Bone Research Protocols (Methods in Molecular Biology, v816, 2nd Ed.)

METHODS IN MOLECULAR BIOLOGY™ Series Editor John M. Walker School of Life Sciences University of Hertfordshire Hatfi...

Author: Miep H. Helfrich | Stuart H. Ralston

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METHODS

IN

MOLECULAR BIOLOGY™

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

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

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Bone Research Protocols Second Edition Edited by

Miep H. Helfrich Musculoskeletal Research Programme, Division of Applied Medicine, Institute of Medical Sciences, University of Aberdeen, Aberdeen, UK

Stuart H. Ralston Rheumatic Diseases Unit, Institute of Genetics and Molecular Medicine, University of Edinburgh, Western General Hospital, Edinburgh, UK

Editors Miep H. Helfrich, PhD Musculoskeletal Research Programme Division of Applied Medicine Institute of Medical Sciences University of Aberdeen Aberdeen, UK [email protected]

Stuart H. Ralston, MD Rheumatic Diseases Unit Institute of Genetics and Molecular Medicine University of Edinburgh Western General Hospital Edinburgh, UK [email protected]

ISSN 1064-3745 e-ISSN 1940-6029 ISBN 978-1-61779-414-8 e-ISBN 978-1-61779-415-5 DOI 10.1007/978-1-61779-415-5 Springer New York Dordrecht Heidelberg London Library of Congress Control Number: 2011940327 © Springer Science+Business Media, LLC 2012 All rights reserved. This work may not be translated or copied in whole or in part without the written permission of the publisher (Humana Press, c/o Springer Science+Business Media, LLC, 233 Spring Street, New York, NY 10013, USA), except for brief excerpts in connection with reviews or scholarly analysis. Use in connection with any form of information storage and retrieval, electronic adaptation, computer software, or by similar or dissimilar methodology now known or hereafter developed is forbidden. The use in this publication of trade names, trademarks, service marks, and similar terms, even if they are not identified as such, is not to be taken as an expression of opinion as to whether or not they are subject to proprietary rights. Printed on acid-free paper Humana Press is part of Springer Science+Business Media (www.springer.com)

Preface The last decade has continued to bring tremendous advances in our understanding of bone biology. The genes responsible for the majority of rare inherited bone disorders have been identified and much progress has been made in the identification of genes in polygenic disorders, such as Paget’s disease and osteoporosis. Studies of genetically modified mice have resulted in the identification of other genes with profound effects on bone. These studies have uncovered many new pathways which form the focus of research by bone cell biologists to understand the mechanisms by which these genes and gene products affect bone mass and bone strength. The second edition of Bone Research Protocols contains a catalogue of protocols to assist researchers in the pursuit of mechanistic studies. In the tradition of the Methods in Molecular Medicine series, the chapters are practical laboratory protocols that should enable the reader to carry out the techniques from scratch. We have concentrated on laboratory techniques, rather than clinical methods of assessment and have tried to tailor the methods to the study of bone cells and bone tissue. For example, there are no differences in the analysis of DNA and RNA from bone or other tissues, but special considerations apply to isolation of DNA and RNA from bone and these are described. Equally, histological and histochemical procedures for soft tissues are often easily adapted to bone and are not specifically covered, apart from those included as analysis tools in various chapters. Tissue fixation, embedding and sectioning of bone, however, present unique problems and such methods are described as part of the chapters dealing with electron microscopy and immunostaining. Much progress has been made in digital image analysis recently and several chapters (dealing with confocal microscopy, bone resorption assays, and histomorphometry) include a detailed description on how to make best use of this powerful technology. New chapters have also been included on the analysis of bone tissue by Fourier transform infrared microscopy and Raman Spectroscopy in view of the increasing interest in these techniques as methods of assessing bone quality. The chapters on bone imaging have been updated and extensively revised with new chapters on quantitative analysis of radiographs and real-time bioluminescent imaging. Generally, one method is given for each technique, with the exception of in vitro osteoclast formation studies, for which several protocols are described, illustrating that many methods, often only subtly different, are described in the literature for different species and for different applications. Increasingly, high-throughput methods are used and some of the culture techniques described are suitable for such studies. Those interested in this field are encouraged to read all methods first before deciding which one is most appropriate for their particular application. The section on osteoblast cultures has been updated and new chapters have been included on primary osteocyte cultures, analysis of osteocyte cell lines, and osteogenic differentiation of bone cells from mesenchymal stem cells.

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Preface

A new section has been added on biochemical and molecular analysis of bone cells to cover topics, such as transfection, analysis of intracellular signaling, promoter reporter assays, gel shift assays, and chromatin immunoprecipitation assays. The section on mechanical loading techniques has been updated from the previous edition and expanded by inclusion of a new chapter on in vivo loading techniques. We hope that Bone Research Protocols will help those entering the bone field to establish new techniques in their laboratories. For those already experienced in bone research, we hope that they will benefit from the detailed description of the methods, in particular the many pointers and pitfalls, which the authors were specifically asked to discuss in the Notes section. We certainly learned a lot! We express our sincere thanks to all authors for their willingness to share their trade secrets and to Prof. John Walker at Humana Press for giving us the opportunity to publish a second edition of Bone Research Protocols; both he and the authors have been most patient during the edits of this volume. Aberdeen, UK Edinburgh, UK

Miep H. Helfrich, PhD Stuart H. Ralston, MD

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

PART I

CULTURE OF OSTEOBLASTS AND OSTEOCYTES

1 Primary Human Osteoblast Cultures. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Jane P. Dillon, Victoria J. Waring-Green, Adam M. Taylor, Peter J.M. Wilson, Mark Birch, Alison Gartland, and James A. Gallagher 2 Osteoblast Isolation from Murine Calvaria and Long Bones . . . . . . . . . . . . . . Astrid D. Bakker and Jenneke Klein-Nulend 3 Rat Osteoblast Cultures . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Isabel R. Orriss, Sarah E.B. Taylor, and Timothy R. Arnett 4 Isolation of Primary Avian Osteocytes . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Cor M. Semeins, Astrid D. Bakker, and Jenneke Klein-Nulend 5 Isolation of Mouse Osteocytes Using Cell Fractionation for Gene Expression Analysis. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Christine Halleux, Ina Kramer, Cyril Allard, and Michaela Kneissel 6 Studying Osteocyte Function Using the Cell Lines MLO-Y4 and MLO-A5. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Jennifer Rosser and Lynda F. Bonewald 7 Isolation, Differentiation, and Characterisation of Skeletal Stem Cells from Human Bone Marrow In Vitro and In Vivo . . . . . . . . . . . . . . . . . . . . . . Rahul S. Tare, Peter D. Mitchell, Janos Kanczler, and Richard O.C. Oreffo

