,1462-1472. Takeo, M., Schroeder, A. C., Mobraaten, L. E., Gunning, K.B., Hanten, G., Fox, R. R., Roderick, T. H., Stewart, C. L., Lilly, F., Hanse, C. T., and Overbeek, P. A. (1991). FVBM: An inbred mouse strain preferable for transgenic analyses. Proc. Nutl. Acud. Sci. USA 88,2065-2069. Wilkinson, D. G. (1992). “In Siru Hybridization. A Practical Approach.” IRL Press at Oxford University Press. Wilkinson, D. G., and Green J. (1990). I n situ hybridization and the three-dimensional reconstruction of serial sections. In: “Postimplantation Mammalian Embryos. A Practical Approach” (A. J. Copp and D. L. Cockroft, eds.), pp. 155-172. IFU Press at Oxford University Press.

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Wilkinson, D. G., and Nieto, M. A. (1993).Detection of messenger RNA by in siru hybridization to tissue sections and whole mounts. I n “Methods in Enzymology: Guide to Techniques in Mouse Development” (P. M. Wassarman and M. L. DePamphilis, eds.), Vol. 225. New York Academic Press. Williams, B. A., and Ordahl, C. P. (1994). Pax-3 expression in segmental mesoderm marks early stages in myogenic cell specification. Development 1u),785-796. Yee, S-P., and Rigby, W.J. (1993).The regulation of myogenin gene expression during the embryonic development of the mouse. Genes & Devel. 7,1277-1289.


CHAPTER 19

DNA Transfection of Cultured Muscle Cells Craig Neville,' Nadia Rosenthal,* and Stephen Hauschkat 'Cardiovascular Research Center Massachusetts General Hospital-East Charlestown, Massachusetts 02129 'Department of Biochemistry University of Washington Seattle, Washngton 98195

I. Introduction 11. Transient Transfections A. Calcium Phosphate Coprecipitation B. Alternative Transient Transfection Protocol for MM14 Cells C. Electroporation 111. Stable Transfections A. Standard Protocol for Stable Transfections B. Alternative Stable Transfection Protocol for MM14 Cells IV. Identification of &-Acting Control Regions with Reporter Assays A. Reporter Gene Assays V. Summary and Future Prospects References

I. Introduction The molecular basis of vertebrate skeletal myogenesis has been elucidated largely by the use of muscle-cell transfection. Myoblasts are fairly straightforward to isolate from embryonic and neonatal muscles (see accompanying chapter), and myogenic cultures are generally amenable to transfection with exogenous DNA. Following introduction of vector DNA into myoblasts by transfection, expression of exogenous genes generally can be measured for up to 72 hr. During this time the transfected plasmids remain as episomal, nonreplicating METHODS IN CELL BIOLOGY, VOL. 52

Copytighr 0 1998 by Academic Press. AU nghU of reproduction in any form reserved. 00YI-h7YX/Y8 125.00

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minichromosomes that are gradually degraded. Assays of gene expression from these transient DNA structures are most useful when rapid testing of numerous constructs is required, as multiple transient transfections can be harvested in parallel. If continuous monitoring of gene expression is required, or if the effects of genes expressed for longer than 72 hr is desired, myoblast clones can be generated in which the expression vector is stably integrated into the host genome. A selectable gene, usually a drug-resistancemarker, is cotransfected into myoblasts together with an excess of the expression vector under study. The cells are grown in the presence of the appropriate drug, producing a drug-resistant subpopulation in which both selected and nonselected genes are integrated into the genome. Stable transformation assays suffer from a number of confounding factors, however. The most serious drawback is that the assay is subject to long-term effects and uncontrolled parameters, such as the influence of variable integration sites of the transfected genes into the host-cell genome. For this reason, gene expression is often measured in a mass culture of stable transformants. Individual clones, corresponding to separate integration events, can also be propagated as myoblasts, analyzed for the number of integrated numbers of test gene copies, and assayed for reporter gene expression in proliferating or differentiated clonal cultures. The popularity of musclecell culture systems for the analysis of cloned gene expression is due principally to the standard techniques involved and to the reproducibility of the assays. Although, in general, basal promoter function has been successfully reconstituted in cell-free systems, distal regulatory elements such as enhancers are still refractory to study in v i m . Introduction of transcription units into the germ line of transgenic mice is still the ultimate test of correct spatial and temporal control of expression, since the introduced genes are subject to cell-specific signals during embryonic development. With the appropriate muscle-cell system in hand, however, much of the preliminary mapping can be completed prior to transgenesis, saying valuable time and resources. Muscle-cell transfection also affords a fine-scale mutational analysis of regulatory elements that would be prohibitive in mice. These considerationsnotwithstanding, extrapolations from muscle culture transfection data to the probable behavior of the same gene in mice should be viewed with caution. Several examples of unanticipated differences have been noted between cell culture and transgenic studies of the mouse M-creatine kinase gene and rat myosin light chain 113 locus.

11. Transient Transfections Following introduction of expression vector DNA into myoblast cell cultures, transient expression of the exogenous genes can usually be measured for up to 72 hr posttransfection. The virtues of transient assays include the convenience of the short time between transfection and harvest. This is ideal for the design

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of expression studies such as deletion or mutation analysis of a DNA regulatory sequence, requiring sequential testing of many constructs. For transient assays, DNA transfection in the presence of facilitating agents such as calcium phosphate is most commonly used, since chemical methods are best suited to adherent cells such as myoblasts. Electroporation,the introduction of DNA molecules by a transient electric impulse passed through suspended myoblast cultures, is also used; it requires no special buffers and provides high transfection efficiency with little variation between samples. Protocols for both procedures are described. In any transfection procedure, efficiencies of vector delivery and expression can vary as a function of myoblast density and condition, which is particularly important for transient assays where multiple DNA constructs are compared. Especially when using rapidly proliferating cells, such as primary cultures or the cell line, it is important to transfect at subconfluent densities to ensure that during subsequent proliferation the cells are not induced to differentiate by reaching confluence. DNA purity can also affect efficiency of vector delivery. For example, RNA contamination significantly reduces the transfection efficiency of the calcium phosphate coprecipitation technique. DNA conformation is another variable. Supercoiled plasmid is more efficiently transfected than linear forms. In addition, it is more resistant to nucleases. On the other hand, if the introduced DNA needs to integrate into a chromosome of the host cell, this is normally done with linearized DNA. A cotransfected transcription unit including a reporter gene driven by a strong regulatory element is a valuable addition to the experimental design, as it can be used to correct for small fluctuations in transfection efficiency, and it alleviates the need for multiple experiments to achieve a statistically significant result. A. Calcium Phosphate Coprecipitation

Calcium-mediatedDNA transfection is a standard method based on the ability of cultured cells to take up DNA that has been complexed with a calcium phosphate coprecipitate. It is appropriate for both transient and stable transfection assays and is popular because in general it has minimal adverse effects on cell viability, and it does not require expensive equipment. Included is a model procedure for the transfection of DNA using calcium phosphate coprecipitate in myoblast cultures. The protocols are modified versions of published techniques (Graham and van der Eb, 1973). 1 . Cells

Approximately 5 X 105 myoblasts are seeded on a 100-mm culture dish 20 hr before transfection. The cells are refed with fresh media 4 hr before transfection.

2. DNA The amount of test plasmid to be transfected is empirically determined and depends to some extent on the relative activity of the regulatory elements driving

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the gene to be expressed. For example, as little as 1 pg of a plasmid carrying a strong muscle-specific regulatory element, such as the myosin light-chain enhancer, or a ubiquitous regulatory element, such as the SV40 enhancer, can yield consistently high expression of the linked gene (Donoghue et aL, 1988). If the strength of a putative regulatory sequence is unknown, it is advisable to start with a maximal amount of test plasmid (10-25 pg per 100-mm dish) and subsequently reduce that amount, always keeping the total amount of transfected DNA constant with a neutral filler plasmid. The filler DNA should be prepared in the same manner as the test DNA. This ensures uniformity in the quality of the calcium phosphate coprecipitate, as it is sensitive to both contaminants and total quantity of DNA. For optimal transfection efficiency with this method, the plasmid DNA should be protein- and RNA-free, and in supercoiledform. Plasmid preparations of consistently high quality can be reliably achieved by two successive CsCl gradient purifications, although certain DNA purification columns currently on the market also reportedly yield consistent results.

3. Precipitate Preparation a. Plasmid DNAs (up to 25 p g total per 100-mm dish of subconfluent myoblasts) are mixed with a fresh, sterile-filtered CaClzsolution to a final concentration of 124 mM in a total volume of 0.5 ml. b. A 0.5-ml solution of sterile filtered 2X HBS (Hepes buffered saline: 50 mM Hepes, 280 mM NaC1, and 1.5 mM Na2P04, pH 7.1) is placed in a separate tube. The pH of the HBS solution is critical for the proper formation of the coprecipitate. It drops over time and should be adjusted before each use. c. The DNA-calcium solution is added dropwise to the HBS solution, which is either continuously vortexed or gently bubbled with a stream of N2 through a l-ml sterile pipet. The solution should become slightly cloudy and opalescent. d. After 30 min at room temperature the DNA-calcium phosphate coprecipitate is applied dropwise to the media covering the myoblasts. If the same DNA precipitation is aliquotted for multiple dishes, a master mix of the precipitate can be prepared, since the formation of the precipitate is not volume dependent. Depending on the muscle cell type, the level of DNA uptake may be increased by leaving the precipitate on the cultures for up to 24 hr. Since the overall viability of the culture can be affected by longer incubation times, optimal conditions for DNA uptake vary for each cell type. e. DNA uptake may also be significantly increased by a glycerol shock 412 hr posttransfection. The glycerol shock solution contains 15% glycerol in 1X HBS and should be sterile filtered. To shock the cells, media containing the DNA precipitateis removed and the cell monolayeris washed once with serum-free medium, then 3 mlglycerol shock solution (room temperature) is added directly to the monolayer. After exactly 3 min the glycerol is removed and replaced by 10 ml serum-free medium, followed by a second wash. The ceUs are then incubated for


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24-48 hr in growth or differentiation medium before harvesting for transient assays, or before applying selective media for long-term transformation. B. Alternative Transient Ttansfection Protocol for M M 1 4 Cells MM14 cells are relatively easy to transfect and their transient expression of test gene constructs is quite reproducible. When transfected with a plasmid constitutively expressing b-galactosidase plasmid according to the protocol described next, MM14 cultures consistently exhibit 5 1 0 % b-galactosidase-positive cells. See protocol for growth and differentiation of MM14 cells in the accompanying chapter. For additional details of the transfection protocol, see Amacher et al. (1993). A harvest protocol may be found in Buskin et al. (1996). The typical transient transfection protocol is as follows: 1. Inoculate 100-mm dishes with 2 X lo5 log phase cells, and feed with a complete GM plus FGF change at about 12 hr. (Transfections are usually carried out 24-28 hr after inoculating the plates with cells. The cultures have then expanded to about 5 X lo5 cells/plate.) 2. About 4 hr prior to the transfection, aspirate and refeed the cultures with 5 ml complete GM plus 2 ng/ml bFGF. 3. Individual culture dishes are transfected by adding CaP04-precipitated DNA directly to the GM. (Typical DNA additions contain 10 pg precipitated DNA in a 500-pl volume. The DNA consists of 8 pg test plasmid and 2 pg of a reference plasmid.) After dispersal of the DNA precipitate, the culture dishes are reincubated. 4. Four hours later, aspirate GM and rinse once with 5 ml Puck’s saline G (room temperature) by rocking gently for 5 sec. 5. Add 3 mll5% glycerol in HBS (room temperature). This should be left on each plate for exactly 2 min; thus, aspiration of the glycerol should begin at 1 min, 45 sec. When handling large numbers of dishes it is best to carry out this step with two persons so that glycerol is not left on any of the dishes for more than 2 min. Longer glycerol treatments lead to cell lysis. 6. Add 5 ml saline G (room temperature) exactly 2 min after adding the glycerol and rock the plate gently for about 5 sec. 7. Aspirate the saline G and add 10 ml DM (37°C) plus insulin for the analysis of gene expression in differentiated cultures, or add GM plus FGF for the analysis of gene expression in proliferating cultures. In the latter case, cultures should be refed with GM plus FGF about every 12 hr. 8. Transient transfection cultures are typically harvested 24-36 hr after the glycerol shock step. C. Electropotation

Electroporation, the use of a high-voltagepulse to introduce exogenous nucleic acids into a cell, has grown greatly in popularity in the past decade. Electropora-

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tion is considerably easier than alternative methods and gives more consistant results. Because the technique is physical rather than biochemical, it may cause less perturbation in the surviving recipient cells than alternatives such as calcium phosphate. Electroporation takes advantage of the fact that the cell can act as an electrical capacitor. The membrane normally acts a barrier and is unable to pass a current, except through ion channels. When the cell is exposed to a highvoltage electrical current, however, its membrane temporarily breaks down and forms pores that allow macromolecules (including DNA, oligonucleotides, and protein) to enter. The holes reseal spontaneously at room temperature, but placing the cells on ice delays the process. The DNA introduced to cells then translocates from the cytoplasm into the nucleus. Electroporation of cells requires a capacitor discharge device. Commercial electroporation devices are available from BioRad, BTX, IBI, Invitrogen, and Life Sciences. Although the procedure has been successfully adapted for animal, bacterial, plant, and yeast cells, optimal conditions should be determined for each cell type studied. Varying transfection buffer and the voltage, capacitance, and duration of the pulse can all greatly affect the efficiency of DNA uptake. Exact conditions for each cell type should be determined for each individual machine, as the pulses they deliver vary considerably. A reporter gene that is easy to assay, such as secreted alkaline phosphatase (SEAP, available from Tropix or Clontech) (Berger et al., 1988) or human growth hormone (Allegro) (Selden et al., 1986), both of which are secreted and whose expression can be measured in a sample of medium, or P-galactosidase (Alam and Cook, 1990) should be used to determine transfection efficiency. Electroporation is normally camed out with a relatively low capacitance and a high voltage setting. Some cell types have poor viability under these conditions and respond better to high-capacitanceflow-voltagesettings (Chu et aL, 1987). The commonly used muscle-cell line CZ falls in the latter category. When electroporating a new cell type for the first time, it is important to establish whether it tolerates high voltage or high conductance better. This can usually be accomplished with as few as two electroporation conditions. A capacitance of 25 pF and voltage of 1200 V is used to test the standard high-voltage conditions, and 960-1080 pF and 250 V for high-capacitance conditions. PBS or Hepes-buffered Saline are most often used as electroporation buffers, but it has been reported that the use of phosphate-bufferedsucrose as an electroporation buffer can be beneficial for sensitive cells because it allows for lower electroporation voltages (Chu et al., 1987). In addition, decreasing the sodium chloride concentration by 20 mM and replacing it with 20 mM Hepes, pH 7.5, to improve the buffering capacity of PBS during electroporationdecreases pH changes due to electrolysis at the electrodes of the cuvette, often improving cell viability (Potter, 1987). Electroporation requires more cells than other methods of inducing DNA uptake, as it causes considerable cell death. Optimal conditions normally involve a survival rate of 30-60%. For this reason myoblasts such as CZ are initially grown on 150-mm tissue-culture plates, to a density of no more than 60-708


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confluence. Typically, one plate per transfection is required. The mixture will be somewhat viscous after electroporation because of the dead cells. One-hundredmillimeter plates are used for replating to ensure that when the myoblasts are shifted to differentiation medium, they will be at a sufficient density to fuse efficiently. In many cases, expression levels will be high enough to allow use of one-half of a 100-mmplate of cells and replating in a 35-mm plate. For secreted reporter proteins, even the 10-mm well of a 24-well plate contains sufficient medium to measure gene activity. The growth medium should be replaced with medium containing 2% horse serum at about 12 hr after transfection to remove dead cells resulting from the transfection process and to initiate differentiation. For secreted reported proteins, it is important that exactly the same amount of medium be added to each plate. C2cells are maximally differentiated 2 to 3 days later and are harvested. 1. Remove cells with PBS and 1mM EDTA (use trypsin if necessary, adding an equal volume of calf serum afterwards to inactivate). Pool in a 50-ml conical tube. 2. Pellet cells at 500g for 10 min. Gently resuspended in ice-cold electroporation buffer. 3. Repellet cells bring up once again in a volume of cold electroporation buffer corresponding to 1 ml per initial 150-mm plate. 4. Add DNA to be transfected (10-40 pg, 25 pg is commonly used) to each cuvet, followed by 0.9 ml of cells. Incubate cells for 5 min at room temperature. 5. Mix cells by flickingthe cuvet on the bottom (without touching the metal surfaces). 6. Electroporate according to instructions of the apparatus’ manufacturer. 7. Place cuvet on ice for 10 minutes. Plate cells on 100-mm tissue culture plate. Rinse cuvet with additional medium and add to the plate.

Once general electroporation conditions have been established, they should be optimized. A nonlimiting pulse length setting of 1000 msec is standard and not normally varied. The efficiency of DNA uptake is very sensitive to small changes in field strength, so an electroporation profile should be performed. As a 25 V/cm change can cause the optimum to be missed entirely (Chu et al., 1987), steps of 10 V for standard cuvets with a 0.4-cm gap are required. This mandates many measurements; hence the importance of an easily assayed reporter system. Voltage ranges of 1OOO-1400 and 100-350 V for high-voltage and highconductance conditions, respectively, are normally assayed for optimal transfection. A new electroporation protocol defines universal conditions that, when tested on 19 different cell lines, reportedly gave a minimum 20% transfection efficiency in survivingcells without optimization (Baum et al., 1994). With voltage optimization, between 20-100% of survivingcells expressed exogenous DNA. This simple protocol works quite well for the muscle cell line G. Optimum voltage for these cells at 1080 pF is between 260 and 280 V. Briefly, cells are harvested and


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resuspended in complete growth medium (with serum) to a density of 2-5 X lo6 cells/ml at room temperature. DNA and 400 p1 of cells are mixed in an electroporation cuvet and pulsed at 1050pF and 260 V. Optimization, if needed, is done at 20-V intervals between 220 and 300 V. Cells are immediately plated in complete medium. Cell number had little effect on efficiency,and DNA uptake was fairly linear with DNA ranges from 1 to 100 pg, with some additional cell death reported at the higher end. Several growth media (DMEM, IMDM, RPMI, Ham's F10) proved suitable, provided they were not of high ionic strength. The suitability of this protocol to muscle cell lines other than C, cells is not known. Some may require experimentation with the precise conditions as described earlier. Phosphate-buffered saline (PBS, without Ca2+or Mg2+): 8.0 g NaCl 0.2 g KCl 0.2 g KHzP04 2.16 g Na2HP04 HzO to 1 liter, pH to 7.3 Hepes-bufferedsaline (HBS): 4.76 g Hepes 8.0 g NaCl 0.4 g KCI 0.18 g NazHPO4 1.08 g glucose H20 to 1 liter, pH to 7.05 Phosphate-buffered sucrose: 93.1 g sucrose 0.2 g MgCl2 1.6 g K2HPO4 HzO to 1 liter, pH to 7.4 with phosphoric acid

111. Stable Transfections If continuous monitoring of gene transcription is required to measure induction or modulation of expression during a period longer than the 72-hr timespan of transient assays, the vectors must first be integrated into the host-cell genome. In many instances, a cell line with a permanently integrated exogenous gene is highly desirable. It allows one to know the exact copy number of the exogenous genes, as well as eliminating the transfection efficiency variations inherent in transient assays. It does, however, entail some initial time and effort. A drugresistant gene whose expression can be selected for is simultaneously cotransfected with the gene of interest or included on the expression vector. Cells that have taken up the selectable marker become drug-resistant. Surviving cells normally have incorporated multiple copies of the nonselected gene and give rise to a poplation of stable transformants.


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The calcium phosphate-mediated transfection procedure causes larger amounts of calcium phosphate coprecipitated DNA to be taken up by fewer recipient cells than electroporation (Potter, 1987). Once within the cell, the DNA is integrated into the genome as long tandem arrays, with up to several hundred copies of the transgene in a single location. This can often lead to aberrant results when examining ectopic expression of a gene product, or the role of cis-regulatory regions in transcription. Electroporation conditions can be easily adjusted to allow for a low copy number per recipient cell (Potter et al., 1984). In addition, electroporated DNA does not integrate in long tandem arrays, instead integrating at multiple random sites throughout the genome. The general protocols for stable transfections are similar to those for transient transfections, although the number of cells and amount of DNA can be scaled down if either are limiting. Conditions found to be optimal for transient transfections can be used for stable transfections. There are several systems available for generating stably transformed mammalian cells. The most popular one utilizes neomycin resistance. Others include eco-gpt and hygromycin. The puromycin resistance gene (Morgenstern and Land, 1990) is ideal for muscle cell cultures. The selection process is quick, inexpensive, and effective. A large fraction of the nonresistant cells die within 24 hr of application of puromycin, and the vast majority are dead in 2 days. The cost of the selection drug puromycin is quite low, and in addition it is used in minute quantities. Spontaneously resistant colonies, a problem frequently seen with other selection systems, are very rarely seen with puromycin. The plasmid pBabePuro (Morgenstern and Land, 1990), which encodes the puromycin resistance gene, is driven by the SV40 early promoter. Another puromycin selection plasmid, pPUR, is available from Clontech.

A. Standard Protocol for Stable Transfections 1. Prepare plasmid DNA by the suggested method for the transfection procedure to be used. Digest the DNA with a restriction endonuclease that does not cleave within the expression cassette. Do the same with the vector providing selection. Uptake of linearized plasmid is perhaps two orders of magnitude less efficient than that for supercoiled DNA, but cleaving it before introducing it into cells greatly facilitates its integration into the genome in an expressible form. Mix the digested DNA containing the expression cassette with the selectable marker at a ratio of 25 :1 to 50 :1. Add sodium chloride to a final concentration of 300 mM. Extract the mixture with a combination of phenokhloroform, then with chloroform alone. Precipitate with ethanol, then rinse the resulting DNA pellet with 70% ethanol. Air-dry on the benchtop. Resuspend in a small volume of sterile water. Use between 1 and 10 pg for each transfection. 2. DNA can be transfected into the recipient cells by either calcium phosphate precipitation or electroporation. Fewer cells are needed for stable transfection, perhaps 5 X lo5 cells for calcium- and 5 X 106cells for electroporation-mediated transfection. In the case of electroporation, replate on 150-mm plates to ensure

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that resistant colonies are well separated. Splitting to as many as six plates may be necessary if transfection efficiencies are high. 3. Twenty-four hours after transfection, change the medium, maintaining the 20% fetal calf serum. The following day, after the cells have gone through at least two cell divisions following transfection, the selective drug is added. For puromycin, 1 p g / d (Sigma Cat. No. P-7255) is usually sufficient, although a range of 1 to 2.5 pg/ml is tolerated. Neomycin selection generally requires 500 pg/ml G418 for the initial selection, but after 2-3 weeks, a maintenance dose of 100-200 pg/d is used. Micophenolic acid at 1pg/mlg 250 pg/ml xanthine, and 15 pg/d hypoxanthine are used for eco-gpt selection. 4. After several days, individuallyresistant cells will have grown to form clonal colonies of 500 to lo00 cells. If colonies are to be pooled, the resistant cells can be trypsinized in situ and replated to form a mixed population. This mitigates much of the effect of uncontrollable parameters such as variable integration sites. It is more difticult, but sometimes essential to maintain and characterize each colony separately. This is particularly important if one requires a low or known copy number or where an aberrant integration site may be deleterious to the experiment. To isolate colonal stably transformed cells requires the use of glass cylinder cloning rings. Spread a thin layer of silicone grease on the inside of a 100-mm glass Petri dish. Arrange the cloning rings upright within the dish and tape the top of the dish on. Autoclave at 20 pounds for 20 minutes, dry heat, to sterilize. Mark on the bottom of the culture plate to indicate the colonies to be harvested. Aspirate the medium and rinse twice with PBS. With a pair of sterile forceps (dip in ethanol and flame), place cloning rings around each colony. Lift the cells off the plate by adding about 400 ~10.25% trypsin to each cylinder. Adding a few milliliters of medium to the plate will prevent the rest of the cells from drylng out. Gently replace the trypsin with medium when the cells start to loosen. Disperse the cells in the medium by gently pipetting up and down. Transfer the cells to a 24-well plate. Rinse the cylinder with additional medium and add to the first. Bring the medium level up to about 1 ml. When the cells grow to fill the plate, transfer them to a larger multiwell dish. B. Alternative Stable Transfection Protocol for MM14 Cells MM14 cells are also an easy and rapid experimental system in which to examine the expression of stably integrated plasmids. When transfected according to the following protocol, less than 0.2-0.5% of the initial cells produce stably transfected colonies following selection for neomycin resistance. The typical protocol for stably transfecting MM14 cells is as follows:

1. The initial steps are identical to those used for transient transfections, except that 100-mm plates are inoculated with 1 X 10s log-phase cells. (Use of a lower


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cell number is helpful with respect to the spatial separation of individual clones during the neomycin selection process. The lower cell density also decreases the percentage of spontaneously differentiating cells during the period in which the initial neomycin-resistant cells are being expanded.) 2. The CaP04-precipitated DNA for stable transfections contains 10 pg of the test plasmid plus 2 pg of the selectable plasmid (typically neomycin resistance). To judge when the selection period is complete it is useful to set up at least one culture that does not receive the selectable plasmid. When taken through the entire transfection and selection process, all cells in this culture should die. 3. Following the glycerol shock treatment, feed cultures with GM plus FGF and continue at 12-hr intervals. 4. Begin the selection for stably integrated plasmids about 18 hr post glycerol shock by adding 100 p1 of 50 mg/ml GeneticidG418 (prepared in distilled water) to each dish (final concentration of G418 in the culture medium is thus 500 pglml). 5. Continue feeding with complete changes of GM plus FGF plus G418 at 12-hr intervals. (When different plasmids are being analyzed, particular care should be taken not to risk cross-contaminating cultures by the inadvertent transfer of cells between dishes.) 6. Passage the entire dish or pick individual subclones after 4 days of G418 selection. (Although most selected clones may have only several hundred cells, they will begin to undergo spontaneous differentiation if they are not passaged and this will decrease the yield of stably selected cells.) 7. Replate passaged cells into G418-free GM plus FGF for the first 12 hr, then continue successive 12-hr feedings using GM plus FGF and G418. 8. Ascertain when the G418 selection period is complete by comparing cell numbers in the test cultures to those in the culture that received no neomycinresistance plasmid. Selection is typically complete by 7-10 days. If cell populations become greater than 5 X lvlplate during the selection process, the cultures should be passaged as described earlier. Expansion of stably selected cultures for experimental analysis and for preparing frozen stocks should always be done in the presence of G418, but improved yields will be obtained if G418 is omitted during the first 12 hr following each passage.

IV.Identification of &-Acting Control Regions with Reporter Assays The current popularity of skeletal muscle as an experimental system has resulted in the characterization of numerous muscle-specific regulatory DNA elements that respond to myogenic differentiation in cell culture by increasing transcription of their cognate genes. From comparative sequence analysis it is

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evident that many of these elements share common consensus sequences present upstream of the transcription start site. These include canonical elements associated with the basal transcription apparatus, such as the TATAA box, an ATrich sequence that sets the initiating nucleotide for transcription 25-30 bp downstream, and GC-rich oligomers or CCAAT motifs that constitute binding sites for other constitutive factors associated with transcription initiation. Numerous studies of both muscle and nonmuscle genes have demonstrated that the position of these promoter elements relative to the transcription start site of a gene is inflexible. Another class of regulatory DNA elements, called enhancers, differ in several ways from promoter-associated control sequences. First, the position of enhancers within a gene locus is unpredictable. Enhancers have been identified in muscle gene introns, and in regions several thousands of bases away from a gene. In functional assays, enhancers can dramatically increase transcription from a linked gene from positionsupstream or downstream,in either orientation relative to the direction of transcription. In practice, muscle-gene promoter and enhancer elements fall into two categories: those that are generally active in a wide variety of muscle and nonmuscle cells, and those whose optimal activity is restricted to the muscle lineage. Although no single element is shared by all muscle regulatory regions, several sequencemotifs in evolutionarily unrelated muscle genes have been characterized as binding sites for common nuclear protein complexes (Rosenthal, 1989). In cases where deletion or mutation of a motif impairs or obliterates the activation of linked gene expression in muscle cultures, many of these sequence motifs have been shown to play a critical role in the function of the regulatory element. In general, transcriptional regulation of muscle genes reflects the situation in constitutive transcriptional control elements, where multiple proteins bind to a complex and often overlapping array of recognition sequences. To assign roles to these regulatory elements, functional assays have been developed in which cloned sequences linked to a reporter gene can be reintroduced into muscle or nonmuscle cultures and tested for transcriptional activity. Although basal promoter function has been successfully analyzed in cell-free systems, attempts to reconstitute tissue-specific enhancer activity in vitro have been largely unsuccessful. The popularity of muscle cell culture systems for the analysis of gene expression derives principally from the relative ease with which myoblasts can be transfected and myogenic differentiation can be subsequently induced. Through modification of the sequence under study, the role of particular nucleotides in the activation of a linked reporter gene can be rapidly characterized in muscle cell culture. As a control for transfection efficiency, a second reporter construct whose transcription is not affected by myogenic differentiation can be included in the transfection. Numerous reporter systems are currently available that can be assayed biochemically or histologically. Although novel expression vectors are constantly being developed, they generally consist of a reporter transcription unit with


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convenient restriction enzyme sites for introduction of test DNA sequences either upstream of the transcription start site (for testing promoter activity) or downstream of the gene (for testing enhancer activity). In the latter case, basal promoter elements are included with the reporter coding region. When generating an expression vector in which promoter elements are to be tested, it is important to sequence the promoterheporter junction, since extra ATG translation initiation codons unintentionally introduced into the transcribed sequence during the cloning process can result in spurious, out-of-frame translation of nonsense peptides, to the exclusionof correct translation initiation of the reporter coding sequence. Several standard assay systems that have been successfully applied to the study of muscle-specific transcriptional regulation are now described.

A. Reporter Gene Assays Reporter gene activity in myotubes is normally assayed 2 days following replacement of growth medium with differentiation medium. Unintegrated DNA is not stable, causing reporter gene activity to decline rapidly after this period. This is not a consideration when measuring reporter activity derived from integrated genes arising from stable transfections, and maximal activity is normally found 3 days subsequent to switching to differentiation medium. Depending upon the reporter system used, either the cells are harvested and a crude wholecell extract is made from which activity is determined, or an aliquot of the medium is taken to determine the amount of reporter protein secreted. In addition to being easier, secreted reporter systems have the added advantages over cytosollocalized reporter systems that many fewer cells are needed and that the medium can be continuously sampled, allowing for a time course of reporter activity to be determined with a single transfection. p-Galactosidase traditionally has not been used as a reporter system to map &-regulatory elements of cellular genes, as some eukaryotic cell lines contain low levels of endogenous activity and the normal spectrophometric-based assay is not as sensitive as alternative methods for other reporters. However, Tropix (Bedford, Massachusetts) has recently developed a sensitive chemiluminescent-based assay that should be considered when designing new experimental protocols. Most commonly, a small amount of a plasmid containing a p-galactosidase or luciferase gene controlled by strong viral regulatory elements is cotransfected to control for various parameters of the assay (e.g., transfection and harvesting efficiency). It is also important always to include a mock transfection, using an equilavent amount of vector DNA, to determine endogenous reporter protein levels. If a p-galactosidase-expressingplasmid is used as a control for transfection efficiency, a whole-cell extract must be prepared as described later. Both pgalactosidase and luciferase are heat labile, so if the extract is to be used for determining CAT activity, an aliquot should be removed before heating to destroy endogenous acetyl transferases. A Bradford assay is preformed first to

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equilibrate protein concentrations, and therefore relative extract concentrations. If the subsequent P-galactosidase assays indicate small variations in transfection efficiencies,the extract concentrationscan be adjusted with buffer before continuing with reporter gene assays.

1. Whole-Cell Extract Preparation a. Remove media and rinse cells with three times with PBS. b. Add 1 ml TEN buffer to the cells and let sit for 5 min. c. Scrape loosened cells into a tube and centrifuge for 1 min. d. Aspirate supernatant and resuspend pellet in 150p10.25 MTris-HC1, pH 8.0. e. Freezekhaw in crushed dry ice/37"C water bath 4 times, vortexing between freezes. f. Centrifuge for 15 min at 4°C. Transfer supernatant to a new tube. TEN buffer: 40 mM Tris-HCI, 8.0 1 mM EDTA 150 rnM NaCl

2. Protein Concentration a. Dilute Bradford reagent (BioRad #500-OOO6) 1 to 4 with water. Aliquot 1 ml per disposable plastic cuvet. b. Add 5 pi cell extract and mix. Make a sample to blank with by adding 5 pl H20 to an additional cuvet containing diluted Bradford reagent. Wait 5 min. c. Read absorbance at 595 nm. A reading between 0.1 and 1.0 is considered to be in the linear range. If the reading is too high, dilute the extract appropriately and read another aliquot. 3. P-Galactosidase Assays a. Make fresh 1OOX Mg Buffer by adding 100 pl 1 M MgC12, 350 p1 @mercaptoethanol, and 550 pl H20. b. Make a master mix containing 300 pl of @-galmix per sample by adding 3.3 p11OOX Mg buffer, 73 p 1 l X ONPG, and 221 p10.1 M NaHP04/Na2P04. c. Aliquot 270 p1 @-galmix per microfuge tube. Add 30 pl of cell extract. (Be sure to include a control extract containing no @-galreporter gene, as some cell types contain considerable endogenous activity.) d. Incubate 1 hr at 37°C. (Positive samples will be slightly yellow.) e. Stop reaction with 500 pl 1M Na2C03.Read absorbance at 410 nm. 1X

ONPG

4 m g / d in 0.1 M NaHPO4/Na2PO4

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0.8 M NaHP0.,/Na2P04, pH 7.5 4.41 g NaHPO., 23.9 g Na2P04 Bring up to 250 ml with water.

4. Chloramphenicol Acetyl Transferase Assay The bacterial gene encoding chloramphenicol acetyltransferase (CAT) is a popular example of a sensitive reporter whose enzymatic activity can be easily assayed in crude cell extracts. With virtually no background activity in eukaryotic cells, the CAT assay is one of the most widely used methods of measuring transcriptional activity derived from regulatory sequences. CAT plasmid-based vector sets with convenient restriction sites upstream and downstream of the CAT transcription unit, and control vectors including basal promoters andor viral enhancers for positive and negative transfection controls, are commercially available from several sources (e.g., Promega). Disadvantages include the relative high cost per assay and the need to dispose of long-lived radioactive waste. Several CAT assays have been described. Readers performing assays using MM14 cells should consult Amacher et al. (1993). The following procedure, modified from Neumann et al. (1987) and Eastman (1987), involves significantly less effort than the more conventional methods to measure acetyltransferase activity. The acetyltransferase reaction takes place in the aqueous phase at the bottom of the vial. A nonradioactive acetyl group is transferred from a I4C-labeled acetyl coenzyme A to a chloramphenicol molecule. The still-labeled coenzyme A moiety involved in the reaction then partitions itself into the organic phase of the vial, allowing its subsequent radioactive decay to be detected by the scintillation counter. The assay involves periodically measuring the rate at which the I4Clabel enters the scintillation fluid. As with all CAT assays, it is run as a pseudofirst-order rate reaction. If accurate results are desired, it is essential that the reaction not be allowed to proceed so far that the substrate becomes rate-limiting. Such a situation is less likely with the assay described here, as it is continuously monitored rather than depending upon a single end-point value. If many reactions are run simultaneously, dilute the cell extracts sufficiently to ensure that the time involved in setup and scintillation counting do not contribute significant error to the relative values obtained. a. Heat the remainder of the cell extract at 65°C for 15 min. b. Centrifuge at 4°C for 10 min. Transfer supernatant to a new tube. The extract can be stored for a few weeks at -20°C. c. Make up a master mix containing the following per sample to be analyzed (include a negative control with Tris buffer and no extract in the assay): 27.5 pl 1 M Tris, pH 7.8, 55 pl 5 mM chloramphenicol (made up in 100% ethanol, 1.6 mg/ml, and stored at -2O"C), 11 p1 14C-acetyl coenzyme A (= 0.1 pCi, DupontNEN Cat. No. NEC3131), 126.5 pl H20.

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d. Aliquot 200 pl master mix into microfuge tubes. Add 50 pl normalized extract and mix well. e. When all extracts have been added to chloramphenicol mix, transfer each to the bottom of a 5 ml scintillation vial and carefully layer 5 ml nonaqueous scintillation fluid (e.g., Dupont Econofluor-2,Cat. No. NEF-969) on top. Incubate at room temperature. f. Samples should be counted immediately, and every 20 to 90 min thereafter, depending upon activity of the extract. Counts above 75,000 cpm are not considered to still be within the linear range of the assay. For extracts with relatively low concentrationsof CAT activity, the reaction can be allowed to proceed overnight.

5. The Luciferase Reporter System The gene encoding firefly luciferase (de Wet et aL, 1987) offers an additional reporter with many advantages over other reporter systems. The luciferase enzyme catalyzes the emission of light in the presence of the substrates luciferin and ATP. Light production is usually measured using an injection luminometer (e.g., Berthoid Lumat LB 9501). As the reaction produces only a brief spike of light, the luminometer is programmed to inject a luciferase substrate-containing buffer and detect activity (photon emission) immediately. Relative to a CAT assay, the luciferase assay is more sensitive, rapid, nonradioactive, inexpensive, and accurate over several orders of magnitude. Mammalian systems also produce almost no background activity. Cloningvectors similar to those for CAT reporters are widely available from Promega and others. The reagents described here are available as a kit from Promega (#E4030). A universal reporter lysis buffer is included that some investigators may find useful. a. The whole-cell extract described earlier can be used for luciferase assays. It should not be heat-treated, as the enzyme is labile. Although the luciferase enzyme is reportly sensitive to an extensive regime for producing whole-cell extracts, the procedure described here gives strong, reproducible activity. b. Dissolve Luciferase Activity Substrate into the supplied buffer. Mount vial in luminometer. Program to inject 200 pl substrate and to count photons for 5 sec. c. Add 30 pl of each cell extract to separate luminometer tubes. d. Insert tubes into luminometer to initiate reaction.

6. Secreted Alkaline Phosphatase Assay Human placental alkaline phosphatase (hAP) has been modified to produce secreted alkaline phosphatase (SEAP) (Berger et al., 1988). hAP is normally a membrane-associatedprotein. By deleting from the hAP gene the DNA encoding the carboxy-terminal24amino acids, a secreted alkaline phosphatase was created. Rather than measuring gene activity within a cellular extract, SEAP activity is


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simply measured in a small aliquot of medium, allowing the monitoring of gene expression over a period of time in a single culture. Unlike other alkaline phosphatases, hAP is heat stable. In addition to a 65°C incubation step, any endogenous alkaline phosphatases are further inactivated by the inclusion of levamisole and L-homoarginine in the assay, producing a system with low background. SEAP assays offer the most sensitive method of measuring reporter gene activity currently available; furthermore, they are also linear over several orders of magnitude, inexpensive, nonradioactive, and non-labor-intensive. Because of the sensitivity of detecting SEAP expression, small populations of transfected cells can be grown and assayed in individual 1 centimeter wells of a 24 multiwell dish. A chemiluminescent-based kit from Tropix (#BP300) is used to quantitate SEAP expression levels. A set of SEAP-containing vectors is available from them as well. a. Transfer 100 pl medium from the tissue-culture dish to a microfuge tube. It is important to have the same number of cells and volume of medium in cell cultures to be compared. b. Add 300 p1 1 X Phospha Dilution Buffer. Heat at 65°C for 15 min to inactivate endogenous alkaline phosphatases. c. Cool to room temperature. Transfer 100pl to a luminometer tube containing 100 pl of Phospha Assay Buffer. Mix and allow to incubate at room temperature for 5 min. d. Add 100 p11X Phospha Reaction Buffer to each luminometer tube. Allow the reaction to procede for 20 min. e. Measure luminescent activity with a luminometer (e.g., Berthoid Lumat LB 9501).

V. Summary and Future Prospects The recent explosion of information concerning the molecular pathways underlying the muscle cell phenotype has been possible largely by the relative ease with which exogenous DNA vectors can be expressed in muscle cell lines. As novel regulatory genes are isolated, their role in the determination and modulation of myogenesis will undoubtedly continue to be tested through muscle-cell transfection. New reporter systems will also broaden the spectrum of experimental designs available to the muscle cell biologist. For example, the green fluorescent protein that acts as a vital dye upon the absorption of blue light has been successfullyused as a reporter to mark living muscle cell cultures, in both transient and stable transfections (Moss et al., 1995). Inducible gene systems, such as those based on the bacterial lac or tetracycline-resistant operons (Gossen et aL, 1993), offer an unprecedented degree of control over transfected gene expression. By providing uniform regulated induction of gene expression before, during, or after the differentiation process, use of these systems in muscle cultures promises to

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inspire new strategies for further dissecting the structural and functional features of the myogenic cell. Acknowledgments Dr. Jean Buskin and DeeAnn Gregory are thanked for their many helpful contributions to protocols provided for MM14 cells; Dr. Mary Pat Wenderoth is thanked for help with the manuscript.

References Alam, J., and Cook, J. (1990).Reporter genes: Application to the study of mammalian gene transcription. Anal. Biochem. 188,245-254. Amacher, S . L., Buskin, J. N., and Hauschka, S. D. (1993).Multiple regulatory elements contribute differentially to muscle creatine kmase enhancer activity in skeletal and cardiac muscle. Mol. Cell. Biol. W, 2753-2764. Baum, C., Forster, P., Hegewisch-Becker, S., and Harbers, K. (1994).An optimized electroporation protocol applicable to a wide range of cell lines. BioTechniques 17,1058-1062. Berger, J., Hauber, R.,Geiger, R., and Cullen, B. (1988).Secreted placental alkaline phosphatase: A powerful new quantitative indicator of gene expression in eukaryotic cells. Gene 61-10. Buskin, J. N., Gregory, D. L., LaFramboise, W. A. and Hauschka, S. D. (1996).A harvest protocol to reduce variability of soluble enzyme yield from cultured cells. BioTechniques, u),92-100. Chu, G., Hayakawa, H., and Berg, P. (1987).Electroporation for the efficient transfection of mammalian cells with DNA. Nucl. Acids Res. 15,1311-1326. de Wet, J. R.,Wood, K. V.,DeLuca, M., Helsinki, D. R. and Subrarnani,S. (1987).Firefly luciferase gene: Structure and expression in mammalian cells. Mol. Cell. Biol. 7,725-737. Donoghue, M., Emst, H., Wentworth, B., Nadal-Ginard, B., and Rosenthal, N. (1988).A musclespecific enhancer is located at the 3’end of the myosin light chain 1/3locus. Gene Dev. 2,1779-1790. Eastman, A. (1987).An improvement to the novel rapid assay for chloramphenicol acetyltransferase gene expression. Bw Techniques 5,730. Gossen, M., Bonin, A. L. and Bujard, H. (1993). Control of gene activity in higher eukaryotic cells by prokaryotic regulatory elements. TIES 18,471-475. Graham, F. L., and van der Eb, A. J. (1973).A new technique for the assay of infectivity of human adenovirus 5 DNA. Virology 52,456-467. Morgenstern, J. P., and Land, H. (1990). Advanced mammalian gene transfer: High titre retroviral vectors with multiple drug selection markers and a complementary helper-free packaging cell line. Nucl. Acids Res. 18,3587-35%. Moss, J. B., Price, A. L., Raz,A., and Rosenthal, N. (1995). The green fluorescent protein marks skeletal muscle in murine cell lines and zebrafish. Gene, 173,89-98. Neumann, J. R., Morey, C.A. and Russian, K.0. (1987). A novel rapid assay for chloramphenicol acetyltransferase gene expression. BioTechniques 5,444. Potter, H. (1987). Transfection by electroporation. In “Current Protocols in Molecular Biology” (F. A. Ausubel, R. Brent, R.E. Kingston, D. D. Moore, J. G. Seidman, J. A. Smith, K. Struhl, eds.), Vol. 1, pp. 9.3.1-9.3.6.Greene Publishing Associates and John Wiley & Sons. Potter, H., Weir, L., and Leder, P. (1984).Enhancer-dependent expression of human kappa immunoglobulin genes introduced into mouse pre-B lymphocytes by electroporation. Proc. Nutl. Acad. Sci. USA 81,7161-7165. Rosenthal, R. (1989).Muscle cell differentiation. Curr. Opin. in Cell Biol. 1,1094-1101. Selden, R. F., Howie, K. B., Rowe, M. E., Goodman, H. M., and Moore, D. D. (1986). Human growth hormone as a reporter gene in regulation studies employing transient gene expression. MoL Cell. Biol. 6, 3173-3179.