PART II

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3

19 31

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CULTURE OF OSTEOCLASTS

8 Rodent Osteoclast Cultures. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Isabel R. Orriss and Timothy R. Arnett 9 Isolation and Culture of Primary Chicken Osteoclasts . . . . . . . . . . . . . . . . . . . Patricia Collin-Osdoby and Philip Osdoby 10 Isolation and Purification of Rabbit Osteoclasts. . . . . . . . . . . . . . . . . . . . . . . . Fraser P. Coxon, Michael J. Rogers, and Julie C. Crockett

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Contents

11 Generation of Human Osteoclasts from Peripheral Blood . . . . . . . . . . . . . . . . Kim Henriksen, Morten A. Karsdal, Adam Taylor, Denise Tosh, and Fraser P. Coxon 12 Osteoclast Formation in Mouse Co-cultures . . . . . . . . . . . . . . . . . . . . . . . . . . Cecile Itzstein and Robert J. van ’t Hof 13 RANKL-Mediated Osteoclast Formation from Murine RAW 264.7 cells . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Patricia Collin-Osdoby and Philip Osdoby

PART III

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BIOCHEMICAL AND MOLECULAR ANALYSIS OF BONE CELLS

14 Transfection of Osteoclasts and Osteoclast Precursors . . . . . . . . . . . . . . . . . . . Julie C. Crockett, David J. Mellis, and Adam Taylor 15 Analysis of Signalling Pathways by Western Blotting and Immunoprecipitation . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Aymen I. Idris 16 Analysis of Transcriptional Regulation in Bone Cells . . . . . . . . . . . . . . . . . . . . Huilin Jin and Stuart H. Ralston 17 Extraction of Nucleic Acids from Bone . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Alun Hughes, Tracy L. Stewart, and Val Mann 18 Analysis of Gene Expression in Bone by Quantitative RT/PCR . . . . . . . . . . . . Alun Hughes

PART IV

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223 233 249 261

MICROSCOPICAL TECHNIQUES

19 Histomorphometry in Rodents . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Reinhold G. Erben and Martin Glösmann 20 Studying Gene Expression in Bone by In Situ Hybridization . . . . . . . . . . . . . . Ina Kramer, Rishard Salie, Mira Susa, and Michaela Kneissel 21 Immunostaining of Skeletal Tissues . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Tobias B. Kurth and Cosimo De Bari 22 Techniques for the Study of Apoptosis in Bone . . . . . . . . . . . . . . . . . . . . . . . . Sudeh Riahi and Brendon Noble 23 Transmission Electron Microscopy of Bone . . . . . . . . . . . . . . . . . . . . . . . . . . . Vincent Everts, Anneke Niehof, Wikky Tigchelaar-Gutter, and Wouter Beertsen 24 Scanning Electron Microscopy of Bone . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Alan Boyde 25 Fluorescence Imaging of Osteoclasts Using Confocal Microscopy . . . . . . . . . . Fraser P. Coxon 26 Live Imaging of Bone Cell and Organ Cultures. . . . . . . . . . . . . . . . . . . . . . . . Sarah L. Dallas and Patricia A. Veno

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321 335 351

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Contents

PART V

IMAGING TECHNIQUES

27 Analysis of Bone Architecture in Rodents Using Microcomputed Tomography . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Robert J. van ‘t Hof 28 Bone Measurements by Peripheral Quantitative Computed Tomography in Rodents . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Jürg A. Gasser and Johannes Willnecker 29 Quantitative X-ray Imaging of Rodent Bone by Faxitron . . . . . . . . . . . . . . . . . J.H. Duncan Bassett, Anne van der Spek, Apostolos Gogakos, and Graham R. Williams 30 Bioluminescence Imaging of Bone Metastasis in Rodents . . . . . . . . . . . . . . . . Thomas J.A. Snoeks, Ermond van Beek, Ivo Que, Eric L. Kaijzel, and Clemens W.G.M. Löwik 31 Fourier Transform Infrared Imaging of Bone . . . . . . . . . . . . . . . . . . . . . . . . . Eleftherios P. Paschalis 32 Raman Microscopy of Bone . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Simon R. Goodyear and Richard M. Aspden

PART VI

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IN VIVO TECHNIQUES

33 The Calvarial Injection Assay. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Robert J. van ‘t Hof 34 Ovariectomy/Orchidectomy in Rodents . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Aymen I. Idris

PART VII

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MECHANICAL LOADING TECHNIQUES

35 Mechanical Properties of Bone Ex Vivo. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Simon R. Goodyear and Richard M. Aspden 36 Mechanical Stimulation of Bone Cells Using Fluid Flow . . . . . . . . . . . . . . . . . Carmen Huesa and Astrid D. Bakker 37 Using Cell and Organ Culture Models to Analyze Responses of Bone Cells to Mechanical Stimulation . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Andrew A. Pitsillides and Simon C.F. Rawlinson 38 In Vivo Mechanical Loading . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Roberto Lopes de Souza and Leanne Saxon Index . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

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Contributors CYRIL ALLARD • Transplantation and Inflammation Department, Novartis Institutes for BioMedical Research, Basel, Switzerland TIMOTHY R. ARNETT • Department of Cell and Developmental Biology, University College London, London, UK RICHARD M. ASPDEN • Musculoskeletal Research Programme, Division of Applied Medicine, Institute of Medical Sciences, University of Aberdeen, Aberdeen, UK ASTRID D. BAKKER • Department of Oral Cell Biology, Academic Centre for Dentistry Amsterdam (ACTA), University of Amsterdam and VU University Amsterdam, Research Institute MOVE, Amsterdam, The Netherlands J.H. DUNCAN BASSETT • Molecular Endocrinology Group, Department of Medicine and MRC Clinical Sciences Centre, Imperial College London, Hammersmith Campus, London, UK ERMOND VAN BEEK • Department of Endocrinology, Leiden University Medical Center, Leiden, The Netherlands WOUTER BEERTSEN • Department of Oral Cell Biology and Dept. Periodontology Academic Centre for Dentistry Amsterdam (ACTA), University of Amsterdam and VU University Amsterdam, Research Institute MOVE, Amsterdam, The Netherlands MARK BIRCH • Musculoskeletal Research Group, Institute of Cellular Medicine, The Medical School, Newcastle University, Newcastle-upon-Tyne, UK LYNDA F. BONEWALD • Department of Oral Biology, University of Missouri at Kansas City, Kansas City, MO, USA ALAN BOYDE • Biophysics Section, Oral Growth and Development, Dental Institute, Barts and The London School of Medicine and Dentistry, Queen Mary University of London, London, UK PATRICIA COLLIN-OSDOBY • Department of Biology, Washington University, St. Louis and Division of Bone and Mineral Metabolism, Washington University Medical School, St. Louis, MO, USA FRASER P. COXON • Musculoskeletal Research Programme, Division of Applied Medicine, Institute of Medical Sciences, University of Aberdeen, Foresterhill, Aberdeen, UK JULIE C. CROCKETT • Musculoskeletal Research Programme, Division of Applied Medicine, Institute of Medical Sciences, University of Aberdeen, Foresterhill, Aberdeen, UK SARAH L. DALLAS • School of Dentistry/Department of Oral Biology, University of Missouri, Kansas City, MO, USA COSIMO DE BARI • Regenerative Medicine Group, Musculoskeletal Research Programme, Institute of Medical Sciences, University of Aberdeen, Aberdeen, UK JANE P. DILLON • Bone and Joint Research Group, Musculoskeletal Biology, Institute of Ageing and Chronic Disease, University of Liverpool, Liverpool, UK

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Contributors

REINHOLD G. ERBEN • Institute of Physiology, Pathophysiology, and Biophysics, Department of Biomedical Sciences, University of Veterinary Medicine, Vienna, Austria VINCENT EVERTS • Department of Oral Cell Biology, Academic Centre for Dentistry Amsterdam (ACTA), University of Amsterdam and VU University Amsterdam, Research Institute MOVE, Amsterdam, The Netherlands JAMES A. GALLAGHER • Bone and Joint Research Group, Musculoskeletal Biology, Institute of Ageing and Chronic Disease, University of Liverpool, Liverpool, UK ALISON GARTLAND • Academic Unit of Bone Biology, Department of Human Metabolism, Mellanby Centre for Bone Research, University of Sheffield, Sheffield, UK JÜRG A. GASSER • Department of Musculoskeletal Diseases, Novartis Institutes for BioMedical Research, Basel, Switzerland MARTIN GLÖSMANN • Department of Biomedical Sciences, Institute of Physiology, Pathophysiology, and Biophysics, University of Veterinary Medicine, Vienna, Austria APOSTOLOS GOGAKOS • Molecular Endocrinology Group, Department of Medicine & MRC Clinical Sciences Centre, Imperial College London, Hammersmith Campus, London, W12 0NN, UK SIMON R. GOODYEAR • Musculoskeletal Research Programme, Division of Applied Medicine, Institute of Medical Sciences, University of Aberdeen, Aberdeen, UK CHRISTINE HALLEUX • Musculoskeletal Disease Department, Novartis Institutes for BioMedical Research, Basel, Switzerland KIM HENRIKSEN • Nordic Bioscience A/S, Herlev, Denmark ROBERT J. VAN ‘T HOF • Institute of Genetics and Molecular Medicine, University of Edinburgh, Edinburgh, UK CARMEN HUESA • Department of Developmental Biology, The Roslin Institute, University of Edinburgh, Edinburgh, UK ALUN HUGHES • Musculoskeletal Research Programme, Division of Applied Medicine, Institute of Medical Sciences, University of Aberdeen, Aberdeen, UK AYMEN I. IDRIS • Bone and Cancer Group, Edinburgh Cancer Research UK Centre and Rheumatic Disease Unit, the Centre of Molecular Medicine, University of Edinburgh, Edinburgh, UK CECILE ITZSTEIN • Musculoskeletal Research Programme, Division of Applied Medicine, Institute of Medical Sciences, University of Aberdeen, Aberdeen, UK HUILIN JIN • Kennedy Institute of Rheumatology, London, UK ERIC L. KAIJZEL • Department of Endocrinology, Leiden University Medical Center, Leiden, The Netherlands JANOS KANCZLER • Bone and Joint Research Group, Centre for Human Development, Stem Cells and Regeneration, Developmental Origins of Health and Disease, Institute of Developmental Sciences, University of Southampton Medical School, Southampton, UK MORTEN A. KARSDAL • Nordic Bioscience A/S, Herlev, Denmark JENNEKE KLEIN-NULEND • Department of Oral Cell Biology, Academic Centre for Dentistry Amsterdam (ACTA), University of Amsterdam and VU University Amsterdam, Research Institute MOVE, Amsterdam, The Netherlands MICHAELA KNEISSEL • Musculoskeletal Disease Department, Novartis Institutes for BioMedical Research, Basel, Switzerland

Contributors

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INA KRAMER • Musculoskeletal Disease Department, Novartis Institutes for BioMedical Research, Basel, Switzerland TOBIAS B. KURTH • Regenerative Medicine Group, Musculoskeletal Research Programme, Institute of Medical Sciences, University of Aberdeen, Aberdeen, UK ROBERTO LOPES DE SOUZA • Department of Veterinary Basic Sciences, Royal Veterinary College, Royal College Street, London, UK CLEMENS W.G.M. LÖWIK • Department of Endocrinology, Leiden University Medical Center, Leiden, The Netherlands VAL MANN • School of Health, Science and Social Care, University Campus Suffolk, Waterfront Building, Ipswich, UK DAVID J. MELLIS • Musculoskeletal Research Programme, Division of Applied Medicine, Institute of Medical Sciences, University of Aberdeen, Aberdeen, UK PETER D. MITCHELL • Bone and Joint Research Group, Centre for Human Development, Stem Cells and Regeneration, Developmental Origins of Health and Disease, Institute of Developmental Sciences, University of Southampton Medical School, Southampton, UK ANNEKE NIEHOF • Department of Periodontology, Academic Centre for Dentistry Amsterdam (ACTA), University of Amsterdam and VU University Amsterdam, Amsterdam, The Netherlands BRENDON NOBLE • University Campus Suffolk, Ipswich, UK RICHARD O.C. OREFFO • Bone and Joint Research Group, Centre for Human Development, Stem Cells and Regeneration, Developmental Origins of Health and Disease, Institute of Developmental Sciences, University of Southampton Medical School, Southampton, UK ISABEL R. ORRISS • Department of Cell and Developmental Biology, University College London, London, UK PHILIP OSDOBY • Department of Biology, Washington University, St. Louis, and Division of Bone and Mineral Metabolism, Washington University Medical School, St. Louis, MO, USA ELEFTHERIOS P. PASCHALIS • Ludwig Boltzmann Institute for Osteology, Vienna, Austria ANDREW A. PITSILLIDES • Department of Veterinary Basic Sciences, The Royal Veterinary College, Royal College Street, London, UK IVO QUE • Department of Endocrinology, Leiden University Medical Center, Leiden, The Netherlands STUART H. RALSTON • Rheumatic Diseases Unit, Western General Hospital, University of Edinburgh, Edinburgh, UK SIMON C.F. RAWLINSON • Queen Mary University of London, Barts & The London School of Medicine and Dentistry, Institute of Dentistry, Turner Street, London, UK SUDEH RIAHI • University Campus Suffolk, Ipswich, UK MICHAEL J. ROGERS • Musculoskeletal Research Programme, Division of Applied Medicine, Institute of Medical Sciences, University of Aberdeen, Aberdeen, UK JENNIFER ROSSER • Department of Oral Biology, University of Missouri at Kansas City, Kansas City, USA RISHARD SALIE • Musculoskeletal Disease Department, Novartis Institutes for BioMedical Research, Basel, Switzerland