CHAPTER 20

DNA- and Adenovirus-Mediated Gene Transfer into Cardiac Muscle Alyson Kass-Eisler* and Leslie A. Leinwandt 'Cold Spring Harbor Laboratory Cold Spring Harbor, New York 11724 +Department of Molecular, Cellular, and Developmental Biology University of Colorado at Boulder Boulder. Colorado 80309

I. Introduction 11. Methods for Cardiac Injection 111. Homogenization of Tissue and Assays IV. Experimental Design V. Future Directions References

I. Introduction In this chapter we will compare two methods of gene transfer into the rodent heart: direct plasmid DNA injection and injection of recombinant adenovirus. The goals of cardiac gene transfer that will be considered here are defining the elements regulating cardiac gene expression and phenotypic modification. Most studies aimed at understanding cardiac gene regulation have used transient transfection of DNA constructs into fetal or neonatal cardiocyte cultures. No transfections of adult cardiocytes have been reported. However, these studies are limited because of the developmental stage difference between the cultured cells and the adult and because of the inability to reproduce complex physiological and pathological processes in a tissue-culture environment. One solution to this problem is the creation of transgenic mice bearing reporter genes whose expression is driven by various promoters and putative regulatory regions. However, the creation of transgenic animals for this purpose is both time-consuming and expenMETHODS IN CELL BIOLOGY. VOL. 52 Copyright Q 1998 by Academic Press. AU rights of reproduction in any form resewed. lN91-67YX/98 125.00

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sive. Direct gene injection using naked plasmid DNA was first described in skeletal muscle by J. Wolff et al. in 1990. Several laboratories have extended the use of this methodology to include heart, which appears to be much more efficient than skeletal muscle in taking up and/or expressing naked DNA (Lin et al., 1990; Ascadi et al., 1991; Buttrick et al., 1992). The predominant use of direct injection into the heart has been to analyze promoter elements in vivo using reporter genes driven by the promoter element of interest (Kitsis et al., 1991; Buttrick et al., 1993). There are several advantageous features of naked DNA-mediated gene transfer. Primary among these are the stable expression resulting from gene transfer (Wolff et al., 1992) and the relative simplicity of the approach, which avoids infectious agents. Reports of gene expression beyond 1 year exist (Wolff et al., 1992). In addition, the DNA remains episomal, avoiding potential risks associated with the integration of DNA into the host genome (Wolff et al., 1992). The major drawback to this approach is its relative inefficiency.The maximum activity that can be obtained from gene expression appears to be restricted to an area immediately surrounding the injection site (Buttrick et al., 1992). Furthermore, the number of cells that take up and express the DNA is relatively small, corresponding to -0.02% of the myocytes in the heart (Kitsis et al., 1993). These drawbacks do not affect the utility of this method for mapping the regulatory elements of genes. Substantial reporter gene activity is obtained even after the introduction of relatively small amounts of DNA. In fact, when contemplating gene regulation studies, it is important to use relatively small quantities of DNA if the promoter is very active. For example, a strong promoter such as the Rous sarcoma virus LTR is out of the linear range of dose responsiveness above -2 pg in a rat heart. The assumption is that there are limiting transcription factors and that the cell rapidly becomes saturated with DNA sequences that bind these factors. Therefore, a DNA dose-response curve should be generated for each promoter under investigation. The second general use of in vivo gene transfer is to examine the consequences of expression of a given gene on the structure or function of the heart. This could involve expression of a mutant protein or ectopic expression of an isoform of a protein normally found in another setting. For these purposes, unless the gene encodes a secreted protein, DNA injection would not be efficient enough for most forms of phenotypic modification. We have therefore turned to the use of recombinant adenoviruses. Adenoviruses are double-stranded linear DNA viruses that have been extensively characterized in clinical and laboratory settings. Adenoviruses efficiently infect a large number of replicating and nonreplicating cell types. Their ability to infect nonreplicating cells makes them ideal for use in the heart, since adult cardiac myocytes are terminally differentiated. The recombinant adenoviruses in current use by most laboratories are replication deficient because they are deleted in a major regulatory region, termed Ela. Vectors currently exist that can accommodate up to 8 kb of a recombinant sequence, which makes them potentially useful for the majority of cDNA se-

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quences (Bett et af., 1994). These viruses can also be produced as extremely high-titer stocks (1011-1012pfdml). Finally, adenoviruseshave an excellent safety profile. The genome rarely integrates into the host genome and has never been associated with human malignancy. Several groups have demonstrated very efficient infection of the cardiovascular system by recombinant adenoviruses (Kass-Eisler et af., 1993; Stratford Perricaudet et al., 1992). Our group has focused on direct intracardiacmuscle injection of viruses. We first determined that both fetal and adult cardiac myocytes can be efficiently infected in culture (Kass-Eisler et af., 1993), achieving virtually 100% infection with a multiplicity of infection (MOI) of 2-3 pfdcell. We then examined the amount and distribution of gene expression resulting from injection of various viral doses into the left ventricular wall of the rat heart. Between 6 X lo6 and 2 X lo9 pfu there is a linear response between viral dose and expression. We have also compared the efficiency of gene transfer of a plasmid and an adenovirus bearing identical transgenes. For these experiments, we have used an adenovirus of the serotype 5 that bears the CAT reporter gene driven by the cytomegaloviruspromoter. The CAT gene was chosen because of its ability to be quantitated as well as its ability to be detected immunohistochemically.At maximal dose responsiveness for virus and plasmid DNA, the amount of CAT activity was approximately lOOOX higher in hearts injected with virus, 5 days following injection (Kass-Eisler et af., 1993), compared with its level in hearts injected with plasmid DNA. We had previously demonstrated in hearts injected with DNA that 98% of the activity of reporter genes was restricted to a small area around the injection site and that the transfection efficiency was low, corresponding to only -2000 cells in the entire heart (Kitsis et al., 1993). We compared both the distribution and the proportion of CAT-positive cells resulting from a single virus injection with results obtained from DNA injection. The quantitative distribution of expression was no different between adenovirus and plasmid DNA injection, in that the majority of expression was restricted to the injection site. However, because the amount of gene expression was so much higher in the virus injections, there was significant gene expression throughout the entire heart. In order to determine the proportion of cells in the heart which were infected with the virus, sections of the heart were stained with an anti-CAT antibody. In the regions of the heart surrounding the injection site, virtually 100% of cells were expressing CAT. In contrast to naked DNA injection, which transfects only cardiac myocytes, all cell types in the heart were able to be infected by the virus. The proportion of CAT-positive cells decreased as a function of distance from the injection site, reaching -5-10% of the cells at the base of the heart, which was the most distal location relative to the injection site. An additional feature of adenovirus infections in the adult heart that must be taken into consideration is the transient expression of genes. A universal observation with adenovirus is that independently of the route of the virus or the transgene it expresses, expression is transient, dropping sharply within -2 weeks, reaching undetectable levels

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by 80 days (Kass-Eisler et al., 1994; Mittal et al., 1993; Quantin et al., 1991). It now seems clear that the long-term expression of genes is limited by the immune response of the host to the virus and to the transgene. This immune response appears to be both humoral and cellular (Kass-Eisler et al., 1994; Yang et al., 1994). Extremely high levels of neutralizing antibodies develop within 5 days following virus administration (Kass-Eisler et al., 1994). Antibodies directed against the transgene also develop, but with a temporal delay compared with those directed against the virus (Kass-Eisler et aL, 1994).Traditionalimmunosuppression by agents such as cyclosporine is ineffective in prolonging gene expression (Kass-Eisler et al., 1994; Engelhardt et al., 1994). Experiments are underway to examine the time course of gene expression in transgenic animals whose immune systems have been modified in a variety of ways. The response of the immune system could be potentially circumvented by introducing virus into the immunologicallyincompetent neonatal animal. Neonatal rodents are immunologically incompetent and therefore do not recognize viral or transgene proteins as foreign. Expression by adenovirus-mediated genes in a neonate appears to be indefinite, reaching a time course of at least 7-8 months (Kass-Eisler et al., 1994; Rosenfeld et al., 1991). Experiments are currently in progress to determine whether neonates can be tolerized to the virus, allowing multiple administration of various recombinant viruses. In the case of injection into neonates, the technique will be described later. Because of the size of the animals, direct cardiac injection is not feasible. Therefore, animals are injected percutaneously into the thoracic cavity. This mode of introducing virus results in effective infection of multiple organs, including heart, diaphragm, lung, and liver, for an indefinite period of time. The amount of CAT activityper heart is approximately 1OOOX lower than that seen from direct cardiac injection. The lower levels of expression in the heart are most likely due to the distribution of virus by this route of administration. Thoracic-cavity injection most likely results in the majority of adenovirus being introduced into the lung. In support of this, 100-fold greater expression levels can be measured in the lungs of neonatally injected animals than in the lungs of directly cardiac-injected adults (Kass-Eisler et al., 1994).

II. Methods for Cardiac Injection Our injection protocol involves a simple surgical procedure to exteriorize the heart followed by injection of the heart muscle with a hypodermic needle and is usually performed by two people. Although other injection protocols have been developed for various species (Hansen et aZ., 1991; von Harsdorf et aL, 1993), the following protocol is for adult rats. This procedure can be used for delivery of naked DNA as well as recombinant adenoviruses. Animals are anesthetized with an intraperitoneal injection of 4% chloral hydrate (made fresh with water) at a dosage 0.75 d l 0 0 g body weight. This


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anesthesia, which takes effect within -10 min, lasts for approximately 30 min of deep sleep followed by approximately 1-2 hr of partial anesthesia. When animals are properly anesthetized there should be a loss of corneal reflexes and a lack of withdrawal to painful stimulation. Once anesthetized the animal is laid in a supine position. A vertical incision is made slightly to the left of midline with either a scalpel or sharp scissors. This can be accomplished through a natural slit in the muscles that runs diagonally across the animal’s left chest. The chest muscles are then blunt-dissected to the level of the ribs. The apical impulse of the heart can then be visualized (avoid the pulsation coming from the nearby great vessels). The rib just below the apical impulse is then cut with a scissors 5 mm to the left of the sternum. An incision too close to the sternum would risk damaging the internal mammary artery. The scissorsshould be held perpendicular and inserted as shallowly as possible to avoid damage to the heart or lungs. A Crile forceps is then inserted into the chest cavity through the rib incision perpendicular to the chest wall. The incision is then gently widened with the opening of the forceps. This opening of the chest cavity will cause the lungs to deflate, preventing respiration; therefore, it is important to continue as quickly as possible. The heart can usually be seen in the chest cavity at this point. If not, gentle pressure the right side of the chest can help move the heart into view. The heart is then gently exteriorized by grasping its apex with the Crile forceps. It is very important not to grip the heart by the atria or the great vessels, or to grasp the lungs. The heart can then be gently held by the operator’s fingers while the second operator performs the injection. A 100-pl Hamilton syringe with a 27-gauge needle containing the injectate is then inserted into the wall of the left ventricle. We have generally used an injectate volume of 50 p1 PBS (137 mM NaCl; 2.7 mM KCl; 10 mM Na2HP04; 2 mM KH2P04,pH 7.4)’ but this volume can be varied from 10 to 100 pl (Wolff et al., 1991) if necessary for a rat heart. The heart is then gently placed back into the chest cavity. The chest of the animal is then gently squeezed with fingers to expel any air from the cavity. The lungs are inflated using a Harvard small-animal respirator with a nose-cone adapter that fits over the snout of the rat. The nose cone is easily constructed by cutting of the bottom 4 cm of a 30-ml syringe and attaching a piece of glove latex over the wide end. The cone is attached to the respirator through tubing placed on the Luer-lock end of the syringe. The need for intubating the trachea is eliminated with the use of the nose cone, which provides positive-pressure ventilation of the animal. Two to four respirations with the respirator should be enough to induce spontaneous respiration; additional use should be kept to a minimum. At this point the muscle layers of the chest are closed using 3-0 silk sutures. Complete evacuation of pneumothorax should be accomplished before final suturing of the chest by gentle squeezing of the chest. The skin can then be stapled closed. Approximately 10%of all animals will succumb during this surgical

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procedure. The two main causes of death are improper anesthetic dosage and the trapping of air in the chest. Unlike rats, mice do not survive this procedure as well without modification. We have recently injected mice with adenovirus directly into the thoracic cavity. To do this, mice are held gently between the thumb and forefinger, and a 26gauge needle containing virus in a volume of 30 pl is introduced into the chest near the site of the heart. Care must be taken to insert and remove the needle slowly and gently. Complete immobilization of the animal is essential. Approximately 10% mortality (usually immediately after injection) can be seen with this procedure. Thoracic-cavity injection is also quite effective in the infection of neonatal animals. Neonatal animals are quite transparent, and if they are held over a light source, the ability to inject near the heart is greatly enhanced. Mortality rates from the injection in neonates are only about 2%.

III. Homogenization of Tissue and Assays For CAT (chloramphenicol acetyl transferase) and luciferase assays, tissues are homogenized. We have also used /3-galactosidase as a reporter gene, in which case whole tissues are harvested for staining. In addition, we have used human growth hormone (Selden et aZ., 1986) as a reporter gene in rats. Although high levels of expression of hGH can be measured in heart following administration of either phGH or AdhGH by direct cardiac injection,there is very little secretion of the hormone from the heart (see Fig. l),although detectable amounts in serum allow for multiple measurements on the same animal over time. In addition, hGH activity can be measured in homogenized tissue as discussed later. Before sacrifice, animals are given a large dose of 4% chloral hydrate (-1.5 ml/lOO g body weight). This is potentially a lethal dose, so care should be taken not to let the animals expire before the removal of tissues. The heart is removed by an abdominal incision through the midline. The ribs on both sides of the xiphoid process of the sternum are cut, revealing the heart in the chest cavity. The heart is then held with forceps by the apex while being cut at the base of the heart with a pair of scissors. The heart is immediately placed in cold PBS. If other tissues are to be harvested, they are also removed at this time and also placed in cold PBS. Collection of serum is facilitated by placing a borosilicate tube into the chest cavity in an angled position so that blood from the chest can drain into the tube. This tube is then placed on ice to allow clotting. Once all of the tissues have been harvested, they are trimmed and blotted dry. The tissues are weighed and transferred to Falcon 2059 tubes. The appropriate homogenization buffer is then added. We generally homogenize our tissues in 1ml buffer/OJg wet tissue weight. For CAT and luciferase assays we homogenize our samples in a buffer, which is prechilled on ice, composed of 25 mM glycylglycine (pH 7.8), 15 mM MgS04, 4 mM ethylene glycol bis(B-aminoethyl ether)-

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Fig. 1 Human growth hormone expression in adult rats followingdirect cardiac injection of plasmid DNA bearing the hGH gene (phGH) driven by the RSV promoter. Human growth hormone was measured by radioimmunoassay in heart homogenate (left ventricle) or serum of the same animal

4 days after injection. N = 5.

N,N,N’,N’-tetraacetic acid (EGTA)(pH 7.8), and 1 mM dithiothreitol (DTT) (added at time of use). For hGH assays we homogenize our tissues in 0.25 M Tris, pH 7.4. Homogenization is performed using a Tissue Tek (Tekmar) homogenizer at the maximal setting for 20 sec. During homogenization, the sample is placed in a beaker of ice to keep it cool. The sample is then returned to an ice bucket while the remaining samples are homogenized. Generally, the homogenization is completed for one animal before the harvesting of tissues from the next animal begins. Once all samples have been homogenized, they are centrifuged at 5000s for 25 min at 4°C.Supernatants are then removed and measured. At this point samples can be placed at -70°C until they are ready to be assayed. We generally assay a fixed percentage of tissue from each sample. This is generally 5% of the lysate from constructs containing strong viral promoters; a larger percentage may need to be assayed when cellular promoters are used. In addition, when adenovirus is used it may be necessary to dilute the samples significantly. When assaying from AdCAT dilutions are made with 0.1 mg/ml BSA (bovine serum albumin).

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CAT and luciferase assays have been described in detail elsewhere (Kitsis et al., 1993). Briefly, when CAT assays are done alone the entire sample is heated to 65°C for 10 min. The lysates are cleared by microfuge centrifugation, and the assay is performed by the standard 14C-chloramphenicol,thin layer chromatography method (Kitsis et al., 1993). When CAT activity is being assayed along with luciferase or hGH, the sample is generally divided. Samples for both luciferase and hGH assays are kept on ice throughout the preparation. Luciferase assays are performed by diluting the sample corresponding to 5% of the lysate in homogenization buffer so that the final volume is 100 p1. Samples are usually done in duplicate; 360 pl of an ice-cold buffer containing 25 mM glycylglycine (pH 7.Q 15 mM MgS04, 4 mM EGTA (pH 7.8), 1 mM DTT, 15 mM potassium phosphate (mono- and dibasic, pH 7.8), 2 mM ATP (pH 7.0), and 0.27% (v/v) Triton X-100 is added to each sample. The final solution containing the substrate luciferin-02 mM luciferin (sodium salt), 25 mM glycylglycine (pH 7.8), 15 mM MgS04,4 mM EGTA (pH 7.8), 2 mM DTT-is added directly to the luminometer (Monolight 2010; Analytical Luminescence Laboratory, San Diego, California). Approximately 5 ml luciferin solution is enough for 30 samples. The luciferin solution should be kept in foil because of the light sensitivity of luciferin. The luminometer is adjusted to measure light production over 20 sec following the addition of the luciferin solution. A control tube containing only buffers should be assayed as well. Generally this background will be approximately 200-600 RLUs (relative light units). Growth hormone assays are done on either serum or homogenized tissue samples by radioimmunoassay. A kit containing standards and all materials is available from Nichols Institute Diagnostics (San Juan Capistrano, California). In our hands, this kit has produced very consistant results without any modification of their directions. Although we have not done quantitativej3-galactosidase assays, we have done j3-gal staining on tissue sections following direct injection. Many other groups have used recombinant adenoviruses with the &gal reporter gene in tissue sections as well (Jaffe et aZ., 1992, Le Gal La Salle et al., 1993). The p-gal staining is performed on frozen tissue sections. We freeze tissue in isopentane (Zmethylbutane) and store it at -70°C. The tissue is tembedded in OTC, and 10-pl sections are cut on a cryostat. The tissues are placed on poly (L-1ysine)coated slides and can be stored at -70°C. The sections are then fixed in 1.5% glutaraldehyde in PBS for 5 min at room temperature. The slides are then rinsed three times in PBS. The sections are then incubated in a solution containing 400 pg/ml X-gal, 5 mM potassium ferriferrocyanide, and 1 mM MgClz in PBS for approximately 6 hr.

IV.Experimental Design To minimize variability in expression from animal to animal, we have routinely used a second plasmid as an internal standard when comparing the strength of


431

promoters. For example, when studying the effects of a cellular promoter driving the expression of luciferase, we have used a viral promoter driving CAT as a reference for injection quality. The promoter strength can then be reported as the ratio of luciferase activity to CAT activity. A twofold range of variability can still be seen following direct cardiac injection; therefore, we generally use five to eight animals per group. The use of two plasmids has brought to our attention a competition effect seen with strong viral promoters and large quantities of DNA. For example, when RSVCAT and RSVLUC are injected and are compared to an injection of RSVCAT and aLUC (alpha myosin heavy chain driving luciferase expression), higher levels of RSVCAT will be seen in the second group because it contains the weaker promoter (Buttrick et al., 1992). This phenomenon is presumably due to competition for transcription factors. We have found that this effect can be minimized when less plasmid is injected. For strong viral promoters, 0.52 p g of plasmid DNA should be sufficient to detect reporter gene expression. For each individual construct it may be necessary to test different doses in order to find the linear range. The minimum amount of plasmid that can be easily detected is best. To choose the best dose of adenovirus to inject, we quantitiated the level of CAT activity following intracardiac injection of various doses of AdCMVCATgD. For this virus, a roughly linear increase in CAT expression could be measured from 6 X lo6 pfu of virus through 2 X lo9 pfu, the highest dose tested. Adenovirus should be stored in a 50% glycerol solution. When highly concentrated virus was injected we saw an increase in mortality at the time of injection, presumably due to the concentration of glycerol in the injectate. We have routinely injected 6 X lo7 pfu of virus per animal. With our viral stocks, this is generally -1-10 pl virus stock, which can be diluted to 50 p1 with PBS. This dose gives us consistent results and limited mortality. Roughly fivefold variablity in expression can be seen in hearts 5 days following direct cardiac injection. Thoracic-cavityinjections generally give less consistent expression levels in a particular tissue, presumably because of the inherent imprecision of the route of administration. Using an ELISA assay for CAT protein (Boehringer-Mannheim), we have estimated that an average 132 pg of CAT protein can be expressed in the left ventricle 5 days following a single administration of adenovirus. In addition, approximately 20 ng/ml human growth hormone can be detected in serum following a single administration of AdhGH (secreted from all infected tissues), and 0.32 ng/ml of serum following hGH plamid injection. These quantities of protein are potentially consistent with a functional modification. Figure 2 shows the relative time course of expression from rats injected with either DNA or adenovirus. Following plasmid DNA injection, expression is maximal at approximately 7 days and remains consistent throughout the experiment. Animals injected with adenovirus show peak expression at approximately 5 days, but expression quickly drops, with no expression seen after 80 days. As

Alyron hs-Eider and Leslie A. Leinwand

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Fig. 2 Time course of CAT expression following cardiac muscle injection of either pRSVCAT or AdCMVCAT. Adult rats were injected with either 100 pg RSVCAT plasmid DNA or 6 X lo7 pfu adenovirus expressing CAT. At various times after injection the left ventricles of each animal were homogenized and assayed for CAT expression. Relative CAT expressionis based on CAT expression in 5% of the tissue lysate multiplied by the dilution of the sample.

seen in Fig. 3, unlike in adults, expression of adenovirus in a neonate is stable with no significant change in expression through at least 7.5 months. The differences in duration of expression between adults and neonates, as well as between adenovirus and naked DNA, can be attributed to the generation of a severe immune response in adult animals following adenovirus administration. Five days after administration of adenovirus, high levels of neutralizing antibodiesare generated against the virus (Kass-Eisler et al., 1994).These neutralizing antibodies prohibit gene transfer from additional injections of adenovirus of the same serotype (Kass-Eisler ef al., 1994), making subsequent injections impossible. In addition to the antibody response, a severe infiltratinglymphocyte response can be seen in adenovirus-injected animals, suggesting T-cell involvement (Kass-Eider et d., 1993;Yang d d., 1994; Engelhardt et al., 1994). We and others are now trying to dissect the immune response to determine which factors

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Fig. 3 Time course of CAT expression in adult vs neonatal rats injected with 6 X lo7 pfu of AdCMVCAT. Adult rats were injected by intracardiac muscle injection. Neonatal rats were injected by thoracic cavity injection. Values are shown as relative CAT expression in the heart as in Fig. 2.

are involved in the transience of adenovirus-mediated gene expression in adults. Since these responses are not seen in neonatal animals given adenovirus, or in animals given plasmid DNA injections, these are currently better models for long-term gene transfer

V. Future Directions The advantageous features of naked DNA injection make it desirable to consider its optimization for research purposes and to consider its potential for therapeutics. This technique has already been hailed as a breakthrough for vaccines, and DNA vaccination has now been demonstrated to be effective in the laboratory setting (Ulmer et al., 1993; Fyran et aZ., 1993). The other system where DNA injection is likely to be effective therapeutically is in the expression of secreted gene products. As mentioned earlier, the use of DNA injection for studying cardiac gene regulation has already been shown to be effective (Kitsis et al., 1991; Buttrick et al., 1993), so for those purposes, no further modification

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Alyson hs-Eider and Lealie A. Leiwand

of the technology needs to be considered. It is undoubtedly possible to increase the efficiency of gene transfer if the need arises. One potential means would be to increase the volume of the injectate. Another would be to test systematically the methods for preparing the DNA and the nature of the vehicle for delivery. A third potential means of optimization would be a mechanism for delivering multiple boluses of DNA over a wider geographic distribution. In contrast, the efficiency of adenovirus-mediated gene transfer is extremely high, and it is not considered to be necessary to improve it dramatically. However, major efforts are underway to develop more effective means of systemic delivery of virus. There are certain circumstances under which it might be desirable to have gene products expressed from multiple sites. Thus far, the most efficient means of achieving widespread gene transfer is to introduce virus into the apex of the left ventricle. This route of administration is even more efficient than when virus is introduced into the cardiac cavity (Kass-Eisler et al., 1994). The mechanism whereby injection of virus into the cardiac muscle is so efficient remains unknown, but is an observation that merits further investigation. The major hurdle that must be overcome with respect to adenovirus is the immune response to the virus. The immune response to the transgene must also be considered, although this would not be specific to adenoviruses as gene-transfer vehicles. The immune response to the virus appears to be both humoral and cellular and recognizes several structural proteins of the virus (Kass-Eisler et al., 1994;Engelhardt et al., 1994).Neutralizing antibodies to the virus develop rapidly after infection (within 5 days) and remain elevated for at least 96 days. In addition, there appears to be a potent cellular immune response, since inflammatory cells have been shown to be present in the area immediately surrounding infected cells (Yang et al., 1994; Engelhardt et al., 1994). Adenovirus DNA is lost as a function of time following infection, suggesting that infected cells are preferentially lost as a function of the immune system (Kass-Eisler et al., 1994). Two lines of experimentation are currently underway to solve this problem. The first relies upon the hypothesis that the immune response is activated primarily from leaky gene expression of late adenovirus genes, which are known to be potent immunogenes (Yang et al., 1994). This hypothesis has received support from the observation made by J. Wilson’s group that a recombinant adenovirus with a temperature-sensitivemutation in the E2b gene (in addition to an Ela deletion), which has a regulatory role in late gene expression, shows a longer duration of gene expression than its Ela-deleted counterpart (Engelhardt et al., 1994). However, it remains to be formally proven that late gene expression is responsible for the transient expression of adenovirus-encodedgenes. Further manipulations of the viral genome are ongoing to eliminate additional viral genes. This will require the creation of cell lines that can supply those functions in trans for the production of viruses. The second line of experimentation that addresses these issues is to define which aspects of the immune system are responsible for the transient expression of genes. For these purposes, our laboratory is determining the duration of


435

expression following injection of the AdCAT virus in mice that have been genetically manipulated to “knock out” various aspects of the immune systems. Preliminary results suggest that the immune system involvement may be different in different tissues (Kass-Eisler, Bloom, and Leinwand, unpublished observation) and that both humoral and cellular immunity are involved in the temporal decline in gene expression. Once the determinants of the immune system are defined, it may be possible to devise local and temporally restricted immunosuppressive agents that allow long-term gene expression to occur. Additional strategies for long-term, efficient gene transfer are also being developed using viral fragments and chemical reagents. To take advantage of the ability of adenovirus to disrupt the endosome, a number of groups have successfullyused adenovirus either chemically coupled or mixed with plasmid DNA (Curiel et af., 1991; Cotten et af., 1992; Yoshimura et af., 1993) in cultured cells. Both fetal and adult cardiac myocytes in culture can be transfected in this manner (Rivera and Leinwand, unpublished observation). These experiments now need to be performed in the intact animal to test whether efficient gene transfer can be accomplished by this method. The major advantage of this technology is that the adenoviruses used in these types of experiments can be psoralen or UVinactivated, therefore reducing the possibility of viral gene expression, and potentially reducing the immune response (Cotten et al., 1992). If the immune response to adenovirus cannot be overcome, then nonviral substitutes must be developed for long-term efficient gene transfer. One promising report (Zhu et al., 1993) has shown that when mixed with cationic liposomes, DNA transfer into adult mice by intravenous injection is relatively efficient and expression can last for at least 9 weeks. The field of gene transfer is one that is receiving a great deal of attention, and substantial progress has been made in a relatively short time. Many of the areas in need of improvement or expansion have now been identified, and it seems clear that in the near future, safe and effective gene transfer will be in common use. References Ascadi, G., Jiao, S. S., Jani, A,, Duke, D., Williams, P., Chong, W., and Wolff, J. A. (1991).Direct gene transfer and expression into rat heart in vivo. New Biol. 3,71-81. Bett, A. J., Haddara, W., Prevec, L., and Graham, F. L. (1994). An efficient and flexible system for construction of adenovirus vectors with insertions or deletions in early regions 1 and 3. Proc. Nafl. Acad. Sci. USA 91,8802-8806. Buttrick, P. M., Kass, A., Kitsis, R. N., Kaplan, M. L., and Leinwand, L. A. (1992). Behavior of genes directly injected into rat heart in vivo. Circ.Res. 70, 193-198. Buttrick, P. M.,Kaplan, M. L., Kitsis, R. N., and Leinwand L. A. (1993). Distinct behavior of cardiac myosin heavy chain gene constructs in vivo-discordance with in vitro results. Circ. Res.