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Contributors

LEANNE SAXON • Department of Veterinary Basic Sciences, Royal Veterinary College, Royal College Street, London, UK COR M. SEMEINS • Department of Oral Cell Biology, Academic Centre for Dentistry Amsterdam (ACTA), University of Amsterdam and VU University Amsterdam, Research Institute MOVE, Amsterdam, The Netherlands THOMAS J.A. SNOEKS • Department of Endocrinology, Leiden University Medical Center, Leiden, The Netherlands ANNE VAN DER SPEK • Molecular Endocrinology Group, Department of Medicine & MRC Clinical Sciences Centre, Imperial College London, Hammersmith Campus, London, UK TRACY L. STEWART • Musculoskeletal Research Programme, Division of Applied Medicine, Institute of Medical Sciences, University of Aberdeen, Aberdeen, UK MIRA SUSA • Oncology Department, Novartis Institutes for BioMedical Research, Basel, Switzerland RAHUL S. TARE • Bone and Joint Research Group, Centre for Human Development, Stem Cells and Regeneration, Developmental Origins of Health and Disease, Institute of Developmental Sciences, University of Southampton Medical School, Southampton, UK ADAM TAYLOR • Musculoskeletal Research Programme, Division of Applied Medicine, Institute of Medical Sciences, University of Aberdeen, Foresterhill, Aberdeen, UK ADAM M. TAYLOR • Lancaster Medical School, Faculty of Health and Medicine, University of Lancaster, Lancaster, UK SARAH E.B. TAYLOR • Department of Cell and Developmental Biology, University of Lancaster, Lancaster, UK WIKKY TIGCHELAAR-GUTTER • Department of Cell Biology and Histology, Academic Medical Centre (AMC), University of Amsterdam, Amsterdam, The Netherlands DENISE TOSH • Musculoskeletal Research Programme, Division of Applied Medicine, Institute of Medical Sciences, University of Aberdeen, Aberdeen, UK PATRICIA A. VENO • School of Dentistry/Department of Oral Biology, University of Missouri, Kansas City, MO, USA VICTORIA J. WARING-GREEN • Royal Veterinary College, London, UK GRAHAM R. WILLIAMS • Molecular Endocrinology Group, Department of Medicine and MRC Clinical Sciences Centre, Imperial College London, Hammersmith Campus, London, UK JOHANNES WILLNECKER • Stratec Medizintechnik GmbH, Pforzheim, Germany PETER J.M. WILSON • Bone and Joint Research Group, Musculoskeletal Biology, Institute of Ageing and Chronic Disease, University of Liverpool, Liverpool, UK

Part I Culture of Osteoblasts and Osteocytes

Chapter 1 Primary Human Osteoblast Cultures Jane P. Dillon, Victoria J. Waring-Green, Adam M. Taylor, Peter J.M. Wilson, Mark Birch, Alison Gartland, and James A. Gallagher Abstract Osteoblast cultures can be used to investigate the mechanisms of bone formation, to probe the cellular and molecular basis of bone disease, and to screen for potential therapeutic agents that affect bone formation. Here, we describe the methods for establishing and characterising primary human osteoblast cultures. Key words: Osteoblast, HOBs, Osteosarcoma cell, Osteogenesis, Bone formation

1. Introduction Osteoblasts are the cells responsible for the formation of bone; they synthesise almost all of the constituents of the bone matrix and direct its subsequent mineralisation. Once bone formation is complete, osteoblasts differentiate into osteocytes or bone lining cells, both of which play major roles in the regulation of calcium homeostasis and bone remodelling. Osteoblast cultures can be used to study the biochemistry and physiology of bone formation, to investigate the molecular and cellular basis of human bone disease, and to study the mechanisms by which bone resorption is coupled to bone formation. In addition, osteoblast cultures can be used to screen for potential anabolic agents, to develop and test new biomaterials, and to provide a source of cells for tissue engineering. The main focus of the present chapter is to describe the methods for establishing primary cultures of human osteoblast-like cells using the explant technique. We also describe methods for cryopreservation and phenotypic characterisation of these osteoblast cultures. In the following sections, we have also reviewed the

Miep H. Helfrich and Stuart H. Ralston (eds.), Bone Research Protocols, Methods in Molecular Biology, vol. 816, DOI 10.1007/978-1-61779-415-5_1, © Springer Science+Business Media, LLC 2012

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characteristics of primary osteoblast cultures in relation to other experimental models, such as osteosarcoma cell lines, mesenchymal stem cells (MSCs), and osteoblasts derived from peripheral blood. 1.1. Human Primary Osteoblast Cultures

Bard and co-workers (1) were the first to isolate living cells from adult human bone, but these had low levels of alkaline phosphatase and produced only small amounts of collagen. Although the cells remained viable for up to 2 weeks, they did not proliferate and it was concluded that osteocytes were the predominant cell type present. Subsequently, Mills et al. were successful in culturing cells from explants of human bone from patients with Paget’s disease. These cultures were responsive to parathyroid hormone and expressed alkaline phosphatase positive cells (2). The methods for isolation, culture, and characterisation of primary human osteoblast-like cells on a large scale were established in Graham Russell’s laboratory at the University of Sheffield in the early 1980s (3, 4). The defining characteristics of these studies were the use of an explant culture technique, which avoided the need for digestion of the tissue and the availability of osteocalcin as a phenotypic marker to confirm identity of the cells in vitro. In addition to producing osteocalcin, the cells grown from explants of human bone express other characteristics of osteoblasts, including the fact that they respond to parathyroid hormone (5, 6) and produce high levels of alkaline phosphatase and type I collagen (7). While several modifications of this technique have been published (8–11) (see Note 1), the vast majority of published reports on human osteoblasts rely on the explant technique as depicted in Fig. 1. The nomenclature used by various research groups to describe these cells has also varied including “human bone cells”, “human osteoblasts in vitro”, “human osteoblast-like cells” (HOBs), and human bone-derived cells (HBDCs). In the previous edition of this book, we used the term human bone-derived cells to describe these cultures, but here we refer to the cells as human osteoblast-like cells since the evidence is now overwhelming that these cells express several characteristics of osteoblasts.