72,1211-1217.

Cotten, M., Wagner, E., Zatloukal, K.,Phillips, S., Curiel, D. T., and Birnstiel, M. L. (1992).Highefficiency receptor-mediated delivery of small and large (48 kilobase) gene constructs using the endosome-disruption activity of defective or chemically inactivated adenovirus particles. Proc. Natl. Acad. Sci. USA 89,6094-6098.

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Curiel,D. T., Agarwal,S., Wagner,E. and Cotten, M. (1991). A d e n o h enhancement of transferrinpolylysine-mediated gene delivery. Proc. NatL Acad. Sci. USA 88,8850-8854. Engelhardt, J. F., Ye, X.,Doranz, B., and Wilson, J. M. (1994). Ablation of E2A in recombinant adenoviruses improves transgene persistence and decreases inflammatory response in mouse liver. Proc. Natl. Acad. Sci. USA 91,61%-6200. Fynan, E. F., Webster, R. G., Fuller, D. H., Haynes, J. R., Santoro, J. C., and Robinson, H. L. (1993). DNA vaccines: Protective immunizations by parenteral, mucosal, and gene-gun innoculations. Proc. Natl. Acad. Sci USA 90,11478-11482. Hansen, E., Fernandes, K., Goldspink, G., Butterworth, P., Umeda, P. K., and Chang, K. C. (1991). Strong expression of foreign genes following direct injection into fish muscle. FEBS Letfers 290, 73-76. Jaffe, H. A., Danel, C., Longenecker, G., Metzger, M., Setouchi, Y., Rosenfeld, M. A., Gant, T. W., Thorgeirsson, S. S., Stratford-Pemcaudet, L. S., Pemcaudet, M., Pavirani, A., Lecocq, J.-P., and Crystal, R. G. (1992). Adenovims-mediated in vivo gene transfer and expression in normal rat liver. Nature Genetics 1,372-378. Kass-Eisler,A,, Falck-Pedersen,E., Alvira, M., Rivera, J., Buttrick, P. M., Wittenberg, B. A., Cipriani, L., and Leinwand, L. A. (1993). Quantitative determination of adenovirus-mediatedgene delivery to rat cardiac myocytes in vitro and in vivo. Proc. Natl. Acad. Sci. USA 90,11498-11502. Kass-Eisler, A., Falck-Pedersen, E., Elfenbein, D. H., Alvira, M., Buttrick, P. M., and Leinwand, L. A. (1994). Distribution, duration, and immune response to adenovirus-mediated gene transfer. Gene Therapy 1,395-402. Kitsis, R. N., Buttrick, P. M., McNally, E. M., Kaplan, M. L., and Leinwand L. A. (1991). Hormonal modulation of a gene injected into rat heart in vivo. Proc. Natl. Acad. Sci. USA 88,4138-4142. Kitsis, R. N., Buttrick, P. M., Kass, A. A., Kaplan, M. L., and Leinwand L. A. (1993). Gene transfer into adult rat heart in vivo. Methods in Molecular Genetics 1,374-392. Le Gal La Salle, G., Robert, J. J., Berrard, S., Ridoux, V.,Stratford-Perricaudet, L. D., Perricaudet, M., and Mallet, J. (1993). An adenovirus vector for gene transfer into neurons and glia in the brain. Science 259, 988-990. Lin, H., Parmacek, M.S., Morle, G., Bollling, S., and Leiden, J. M. (1990). Expression of recombinant genes in myocardium in vivo after direct injection of DNA. Circulation 82,2217-2221. Mittal, S . K., McDermott, M. R., Johnson, D. C., Prevec, L., and Graham, F. L. (1993). Monitoring foreign gene expression by a human adenovirus-based vector usning the firefly luciferase gene as a reporter. Virus Research 28,67-90. Quantin, B., Pemcaudet, L. D., Tajbakhsh, S., and Mandel, J-L. (1991). Adenovirus as an expression vector in muscle cells in vivo. Proc. Natl. Acad. Sci. USA 89,2581-2584. Rosenfeld, M. A., Siegfried, W., Yoshiura, K., Yoneyama, K., Fukayama, M., Stier, L. E., Paakko, P. K., Gilardi, P., Stratford-Pemcaudet, L. D., Pemcaudet, M., Jallat, S., Pavirani, A., Lecocq, J.-P., and Crystal, R. G. (1991). Adenovirus-mediated transfer of a recombinant al-antitrypsin gene to the lung epithelium in vivo. Science 252,431-434. Selden, R. F., Howie, K. B., Rowe, M. E., Goodman, H. M., and Moore, D. D. (1986). Human growth hormone as a reporter gene in regulation studies employing transient gene expression. Mol. Cell Biol. 6,3173-3179. Stratford-Pemcaudet, L. D., Makeh, I., Pemcaudet, M., and Briand, P. (1992). Widespread longterm gene transfer to mouse skeletal muscles and heart. J. Clin. Invest. 90, 626-630. Ulmer, J. B., Donelly, J. J., Parker, S. E., Rhodes, G. H., Felgner, P. L., Dwarki, V.J., Gromkowski, S. H., Deck, R. R., De Witt, C. M., Friedman, A., Hawe, L. A., Leander, K. R., Martinez, D., Perry, H. C., Shirer, J. W., Montgomery, D. C., and Liu, M. A. (1993). Heterologous protection against influenza by injection of DNA encoding a viral protein. Science 259,1745-1749. von Harsdorf, R., Schott, R. J., Shen, Y. T., Vatner, S. F., Mahdavi, V.,and Nadal-Ginard, B. (1993). Gene injection into canine myocardium as a useful model for studying gene expression in the heart of large mammals. Circ. Res. 72,688-695. Wolff, J. A., Malone, R. W., Williams, P., Chong, W., Ascadi, G., Jani, A., and Felgner, P. L. (1990). Direct gene transfer into mouse muscle in vivo. Science 247, 1465-1468.


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Wolff, J. A., Williams, P., Ascadi, G., Jiao, S.,Jani, A., and Chong, W. (1991). Conditions affecting direct gene transfer into rodent muscles in vivo. Biorechniques 11,474-485. Wolff, J. A., Ludtke, J. J., Acsadi, G., Williams, P., and Jani, A. (1992). Long-term persistence of plasmid DNA and foreign gene expression in mouse muscle. Human Molecular Genetics 1,363-369. Yang, Y.,Nunes, F. A., Berenni, K., Furth, E. E., Gonnol, E., and Wilson,J. M. (1994). Cellular immunity to viral antigens limits El-deleted adenoviruses for gene therapy. Proc. Natl. Acad. Sci. USA 91,4407-4411. Yoshimura, K., Rosenfeld, M. A., Seth, P., and Crystal, R. G. (1993). Adenovirus-mediated augmentation of cell transfection with unmodified plamid vectors. J. B i d Chem. 268,2300-2303. Zhu, N., Liggitt, D., Liu, Y.,and Debs, R. (1993). Systemic gene expression after intravenous DNA delivery into adult mice. Science 261,209-211.


CHAPTER 21

Nuclear DNA-Binchng Proteins Kristen L. Kucharczuk and David J. Goldhamer Department of Cell and Developmentd Biology University of Pennsylvania School of Medicine Philadelphia, Pennsylvania 19104

I. Introduction 11. Nuclear Extract Preparation: Overview and Important Parameters 111. Nuclear Extract Protocol A. Protocol B. Reagents C. Notes IV. Electromobility Shift Assays: Overview and Important Parameters A. Binding Reaction Conditions B. Electrophoresis Conditions C. Sensitivity V. EMSA Protocol A. Probe Preparation B. EMSA Protocol: High-Ionic-Strength Conditions C. EMSA Protocol: Low-Ionic-Strength Conditions D. Reagents E. Competition Experiments F. Antibody Supershifts VI. DNase I Protection Assays: Overview and Important Parameters A. CarrierDNA B. DNase I C. Binding Conditions D. Probe VII. DNase I Protection Assay Protocol A. Probe Preparation B. DNase I Protection Protocol C. Preparation of G A Ladder: Maxam and Gilbert Reactions D. Reagents E. Notes

+

METHODS IN CELL BIOLOGY. VOL. 52 Copyright 0 1998 by Academic Press. MI rights of reproduction in any form mewed. w91-679X1981225.00

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VIII. Relevance of the EMSA and DNase I Protection Assay to Muscle Research A. I n Vitro Techniques using Crude Nuclear Extracts B. In Vitm Techniques using Purified Proteins C. Concluding Comments References

I. Introduction Much of our understanding of the molecular control of myogenesis has come from the study of DNA-binding proteins that bind to enhancers and promoters of muscle-specific genes. Techniques that allow the detection of DNA-protein interactions in vitro, notably the electromobility shift assay (EMSA) and the DNase I protection assay, have been used extensively in muscle research both to detect novel DNA-binding proteins in muscle cell nuclear extracts and to define fundamental properties of previously identified factors produced in bacterial or reticulocyte lysate expression systems. In this regard, it should be noted that the great utility of EMSAs and DNase I protection assays in muscle research is due, to a large extent, to the availability of skeletal muscle cell culture model systems (Chapters 5 and 8). The muscle regulatory proteins MyoD and MEF-2, for example, were first identified by EMSAs as muscle-specific DNA-binding activities present in cultured myocyte nuclear extracts (Buskin and Hauschka, 1989; Gossett et aL, 1989). For cloned factors, identification of DNA sequence requirements for binding, characterization of protein domains for DNA binding and heteroligomerization, identification of dimerization partners, and characterization of protein-protein interactions that enhance or inhibit DNA binding are just some of the applications these techniques have found in muscle research. In conjunction with mutagenesis and functional assays (Chapters 18 and 19), both EMSAs and DNase I protection assays have been instrumental in defining the anatomy of muscle gene regulatory regions. In short, these techniques have served a fundamental role in dissecting gene regulatory pathways in vertebrate species where classical genetics is not available. Both EMSAs and DNase I protection assays rely on the ability of proteins to bind radiolabeled DNA in vitro. In EMSAs, DNA-protein complexes are separated from unbound DNA fragments by nondenaturing polyacrylamide gel electrophoresis. In DNase I protection assays, protein binding protects regions of DNA from DNase I cleavage. This protection, or “footprinting,” results in gaps in DNase I-generated fragment ladders. Although both EMSAs and DNase I protection assays can be used to detect and characterize DNA-bindingactivities and to map protein-binding sites within muscle enhancers and promoters, differences in sensitivity, type of information gained, and facility of use typically mandate the use of one technique over the other for a particular application. The advantages and particular utility of each technique will be presented within the EMSA and DNase I protection sections of this chapter.

21. Nuclear DNA-Binding Proteins

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In this chapter, we discuss and provide protocols for EMSAs and DNase I protection assays, and present particular experiments that exemplify important biological paradigms or methodologies in muscle research. In addition, we describe a detailed method for nuclear extract preparation, as preparation of highquality nuclear extracts is paramount in the success of these assays. We will focus on skeletal muscle, but it should be emphasized that similar methods have yielded considerable progress in cardiac and smooth muscle research. As this is intended as a methodological guide, practical considerations will be emphasized.

11. Nuclear Extract Preparation: Overview and Important Parameters Nuclear extract production, originally described by Dignam and colleagues (1983), exploits the observation that transcription factors can be selectively extracted from nuclei that have been isolated under low-ionic-strength conditions. Briefly, tissue culture cells are harvested, washed, and resuspended in hypotonic buffer. Once the cells are adequately swollen, they are mechanically homogenized, often in the presence of mild detergent, to achieve lysis and release of cytoplasmicproteins. Nuclei are then pelleted and resuspended in high-salt buffer to release soluble nuclear proteins (e.g., transcription factors). Centrifugation yields an extract containing the nuclear proteins of interest (supernatant). Ultimately, the extract is dialyzed into an intermediate-salt buffer that is compatible with most EMSA and DNase I protection assay conditions. Several parameters are critical to the production of active nuclear extracts. First, protein denaturation and degradation must be avoided. Once tissue culture cells have been harvested, all subsequent steps should be performed on ice, preferably in the cold room. Precool all centrifuges, rotors, rubber adapters, tubes, pipets, and buffers. Phenylmethylsulfonyl fluoride (PMSF), a serine-threonine protease inhibitor, is typically the only protease inhibitor required for the isolation of mammalian nuclear extracts. Although the original Dignam protocol demonstrated that nuclear extracts prepared in the presence of protease inhibitors were not appreciably more transcriptionally active than extracts prepared in the absence of protease inhibitors (Dignam et al., 1983), the potential deleterious effect of proteases on subsequent assays has led to the routine inclusion of at least PMSF in all preparatory buffers. Some investigators routinely use additional protease inhibitors (leupeptin, pepstatin A, soybean trypsin inhibitor, antipain). Each should be added just prior to buffer use. Second, a balance between efficient extraction of soluble nuclear proteins and extraction of nonspecific chromatin-binding proteins must be achieved. Typically, a salt concentration of 420 mM, empirically determined by Dignam et al. (1983), is used for the extraction buffer. This relatively high salt concentration allows adequate extraction of the proteins of interest while avoiding release of histones and other nonspecificchromatin-binding proteins. Any contaminating nonspecific

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DNA-binding proteins are usually titrated out in EMSAs and DNase I protection assays by excess nonspecific competitor DNA.

In. Nuclear Extract Protocol A. Protocol The following is a modification of a protocol developed in the laboratory of

K.S. Zaret (Milos and Zaret, 1992, Zaret et al., 1992). Nuclear extracts prepared in the following manner have been successfully used in our lab for both EMSA and DNase I protection analyses. Forty 15-cm plates of C2C12 cells harvested at 5040% confluence should yield a minimum of 2 mg of nuclear protein (12 pg/p1 final concentration).Yield will vary with cell type and degree of confluence. Approximately 5 to 6 hr are required for completion of the protocol. It is important to accomplish the entire procedure in 1 day.

1. Rinse cells twice with calcium- and magnesium-free phosphate buffered saline (PBS) at room temperature. Typically, 20 to 40 15-cm plates of cells at 50 to 80%confluence are used for each nuclear extract preparation. Approximately 5 ml PBS/plate is sufficient for adequate rinsing. Pipet PBS onto each plate, swirl gently, and aspirate with a Pasteur pipet attached to a vacuum source. 2. Add 5ml of ice-cold PBSlsucroselDTTPMSFIleupeptin (PSDP) to each plate. Scrape cells into 50-ml disposable conical tubes on ice. Pellet cells at 650g for 5 min at 4°C. All subsequent steps should be performed on ice in the cold room. 3. Decant supernatant and resuspend cell pellet in 10 ml PSDP. Pool 10-ml fractions into one 50-ml conical tube and pellet cells at 650g for 5 min at 4°C. Resuspension works best if the cell pellet is loosened prior to addition of buffer. 4. Decant supernatant and resuspend cell pellet in 10 ml PSDP. Transfer suspension to a 15-ml polypropylene snap-cap tube. Pellet cells at 650g for 5 min at 4°C. 5. Decant supernatant. Resuspend the packed cells in 2 ml hypotonic buffer A and allow to swell on ice for 5 min. Centrifuge the cells at 650g for 5 min at 4°C and discard supernatant. This step removes residual salt from the PSDP solution. 6. Resuspend the cell pellet in 3 ml hypotonic buffer A containing 0.5% NP40. Allow to swell for an additional 5 min on ice. Cells should swell at least twofold. The added detergent will help to lyse the cell membrane while leaving the nuclear membrane intact. See Notes, Section 111.C, for preparation of wholecell and cytoplasmic extracts. 7. Lyse cells with a Dounce homogenizer. Use 10 to 15 up-and-down strokes with a type A pestle. Homogenize with slow strokes to avoid foaming. Check under a microscope for cell lysis > 95%. If lysis is incomplete, dounce several more times and check again. If not using detergent, a type B pestle should be used to give more complete cell lysis.


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8. Transfer homogenate to a fresh 15-ml tube, rinse Dounce with 4 ml buffer B, add rinse to the homogenate, and centrifuge at 650s for 5 min at 4°C. Discard supernatant. 9. Add an additional 2 ml buffer B. Again, centrifuge at 650g for 5 min at 4°C. Discard supernatant. 10. Gently resuspend the nuclear pellet in 2 ml hypertonic buffer C. Tap tube to loosen the nuclear pellet, then gently pipet nuclei up-and-down with a Pl000 Pipetman to resuspend. 11. Transfer the nuclear suspension to 1.5-ml microfuge tubes (1 ml/tube) and incubate on ice for 30 min. Gently invert the tubes every 5 min while they are on ice. 12. Microfuge at 3000 rpm at 4°C for 5 min. Transfer supernatant to a new microfuge tube that has had its cap cut off cleanly. Place a small square of dialysis tubing (6000-8000 M W cutoff) over the top of the filled tube and secure with a rubber band wrapped many times around the body of the tube. Place inverted tubes securely into a floating rack and float on Buffer D in a beaker containing a stir bar. Dialyze twice at 4°C against 500-ml volumes of Buffer D for 1hr each, inverting tubes once or twice during each dialysis to rinse residual salt from the bottom of the tubes. Follow manufacturer’s instructions for preparation of the dialysis tubing. 13. Centrifuge the dialyzed extract at full speed for 5 min at 4°C. This final spin removes protein that has precipitated during dialysis. 14. Aliquot supernatant into 20 pl (EMSA) or 50 pl (DNase I protection) portions in precooled 0.5-ml tubes, freeze in liquid nitrogen, and transfer to a -80°C freezer. Thaw extracts only once. 15. Determine protein concentration of the extract. Protein concentration is easily determined by the modified Bradford assay (BioRad) using bovine serum albumin as a standard. B. Reagents

Stocks of the following solutions (minus DTT and protease inhibitors) may be kept at 4°C for several weeks. Add DTT and protease inhibitors to buffers immediately before use, as they are unstable in aqueous solutions. Be sure to wear gloves when handling protease inhibitors. 1. lOOmM PMSF

Dissolve the appropriate amount of (17.4 mg/ml) of PMSF in anhydrous isopropanol. Store at -20°C. Because PMSF is rapidly inactivated in aqueous solutions, add it to buffers just prior to use. 2. Leupeptin Make as an aqueous 10 mg/ml stock. Store at -20°C. Avoid repeated freeze-thawing.


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3. PSDP 0.15 M NaCl 20 mM sodium phosphate, pH 7.4 0.35 M sucrose 0.5 mM dithiothreitol (DTT) 1mM phenylmethylsulfonyl fluoride (PMSF) 5 p g / d leupeptin.

4. Hypotonic Buffer A 10 mM KC1 10 mM Hepes, pH 7.9 1.5 mM MgC12 0.5 mM DTT 1 mM PMSF 5 pg/d leupeptin

5. Buffer B 0.35 M sucrose 60 mM KC1 15 mM NaCl 15 mM Tris-C1, pH 7.4 0.2 mM EDTA 0.2 mM EGTA 0.5 mM spermine 0.15 mM spermidine 1 mMDTT 2 mM PMSF 5 pg/d leupeptin

6. Hypertonic Buffer C 0.42 M NaCl 20 mM Hepes, pH 7.9 1.5 mM MgC12 0.2 mM EDTA 25%glycerol 1 mM DTT


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2 mM PMSF 5 pg/ml leupeptin

7. Dialysis Buffer D 60 mM KCI 20 mM Hepes, pH 7.9 20% glycerol 0.2 mM EDTA 0.5 mM DTT 1 mM PMSF C . Notes

1. Situations exist where proteins contained within the cytoplasmic fraction are required for analysis of protein-DNA interactions. For example, the nuclearlocalized inhibitory factor Id is quantitatively extracted from proliferating myoblasts under low-ionic-strength conditions ( Jen et al., 1992). Similarly, the transcription factor NF-KBis retained in the cytoplasm by the inhibitory protein IKB until cytokine or mitogen induction stimulates its release and translocation to the nucleus (Grilli et al., 1993). Investigators wishing to prepare whole-cell extracts from tissue culture cells should consult Jen et al. (1992). For a detailed description of cytoplasmic fraction preparation, refer to Ausubel et al. (1995). 2. Working with limited cell numbers (e.g., early embryonic tissues) may necessitate the use of whole-cell rather than nuclear extracts for in vitro binding assays. Whole-cell extracts require fewer manipulations in their preparation and, as such, result in less protein loss. Once a tissue sample has been frozen, it becomes necessary to prepare whole-cell rather than nuclear extracts; freezing damages the nuclear membrane, permitting nuclear components to escape into the cytoplasmic fraction during nuclear-extract preparation (Dent and Latchman, 1993). For a description of whole-cell extract preparation from Xenopus early embryos and somites, refer to Taylor et al. (1991). 3. Occasionally,inhibitory proteins present in crude nuclear extracts can destabilize or preclude particular protein-DNA interactions under in vitro conditions (Lichtensteiner eta/., 1987). If this is the case, nuclear extracts can be fractionated over heparin-agarose columns to isolate the activity of interest. Once bound to the column, nuclear proteins can be eluted in stepwise fashion with increasing salt concentrations. These eluates containing subsets of the nuclear protein pool can be aliquoted and assayed individually for DNA-binding activity. See Lichtensteiner et al. (1987) for fractionation protocol. 4. The basic protocol just given may be scaled up or down, depending on the desired yield and cell density. Larger-scale preparations of nuclear extracts may


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be dialyzed in dialysis tubing rather than in the modified Eppendorf tubes described earlier. Refer to Ausubel et al. (1995) for dialysis times and conditions.

IV.Electromobility Shift Assays: Overview and Important Parameters The electromobility shift assay (EMSA) has been invaluable to the study of protein-DNA interactions in myogenic cells. First described in 1981 by Garner and Revzin, the EMSA was originally used to characterize the binding kinetics of prokaryotic regulatory proteins. Since that time, various modifications involving binding conditions, amount and type of carrier DNA, and electrophoresis conditions have made the EMSA one of the simplest, fastest, and most sensitive methods available for detecting protein-DNA interactions. Electromobility shift assays rely on the ability of proteins to bind radiolabeled DNA fragments in v i m . Once binding occurs, DNA-protein complexes can be separated from unbound DNA fragments by nondenatwing polyacrylamide gel electrophoresis (Fig. 1). In general, the larger the protein or protein complex bound to the DNA, the greater the extent of retardation of mobility of the radiolabeled DNA fragment. The resulting protein-DNA complexes are visualized by autoradiography. Various assay conditions can affect the detection and resolution of proteinDNA complexes. Next, we discuss several parameters that should be optimized.

no competitor salmon sperm DNA footprint # 1 footprint #2 footprints #1

+

#2

Fig. 1 Electromobility shift assay using 5 pg of nuclear extract prepared from C2C12 cells and a labeled synthetic oligonucleotide corresponding to two adjacent footprinted regions (footprints 1 + 2) of the human myOD core enhancer (see Fig. 3). Binding reactions were performed both in the absence of unlabeled competitor, and in the presence of a 1OOO-fold excess of salmon-sperm DNA, unlabeled footprint 1,2, or 1+2 (unlabeled probe) oligos. The results indicate that (i) the lowest of the three bands is a result of nonspecific complex formation (the complex can be eliminated by the addition of excess salmon sperm DNA), and (ii) the upper two complexes bind specifically to the footprint 1+2 probe, in particular to sequences within footprint 2.