1.2. Osteosarcoma Cell Lines

Osteosarcoma cell lines are also widely used as an experimental model of osteoblasts. These cells proliferate more rapidly than primary cells and are immortal which makes them easy to work with. Osteosarcoma cells synthesise bone matrix proteins, respond to various calciotropic hormones, including parathyroid hormone, and express a wide spectrum of osteoblastic marker genes (12, 13). The osteosarcoma cell lines that are most widely used as experimental models are SaOS-2, MG-63, and TE85 cells (sometimes referred as HOS TE85). These cell lines differ with regard to their expression of osteoblastic markers and some researchers feel that

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Primary Human Osteoblast Cultures

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Fig. 1. Technique used to isolate cells expressing osteoblastic characteristics (HOBs) from explanted trabecular bone.

this reflects different stages of maturity with MG-63 being least differentiated and SaOS-2 the most differentiated cell line. All three lines are responsive to PTH and are available from ATCC (http://www.atcc.org) or ECACC (http://www.hpacultures.org). The characteristics of these cells are summarised in Table 1. 1.3. Osteoblasts from MSCs

It is possible to generate human osteoblasts from bone marrow by isolating MSCs and culturing these in osteogenic medium. The methods for this is not described here, but can be found in Chapter 7, this volume. Although some researchers distinguish between these cells and cells grown from explants of human bone, it is likely that there is a significant overlap in their origin and phenotype, including their potential for osteogenesis (19, 20). MSCs are usually isolated on the basis of expression of STRO-1, an uncharacterised antigen associated with the stromal lineage (21) which is also expressed in cells derived from explant culture (11).

Reference

Billiau et al. (14)

Mulkins et al. (15)

Rodan et al. (16)

Cell line

MG-63

TE85

SaOS-2

Osteosarcoma of an 11-year-old female Caucasian

Osteosarcoma of a 13-year-old female Caucasian

Osteosarcoma of a 14-year-old male Caucasian

Origin

Table 1 Characteristics of osteosarcoma cell lines

High levels of ALP, low levels of osteocalcin. Highly sensitive to PTH. Easy to transfect

Higher levels of ALP than MG-63, but lower levels of osteocalcin. Responsive to PTH

High growth rate; high production of type VI collagen, low levels of ALP. Responsive to PTH.

Phenotype

Produces mineralised matrix resembling bone in nude mice (16). Has been used to establish stable reporter cell lines, (17, 18)

Have been used for biological evaluation of tissue scaffolds for orthopaedic applications

Recently has been used in biomaterials research to study the effects of substrates on osteogenic differentiation

Comments

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1.4. Human Osteoblasts from Peripheral Blood


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Recently, there have been several reports describing the isolation of osteoblastic precursor cells from peripheral blood or cord blood (22–24). These cells have been isolated and enriched for osteoblastic characteristics using antibodies against osteocalcin (22), differential attachment on tissue culture plastic, and Ficoll-Paque density gradient centrifugation. Although there are only a few publications using these techniques, if they can be applied reproducibly, it will open up new opportunities to investigate osteoblastic activity especially in genetic diseases.

2. Materials 2.1. Equipment

1. Bone cutters. 2. Solid stainless steel scalpels with integral handles. 3. Forceps, assorted sizes. 4. Class II tissue culture facilities. 5. Incubator. 6. C-Chip haemocytometer (LabTech International) or Neubauer Haemocytometer (VWR). 7. 70 μm “cell strainer” (Becton Dickinson). 8. Cell freezing container, such as Mr Frosty (Nalgene). 9. Cryoampoules (Nalgene).

2.2. Cell Isolation and Culture

1. Dulbecco’s modification of minimum essential medium (DMEM). 2. Complete Culture medium DMEM with 10% FCS, 2 mM Lglutamine, 50 U/ml penicillin, and 50 μg/ml streptomycin (see Note 2). 3. Phosphate-buffered saline (PBS) without calcium and magnesium. 4. Trypsin/EDTA (0.05% trypsin and 0.02% EDTA in Ca2+- and Mg2+-free Hank’s BSS, pH 7.4). 5. L-ascorbic acid-2- phosphate (see Note 3). 6. Tissue-culture flasks (75 cm2) or Petri dishes (100 mm diameter) (see Note 4). 7. 0.4% trypan blue in 0.85% NaC1. 8. Collagenase type VII (Sigma-Aldrich). 9. DNAse I (Sigma-Aldrich). 10. 1,25(OH)2D3. (Sigma-Aldrich). 11. Dexamethasone (Sigma-Aldrich). 12. 5 mM β-glycerophosphate (Sigma-Aldrich).

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13. Inorganic phosphate solution (mix 500 mM solutions of Na2HPO4 and NaH2PO4 in a 4:1 (v/v) ratio. Sterile filter and store at 4°C prior to use). 2.3. Phenotypic Characterisation

1. Alkaline phosphatase assay kit (Sigma-Aldrich). 2. Alkaline phosphatase staining kit (Sigma-Aldrich). 3. Osteocalcin radioimmunassay (IDS Ltd., Boldon, UK or other supplier). 4. Alizarin red (Sigma-Aldrich). 5. qPCR primers and reagents for a panel of osteoblastic markers, including osteocalcin (see Note 5).

3. Methods 3.1. Establishing Primary Explant Cultures

This section describes the method for establishing an initial culture (termed explant 1 or E1) from trabecular bone chips that have been freshly obtained from the donor (see Note 6). An overview of the technique is shown in Fig. 1. 1. Transfer the bone tissue removed at surgery or biopsy (see Note 7) into a sterile container with PBS or serum-free medium (SFM) for transport to the laboratory with minimal delay, preferably on the same day (see Note 8). 2. Remove soft connective tissue from the outer surfaces of the bone by scraping with a sterile scalpel blade. 3. Rinse the tissue in sterile PBS and transfer to a sterile Petri dish containing a 5–20 ml PBS, depending on the size of the sample. 4. Isolate the trabecular bone fragments from the tissue using sterile bone cutters or a solid stainless steel scalpel blade as in Fig. 2 (see Note 9). With some bone samples, it may be necessary to gain access to the trabecular bone by breaking through the cortex with the aid of sterile surgical bone cutters. 5. Transfer the trabecular bone fragments to a clean Petri dish containing 2–3 ml of PBS and divide into pieces 3–5 mm in diameter with a scalpel blade and/or scissors. 6. Decant the PBS and transfer the bone chips to a sterile Universal container containing 15–20 ml PBS. 7. Vortex the tube vigorously three times for 10 s and then leave to stand for 30 s to allow the bone fragments to settle. 8. Carefully decant off the supernatant containing haematopoietic tissue and dislodged cells, add an additional 15–20 ml of PBS, and vortex the bone fragments as before.