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A. Binding Reaction Conditions 1. Type of Carrier DNA Although the first EMSAs did not include unlabeled, or “carrier,” DNA to compete out nonspecific DNA-binding proteins, subsequent studies have shown that the concentration and type of carrier DNA included in binding assays can have a dramatic effect on the number and composition of DNA-protein complexes detected. Typically, carrier DNA utilized in EMSAs is one of two types: complex heterologous or synthetic alternating copolymer. The first experiments to use carrier DNA employed E. coli DNA; since then, additional forms of complex heterologous DNA, including salmon-sperm and calf-thymus DNA, have been used successfully.The second type of carrier DNA, synthetic alternating copolymer DNA, was designed to increase the sensitivity of EMSAs, but still sequester nonspecific DNA-binding proteins. Synthetic copolymers poly(d1dC), poly (dG-dC), and poly (dA-dT) are thought to present a large number of sites for binding of these nonspecific binding proteins, but to compete less effectively than complex heterologous DNA for binding of sequence-specific factors (Imbalzano et al., 1994). The type of competitor used can determine which binding activities are detectable by EMSA. In a study of the skeletal a-actin promoter, Lee and Schwartz (1992) found that, using multiple polyanion polymers and a single DNA probe, they were able to detect three distinct DNA-binding activities in myocyte nuclear extracts. Using only poly(d1-dC) carrier they were able to detect just one of these activities, F-ACT1; using poly(dG-dC) and calf thymus DNA they unmasked an additional serum response element-bindingcomplex. Use of the polyanion heparin revealed a third binding activity, F-ACT;?. All three complexes were subsequently shown to be sequence specific. Of note, the serum response element (SRE) sequence contains an A + T-rich core. Since the more similar the sequence of a given carrier DNA is to the binding site of interest, the more effectively it will compete for sequence-specificbinding proteins and mask important binding interactions, one would correctly predict that SRE-binding activities would be most easily identified using a nonhomologous polyanion competitor such as poly(dG-dC) (Lee and Schwartz, 1992). We suggest that a set of dissimilar polyanion competitors be used to screen all cell extracts for novel DNAbinding activities.

2. Amount of Carrier DNA The amount of carrier DNA included in an EMSA can greatly influence protein-DNA interactions. If too little carrier is used, much of the labeled probe will be bound by nonspecific DNA-binding proteins; these multiple nonspecific protein-DNA complexes will be retained at the top of the gel. If, on the other hand, too much carrier DNA is included, even sequence-specific interactions may be disrupted and only free probe will be observed. In practice, a range of

Kristen L. Kucharczuk and David J. Gqldhamer

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carrier DNA concentrations permits detection of most sequence-specificinteractions-0.5 to 2 pg of carrier DNNreaction is typical.

3. DNA Probe and Cold Competitors The type and length of the DNA probe used for EMSAs depends on the experimental situation at hand. Once relevant protein-binding sites have been identified, synthetic oligonucleotide probes are typically the probes of choice. The use of short, synthetic oligo probes isolates the binding site of interest from other adjacent sites, greatly simplifying the analysis and interpretation of EMSA results. In some cases, however, protein interactions involving adjacent sites may enhance or inhibit binding at the site of interest (Lee et al., 1991). By analyzing protein binding in the absence of these interacting sequences, important control mechanisms may be overlooked. Often, regulatory sequences are identified by functional analyses (transfections, transgenic animals), and EMSAs are used to map important sites within these larger promoter/enhancer fragments. In this case, restriction or deletion fragments up to 250 bp (fragments larger than 250 bp will comigrate with bound probe) may be used as probes (see Buskin and Hauschka, 1989). The complexity of the resulting mobility-shift patterns, however, can make these experiments difficult to interpret. If convenient restriction sites for generating smaller, overlapping subfragments of these larger regulatory regions are not available, overlapping synthetic oligonucleotides spanning the region of interest can be utilized instead. Often, however, the cost of generating overlapping oligonucleotides is prohibitive and these binding sites are best mapped by DNase I protection experiments. The facility of the EMSA allows several competitor oligos to be tested for protein-binding ability within the same experiment, making it the technique of choice to determine binding specificity and further refine binding-site sequences once sites have been identified by sequence comparison, DNase I footprinting, or functional assays (Fig. 1). Typically, specific and nonspecific unlabeled DNA fragments are introduced into the EMSA assay in molar excess of the radiolabeled binding site of interest; the ability of specific competitor and failure of nonspecific competitor DNA to compete for protein binding provides proof that the proteinDNA complex in question is sequence specific (specific competitor is typically identical to the DNA used as a probe).

4. Salt Concentration The salt concentration of the binding reaction can have a dramatic effect on the stability of protein-DNA complexes. High salt concentrations may disrupt weak, but specific, protein-DNA interactions, whereas low-salt binding buffers may allow nonspecific binding interactions to obscure sequence-specific complexes. A compromise between these two extremes must be reached. Generally,


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in the presence of the appropriate type and amount of carrier DNA, salt concentrations between 50 and 100 mM allow detection of most specific protein-DNA complexes while minimizing interference by nonspecific DNA-binding proteins.

5. Temperature Incubation temperatures for DNA-protein binding vary between research groups. We typically perform all incubations on ice to minimize protein degradation and optimize reproducibility; other investigators prefer incubations at room temperature to increase Brownian motion and mixing of reagents. Both conditions have been used successfully. B. Electrophoresis Conditions

Gel electrophoresis conditions can affect detection and resolution of proteinDNA complexes. The mobility of a given complex through a native polyacrylamide gel is dictated by its size, charge, and conformation; individual complexes migrate as distinct bands. Altering the acrylamide percentage, the acrylamide :bis ratio, and the pH can improve the resolution of two closely migrating complexes. Typically, high-ionic-strength buffer systems give higher resolution than do lowionic-strength systems, although some protein-DNA interactions will not survive high-ionic conditions. It is helpful to try both high- and low-ionic buffer systems, particularly when identifying novel activities from crude cell extracts. C . Sensitivity

The kinetic stability of an individual protein-DNA complex will affect its ability to be detected by EMSAs. Great kinetic stability, however, is not required for detection. Although gels are typically run for 2 to 3 hr, protein-DNA interactions lasting even less than 1 min can be detected by this method (Ausubel et aZ., 1995). Several factors are known to enhance the detection of protein-DNA complexes. First, EMSAs rely on direct visualization of proteins bound to radiolabeled probe. Second, as mentioned earlier, low-ionic-strength buffer systems increase the stability of most nucleic acid-protein interactions, resulting in longerlived, more easily detected complexes. Third, a “caging effect” provided by the gel matrix itself increases the apparent kinetic stability of complexes; the polyacrylamide gel matrix physically prevents diffusion of dissociated protein and DNA molecules, thereby increasing their propensity to reassociate (Fried and Crothers, 1981; Garner and Revzin, 1981). As such, detection of protein-DNA interactions depends only on the kinetics of interaction as complexes enter the gel matrix. Together these factors give the EMSA its great sensitivity and allow femtomolar amounts of even low-affinity DNA-binding proteins to be detected routinely.


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V. EMSA Protocol A. Probe Preparation The simplest method for constructing EMSA probes involves end-labeling synthetic oligonucleotides using T4 polynucleotide kinase. Single stranded oligonucleotides to be labeled in this manner should form blunt-ended DNA duplexes when annealed. If restriction fragments or oligonucleotides with 5’ overhangs are to be used as probes, refer to Ausubel et al. (1995) for protocols utilizing [a-32P]dCTPand the Klenow fragment of E. coli polymerase I. An end-labeling method is given here.

.

1. In order, mix together (40 pl final reaction volume): 0.5 pg forward strand oligonucleotide (for a 25 nt oligomer) 4 pl1OX T4 kinase buffer dH2O to 33 pl 5 pl [Y-~~PIATP (3,000 Ci/mmol; approximately 10 pCiIp1) 2 p1 T4 polynucleotide kinase (20 U/pl) Recently, a higher resolution, lower energy alternative to [y3*P]ATPhas become available. [Y-~~PIATP emissions give less scatter, allowing better resolution of closely migrating complexes. [Y-~~PIATP radiographs may require longer exposure times or use of a Phosphorimager for band detection. 2. Incubate for 1 hr at 37°C. 3. Heat inactivate kinase for 15 min at 75°C. 4. Add 5 pg reverse strand oligonucleotide. Heat reaction and allow to cool to room temperature over several hours. A 10-fold excess of cold second-strand oligonucleotide is used to drive most of the labeled single-stranded DNA into double-stranded probe. Annealing is best accomplished in a beaker containing approximately 700 ml of water. Heat water to near boiling on a hot plate. Add sample, turn off the heat, cover beaker with a Plexiglas shield, and allow to cool to room temperature over several hours. 5. Remove free nucleotides. We find it convenient to use premade Sephadex G50-80spin columns to remove free nucleotides at this step. A number of these columns are commercially available. Follow manufacturer’s directions for purification. 6. Count 1pl of the probe and determine labeling efficiency. Compare actual labeling to theoretical labeling using pmol DNA ends and specific activity of the isotope to calculate the theoretical yield. (See p. 5.68 of Sambrook et al., 1989, for conversion factors.) 7. Store at 4°C until ready to use. 8. Prior to use, dilute with water to 30,000 cpdpl.


45 1

B. EMSA Protocol: High-Ionic-Strength Conditions The following is a protocol used successfully in our lab. The binding conditions are compatible with the nuclear extracts described earlier, as well as with most bacterially expressed or in vitro translated proteins. The entire procedure takes approximately 5-6 hr from the time the binding reaction is set up to the time the gel is placed on film. As most of this time is incubation and gel-running time, it is feasible to perform several EMSAs in one day. Note: All steps should be performed on ice unless noted otherwise.

1. On ice, premix 10 p l 2 X binding buffer, 1 p1 of 1 pg/p1 poly(dYdC), and 1pl of 20 pg/pl bovine serum albumin (BSA) per reaction. As mentioned earlier, the appropriate camer DNA type and concentration may vary for each binding interaction being examined. When searching for novel binding interactions, be sure to employ a variety of camer DNAs. 3. To this 12 pl, add 5 pg nuclear extract or 3 pl reticulocyte lysate. Quantities of nuclear extract and in vitro translated protein are approximate and should be optimized. If using reticulocyte lysates, be sure to include an unprogrammed lysate as a control. Thaw extracts on ice just prior to use. 4. Bring volume up to 19 pl with sterile deionized water. It is easiest to make up a master mix containing the common ingredients (i.e., make enough for n + 1 reactions and then aliquot to individual tubes). If you intend to add unlabeled competitor DNA, this is the place to do it (adjust water accordingly). A 100- to 1000-fold molar excess of cold competitor is typical. 5. Finger-tap tubes to mix, then centrifuge briefly to collect all components in the bottom of the tubes. 6. Incubate on ice for 30 min to 1hr. A full hour of incubation helps to reduce background by allowing camer DNA to prebind nonspecific DNA-binding proteins. If further incubations (e.g., antibody binding) are required or background is not a problem, use the shorter 30-min incubation. 7. Add 1 p1 (approximately 30,000 cpms) of labeled probe. Finger-mix, spin briefly and incubate on ice for an additional hour. Specific activity of the probe should be approximately 30,000 cpmshg of labeled 20-mer. Using an excess of probe is important to allow detection of low-affinity interactions. Otherwise, most of the probe will be sequestered in high-affinity complexes and loweraffinity interactions may be missed. 8. During incubation, prerun a 5% nondenaturing acrylamide gel in 0.25X TBE in the cold room for 1 hr at 100V. 14 cm X 14 cm X 1 mm vertical gels are convenient for EMSAs. The polyacrylamide percentage of the gel may be varied to optimize resolution between two or more DNA-protein complexes. 9. Rinse wells with a syringe or Pasteur pipet to clear unpolymerized acrylamide. Load dye into two outside wells to allow progress of electrophoresis to


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be monitored. Load samples. Do not add loading dye to samples. There is enough glycerol in the samples to sink them. 10. Run gel in the cold room for approximately 2.5 hr at 250 V. Gel should be run until the bromophenol blue is 4 cm from the bottom of the gel. To allow comparison of binding complexes between gels, be consistent with run length once the optimal time is established. There is no need to recirculate high-ionicstrength buffer. Running the gel in the cold room permits the use of higher running voltages. In addition, colder temperatures cause a contraction of the gel and an increase its sieving properties. This may result in sharper bands. 11. Dry gel and place on film. To prevent gel from cracking, prewet filter paper before lifting gel off of the glass plates. Fixing in methanoYacetic acid is not necessary.

C.EMSA Protocol: Low-Ionic-Strength Conditions The procedure for performing EMSAs under low-ionic-strengthconditions is essentially the same as the preceding protocol with the following exceptions: 1. Use low-ionic-strengthgel and buffer. 2. Recirculate the buffer during prerun and run. This may be done with a pump or manually. Recirculation prevents polarization of low-ionic-strength buffers.

D. Reagents 1. 1OX T4 Kinase Buffer

500 mM Tris-C1, pH 7.6 100 mM MgC12 50 mM D l T 1 mM spermidine HC1 Make fresh prior to each use. 2. 2X Binding Buffer (2X BB)

40 mM Hepes, pH 7.9 100 m M KC1 2 mM P-mercaptoethanol 20% glycerol Important: Stock solutions of 2 X BB without P-mercaptoethanol (P-ME) may be stored at room temperature. Add p-ME just prior to use.


3. 5% Native Polyacrylamide Gel (High-Ionic-Strength) Mix: 5 ml40% acrylamide (29 acrylamide:1 bisacrylamide) 4 d 5 0 % glycerol 2 ml5X TBE (0.25X final) dHzO to 40 ml

Then add: 250 p1 10% ammonium persulfate 30 pl TEMED

Swirl and pour. No need to degas.

4. High-Ionic Running Buffer (0.25X TBE) 22.5 mM Tris-borate (pH 8.3)

0.5 mM EDTA

5. 5% Native Polyacrylamide Gel (Low-Ionic-Strength) Mix: 5 d 4 0 % acrylamide (29 acrylamide:1bisacrylamide) 4 ml50% glycerol 270 pl 1M TriS-C1, pH 7.9 80 pl0.5 M EDTA 132 pl 1 M sodium acetate, pH 7.9 dHzO to 40 ml

Then add: 250 pl 10% ammonium persulfate 30 pl TEMED

Swirl and pour. No need to degas.

6. Low-Ionic Running Buffer 26.9 ml 1 M Tris-C1, pH 7.9 13.2 ml 1 M sodium acetate, pH 7.9

8 m10.5 M EDTA, pH 8.0 dHzO to 4 liters

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E. Competition Experiments The use of binding site oligos containing single point mutations offers a rapid means to determine which nucleotides within the site of interest are essential for recognition and binding. By varying the amount of cold competitor added to a constant amount of labeled binding site probe, the relative affinity of each binding protein for the binding site in question can be determined. Relative affinity is often measured as the molar excess of cold competitor required to reduce binding of the labeled probe by 50% (Ausubel et al., 1995). F. Antibody Supershifts Another variation on the EMSA employs specific antibodies to determine the composition of protein-DNA complexes. Binding site sequence may suggest that a complex contains one or more previously identified factors. Antisera against these known factors can be used to determine whether certain mobility shifts are the result of binding of these candidate proteins. The effect of an antiserum on the mobility of a given protein-DNA complex depends on the epitope recognized by the antibody. If the antibody binds to a region of the protein required for DNA binding, DNA binding will be blocked and the complex will be eliminated. If, however, the antibody recognizes a protein epitope not involved in DNA binding, the mobility of the resulting antibody-protein-DNA complex will be less than that of the original protein-DNA complex, supershifting the original complex to a higher position on the gel (Fig. 2).

1. Protocol for Antibody Supershib Antibody supershift experiments follow the general EMSA protocol outlined earlier, with the following modifications: a. Perform prebinding incubation (carrier DNA, protein, buffer, BSA and water) on ice for 30 min. b. Once prebinding is complete, add 1 pl of the appropriate antiserum and continue incubation on ice for an additional 45 min. Typically 1 p1 of polyclonal antiserum or monoclonal antibody is used per binding reaction. Optimal antibody concentrationsshould be determined empirically.Prior to use with crude extracts, specificity of the antibody should be determined. Figure 2 shows a supershift experiment demonstrating the specificity of a MyoD anti-peptide antibody. c. Continue as in general protocol.

VI. DNase I Protection Assays: Overview and Important Parameters The DNase I protection assay was originally developed as a qualitative technique to localize specific protein-binding sites within transcriptional regulatory regions

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MvoDElPR mvown:El2R M : E 1 2 R

Fig. 2 Antibody supenhift experiment demonstrating the specificity of an anti-peptide (amino acids 197-211) MyoD antiserum. The designated proteins were produced in reticulocyte lysates and subjected to EMSA using an E-box oligonucleotide probe. The MyoD antibody quantitatively supershifts the MyoDE12RDNA complex but does not recognize Myf5:ElZR or Myogenin:E12R complexes. Note that under the conditions used, neither the MRFs alone nor E12R alone bind to the E-box probe. pre, preimmune serum. E12R, a truncated f o p of El2 (Murre et al., 1989a).

(Jackson et al., 1993). Since that time it has been further refined to allow determination of individual and cooperative site-bindingcurves for site-specificproteinDNA interactions. Such quantitative analyses are beyond the scope of this chapter. Interested readers should consult Ausubel et al. (1995) for a detailed description of both the theory and techniques involved. The DNase I protection, or footprinting assay, is based on the ability of DNase I to cleave unbound DNA but not regions of DNA that are bound (protected) by proteins. One of the two strands of a double-stranded DNA fragment is radiolabeled at one end only. Limited degradation of unbound probe by DNase I (each molecule on average is cleaved only once) results in a ladder of DNA fragmentsthat can be visualized by denaturing polyacrylamide gel electrophoresis


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and subsequent autoradiography (Fig. 3). This protection results in a gap in the DNA fragment ladder seen by autoradiography. Comparison of DNase I cleavage patterns with control samples (BSA in place of nuclear extract) and with Maxam and Gilbert sequencing ladders allows accurate determination of binding site locations. When performing DNase I protection assays using nuclear extracts, it is first necessary to establish assay conditions that allow maximal protection of proteinbound sequences. As described in the section on EMSAs, these conditions must allow detection of specificprotein-DNA interactions without permitting substantial nonspecific binding to occur. As the strength of protein-DNA interactions will vary between sites within the fragment of interest, it is not realistic to expect complete footprinting of all sites under one set of assay conditions. Several of the important variables involved in DNase I footprinting are examined next. A. Carrier DNA

All of the aforementioned camers, both complex heterologous DNA (E. coli, salmon sperm, calf thymus) and synthetic alternating copolymer DNA (poly(dI/ dC), poly(dA/dT), poly(dG/dC)) have been employed successfullyin footprinting experiments using nuclear extracts. In addition to blocking nonspecific probeprotein interactions, the addition of excess carrier DNA serves to keep the concentration of DNase I substrate constant. Carrier DNA concentrations that are too high, however, can interfere with specific protein-DNA interactions. Typically between 0.5 and 2 pg of carrier is included in each binding reaction. It is best to titrate carrier DNA concentration for each regulatory region in question. Again, as the carrier requirement may vary from site to site within the region of interest, the eventual carrier type and concentration chosen will likely represent a compromise between individual binding sites. B. DNase I

Ideally, the fragments generated by DNase I cleavage should be present in equimolar amounts. This corresponds to an average of one single-stranded nick per DNA molecule. To achieve single-hit nicking, it will be necessary to vary the DNase I concentration severalfold. DNase I activities will vary from company to company and from lot to lot, so titration should be performed for each DNase preparation to be used. Use one lot of enzyme for a series of experiments. Ideally, the enzyme used in footprinting assays should cleave naked DNA at random. DNase I, however, shows some sequence preferences. This makes it essential to include BSA controls (no nuclear extract) in each footprinting assay. Comparison of experimental to control lanes will indicate which footprints represent actual bound DNA (only experimental DNA is footprinted), and which apparent footprints may have arisen from nonrandom DNase I cleavage (both experimental and control lanes show gaps in the DNA cleavage ladder). In the


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EQwmbmd

Fig. 3 DNase I protection assay of the 258-bp human myoD core enhancer using 20 pg of nuclear extract from myogenic (C2C12, HMP8) and nonmyogenic (10T112, FC1010, JEG-3) cell types. Five protected regions (boxes) and multiple hypersensitive sites (asterisks; the JEG-3-specific hypersensitive site is denoted by a +) were detected. JEG-3 extract does not protect region 2 and yields partial protection of region 1 (most apparent on forward strand). FClOlO and JEG-3 extracts exhibit substantially different patterns of DNase I hypersensitive sites compared to the other cell types. 10T1/2 nuclear extract yields the same footprinting profile as C2C12 and HMP8 myoblast nuclear extracts. Protected region 1 on the reverse strand is not very apparent due to band compression near the top of the gel. BSA, control lanes in which bovine serum albumin replaced nuclear extract. G+A, purine-specific Maxam and Gilbert sequence ladder. From Goldhamer er al. (1995). Development 121,637-649. Reproduced by permission, Company of Biologists LTD.

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latter case, DNase I sequence preferences may mask true footprints. Other protection methods that do not show sequence preferences (e.g., chemical cleavage by hydroxyl radicals or methidiumpropyl-EDTA-Fe( 11)) may be used to study these regions (Lavoie et d.,1991). C. Binding Conditions Buffer conditions, including pH and salt concentration, affect the binding kinetics of protein-DNA complexes. Please refer to the EMSA binding conditions section for a discussion of these variables.

D. Probe

DNase I protection assays are typically conducted with relatively large DNA fragmentsthat may span an entire putative regulatory region, affording an overall view of protein binding sites in large, potentially complex regulatory regions. The use of long probes can be of particular importance when interactions between adjacent sites and their respective bound factors are required for stable DNAprotein interactions. In DNase I protection assays, the DNA to be footprinted is end-labeled at either the 5' or 3' end of the fragment. The positions of protected regions in relation to the labeled end can then be determined with appropriate standards. Ideally, binding sites should be at least 25 to 100bp from the fragment termini to allow for adequate resolution of the DNase I-generated fragments. Analyzing both the forward and reverse strands of the fragment of interest will allow differences between protein interactions involving forward and reverse strands to be identified. In contrast to EMSAs, where probe should be in excess, the amount of probe added to DNase I protection assays should be limiting. Binding sites present on excess probe will be cleaved by DNase I, obscuring potential protein-DNA footprints. The high level of binding site occupancy required for visualization of footprints makes DNase I footprinting a relatively insensitive assay.

VII. DNase I Protection Assay Protocol A. Probe Preparation

It is important to achieve the highest specific activity possible, as unlabeled DNA will compete with labeled probe for protein binding. Efforts should be made to use highly purified DNA and fresh enzymes and buffers. We routinely purify plasmid DNA containing the fragment of interest through two sequential cesium chloride gradients. Avoid exposure to U V light to minimize nicking of the DNA fragment.


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1. DNA Preparation It is best to prepare enough plasmid DNA to allow multiple kinasing reactions. a. Digest 20pg of plasmid DNA with a restriction enzyme that cuts either at the 5’ end (for labeling the forward strand) or the 3’ end (for labeling the reverse strand) of the insert to be footprinted. Digest overnight to ensure quantitative digestion. Use enzymes that give 5‘ overhangs, as these are the most efficiently labeled. Use excess enzyme to ensure complete digestion. A typical 50-p1 digestion might include 20pg DNA, 5 p11OX restriction buffer, and 50-100 U restriction enzyme. b. Add 2 p1 of 1U/p1 calf intestinal alkaline phosphatase (CIAP). Incubate for 2 hr at 37°C. As CIAP is active in most restriction buffers, it is not necessary to phenolkhloroform extract or exchange reaction buffers prior to adding phosphatase. c. Add an additional 2 p1 of CIAP and again incubate for 2 hr at 37°C. Complete phosphatasing is extremelyimportant, as unphosphatased DNA cannot be kinased and will behave as competitor during the footprinting assay. d. To completely inactivate the CIAP, add 5 pl 0.5 M EDTA and incubate at 70°C for 20 min. e. Purify DNA by phenokhloroform extraction followed by chloroform-only extraction. Precipitate with 3 M sodium acetate and 100% EtOH. Precipitate several hours at -20°C to ensure quantitative recovery. Alternatively, DNA can be purified with a cleanup column such as the Wizard column (Promega). f. Rinse pellet several times with 70% EtOH, dry, and resuspend in 40 pl TE. Final concentration should be approximately0.5 pglpl. Run 1-2 pl on an agarose gel to confirm complete digestion.