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Fig. 2. Equipment required to dissect human trabecular bone prior to the establishment of explant cultures and a sample of bone obtained at surgery.

9. Repeat steps 7 and 8 at least three times, or until no remaining haematopoietic marrow is visible and the bone fragments have assumed a white, ivory-like appearance. 10. Place the washed bone fragments as explants at a density of 0.2–0.6 g of tissue/100-mm diameter Petri dish (Fig. 3). (see Note 10). 11. Add 10 ml of complete medium to each dish and culture at 37°C in humidified atmosphere with 5–7% CO2. 12. Culture the explants undisturbed for 7 days and replace the medium taking care not to dislodge the explants. 13. Check for outgrowth of cells at 7–10 days (Figs. 4 and 5) (see Note 11). 14. Replace the medium after 2 weeks and twice weekly thereafter for 4–6 weeks until the desired density has been attained (see Note 12). 3.2. Secondary Explant Cultures

This section describes the method for culturing HOBs from trabecular bone chips that have already been used to establish an E1 culture. These are termed explant 2 or E2 cultures (see Note 13). 1. Remove bone fragments from the tissue culture plate containing the HOBs using a pair of sterile fine forceps and place in a new Petri dish with fresh medium.

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Fig. 3. Culture dish containing explanted trabecular bone.

Fig. 4. Migration of cells expressing osteoblastic characteristics (HOBs) from explanted trabecular bone.

2. Chop the fragments using a scalpel to stimulate fresh cell outgrowth. 3. Leave for 7–10 days without changing the medium. 4. Follow steps 13–14 of Subheading 3.1 until the desired cell density has been obtained.

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Fig. 5. Typical morphology of cells expressing osteoblastic characteristics (HOBs).

3.3. Passaging Explant Cultures

This section describes the method for passaging HOBs which have already reached confluence or near confluence. This is an appropriate stage to confirm that the cells are expressing osteoblastic characteristics by histochemical or biochemical analysis of alkaline phosphatase activity and/or the measurement of osteocalcin in conditioned medium, following incubation with 10−9 M 1,25(OH)2D3 for 48 h (see Note 5). 1. Remove and discard the spent medium. 2. Gently wash the cell layers two times with 10 ml of PBS without Ca2+ and Mg2+ discarding PBS after each wash. 3. Add 5 ml of freshly thawed trypsin–EDTA solution at room temperature to each dish (20°C) and incubate for 5 min at 37°C with gentle rocking every 30 s to ensure that the entire surface area of the flask and explants is exposed to the solution. 4. Remove the dishes from the incubator and examine under the microscope. Look for the presence of rounded, highly refractile cell bodies floating in the trypsin–EDTA solution. If none, or only a few, are visible tap the base of the dish sharply on the bench top in an effort to dislodge the cells. If this is not effective, incubate the cells for a further 5 min at 37°C. 5. When most of the cells have become detached from the culture substratum, transfer to a Universal container containing 5 ml of culture medium. 6. Wash the flask two to three times with 5 ml of serum-free DMEM and pool the washings with the original cell isolate.

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7. Centrifuge at 250 × g for 5 min at room temperature to pellet the cells. 8. Remove and discard the supernatant, invert the tube, and allow the medium to drain briefly. 9. Resuspend the cells in 2 ml of serum-free DMEM. If the cells are clumping, filter the cell suspension through a 70-μm cell strainer into a 50 ml polypropylene tube (see Note 14). 10. Wash the filter with 2–3 ml serum-free DMEM and add the filtrate to the cells. 11. Take 20 μl of the mixed cell suspension and dilute to 80 μL with serum-free DMEM. Add 5 μl of trypan blue solution, mix, and leave for 1 min before counting viable (round and refractile), and nonviable (blue) cells in a haemocytometer. 12. Plate the harvested cells at a cell density suitable for the intended analysis (see Note 15). 3.4. Ascorbate Containing Cultures

Culturing cells in ascorbate results in the secretion of an abundant collagen-rich extracellular matrix and because of this, difficulty can be encountered in passaging the cells using trypsin/EDTA alone. Here, we describe a method for passage of these cells with collagenase. 1. Rinse the cell layers twice with serum-free DMEM. 2. Incubate the cells for 2 h at 37°C in 10 ml serum-free DMEM containing 25 U/ml purified collagenase type VII and 2 mM CaC12. 3. Gently agitate the flask for 10–15 s every 30 min. 4. Terminate the collagenase digestion by discarding the medium (check that there is no evidence of cell detachment at this stage). 5. Gently rinse the cell layer twice with 10 ml calcium and magnesium-free PBS. 6. Add 5 ml of freshly thawed trypsin–EDTA solution pH 7.4 at room temperature to each flask (20°C). 7. Follow steps 4–12 of Subheading 3.3.

3.5. Mineralising Cultures

Mineralised structures resembling the nodules that form in cultures of foetal or embryonic animal bone-derived cells can be obtained by culturing HOBs in the presence of ascorbate and either β-glycerol phosphate (β-GP) or another source of inorganic phosphate (Fig. 6). These cells can also be shown to make bone in diffusion chambers or on supports following transplantation in vivo (see Note 16). Here, we describe a method for generating mineralised structures from HOBs in vitro.

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Fig. 6. HOBs cultured under osteogenic conditions for 28 days. Alkaline phophatase positive cells are stained dark grey while alizarin red stained mineralised deposits appear black.

1. Prepare fragments of human trabecular bone as described in Subheading 3.1, steps 1–10. 2. Culture the washed bone fragments in medium supplemented with 100 μM L-ascorbic acid 2-phosphate and 10 nM dexamethasone (see Note 17). 3. Culture the cells for 4–5 weeks until they have attained confluence with medium changes twice weekly. 4. When the cells have synthesised a dense extracellular matrix, subculture using the sequential collagenase/trypsin–EDTA protocol described in Subheading 3.4 and plate the cells in 25-cm2 flasks at a density of 104 viable cells/cm2. 5. After a further 14 days, supplement the medium with 0.01% phosphate solution. 6. After 48–72 h, wash the cell layers two or three times with 10 ml of serum-free DMEM. 7. Aspirate the medium and fix with 95% ethanol at 4°C. 8. To identify mineralised bone nodules, wash the cell layer in PBS and stain with alizarin red (1% solution in water) for 20 min in the dark at room temperature. Rinse with 50% ethanol to remove excess stain, before a final wash in distilled water then air-dry. Cells can be counterstained for alkaline phosphatase activity (see Fig. 6).

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3.6. Cryopreservation

If required, HOBs can be stored frozen for extended periods in liquid nitrogen or in ultra-low temperature (−135°C) cell freezer banks. Here, we present a protocol for cryopreservation of HOBs. We recommend that this is performed before full confluence is reached. 1. Passage the cells using trypsin/ETDA as described in Subheading 3.3, steps 1–6. 2. Pellet the cells by centrifugation at 250 × g for 5 min and pour off the supernatant. 3. Resuspend the cell pellet in FCS, bring up to a volume of 900 μl and transfer to a cryoampoule. 4. Swirl the ampoule in an ice water bath. 5. Add 100 μl DMSO gradually while holding the ampoule in the iced water. 6. Close ampoules tightly and freeze at 1°C/min to −80°C in a cell freezing container, such as Mr Frosty (Nalgene). 7. Transfer the cells to liquid nitrogen for long-term storage.

3.7. Retrieval of Cryopreserved Cells

1. Remove the cells from liquid nitrogen and place in a water bath set at 37°C. 2. Transfer the cells to a universal containing at least 20 ml of preheated culture medium. 3. Centrifuge at 250 × g for 2 min to pellet the cells and pour off the supernatant. 4. Resuspend the cells in about 10 ml of the medium and place into culture for 24 h. 5. Replace the medium after 24 h and culture for 2–3 weeks.

4. Notes 1. Although most investigators have used the original explant method with only minor modifications, others have developed alternative techniques for the isolation and culture of HOBs. Gehron-Robey and Termine used prior digestion of minced bone with Clostridial collagenase and subsequent culture of explants in medium with reduced calcium concentrations (8). In contrast, Wergedal and Baylink have used collagenase digestion to directly liberate cells (9). Marie and co-workers have used a method in which explants are first cultured on a nylon mesh (10). These alternative methods are described in greater detail in ref. 11 and readers are encouraged to consult the original publications and other references by these authors.

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2. Batches of serum vary in their ability to support the growth of HOBs. It is advisable to screen batches and reserve a large quantity of serum once a suitable batch has been identified. HOBs will grow in autologous and heterologous human serum, but as yet no comprehensive studies have been performed to identify the effects on the growth and differentiation. 3. Beresford and co-workers introduced the use of this stable analogue of L-ascorbic acid for the generation of mineralising cultures, since it is stable and unlike L-ascorbic acid, does not have to be added to the cultures on a daily basis. 4. The authors have obtained consistent results with plasticware from Sarstedt and Becton Dickinson. Smaller flasks or dishes can be used if the amount of bone available is 1 h). Place lids on dried plates, wrap in plastic, and keep at 4°C for 1–2 months. Before using dried plates, rinse with PBS. 3.3. Fluid Flow using Flexcell ® Streamer ® Shear Stress Device (Flexcell)

1. Plate 2−4 × 105 cells/slide on collagen-coated slides. 2. The next day (day 1), change the medium to include test compounds, or to transfect with siRNA (see Subheading 3.5) if applicable. 3. On day 2, MLO-Y4 cells should be 70–80% confluent and ready for fluid-flow (see Note 5). Wash hands thoroughly before assembling the fluid flow instrumentation. Handle all tubes, parts of the system, and pipette tips carefully making sure they do not come into contact with lab surfaces or skin. When performing the sterilization steps and filling the system with medium, either start the pump from its hardware switch or use the software to choose the manual configuration. The latter is preferable.

3.3.1. To Set Up the Computer System

1. Turn on the computer. 2. Open Stream Gold v1. The Streamer is now under control of the computer and it will read as PO1. 3. Select the “Operate” menu. Add your name as a User by clicking “Add User,” then click “Return.” 4. Select “Operate” menu again, then select “Configure Regimes.” Create procedure by entering values in each step of the new regimen. Click on “Save Step” to save each step, then click “Save Regime” to save the entire procedure. Select the “Visualization” of “Test Profile” tab to verify that the regimen is as expected. Typically we build regimens with a 1 min ramp up to the desired flow intensity, then the entire flow pattern, followed by a 1 min ramp down to zero. This protects cells from undesired shock. 5. On the main screen, select “System 1” tab. Click on “Configure”: this will open a new window. Select the appropriate User, Regime and Hardware (Streamer), and then click “Update.” The regime is now ready to start.

3.3.2. To Set Up the Flow Loop

1. Sterilize the assembled Streamer apparatus with 300 ml of 70% ethanol added to the glass bottle and pumped through the system for 30 min. Check for leaks and if necessary, change rubber seals on the tube connections.

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2. Collect ethanol in the bottle by reverse flow and move it to a used-ethanol container. 3. Rinse the system with 300–400 ml of sterile PBS and pump through the system for at least 5 min and collect in a waste container. 4. Repeat step 3. 5. Place the bottle containing the culture medium (~500 ml) in the FF incubator next to the flow system. Loosen the cap. Take the cap with tube fittings off the empty glass bottle and carefully place the inner tube fittings inside the media bottle without touching anything around it, to maintain optimal sterility. 6. Start pumping the medium through the system to push the remaining PBS out of the flow loop. When the medium reaches the drop off into the bottle and it has pushed most of the PBS out, stop the pump and discard the PBS. 7. Add the rest of the medium. It should be ~500 ml. Cap the bottle again and make sure that it is closed properly. 8. Start pumping the rest of the medium through the system (see Note 6). To create a trap for any excess air that may enter the system, fill each of the pulse dampeners with media by tilting them perpendicular to the flow so that the medium pumping through the system begins to collect there. Start with the pulse dampener closest to the pump and then do the other one. The medium level should be about 1–1.5 inches above the tubing inlets. Check for any bubbles in the tubing and move them out of the system if they are trapped. Also be aware that the Streamer chamber has an air gap spot that will collect bubbles, you need to tilt the chamber on a slant to fill this gap. This is your last chance to check for bubbles and/or leaks. 9. Disconnect the Streamer chamber inlet and outlet tubing and carefully move the system to the tissue culture hood. 10. Remove the screws and open the hinged top. 11. Using sterile forceps and sterile gloves (spray with 70% ethanol), pick up each slide with cells and place it into each one of the slots on the flow device. Make sure the side of the slides with cells is facing the smaller slot, which is the area of flow on the chamber. Gently slide each of them into place and try not to force them in or scratch the sides of the chamber with the glass. All six slots must be filled to ensure proper and consistent flow. If you have less than six slides for an experiment, fill the rest of the slots with empty sterilized slides. 12. Close the chamber, tighten the bolts by hand, and then tighten with the hex wrench. 13. Move the chamber back to the incubator and reconnect the inlet and outlet tubing.