2. Kinasing Reaction Use high-specific-activity [y-32P]ATP(6 to 7000 Ci/mmol) and a minimum of a 5- to 7-fold molar excess of [y-”P]ATP to DNA ends to ensure highefficiency labeling. a. Add in the following order (30 p1 final reaction volume): 4 pl DNA (2 pg DNA) 3 p1 1OX T4 polynucleotide kinase buffer 18 pl dH’0 2 p1 [y-32P]ATP(6 to 7000 Ci/mmol; approximately 150 pCgp1) 3 pl T4 polynucleotide kinase (20 U/pl) b. Incubate for 45 min at 37°C. Stop reaction by adding 1p1 of 0.5 M EDTA. c. Remove free nucleotides and exchange buffer. We find it convenient to use premade Sephacryl spin columns (Pharmacia S-200 Microspin Sephacryl HR gel filtration columns). Follow manufacturer’s directions for probe purification.

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d. Count 1 p1 of the purified probe and determine the efficiency of labeling. Compare actual labeling to theoretical labeling using pmol of DNA ends and specific activity of the isotope to determine the theoretical yield. (See p. 5.68 of Sambrook et al., 1989, for conversion factors.) If labeling worked well, at least 80 to 90% of the ends should be labeled.

3. Probe Liberation To liberate the labeled probe, digest the linearized plasmid just produced with a second enzyme that cuts at the opposite end of the fragment of interest. a. To the purified probe, add: 10 p1 appropriate 1OX restriction buffer 50-100 U restriction enzyme dH20 to 100 pl final volume b. Digest for several hours at 37°C.

4. Probe Purification a. Add 20 p16X standard nondenaturing loading buffer. b. Separate labeled probe from labeled vector on a 5% nondenaturing polyacrylamide gel. Electrophorese in 1 X TBE at 100 V. A 8.5 cm X 10 cm minigel works well here, allowing adequate separation of probe from vector. The entire digest volume (120 pl) will fit in one 1.5-mm well of a five-well comb. For a 250 bp fragment, electrophorese at room temperature for approximately 1hr at loo v. 5. Probe Elution a. Disassemble gel apparatus. Remove top plate of gel sandwich, allowing gel to remain on the longer bottom plate. Wrap gellplate with Saran Wrap. b. Visualize the labeled fragments by autoradiography (a 30-sec to l-min exposure should be sufficient), align the autoradiogram with the gel, and excise the appropriate band with a clean razor blade. c. Electroelute DNA in 1X TBE using either an electroeluter or dialysis tubing placed in a horizontal gel box. Electroelution yields are much better than yields that employ overnight elution in typical ammonium acetate buffers. If using a commercial electroeluter, follow manufacturer’s instructions. To electroelute using dialysis tubing, place gel into 0.5-inch-diameter dialysis tubing with forceps (prepare dialysis tubing according to manufacture’s instructions). Add 0.5 ml of 1 X TBE to tubing and clamp open end. Place dialysis tubing into a horizontal minigel box, perpendicular to the direction of current flow. Electroelute at 100 V for approximately 3 hr. Reverse current for 1 min to release DNA from

46 1


the membrane, then transfer the buffer containing the eluted probe to a fresh 1.5-ml microfuge tube. Rinse the dialysis tubing with an additional 0.5 ml of 1X TBE, and add this rinse volume to the original eluate. Verify complete electroelution by determining the counts remaining in the gel slice and dialysis tubing. Greater than 90% elution should be achieved by this method. d. Microcentrifuge at top speed to pellet any acrylamide particles. Transfer supernatant to a fresh tube. e. Precipitate labeled DNA with 100%EtOH and 3 M sodium acetate. Precipitate either at -20°C overnight or at -70°C for several hours. Follow radioactivity with a Geiger counter to ensure that the probe pellet is not lost during the ensuing wash steps. f. Wash pellet twice with 70% EtOH. Dry in a Speed-Vac for several minutes. g. Resuspend pellet in TE (10 mM Tris, pH 8.0, 1 mM EDTA, pH 8.0) to a final activity of lO,OOo/pl. Store at 4°C. h. Use 1 pl of probe (approximately 1.4 fmol for a 250-bp fragment) per protection assay. We use probes that are up to one half-life old. Continue to use 1 p1 of probe per protection assay despite the decrease in specific activity that occurs. As the amount of protein used per protection assay is determined by titration against a fixed molar amount of probe, do not increase the amount of probe used. B. DNase I Protection Protocol

The following protocol has been used successfully in our laboratory to yield footprints of the 258-bp myoD core enhancer by nuclear extracts (Goldhamer et al., 1995). We give concentrations and conditions that have proved successful in our hands. Optimize protein, competitor, salt and DNase I concentrations by varying each component individually. 1. Assemble 50 p1 binding reactions: BSA control dHzO to 50 p1 5 pl lox binding buffer 5 p1 600 mM KCI 1 pl 1 mg/ml poly(dI/dC) 10 p1 10%polyvinyl alcohol (PVA) 5 pl 1 mg/ml bovine serum albumin (BSA) 1 pl end-labeled probe

Experimental dHzO to 50 pI 5 1 1 1OX binding buffer 4 p1600 mM KCI 1 pl 2 mg/ml poly(dI/dC) 10 p1 10%PVA 20 pg nuclear extract 1 pI end-labeled probe

Protein and carrier concentrations should be tested in preliminary experiments. Thaw extracts on ice just prior to use. We routinely set up three reactions per experimental sample (to allow for three different DNase I concentrations) and a total of three control reactions (without nuclear extract) per experiment. Therefore, for n different experimental

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&isten L. Kucharczuk and David J. Goldhemer

samples you will have 3n + 3 reactions. It is easiest to make up experimental and control master mixes. 2. Incubate binding reactions on ice for 1 hr. 3. Just before beginning digests, dilute DNase I stock (Worthington-1 mgl ml) into ice-cold 50%glycerol. DNase I concentration should be titrated for each new batch of extract. We routinely use three DNase I concentrations with the DNA alone controls (e.g., 1/200,1/400,1/800)and three DNase I concentrations with each experimental sample (e.g., 1/10, 1/20, 1/40). Dilutions given are stock dilutions, not final DNase I concentrations. Samples containing nuclear extract will require more DNase I than controls because of inhibitors present in the extract. The use of high-quality DNase I is critical. Store at -20°C in 50-p1 (lmgl ml) aliquots. Do not reuse DNase Z once thawed. Pipet gently to mix. 4. Equilibrate two reactions at a time to room temperature (1 min) and add an additional 50 p1 of binding buffer (room temperature with fresh DTT) to each reaction. After exactly 1min, add 1pl dilute DNase I to each tube. Pipet gently to mix.After exactly 2 min, terminate the reactions by adding 100 p1 stop buffer to each. Repeat for the remaining tubes. 5. Add 5 pl of 3 m g / d proteinase K to each reaction. Incubate tubes at 37°C for 30 min. 6. Extract samples with an equal volume of cold phenol. To keep the aqueous layer clear to facilitate extraction, add cold phenol, vortex, and incubate on ice for 5 min. Microfuge at 4"C, then place at room temperature until ready to transfer aqueous layer. Be sure to avoid the aqueous/organic interface. The use of phenol :chloroform at this step is not advised, as it yields a thick flocculent interface that traps much of the radioactivity. 7. Ethanol precipitate the DNA fragments at room temperature for 5 min. Do not precipitate at -20°C.The salts which come out of solution at -20°C will cause the sample to run anomalously on the gel. 8. After centrifugation,wash the DNA pellet twice in room-temperature 70% EtOH and dry in a Speed-Vac. 9. Resuspend in 8 pl of formamide/EDTA loading buffer. 10. Prerun a 0.4-mm 6% polyacrylamide sequencing gel for 30 min at 60 W. Flush wells with a syringe or Pasteur pipet before prerunning and again prior to loading samples. Up to approximately 250 nucleotides can be visualized on 6% nongradient sequencing gels. The optimal acrylamide percentage should be determined empirically as it will depend on the positions of the footprints and the size of the DNA fragment. Gradient gels may be used to expand the useful range of the gel. 11. Heat samples to 80°C for 3 min and load 4 pl of each reaction on a 6% denaturing polyacrylamide sequencing gel. Load 2 to 3 pl of G + A sequencing ladder in one lane. 12. Electrophorese at 60 W for approximately 1.5 hr. Dry gel and expose to film with intensifier screens for 2 to 4 days. Optimal run time will, of course,


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depend on the length of the fragment, amount of polylinker, and position of the footprints. Adjust as necessary. C. Preparation of G

+ A Ladder: Maxam and Gilbert Reactions

Maxam and Gilbert sequencing reactions of the footprinting probe should be run alongside of DNase I cleavage reactions to allow the specific nucleotide represented by each band in the cleavage lanes to be determined. Comparison of cleavage reactions with a G + A sequencing ladder is essential for precise mapping of regions of DNase I protection. The following produces enough G + A ladder to use as a standard for 6 to 10 experiments. 1. Mix together: 10 p1 end-labeled probe (l00,OOO cpm) 4 pl salmon sperm DNA (1 pg/p1) 11 pl dHzO 2. Add 4 p1 1 M sodium formate. Incubate for 15 rnin at 37°C. 3. Add 240 p1 stop solution and place reaction on ice. 4. Add 750 p1 of -20°C 100%EtOH and incubate for 15 min on ice. 5. Microcentrifuge at top speed for 5 min at room temperature. Remove as much supernatant as possible without disturbing the pellet. Do not dry pellet. 6. Resuspend pellet in 300 p1 0.3 M sodium acetate, pH 5.2. 7. Add 600 pl 100% EtOH and incubate for 15 min on ice. 8. Centrifuge at top speed for 5 min and discard supernatant. 9. Wash pellet twice in 70% EtOH. Dry pellet. This is a convenient stopping point. Pellet can be frozen overnight at -20°C. 10. Resuspend pellet in 100 pl 1 M piperidine and transfer to a screw-top 1.5-ml tube. Incubate for 30 min at 90°C. 11. Cool reaction to room temperature. Spin briefly in a microfuge to collect all of the reaction in the bottom of the tube. Evaporate to dryness in a Speed-Vac. 12. Add 100 pl dH20 to dry the pellet, vortex, and gain evaporate to dryness in a Speed-Vac. 13. Add 50 pl dH20, vortex, and again evaporate in a Speed-Vac. 14. Resuspend in 20 p1 of formamideEDTA loading buffer. 15. Load 2 to 3 p1 of completed G + A sequencing reaction. D. Reagents 1. 1OX Kinase Buffer Make fresh from stock solutions just before use. 0.5 M Tris-C1, pH 7.6 0.1 M MgC12


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50 mM dithiothreitol 1 mM spermidine HCl 1 mM EDTA, pH 8.0

2. 5% Polyacrylamide Gel This is enough for 1 mini-gel for probe purification. 2.5 ml 1OX TBE 3.12 ml40% acrylamide (19 acrylamide:1bisacrylamide) 19.25 ml dH20 125 p1 10% A P S 25 p1 TEMED

3. 1 mg/ml Salmon-Sperm DNA Resuspend salmon-spermDNA (Sigma) in water at 10mglml for stock solution. Shear DNA by autoclaving for 15 min [at 21 psi]. Remove 0.5 mg of this 10 mg/ ml stock, extract twice in phenol :chloroform (1:l), extract again in chloroform only, ethanol precipitate, wash twice in 70% EtOH, and dry. Resuspend DNA pellet in dH20, quantitate by spectrophotometry, and adjust concentration to 1mg/ml.

4. 1M Sodium Formate Add: 435 pl 100% formic acid (Fisher) 2 ml dHzO

pH to 2.0 with 1M NaOH (approximately 4 ml). Bring volume to 10 ml with dH20.

5. 1 M Piperidine Dilute 100% piperidine (10 M) with dH20.Piperidine is very toxic. Use caution when handling.

6. Stop Solution for G

+ A Sequencing

0.3 M sodium acetate, pH 7.0 0.1 mM EDTA 100 pg/ml yeast tRNA


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7. DNase I Stock Resuspend lyophilizedpancreatic DNase I (Worthington)in dHzO to a concentration of 1 mg/ml. DNase I is unusually sensitive to physical denaturation. Mix by gentle inversion. 8. 10 Binding Buffer 120 mM Tris-C1, pH 8.0 10 mM MgC12 50 mM NaCl 10 mM CaC12 1 mg/ml bovine serum albumin (BSA) 1 mM dithiothreitol (D'IT) 50% glycerol

Add D'IT and BSA just before use.

9. Stop Buffer for DNase I Protection 50 mM EDTA 0.2% SDS 100 pg/ml yeast tRNA

10. Formamide/EDTA Loading Buffer 38% formamide 8 mM EDTA 0.02% bromphenol blue 0.02% xylene cyanol

E. Notes 1. Although the nuclear extract preparation protocol just given generates reproducible preparations of nuclear extracts, extractsprepared at different times from different batches of cells will vary somewhat. To venfy reproducibility of results, we suggest that at least two independently prepared nuclear extracts of each cell type be tested. 2. The binding of proteins to a given DNA sequence can enhance as well as prohibit cleavage by DNase I. This enhancement, known as DNase I hypersensitivity, likely results from a change in DNA conformation brought about by


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protein binding. Although hypersensitive sites often lie just outside of footprinted regions, other hypersensitive sites are unassociated with detectable protection. These sites may reflect interactions with proteins of lower abundance or affinity. Comparison with control digests (no nuclear extract) allows novel hypersensitive sites to be distinguished from cleavage sites preferred by DNase I in the absence of protein binding.

VIII. Relevance of the EMSA and DNase I Protection Assay to Muscle Research

Cis and trans analyses of muscle promoters and enhancers offer a powerful means to define upstream signaling pathways and transcriptional events that govern muscle gene expression in cell culture and the developing embryo. As a first step towards dissecting these transcriptional regulatory pathways, cis-acting transcriptionalcontrol regions must be functionally identified. In muscle research, this identification is predominantly done in muscle cell culture systems (see Chapters 5-8 and 19) and, more recently, in transgenic mice (see Chapter 18). Once relevant sequences have been identified, the in v i m techniques described in this chapter can be used to characterize these regulatory sequences and the proteins that bind to them. We present a few examples from the wealth of muscle research to illustrate how these in vitro techniques may be used to further our understanding of the transcriptional mechanisms controlling myogenesis. A. In vitro Techniques using Crude Nuclear Extracts

1. EnhancedPromoter Mapping Once relevant sequences have been at least broadly defined by functional assays, the in vitro techniques described in this chapter can be used to further refine important regulatory sequences. Together with sequence comparison between closely and more distantly related species, EMSAs and DNase I protection assays can delineate potential transcription factor binding sites. We have utilized all three of these approaches in our analysis of the distal myoD core enhancer. Transfection and transgenic analyses originally identified a 4 kb enhancer fragment located 20 kb upstream of the myoD gene body (Goldhamer et aZ., 1992). Deletion analysis localized this activity to a 258 bp fragment that recapitulates tissue- and stage-specific expression of the endogenous myoD gene within the developing mouse embryo (Goldhamer et aZ., 1995). In the hopes of finding discrete regions of homology within this region, and thus identifying potential transcription factor binding sites, a sequence comparison between mouse and human enhancer sequences was undertaken. Surprisingly,overall sequence identity between these two species is 89% (Goldhamer et aZ., 1995). Because sequence similarity across the entire enhancer sequence is high, an unbiased DNase I protection approach was chosen to map protein-binding sites


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within the enhancer (see Fig. 3). Experiments involving both muscle and nonmuscle nuclear extracts revealed multiple nuclear protein-binding sites in the core enhancer, none strictly muscle-specific (Goldhamer et al., 1995). To gain additional information regarding the number, relative size, and tissue specificity of the protein complexes binding to these sequences, EMSAs employing synthetic oligonucleotides corresponding to these protected regions have been performed. To date, these experiments have shown that the binding activities present in muscle and nonmuscle nuclear extracts exhibit identical migration behaviors (Kucharczuk and Goldhamer, unpublished observations). Further studies examining the binding sequence requirements of each of these activities will help to determine their relatedness. To test the relevance of these binding sites and activities, enhancer constructs harboring mutations spanning these bound regions have been linked to lacZ reporter genes. These constructs are currently being tested for activity in transgenic mice. Some of these mutations have profound general effects on reporter gene expression in vivo, whereas others have quite limited, lineage-specificeffects (Kucharczuk, Dougherty, and Goldhamer, unpublished observations).

2. Dissection of Regulatory Pathways using Nuclear Extracts Using nuclear extracts and the in vitro techniques described earlier, it is possible to dissect transcriptional regulatory pathways without actually having the purified transcription factors in hand. Much has been learned about muscle gene regulation through in vitro analyses of cell specificity and developmental-stage specificity of protein-DNA complex formation. In the original MEF-2 paper, Gossett et al. (1989) used EMSAs to identify and characterize a myocyte-specific enhancerbinding factor (MEF-2) that binds to the 5’ muscle-specific MCK enhancer. This activity was undetectable in myoblasts, but became detectable within 2 hr of exposure to differentiation medium. Treatment of myoblasts with various mitogens (FGF, TGF-P) known to block induction of muscle-specific genes blocked MEF-2 binding. MEF-2 subsequently has been cloned and shown to regulate the muscle regulatory gene, myogenin (Cheng et al., 1993; Edmondson et al., 1992), which is required for muscle differentiation in vivo (Hasty et al., 1993; Nabeshima et al., 1993). In vitro assays using nuclear extracts can also be used to determine the molecular mechanisms governing transcription of muscle-specific genes. One example can be seen in the study of F-ACT1 and serum response factor (SRF), nuclear activities that bind serum response elements (SREs) present in the chicken skeletal a-actin promoter (Lee et al., 1991). Nuclear extracts taken from different stages of myogenic and nonmyogenic cell cultures were used in EMSAs to assess relative levels of F-ACT1; F-ACT1 was found to be enriched in nonmuscle tissues and replicating undifferentiated myoblasts, and reduced as myoblasts fuse and become differentiated myotubes. In contrast, skeletal cu-actin expression is upregulated following fusion, Together, these observations led to the proposal that

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Kriaten L. Euchprczuk and David J. Goldhamer

F-ACT1 functions as a repressor of skeletal a-actin gene expression. To determine whether F-ACT1 and SRF binding to SREs are mutually exclusive, EMSAs were performed under limiting probe conditions (Lee et al., 1991). By gradually changing the protein ratio in favor of one factor, the binding of the second factor could be titrated out. This observation led to the hypothesis that the proposed repressor F-ACT1 and the transcriptional activator SRF compete for binding of the SRE,and that the ratio of F-ACT1 activity to SRF activity modulates skeletal a-actin expression during myogenesis. F-ACT1 has subsequently been identified as the transcriptional repressor IT1 (Lee et al., 1992), confirming this original proposal.

3. Cloning Transcription Factors using Data &om In Vitro Assays Binding sites identified by EMSA and DNase I protection assays have been used to clone novel DNA-bindingproteins through a technique known as expression cloning. This technique entails screening cDNA libraries for cDNAs that encode proteins able to recognize and bind to a radiolabeled binding site of interest. The mesodermally restricted homeobox protein, MHox, was cloned in this manner (Cserjesi et al., 1992). MHox was originally identified by EMSAs as a binding activity that bound an A+T-rich element within the MCK enhancer. To identify the factors that interacted with this motif, Cserjesi and colleagues screened a C2 myotube cDNA library for clones that encode proteins able to recognize the A+T-rich element, but not the closely related MEF-2 binding site, nor a mutant A+T-rich element not bound by nuclear extract proteins. Once identified, the M H o x cDNA was shown by EMSA to encode a protein with the same DNA-binding properties observed for the A+T-rich binding factor present in nuclear extracts. For a detailed expression cloning protocol, refer to Singh et al. (1989). B. In vitro Techniques using Purified Proteins In addition to facilitating the identification of novel DNA-binding activities in cell extracts,EMSAs and DNase I protection assays have been used extensively to characterize previously cloned transcription factors and the DNA sequences to which they bind. I n vitro binding techniques have been of particular value to the muscle field for the definition of consensus binding sequences within muscle promoters and enhancers, and for the mapping and characterization of protein dimerization and DNA-binding domains.

1. Consensus Sequence Definition Sequence requirements for binding of purified proteins are typically determined in one of two ways. Most often, a combination of binding-sitemutagenesis and EMSAs is used. Syntheticoligonucleotides in which one or more nucleotides


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diverge from the original sequence are generated and are used in EMSA competition experiments. To define MHox DNA-bindingproperties and determine DNA sequence requirements for MHox binding, bacterially expressed MHox protein and the labeled A+T-rich element used to identify the MHox protein were subjected to a series of EMSA competition experiments in the presence of a variety of A+T-rich oligonucleotide competitors (Cserjesi et al., 1992). This competition analysis demonstrated that the bacterially expressed MHox protein bound to a nine-nucleotideDNA sequence composed solely of A and T residues. The observation was made that, although the MCK MHox binding site is distinct from the similar, A+T-rich MCK MEF-2 binding site, a subset of binding sequences can bind both MHox and MEF-2. This finding has fueled an ongoing investigation of the combined effect of MHox and MEF-2 factors on MCK enhancer activity (Cserjesi et aL, 1994) and has provided a mechanism through which a single DNA sequence can mediate transcriptional regulation by more than one class of DNA-binding proteins. When little or nothing is known about the recognition site of a transcription factor, unbiased approaches beginning with degenerate oligonucleotides can be used to determine consensus binding sites (Wright et al., 1991). In one approach, oligonucleotides bound by the factor of interest are detected in EMSAs, excised and purified from the acrylamide gel, amplified by PCR, and subjected to several more cycles of binding and amplification (Blackwell and Weintraub, 1990). Ultimately, these oligos are sequenced as a pool. The resulting consensus sequence can be used to compare the binding site preferences of one or more factors.

2. Protein Domain Mapping EMSAs enable the dissection of protein domains involved in DNA binding, transcriptional activation, and protein-protein interaction. The ability of diverse HLH proteins to recognize a consensus DNA sequence suggested to early investigators that certain conserved amino acids within the muscle regulatory factors (MRFs) would be required for recognition of consensus nucleotides, while other divergent amino acids would confer unique roles in myogenesis. To pinpoint the residues required for DNA binding and muscle-specific gene activation, Davis et al. (1990) examined the effects of site-directed mutations within the MyoD basic domain on DNA binding and oligomerization with E12. Using EMSAs, two basic clusters of amino acids within this basic region were shown to be required for binding to a MCK enhancer E-box. By accompanying the EMSA analysis with immunoprecipitation experiments, they were able to dissociate requirements for DNA binding from requirements for heteroligomerization with the ubiquitous factor E12.

3. Identification of Dimerization Partners EMSAs can be used to study protein-protein interactions as well as proteinDNA interactions. For example, MyoDLE12 heterodimers were first identified

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by EMSAs (Murre et al., 1989a). Earlier studies restricted to El2 and E47 had demonstrated that the HLH region of these proteins was sufficient for dimerization (Murre et aZ., 1989b). As the MRFs, MyoD, Myf-5, MRF4, and Myogenin share this protein motif, the ability of one of these factors, MyoD, to form functional heterodimers with El2 and E47 was examined. Reticulocyte lysate translated MyoD, El2 and E47 were analyzed by EMSAs for their ability to bind an E-box contained within the kappa immunoglobulin enhancer (Murre et al., 1989a). Visualization of novel, slower-migrating MyoD/E-proteidprobe complexes in gel-shift assays confirmed the suspicion that the amphipathic helices of both the muscle-specific MRFs and the ubiquitous E-proteins could mediate heterodimerization.

C. Concluding Comments The protocols and examples given in this chapter demonstrate that the electromobility shift assay and DNase I protection assay are powerful tools for the identification and study of the DNA-binding proteins involved in myogenesis. When used in conjunction with nuclear extracts, these techniques can map muscle-specificpromoters and enhancers, determine precise DNA sequence requirements for protein binding, yield information concerning the developmentalstage and tissue specificity of DNA-binding activities, and facilitate the cloning of novel DNA-binding factors. Once DNA-binding proteins have been cloned, these same techniquescan be used to characterize both the proteins and the DNA sequences to which they bind. When interpreted in the light of corroborating functional studies, EMSAs and DNase I protection assays provide a strong foundation for our understanding of the transcriptional mechanisms driving muscle differentiation and determination. Although EMSAs and DNase I protection assays are extremely informative, the in vivo relevance of in vitro data should be confumed by mutagenesis, transfections, and transgenic analysis (Chapters 18 and 19). In vttro conditions may allow transcription factors to bind to sites they would not normally occupy in vivo. MyoDE12 dimers, for example, bind to the kappa immunoglobulin enhancer E-box in vitro (Murre et al., 1989a), but do not regulate the kappa gene in vivo. Alternatively, in vitro assays may not reflect developmentalstage-specific or cell-type-specific regulation of binding site accessibility. In addition to these in vitrohn vivo discrepancies, significant differences can exist between transcriptional regulation in cell culture systems and within the developing organism (Kucharczuk and Goldhamer, unpublished observations). Greater utilization of embryonic tissues for both in vitro and in vivo studies, and widespread application of in vivo techniques that examine protein-DNA interactions in the context of living cells (Mueller and Wold, 1989) will help to address these issues.


47 1

Acknowledgment We thank Dr. Youngwon Lee-Nham for conducting the experiment shown in Fig. 2.

References Ausubel, F. M., Brent, R., Kingston, R. E., Moore, D. D., Seidman, J. G., Smith, J. S., and Struhl, K. (1995). In “Current Protocols in Molecular Biology” (K. Janssen, ed.). New York John Wiley & Sons. Blackwell,T. K., and Weintraub, H. (1990). Differences and similaritiesin DNA-binding preferences of MyoD and E2A protein complexes revealed by binding site selection. Science 250,1104-1110. Buskin, J. N., and Hauschka, S. D. (1989).Identification of a myocyte nuclear factor that binds to the muscle-specific enhancer of the mouse muscle creatine kinase gene. Molecular and Cellular Biology 9,2627-2640. Cheng, T.-C., Wallace, M. C., Merlie, J. P., and Olson, E. N. (1993). Separable regulatory elements governing myogenin transcription in mouse embryogenesis. Science 26% 215-218. Cserjesi, P., Lilly, B., Bryson, L., Wang, Y., Sassoon, D., and Olson, E. N. (1992). MHox: A mesodermally restricted homeodomain protein that binds an essential site in the muscle creatine kinase enhancer. Devel. 115,1087-1101. Cserjesi, P., Lay, B., Hinkley, C., Perry, M., and Olson, E. N. (1994). Homeodomain protein MHox and MADS protein Myocyte Enhancer-binding Factor-2 converge on a common element in the muscle creatine b a s e enhancer. J. Biol. Chem. 269, 16740-16745. Davis, R. L., Cheng, P-F., Lassar, A. B., and Weintraub, H. (1990).The Myod DNA binding domain contains a recognition code for muscle-specificgene activation. Cell 60,733-746. Dent, C. L. and Latchman, D. S. (1993). The DNA mobility shift assay. In “Transcription Factors: A Practical Approach” (D. S. Latchman, ed.), pp. 1-26. New York Oxford University Press Inc. Dignam, J. D., Lebovitz, R. M., and Roeder, R. G. (1983). Accurate transcription initiation by RNA polymerase I1 in a soluble extract from isolated mammalian nuclei. Nucl. Acids Res. 11,1475-1489. Edmondson, D. G., Cheng, T. C., Cserjesi, P., Chakraborty, T., and Olson, E. N. (1992). Analysis of the myogenin promoter reveals an indirect pathway for positive autoregulation mediated by the muscle-specific enhancer factor MEF-2. Mol. CelL Biol. 12,3665-3677. Fried, M. and Crothers, D. M. (1981). Equilibrium and kinetics of lac repressor-operator interactions by polyacrylamide gel electrophoresis. Nucl. Acids Res. 5,6505-6525. Garner, M. M., and 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. Nucl. Acids Res. 9, 3047-3060. Goldhamer, D. J., Faerman, A., Shani, M., and Emerson, C. P.Jr. (1992). Regulatory elements that control the lineage-specificexpression of myoD. Science 256,538-542. Goldhamer, D. J., BN&, B. P., Faerman, A., King, A., Shani, M., and Emerson, C. P. Jr. (1995). Embryonic activation of the myoD gene is regulated by a highly conserved distal control element. Development 12%637-649. Gossett, L. A., Kelvin, D. J., Sternberg, E. A., and Olson, E. N. (1989). A new myocyte-specific enhancer-binding factor that recognizes a conserved element associated with multiple musclespecific genes. Mol. Cell. Biol. 9, 5022-5033. Grilli, M., Chiu, J. S.,and Lenardo, M. J. (1993). NF-KB and Rel-participants in a multiform transcriptional regulatory system. Int. Rev. Cytol. 143,l-62. Hasty, P., Bradley, A., Moms, J. H., Edmondson, D. G., Venuti, J. M., Olson, E. N., and Klein, W. H. (1993). Muscle deficiency and neonatal death in mice with a targeted mutation in the myogenin gene. Nature 364,501-506. Imbalzano, A. N., Zaret, K. S.,and Kingston, R. E. (1994). Transcription Factor (TF) IIB and TFIIA can independently increase the a m i t y of the TATA-binding protein for DNA. J. Biol. Chem. 269,8280-8286.

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Jackson, D. A., Rowader, K. E., Stevens, K., Jiang, C., Milos, P.,and Zaret, K. S. (1993). Modulation of liver-specific transcription by interactions between hepatocyte nuclear factor 3 and nuclear factor 1binding DNA in close apposition. MoL CelL BWL l3,2401-2410. Jen, Y.,Weintraub, H.,and Benezra, R. (1992). Overexpression of Id protein inhibits the muscle differentiationprogram: I n vivo association of Id with E2A proteins. Genes and Dev. 6,1466-1479. Lavoie, B. D., Chan, B. S., AUison, R. G., and Chaconas, G. (1991). Structural aspects of a higher order nucleoprotein complex: Induction of an altered DNA structure at the Mu-host junction of the Mu Type I transpososome. EMBO J. 10,3051-3059. Lee, T. C., and Schwartz, R. J. (1992). Differential detection of multiple DNA-binding complexes using dissimilar polyanion competitors. NucL Acid Res. 20,140. Lee. T.-C., Chow, K.-L., Fang, P., and Schwartz,R. J. (1991). Activation of skeletal alpha-actin gene transcription: The cooperative formation of serum response factor-bindingcomplexes over positive cis-acting promoter serum response elements displaces a negative-acting nuclear factor enriched in replicating myoblasts and non-myogenic cells. MoL Cell. BioL ll,5090-5100. Lee, T.-C., Shi, Y.,and Schwartz, R. J. (1992). Displacement of BrdUrd-induced YY1 by serum response factor activates skeletal alpha-actin transcription in embryonic myoblasts. Proc. Narl. Acad. Sci. USA 89,9814-9818. Lichtensteiner, S., Wuarin, J., and Schibler, U. (1987). The interplay of DNA-binding proteins on the promoter of the mouse albumin gene. Cell Sl, 963-973. Milos, P.M., and Zaret, K. S. (1992). A ubiquitous factor is required for UEBP-related proteins to form stable transcription complexes on an albumin promoter segment in vitro. Genes & Dev. 6,991-1004. Mueller, P. R. and Wold, B. (1989). I n vivo footprinting of a muscle specific enhancer by ligation mediated PCR. Science 246,780-786. Murre, C., McCaw, P. S., Vaessin, H., Caudy, M., Jan, L.Y.,Jan, Y.N., Cabrera, C. V.,Buskin, J. N., Hauschka, S. D., Lassar,A. B., Weintraub, H., and Baltimore, D. (1989a). Interactions between heterologous helix-loop-helix proteins generate complexes that bind specifically to a common DNA sequence. Cell SS, 537-544. Murre, C., McGaw, P. S., and Baltimore, D. (1989b). A new DNA binding and dimerization motif in immunoglobulin enhancer binding, daughterless, MyoD and rnyc proteins. Cell 56,777-183. Nabeshima, Y.,Hanaoka, K.,Hayasaka, M.,Esumi, E.,Li, S., Nonaka, I., and Nabeshima, Y.(1993). Myogenin gene disruption results in perinatal lethality because of severe muscle defects. Nature 3aq 532-535. Sambrook, I., Fritsch, E. F.,and Maniatis, T. (1989). “Molecular Cloning: A Laboratory Manual,” 2nd Ed. Cold Spring Harbor: Cold Spring Harbor Laboratory Press. Singh, H., Clerc, R. G., and LeBowitz, J. H. (1989). Molecular cloning of sequence specific DNA binding proteins using recognition site probes. BioTech. 7,252-261. Taylor, M. V., Gurdon, J. B., Hopwood, N. D., Towers, N., and Mohun, T. J. (1991). Xenopus embryos contain a somite-specific, MyoD-like protein that binds to a promoter site required for muscle actin expression. Genes & Dev. 5,1149-1160. Wright, W. E., Binder, M. and Funk, W. (1991). Cyclic amplificationand selection of targets (CASTing) for the myogenin consensus binding site. Mol. Cell BioL ll,4104-4110. Zaret, K. S., Milos, P., Lia, M.,Bali, D., and Gluecksohn-Waelsch, S. (1992). Selective loss of a DNase I hypersensitive site upstream of the tyrosine amhotransferase gene in mice homozygous for lethal albmo deletions. Proc. Nut. Acad. Sci. USA 89,6540-6544.

INDEX

A Actin, for VSMC characterization, 141-143 Adenoviruses augmented gene transfer, 253-254 for gene therapy cytotoxicity, 249 delivery to muscle, 251-253 in host immune response, 250-251 human safety problems, 248-249 inflammation, 249 safety improvements, 249-250 vector capacity, 249-250 gene transfer to muscle in adult mouse, 242-244 in cardiocytes and myocardium, 247-248 human minidystrophin to mdx mouse, 244-246 in newborn mouse, 241-242 promoters, 244 secreted protein production, 246-247 infected ventricular myocytes, myofilament incorporation indirect immunofluorescence with antibodies, 318-320 Western blot analysis, 317-318 mediated gene transfer into adult cardiac myocytes, 309-313 into cardiac muscle experimental design, 430-433 general considerations, 423-426 ex vivo transfer, 254 recombinant construction, 314-316 design cloning site selection, 235-236 replication-deficient vectors, 235 replicative vectors, 234 transcription signal selection, 235-236 DNA preparation, 238-239 general solutions, 314 infection of ventricular myocytes, 313 isolation, 240-241 plaque assay, 316-317 preparation, 238-239 production, 313-314 propagation, 241

storage, 241 titration, 239-240 vector generation, 236-237 for in vivo gene transfer, 231-234 Agar, coated dishes, for embryo culturing, 125 Age related alterations in synaptic maintenance, 343-346 satellite cell animal models, 157 Albumin, removal, in chicken embryo lowering, 19 Alkaline phosphatase assay, &-acting control region, 420-421 ALV, see Avian leukosis virus Anatomical assay, myoblast transplantation, 269 Antibodies monoclonal, in indirect immunofluorescence of myofilaments, 318-320 supershifts in electromobility shift assay, 454 Autoradiography, transgenic mouse tissues, 390 Avian leukosis virus -receptor mouse model amphotropic vectors, 207 generation in vivo, 205-207 in vitro model, 201-202 in vivo targeting to muscle, 202-205 retroviral system based on, 189-193 in vitro model, 196-198 Avian spleen necrosis virus, for cell lineage studies, 218

B Basement membranes, 3D gel, 151-152 Biochemical assay, myoblast transplantation, 270-271 C Calcium tolerance induction in cells, 311-312 transients in myotubes, 287-289 Calcium currents, in myotubes, measurement, 286-287 473

474

Index

Calcium phosphate, mediation of DNA transfection, 407-409 Calponin, for VSMC characterization, 144 Candling, chick embryo, 5 Cardiac progenitors postgastrulation location, 119-120 pregastrulation location, 119 removal, 121-122 Cardiomyocytes adult, adenovirus-mediatedgene transfer, 309-313 Culturing, 126-127 gene transfer into, 247-248 CAT assay, see Chloramphenicol acetyltransferase assay Cell cultures and adenovirus-mediatedgene transfer, 312-313 chick conditioned media for, 100-101 embryo extract preparation, 90-91 primary muscle cells, 96-101 common solutions, 137-138 establishment, 136 freeze-fracture, 303-304 human tissue atheromatous, 149-150 normal tissue, 147-149 mass transfection, 301-303 mouse C, cells conditions, 111 freezing protocols, 111-112 techniques, 111 thawing protocols, 112 muscle mature, 86-87 mouse, 88-90 rat, 95-96 myoblasts collagen-coated petri dish preparation, 277 culture protocol, 277-278 functional assessment, 279 growth and enrichment, 263 matrigel-coated dish preparation, 277 matrigel preparation, 277 preparation, 262-263 quail growth medium, 277 on rat tail collagen, 276-277 structural visualization via immunostaining, 278-279 transfection and selection, 278 quail primary muscle cells,96-101

skeletal muscle satellite cells activation and proliferation, 167 cell preparation, 161-163 culture homogeneity, 163-167 differentiation, 168-170 medium, 159-161 monolayer mass cultures, 158-159, 172-173 single fiber cultures, 174 characteristics, 172-173 fiber isolation, 171-172 substrates and media, 170-171 substrates, 159 ventricular myocytes, maintenance, 313 Cell differentiation defective MM14 cells, 109-110 media for, 104 MM14 cells, 109 models, 150-151 mouse MM14 line culture dishes, 102 differentiation medium, 104 fibroblast growth factor, 105-106 gelatincoated culture dishes, 102-103 growth medium, 104 horse serum pretesting, 104-105 nutrient medium, 103-104 rat satellite cells, 168-170 specific markers for VSMC actin, 141-143 calponin, 144 myosin-heavy chain, 143 vincuWmeta-vinculin, 143-144 Cell expansion, exponential, MM14 cells, 106-107 Cell growth with fibroblast growth factor, 105-106 media for, 104 myoblasts, 263 Cell lineage studies with retroviruses replication-defective retrovirus, 217-218 safety, 222-223 viral harvest and concentration, 221-222 virus-producing cells Cloning, 219-221 frozen stock, 221 skeletal muscle, with retroviruses, in chicken embryos marking muscles, 224-225 precursor tagging, 223-224 vector plasmid for, 218-219

475

Index

Cell lines clonal, mouse myoblasts, 91-95 HEK293, cotransfection, 315-316 MM14 availability, 113 cell differentiation, 102-106 cell propagation, 102-106 differentiation-defectivecells, 109-110 differentiation protocol, 109 exponential expansion, 106-107 freezing, 108-109 passaging cells, 107-108 stable transfection, 413-414 thawing, 108-109 transient transfection, 409 MM14-DD, availability, 113 rat BC3H1 line, 112-113 rat L6 line, 112 skeletal muscle established lines, 101 mouse C, cell line, establishment, 110 mouse MM14 line, derivation, 101-102 virus-producing cloning, 219-221 frozen stock, 221 Cell migration, VSMC, 144-147 Cell passaging, MM14 cells, 107-108 Cell propagation, mouse MM14 line culture dishes, 102 differentiation medium, 104 fibroblast growth factor, 105-106 gelatin-coated culture dishes, 102-103 growth medium, 104 horse serum pretesting, 104-105 nutrient medium, 103-104 Cell signaling, in embryo, 57 Cell surface receptor, interaction with env glycoprotein, 181-184 Charge, movement in myotube, 286-287 Chicken embryo extract preparation, 90-91 preparation for somite implantation, 18-21 muscle cultures conditioned media for, 100-101 primary cultures, 96-101 Chloramphenicol acetyltransferase assay cis-acting control region, 419-420 protein level in transgenic mouse, 397-398 tissue homogenization for, 428-430

cis-acting control region alkaline phosphatase assay, 420-421 CAT assay, 419-420 8-galactosidase assay, 418-419 luciferase reporter system, 420 reporter gene assay identification, 415-418 Cloning site selection for adenovirus design, 235-236 transcription factors, 468 virus-producing cells, 219-221 Collagen coated petri dish, preparation, 277 coating, for cardiogenic mesoderm explant culturing, 126 rat tail, for myoblast culture, 276-277 Coverslips, coating with laminin, 312-313 Culturing cardiogenic mesoderm explants collagen coating, 126 fibronectin coating, 125-126 media for, 126 cell, see Cell cultures embryos on agar-coated dishes, 125 on uncoated dishes, 124-125 primary cardiomyocytes, 126-127 vascular smooth muscle, 133-135 VSMC and endothelial cells, 151 Cytopathic effect assay, end-point, adenovirus, 240 Cytotoxicity, adenoviral vectors, 249 D

Degeneration, muscle, long-term correction, 245-246 Differentiation cardiac immunohistochemicalanalysis, 127-128 in situ hybridization explants, 130 with riboprobes, 128-129 with 3sS-labeled probes, 131 whole-mount hybridization, 129-130 cell, see Cell differentiation muscle, molecular markers analytical methods, 60 DNA-binding proteins, 61-62 transcript markers, 59 Digestion, see Enzymatic digestion Digital imaging, skeletal muscle and neuromuscular innervation, 335-337

476

Index

Digoxigenin, for in situ hybridization labeled probes, 391-395 labeled riboprobes, 128-129 Disease neuromuscular, adenoviral gene therapy, 255 satellite cell animal models, 157-158 Dishes agar-coated, for embryo culturing, 125 culture gelatin coating, 102-103 for MM14 cells, 102 for embryo microdissection, 8-10 holding, somite transfer to, 17-18 matrigel-coated, preparation, 277 for tissue holding, 10 uncoated, for embryo culturing, 124-125 Dissection microdissection, dishes for embryo surgery, 8-10 mouse, 89-90 rat, 89-90 Xenopus embryo, 55-56 DNA adenovirus preparation, 238-239 purification, 239 carrier in DNase I protection assay, 456 in electromobility shift assay, 447-448 complementary,injection, in dysgenic muscle transfection, 297-301 mediated gene transfer into cardiac muscle experimental design, 430-433 general considerations, 423-426 transfection of muscle cells calcium phosphate coprecipitation, 407-409 by electroporation, 409-412 MM14 cells, 409 stable transfection, 412-413 stable transfection protocol, 413-414 transgenic fsh, 69-70 transgenic mouse, preparation for microinjection, 375-376 transgenic mouse tail, 383 zebrafish, mutagenesis, 68-69 DNA-binding proteins, in Xenopus embryos, 61-62 DNase I, fragments in DNase I protection assay, 456-458 DNase I protection assay binding conditions, 458

carrier DNA, 456 DNase I fragments, 456-458 G + A ladder, 463 parameters, 454-456 probe, 458 probe preparation, 458-461 protection protocol, 461-463 reagents, 463-465 relevance to muscle research dissection of regulatory pathways with nuclear extracts, 467-468 in vitro techniques with nuclear extracts, 466-467 in vitro techniques with proteins, 468-470 Duchenne muscular dystrophy adenoviral gene therapy, 255 gene therapy, 230-231 Dysgenic muscle model general considerations, 296-297 mass transfection of cultures, 301-303 transfection by cDNA injection, 297-301

E Ectoderm, dermal, removal, 15-16 Egg, embryo removal filter-paper method, 120-121 new method, 121 Electromobility shift assay antibody supershifts, 454 binding reaction conditions, 447-449 competition experiments, 454 conditions, 449 high ionic strength conditions, 451-452 low ionic strength conditions, 452 parameters, 446 probe preparation, 450 reagents, 452-453 relevance to muscle research dissection of regulatory pathways with nuclear extracts, 467-468 in v i m techniques with nuclear extracts, 466-467 in virro techniques with proteins, 468-470 sensitivity, 449 Electron microscopy, thin section, muscle preparation for, 292-293 Electrophoresis gel, see Gel electrophoresis myofibrillar protein isoforms, 356-357 myosin isoforms, 357-359 Electroporation, for DNA transfection of muscle cells, 409-412

477

Index

EM, see Electron microscopy Embryology cellular zebrafish heart, 71-72 zebrafish skeletal muscle, 73-74 experimental mammalian embryos, 43-45 somitic cell culturing, 45-48 Embryonic axis, establishment, 118 Embryos animal cap assay, 56-57 avian donor, isolation and preparation, 11 micropipets for solution delivery, 6-8 in muscle research, 4-5 solutions for, 5-6 surgery microdissection dishes for, 8-10 microneedles for, 8 microscalpels for, 8 tissue holding dishes for, 10 tissue markers for, 10-11 chicken acquisition, 5 candling, 5 care for surgery, 5 chimeric harvesting, 23-25 histology, 25 donor somite implantation, 21-22 extract preparation, 90-91 preparation for somite implantation, 18-21 raised, small window with, 20-21 retroviruses for muscle lineage studies marking muscles, 224-225 precursor tagging, 223-224 staging, 11-13 chick muscle cells, 96-101 culturing on agar-coated dishes, 125 on uncoated dishes, 124-125 early, generation in vivo, 205-207 fixation and storage, 128 mouse gene expression analysis, 38-40 immunocytochemistry,41-43 in situ hybridization, 40-41 myogenesis, 29-31 Nieuwkoop conjugate, 56-57 quail

acquisition, 5 care for surgery, 5 muscle cells, 96-101 removal from egg filter-paper method, 120-121 new method, 121 signaling molecules, 57-59 signaling sources, 57 transgenic mouse recovery, 377-378 transfer, 381-382 whole-mount in sihr hybridization, 395-397 Xenopus dissection, 55-56 DNA-binding proteins, 61-62 lithium treatment, 56 UV treatment, 56 zebrafish, skeletal muscle cellular embryology, 73-74 genetics, 75 innervation, 74-75 morphology, 72-73 Endocrine system, satellite cell animal models, 158 Endoderm enzymatic digestion, 124 mechanical removal, 122-123 sodium citrate removal, 123 Endothelial cells, and VSMC,cocultivation, 151 Enzymatic digestion, in endoderm removal, 124 Enzymes in donor somite isolation, 14-18, 18 in medical VSMC isolation, 138-140 in neointimal cell isolation, 140-141 Excision, somites, 16-17 Excitation-contraction coupling, component identification calcium currents, 286-287 calcium transients in myotubes, 287-289 charge movement, 286-287 membrane freeze-fracture, 294-296 muscle preparation for thin section EM, 292-293 muscle preparation for voltage clamp, 285-286 physiological approaches, 284-285 structural approaches, 289-291 vesicles and macromolecules, 293-294 Extracellular matrix, role in phenotypic modulation, 151

478

Index

F

Fate maps axial muscle, 54 cardiac muscle, 54-55 limb and head muscle, 53-54 Fibers muscle, manipulation in vivo, 337-339 single cultures characteristics, 172-173 fiber isolation, 171-172 substrates and media, 170-171 skeletal muscle, analysis, 364-365 Fibroblast growth factor, for MM14 cell growth, 105-106 Fibroblasts, chicken embryo infected, for gene delivery, 203-205 virus stock production, 194-196 Fibronectin coating, for cardiogenic mesoderm explant culturing, 125-126 phenotypic modulation, 151 Filter paper, for embryo removal from egg, 120-121 Fixation embryos, for in situ hybridization with riboprobes, 128 mouse embryos, 395-3% Freeze-drying, vesicles and macromolecules, 293-294 Freeze-fracturing cultured cells, 303-304 membranes in excitation-contraction coupling, 294-2% Freezing MM14 cells, 108-109 mouse C, cell line, 111-112 G

@-Galactosidaseassay, cis-acting control region, 418-419 Gastrulation, cardiac progenitors postgastrulation location, 119-120 pregastrulation location, 119 Gelatin, coating of culture dishes, 102-103 Gel electrophoresis, SDS-PAGE, and immunoblotting, integration with immunohistochemistry, 352-353 with mass proiile fingerprinting,353-354 with microsequencing, 353-354 Gels, 3D, basement membrane, 151-152

Gene delivery to skeletal muscle, 230-231 by somatic infection, 198-199 Genes expression in mouse embryo, 38-43 manipulation in mouse, 31-34 retroviral expression, 185 targeted expression in vivo, 186-187 tyrosinase marker, 382-383 Gene targeting to muscle in vivo, 202-205 to skeletal muscle in vivo infected chicken embryo fibroblasts, 203-205 long-term expression, 203 mitotic requirements, 203 virus stock, 202 Gene therapy with adenoviruses cytotoxicity, 249 delivery to muscle, 251-253 in host immune response, 250-251 humans, safety problems, 248-249 inflammation from, 249 safety improvements, 249-250 vector capacity, 249-250 Duchenne muscular dystrophy, 230-231 Gene transfer adenovirus-mediated, into adult cardiac myocytes primary culture, 312-313 rat, 309-312 adenovirus to muscle in adult mouse, 242-244 in cardiocytes and myocardium, 247-248 in newborn mouse, 241-242 promoters, 244 secreted protein production, 246-247 into cardiac muscle experimental design, 430-433 general considerations, 423-426 human minidystrophin to mdx mouse efficiency and stability, 244-245 long-term correction of degeneration, 245-246 retroviral-mediated, in myoblast labeling in vitro labeled cell selection, 265-266 labeling efficiency, 264-265 retroviral infection, 264 retroviruses, efficiency, 187-189

Index

479 targeting, 253-254 in vivo, with adenovirus, 231-234 Gilbert reaction, for DNase I protection assay, 463 Glycoproteins, env, interaction with receptor, 181-184 Growth, see Cell growth

H Harvesting chimeric chicken embryos, 23-25 viral DNA, 238 viral particles, 221-222 Heart injection protocol, 426-428 Xenopus, formation, 62-63 zebrafish cellular embryology, 71-72 morphology, 70-71 Histology, chimeric embryos, 25 Homogenization, tissue for assays, 428-430 Horse, serum, pretesting, 104-105 Hybridization, in situ explants, 130 in mouse embryo, 40-41 myofibrillar protein, 359-361 with riboprobes, 128-129 with "S-labeled probes, 131 transgenic mouse tissue sections nonradioactive hybridization, 391-395 radioactive hybridization, 387-391 whole-mount hybridization, 129-130, 395-397 I

Imaging, digital, skeletal muscle and neuromuscular innervation, 335-337 Immune response, host, from adenoviral gene therapy, 250-251 Immunoblotting myofibrillar protein isoforms, 356-357 myosin isoforms, 357-359 and SDS-PAGE, integration with immunohistochemistry, 352-353 with mass profile fingerprinting,353-354 with microsequencing, 353-354 Immunocytochemistry, mouse embryo, 41-43 Immunodetection, mouse embryo, 396-397 Immunofluorescence, indirect, myofilarnents, 318-320

Immunohistochemistry in analysis of differentiation, 127-128 integration with SDS-PAGE and immunoblotting, 352-353 myofibrillar protein, 359-361 Immunostaining, in structural visualization of cultures, 278-279 Implantation donor somites into host, 21-22 somite in chicken embryo, 18-21 Incision in separation of somite from neural tube, 16-17 skin, 20 somites, 20 Infection retroviral infection, 264 somatic, in gene delivery, 198-199 ventricular myocytes, 313 with viral DNA, 238 Inflammation, from adenoviral vectors, 249 Injection cDNA, in dysgenic muscle transfection, 2W-301 into heart, protocol, 426-428 microinjection DNA preparation for, 375-376 equipment and setup, 379-381 pipet for, preparation, 378 Injury, satellite cell animal models, 157-158 Inking, chicken embryo, 19-20 Innervation neuromuscular, in vivo analysis adult mouse, 328-329 development, 339-341 digital imaging, 335-337 embryonic mouse, 330 manipulations, 338-339 maturation, 341-343 neonatal mouse, 329-330 nonvital staining, 331-335 with optical microscope, 325-326 stage and animal plate, 327 vital staining, 330-331 zebratish embryo, 74-75 Ionic strength, conditions for electromobility shift assay, 451-452

L Labeling, myoblasts in vitro, by retroviralmediated gene transfer labeled cell selection, 265-266

480

Index

labeling efficiency, 264-265 retroviral infection, 264 Laminin coating of coverslips, 312-313 phenotypic modulation, 151 Life cycle retroviruses, 180-181 wild-type retrovirus, 216-217 Light, UV, treatment of early embryos, 56 Lithium, Xenopus embryo treatment, 56 Long terminal repeats, in vector plasmid, 218-219 Luciferase assay, tissue homogenization for, 428-430 Luciferase reporter system, cis-acting control region, 420

M Macromolecules, freeze-drying and replication, 293-294 Markers for cardiogenic mesoderm, 224-225 for differentiated cardiac muscle, 224-225 differentiation-specific,for VSMC characterization actin, 141-143 calponin, 144 myosin-heavy chain, 143 vinculidmeta-vinculin,143-144 molecular, muscle differentiation DNA-binding proteins, 61-62 transcript markers, 59 for tissues, 10-11 transcript, in muscle differentiation, 59 tyrosinase gene, 382-383 Matrigel, preparation, 277 Maturation, neuromuscular innervation, 341-343 Maxam reaction, for DNase I protection assay, 463 Media for cardiogenic mesoderm explant culturing, 126 conditioned, 100-101 differentiation media, 104 growth media, 104 monolayer mass cultures, 159-161 nutrient, 103-104 preparation for transgenic mouse, 376-377 quail growth, 277 for single fiber cultures, 170-171

Membranes basement, 3D gel, 151-152 in excitation-contraction coupling, freezefracture, 294-296 Mesoderm cardiogenic collagen coating, 126 fibronectin coating, 125-126 marking, 224-225 media for, 126 formation and patterning, role of signaling molecules, 57-59 incision, 17 zebrafish, 76-78 Messenger RNA analysis techniques, 361-364 myofibrillar protein isoform, 354-355 transgenic mouse isolation, 384 nonradioactive in situ hybridization, 391-395 Northern blots, 384-385 PCR, 386 radioactive in situ hybridization, 387-391 RNase protection assay, 385-386 Meta-vinculin, for VSMC characterization, 143-144 Microdissection, dishes, for embryo surgery, 8-10 Microinjection DNA preparation for, 375-376 equipment and setup, 379-381 pipet for, preparation, 378 Microneedles, for avian embryo surgery, 8 Micropipets, for solution delivery, 6-8 Microscalpels for avian embryo surgery, 8 in donor somite isolation, 14-18 Microscopy electron, thin section, muscle preparation for, 292-293 optical, skeletal muscle and neuromuscular innervation, 325-326 Microsurgery, donor embryos, 11 Migration, cell, VSMC, 144-147 Minidystrophin, human, gene transfer to mdr mouse efficiency and stability, 244-245 long-term comxtion of degeneration, 245-246

481

Index

Models animal satellite cells, 157-158 therapeutic effects in, 254-255 avian ALV in vitro, 196-198 in vivo models gene delivery by somatic infection, 198-199 replication-defective retroviral vectors, 199-201 tissue-specificexpression, 199 cell differentiation, 150-151 dysgenic muscle general considerations, 2%-297 mass transfection of cultures, 301-303 transfection by cDNA injection, 297-301 mouse, see Mouse models Molecular biology, zebrafish skeletal muscle, 75-78 Molecular markers, muscle differentiation DNA-binding proteins, 61-62 transcript markers, 59 Molecules macromolecules, freeze-drying and replication, 293-294 signaling, role in mesoderm formation and patterning, 57-59 Monoclonal antibodies, in immunofluorescence of myofilaments, 318-320 Morphology zebrafish heart, 70-71 zebrafish skeletal muscle, 72-73 Motor neurons, manipulation in vivo, 337-338 Mouse models adult adenoviral gene transfer, 242-244 preparation for surgery, 328-329 ALVIreceptor amphotropic vectors, 207 generation in vivo, 205-207 in vitro model, 201-202 in vivo targeting to muscle, 202-205 BCsHl cell line, 112-113 C2 cell line culture conditions, 111 culture techniques, 111 establishment, 110 freezing protocols, 111-112 thawing protocols, 112 embryo, gene expression analysis, 38-40

immunocytochemistry, 41-43 in situ hybridization, 40-41 embryonic, preparation for surgery, 330 experimental embryology, 44-45 ma'x

human minidystrophin gene transfer to efficiency and stability, 244-245 long-term correction of degeneration, 245-246 therapeutic effects in, 254-255 MM14 cell line cell differentiation, 102-106 cell propagation, 102-106 derivation, 101-102 differentiation-defective cells, 109-110 differentiation protocol, 109 exponential expansion, 106-107 freezing, 108-109 passaging cells, 107-108 thawing, 108-109 MMlCDD, availability, 113 muscle, cell cultures, 88-90 myoblast clonal cell lines, 91-95 neonatal, preparation for surgery, 329-330 newborn, adenoviral gene transfer, 241-242 somitic cell culturing, 46 Murine leukemia virus, for cell lineage studies, 218 Muscle differentiation, molecular markers analytical methods, 60 DNA-binding proteins, 61-62 transcript markers, 59 DNase I protection assay, 466-470 dysgenic model general considerations, 296-297 mass transfection of cultures, 301-303 transfection by cDNA injection, 297-301 electromobility shift assay, 466-470 fate maps axial muscle, 54 cardiac muscle, 54-55 limb and head muscle, 53-54 preparation for thin section EM, 292-293 research, with avian embryos, 4-5 skeletal, see Skeletal muscle smooth, see Smooth muscle Mutagenesis, zebrafish, 68-69 Myoblasts autologous, in adenovirus-mediated ex vivo gene transfer, 254 calcium phosphate-mediated DNA transfection, 407-409

Index cell cultures collagen-coated petri dish preparation, 277 culture protocol, 277-278 functional assessment, 279 growth, 263 matrigel-coated dish preparation, 277 matrigel preparation, 277 preparation, 262-263 quail growth medium, 277 on rat tail collagen, 276-277 structural visualization via immunostaining, 278-279 transfection and selection, 278 DNA transfection by electroporation, 409-412 stable transfection, 412-413 stable transfection protocol, 413-414 isolation from rat, 90 labeling in vitro, by retroviral-mediated gene transfer labeled cell selection, 265-266 labeling efficiency, 264-265 retroviral infection, 264 mouse, clonal cell lines, 91-95 transplantation anatomical assay, 269 biochemical assay, 270-271 evaluation, 268-269 techniques, 266-268 Myocardium, gene transfer into, 247-248 Myocytes cardiac, see Cardiomyocytes isolation, 127 ventricular indirect immunofluorescence with antibodies, 318-320 infection with recombinant adenovirus, 313 maintenance in culture, 313 plating, 313 Western blot analysis, 317-318 Myofibrillar protein electrophoresis, 356-357,359 immUnOblOtting, 356-357,359 immunohistochemistry,359-361 isofom in muscle tissue analysis at protein and mRNA level, 354-355 approaches to analysis, 350-352 distribution, 349-350 multiple analysis, 355-356 SDS-PAGE, immunoblotting

and immunohistochemistry,352-353 microsequencing, and mass profile fingerprinting,353-354 mRNA, analysis techniques, 361-364 in situ hybridization, 359-361 Myofilament protein, incorporation in infected ventricular myocytes indirect immunofluorescence with antibodies, 318-320 Western blot analysis, 317-318 Myogenesis, in mouse embryo, 29-31 Myogenic determination factors, in skeletal muscle, 86 Myosin, isofom analysis, 357-359 Myosin-heavy chain, for VSMC characterization, 143 Myotubes calcium currents, 286-287 calcium transients in, 287-289 charge movements, 286-287

N Neointimal cells, enzymatic isolation, 140-141 Neural tubes, separation from somites, 16-17 Neuromuscular disease, adenoviral gene therapy, 255 Neuromuscular system, innervation, in vivo analysis adult mouse, 328-329 development, 339-341 digital imaging, 335-337 embryonic mouse, 330 manipulations, 338-339 maturation, 341-343 neonatal mouse, 329-330 nonvital staining, 331-335 with optical microscope, 325-326 stage and animal plate, 327 vital staining, 330-331 Neurons, motor, manipulation in vivo, 337-338 Northern blots, transgenic mouse RNA, 384-385 Nuclear extract crude, in virro techniques with, 466-467 in dissection of regulatory pathways, 467-468 preparation, 441-442 protocol, 442-446 in virro techniques with proteins, 468-470 Nutrients, in culture media, 103-104

483

Index

0

Q

Optical microscopy, skeletal muscle and neuromuscular innervation, 325-326 Organoids, in adenovirus-mediated ex vivo gene transfer, 254 Ovulation, superovulation, transgenic mouse, 376

Quail primary muscle cell cultures, 96-101 somite isolation, 14-18

P Pancreatin, for avian embryos, 6 Perfusion, Langendorff, for myocyte isolation, 310-311 Petri dish, collagen-coated,preparation, 277 Phenotypes modulation of fibronectin and laminin, 151 in VSMC modulation, 135-136 Pipets, holding and microinjection, preparation, 378 Plaque assay adenovirus, 239-240 recombinant adenoviruses, 316-317 Plasmids adaptor construction, 191-193 propagation, 191-193 for retrovirus generation, 218-219 Plasticity, neuromuscular junction, 339-341 Polymerase chain reaction, transgenic mouse RNA, 386 Probes digoxigenin-labeled, for in situ hybridization, 391-395 for DNase I protection assay, 458-461 for electromobility shift assay, 450 35S-labeled,for in situ hybridization, 131 Promoters, for adenoviral gene transfer, 244 Propagation, viral adaptor plasmids, 191-193 ALV-based retroviral vectors, 191-193 Protection assay, see DNase I protection assay; RNase protection assay Protein domain mapping, with electromobility shift assay, 469 Proteins exogenous expression, 280-281 level in transgenic mouse CAT assay, 397-398 LacZ staining, 398-400 reporter gene assay, 418 secreted, production in skeletal muscle, 246-247

R Recombination, homologous, transgenic mouse, 34-38 Regulatory pathways, dissection with nuclear extracts, 467-468 Replication defective retroviral vectors, 199-201, 217-218 deficient adenovirus vectors, 235 vesicles and macromolecules, 293-294 Reporter gene assay, in cis-acting control region identification, 415-418 Retroviruses ALV-based vector system, 189-190 construction, 191-193 propagation, 191-193 for cell lineage studies helper virus test, 222 safety, 222-223 viral harvest and concentration, 221-222 virus-producing cells Cloning, 219-221 frozen stock, 221 env glycoprotein interaction with receptor, 181-184 gene expression, 185 gene transfer efficiency, 187-189 life cycle, 180-181 mediated gene transfer, in myoblast labeling in vitro labeled cell selection, 265-266 labeling efficiency, 264-265 retroviral infection, 264 for muscle lineage analysis in chicken embryos marking muscles, 224-225 precursor tagging, 223-224 replication-defectivevectors, 199-201, 217-218 targeted expression in vivo, 186-187 vector plasmid for generation, 218-219 vector system, 185-186 wild-type, life cycle, 216-217 Riboprobes digoxigenin-labeled, preparation, 128-129 preparation, 388

484

Index

RNA, see Messenger RNA RNase protection assay, transgenic mouse, 385-386 Rous sarcoma virus, for cell lineage studies, 218

S Salt, concentration in electromobility shift assay, 448-449 Satellite cells, skeletal muscle activation and proliferation, 167 animal models, 157-158 cell preparation, 161-163 culture homogeneity, 163-167 differentiation, 168-170 early identification, 155-156 medium, 159-161 monolayer mass cultures, 158-159,173-174 single fiber cultures, 174 characteristics, 172-173 fiber isolation, 171-172 substrates and media, 170-171 substrates, 159 SDS-PAGE, see Gel electrophoresis, SDS-PAGE Sealing tape, embryo surgery, 11 Serum, equine, pretesting, 104-105 Signaling molecules, role in mesoderm formation and patterning, 57-59 Skeletal muscle adenoviral gene transfer, 242-244 cell cultures chick embryo extract preparation, 90-91 mature, 86-87 mouse primary cultures, 88-89 dissections, 89-90 myoblast isolation, 90 primary muscle, 87-88 cell lines, 101 chick primary muscle cell culture, 96-101 fibers, manipulation in vivo, 337-339 gene delivery to, 230-231 gene targeting in vivo infected chicken embryo fibroblasts, 203-205 long-term expression, 203 mitotic requirements, 203 virus stock, 202 gene therapy deliveries, 251-253 lineage studies with retroviruses, in chicken embryos marking muscles, 224-225

precursor tagging, 223-224 mouse cell line culture conditions, 111 culture techniques, 111 establishment, 110 freezing protocols, 111-112 thawing protocols, 112 mouse Mh414 cell line availability, 113 cell differentiation, 102-106 cell propagation, 102-106 derivation, 101-102 differentiationdefective cells, 109-1 10 differentiation protocol, 109 exponential expansion, 106-107 freezing, 108-109 passaging cells, 107-108 thawing, 108-109 mouse MMl4-DD cell Line, availability, 113 myofibrillar protein isoforms analysis at protein and mRNA level, 354-355 approaches to analysis, 350-352 distribution, 349-350 multiple analysis, 355-356 SDS-PAGE, immunoblotting and immunohistochemistry,352-353 microsequencing, and mass profile fingerprinting, 353-354 myogenic determination factors, 86 preparation for thin section EM, 292-293 preparation for voltage clamp, 285-286 quail primary muscle cell culture, 96-101 rat primary cultures, 95-96 satellite cell cultures activation and proliferation, 167 animal models, 157-158 cell preparation, 161-163 culture homogeneity, 163-167 differentiation, 168-170 early identification, 155-156 medium, 159-161 monolayer mass cultures, 158-159, 173-174 single fiber cultures, 174 fiber isolation, 171-172 substrates and media, 170-171 substrates, 159 secreted protein production, 246-247 single fiber analysis, 364-365 targeted gene expression, 253-254 in vivo analysis

Index

485 adult mouse, 328-329 digital imaging, 335-337 embryonic mouse, 330 neonatal mouse, 329-330 nonvital staining, 331-335 with optical microscope, 325-326 stage and animal plate, 327 vital staining, 330-331 zebrafish cellular embryology, 73-74 genetics, 75 innervation, 74-75 molecular biology, 75-78 morphology, 72-73 Skin ectoderm, removal, 15-16 incision, 20 Smooth muscle adenoviral gene transfer, 241-242, 244 cardiac differentiated, marking, 224-225 gene transfer experimental design, 430-433 general considerations, 423-426 gene therapy deliveries, 251-253 myofibrillar protein isoforms analysis at protein and mRNA level, 354-355 approaches to analysis, 350-352 distribution, 349-350 multiple analysis, 355-356 SDS-PAGE, immunoblotting and immunohistochemistry, 352-353 microsequencing, and mass profile fingerprinting, 353-354 targeted gene expression, 253-254 vascular cells, see Vascular smooth muscle cells Sodium citrate, in endoderm removal, 123 Solutions for avian embryos, 5-6 for cell culture, 137-138 micropipets for delivery, 6-8 for myocyte isolation, 312 for recombinant adenovirus production, 314 Somites counting, 11-13 culturing, 42, 45-48 donor, implantation into host, 21-22 excision, 16-17 half-somites, transplantation cranial-caudal somite, 23

dorsal-ventral somite, 23 medial-lateral somite, 22 implantation, 18-21 incision, 20 molecular biology, 75-76 quail, isolation, 14 without enzymes, 18 with microneedles, 18 with microscalpels and enzymes, 14-18 separation from neural tube, 16-17 skeletal muscle precursor tagging, 223-224 stages, 14 target somites, 15-16 transfer to holding dish, 17-18 Staining immunostaining, in structural visualization of cultures, 278-279 with LacZ, protein in transgenic mouse, 398-400 neuromuscular junction synaptic components nonvital staining, 331-335 vital staining, 330-331 transgenic mouse tissues, 390-391 Stem cells, myogenic, early identification, 155-156 Storage embryos, for in situ hybridization with riboprobes, 128 recombinant adenovirus stock, 241 Superovulation, transgenic mouse, 376 Surgery adult mouse preparation for, 328-329 adult rat, 309-310 avian embryos, 5 microsurgery, donor embryos, 11 Synapses age-related maintenance alterations, 343-346 components of neuromuscular junctions nonvital staining, 331-335 vital staining, 330-331 manipulation in v i v q 337-338

T Temperature, in electromobility shift assay, 449 Thawing MM14 cells, 108-109 mouse C, cell line, 112 Tissues chicken, specific expression, 199 holding dishes, 10 homogenization for assays, 428-430

486

Index

human, culturing atheromatous tissue, 149-150 normal tissue, 147-149 markers, 10-11 muscle, myofibrillar protein isoforms analysis at protein and &A level, 354-355 approaches to analysis, 350-352 distribution, 349-350 multiple analysis, 355-356 SDS-PAGE, immunoblotting and immunohistochemistry, 352-353 microsequencing, and mass profile fingerprinting, 353-354 transgenic mouse nonradioactive in situ hybridization, 391-395 radioactive in situ hybridization, 387-391 Titration, adenovirus, plaque assay, 239-240 Transcription factors, cloning, 468 Transcription signals, selection for adenovirus design, 235-236 Transcript markers, in muscle differentiation, 59 Transfection DNA, in muscle cells calcium phosphate coprecipitation,

407-409

by electroporation, 409-412 M M 1 4 cells, 409 stable transfection, 412-413 stable transfection protocol, 413-414 dysgenic muscle by cDNA injection, 297-301 mass, primary cultures, 301-303 muscle culture, 278 Transgenes, manipulaticq in mouse, 31-34 Transgenic fish, DNA, 69-70 Transgenic mouse embryo myogenesis, 29-31 whole-mount in situ hybridization, 395-397 gene manipulation, 31-34 generation in vivo, 205-207 homologous recombination, 34-38 identification, tyrosinase gene marker, 382-383 production DNA preparation for microinjection, 375-376 embryo recovery, 377-378

embryo transfer, 381-382 medium preparation, 376-377 microinjection equipment, 379-381 pipet preparation, 378 superovulation, 376 vasectomy, 381-382 protein level CAT assay, 397-398 IACZ Staining, 398-400 RNA isolation, 384 nonradioactive in sihr hybridization, 391-395 Northern blots, 384-385 PCR,386 radioactive in situ hybridization, 387-391 RNase protection assay, 385-386 transient transgenics, 383-384 Transplantation half-somites cranial-caudal somite, 23 dorsal-ventral somite, 23 medial-lateral somite, 22 myoblast anatomical assay, 269 biochemical assay, 270-271 evaluation, 268-269 techniques, 266-268 Tyrosinase, gene marker, 382-383 U Ultraviolet light, treatment of early embryos, 56

V Vascular smooth muscle cells cell migration, 144-147 differentiated state, 150-151 differentiation-spec& markers actin, 141-143 calponin, 144 myosin-heavy chain, 143 vinculin/meta-vinculin, 143-144 and endothelial cells, cocultivation, 151 medical, enzymatic isolation, 138-140 phenotypic modulation, 135-136 sources for culture, 133-135 Vasectomy, transgenic mouse, 381-382 Vesicles, freeze-drying and replication, 293-294

Index

487 Vinculin, for VSMC characterization, 143-144 Virus propagation adaptor plasmids, 191-193 ALV-based retroviral vectors, 191-1 93 stock production on chicken embryo fibroblasts, 194-196 Visualization neuromuscular junction synaptic components in vivo nonvital staining, 331-335 vital staining, 330-331 structural, cultures, via immunostaining, 278-279 Voltage clamp, muscle preparation for, 285-286 VSMC, see Vascular smooth muscle cells

W Western blot analysis, myofilament protein expression, 317-318

Windowing chicken embryo, 19 with raised embryos, 20-21

X Xenopus, embryo dissection, 55-56 DNA-binding proteins, 61-62 heart formation, 62-63 Z

Zebrafish genetics, 68 heart cellular embryology, 71-72 morphology, 70-71 mutagenesis, 68-69 skeletal muscle cellular embryology, 73-74 genetics, 75 innervation, 74-75 molecular biology, 75-78 morphology, 72-73


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Cell Imaging Techniques (Methods in Molecular Biology)

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Microtubules: in vivo, Volume 97 (Methods in Cell Biology)

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Experimental Methods in Biology

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Methods in Cell Biology Volume 68 Atomic Force Microscopy in Cell Biology

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Cell-Penetrating Peptides: Methods and Protocols (Methods in Molecular Biology)

Cell Fusion: Overviews and Methods (Methods in Molecular Biology)

Methods in Cell Biology Volume 61 Mitosis and Meiosis

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