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14. Click start on the software. The profile should already be loaded. A “save as” window will come up, click “cancel.” The flow will start. 15. As the flow is ramping up, tilt the chamber on a slant to help the bubbles in the air gap inside the chamber clear. This should take a few seconds. 16. You will see the shear stress graph building at real time on the software. Periodically monitor the flow system to check for leaks or bubbles. 17. When the program ends, remove the chamber from the incubator and take it to the tissue culture hood. Open the chamber and remove slides for immediate processing, or return to sterile culture conditions for further incubation. Process the cells for example for Western blotting (see Note 7), for RNA isolation, or for analysis of dendrite length (see Subheading 3.4). 18. When you have collected the cells, bring the chamber back to the flow system. Plug in the inlet and outlet tubing and collect all the media into a different container by starting reverse flow. 19. Rinse system with deionized water for 10 min, collect, and discard. 20. Repeat step 18 another two times. 21. Rinse the system with 300 ml of 70% ethanol for 10 min and collect ethanol and discard. 22. Disconnect the tubing, bottle, chamber, and baffles. Open the chamber and clean it with water and ethanol. Make sure that there is no medium residue left on any of the parts, as this will corrode the system. 23. Let the system air-dry until next use. 3.4. Staining and Quantitation of Dendrite Length

Crystal Violet staining is a useful method to enhance the contrast of the dendrites and cell body. See Fig. 2 for an illustration of the method. 1. Fix cell cultures in 10% buffered formalin for 10 min. and wash with H2O. 2. Stain cultures with 0.1% Crystal Violet for 10 min; aspirate and save solution. 3. Gently rinse the cell layer with H2O several times until the solution becomes clear. 4. Drain slides at an angle, or turn plates upside down to air-dry before photographing. Quantitation of dendrite length using the AnalySIS Image Software. 1. Using AnalySIS Image Software (Soft Imaging Systems Corp.), open the photo file to be quantitated.

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Fig. 2. Stained with Crystal Violet, the red line shows the tracing of the cell body, and the green lines, the tracing of the cell extensions. In this manner, the area of the cell body can be quantitated as can the length and number of dendritic processes.

2. Set the “M” magnification at which the original photo was taken. It is very important to make sure that the units are “μm”. Even if there is a “√” next to automated, click on it, then recheck “√” it. This procedure is necessary each time you open a new photo file, so that the dendrite measurements are recorded in “μm” and not pixels. 3. Choose “Measure → Arbitrary area” from the tool bar. Outline the cell body with the cursor (by depressing the left button on the mouse). Double right click the mouse to end the measurement. The software program will automatically generate a data sheet for the cell body area and record the first set of data in column “A.” 4. Outline the dendrites of the same cell (choose “Measure → Polygon length”) using the cursor and using the previously drawn “cell body” as start/stop point. Double right click to end measurement of all dendrites of that cell. The program generates a separate data sheet for the polygon length, and records this first set of data in column “A” also. Double check to make sure that the data measurements are in “μm.” 5. To start measurements on the second cell, click on the appropriate data sheet, and click onto column “B”; otherwise, the program will overwrite or add the new data to the previous measurement. Each individual cell should have a separate column, but this column letter should be the same for both data measurements. (i.e., third cell “C,” fourth cell “D,” etc.).

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6. To erase and redraw, double right click to end the current tracing. Manually delete the numbers from the data sheet. Then click on the picture of the eraser with an “X,” and this will delete the previously drawn layers. 7. To obtain the dendrite Mean for each cell, click on the Polygon length data sheet and choose “Measure → Define Statistics → Mean,” then choose “Measure → Statistics” We combine the dendrite Means from individual cells into any stats program, to get an overall Mean and Std. Error from multiple cells within each sample group. 3.5. E11 siRNA Transfection of MLO-Y4

1. The day before transfection, plate cells at 4 × 104 cells/well in 0.5 ml of growth medium without antibiotics in collagencoated 48 well plates so that they will be 50% confluent at the time of transfection. 2. For each sample, prepare siRNA–Oligofectamine complexes as follows: (a) Dilute 25 nM of each of the 3 siRNA in 40 μl of OptiMEM® I Reduced Serum Medium without serum (or use other medium without serum) (see Note 10). Mix gently. (b) Mix Oligofectamine gently before use, then dilute 1 μl in 9 μl of Opti-MEM® I Medium (or other medium without serum) (see Note 8). Mix gently and incubate for 5 min at room temperature. (c) After the 5 min incubation, combine the diluted siRNA with the diluted Oligofectamine (total volume is 50 μl). Mix gently and incubate for 20 min at room temperature to allow the siRNA–Oligofectamine complexes to form. 3. Add the 50 μl of siRNA–Oligofectamine complexes to each well. Mix gently by rocking the plate back and forth containing 200 μl growth medium without serum. 4. Incubate the cells at 37°C in a CO2 incubator for 24 h until they are ready to assay for gene knockdown. It is generally not necessary to remove the complexes; however, growth medium (1, 2.5, or 5% 50:50 FBS:CS) may be added after 4–6 h without loss of transfection activity, but lower serum may enhance the effect of the siRNA (see Fig. 3). 5. To perform fluid flow on siRNA transfected cells, transfect the cells 24 h after seeding onto collagen-coated slides, then perform fluid flow 24 h later. (a) After 2 h of FF at 16 dynes/cm2, the slides were transferred back to a petri dish with medium containing 2.5% FBS and 2.5% CS. The slides were incubated for another 24 h before assessing dendrite length and/or protein expression.

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Fig. 3. Decreasing serum concentration can enhance the effects of siRNA. Concentrations of serum are indicated above the panels. The Y-axis is percent reduction in protein expression. siRNA = siRNA to E11, RF = Risc free control, Vehicle = no RNA, Cells only = no treatment.

Results: Transfection with siRNA to E11 can inhibit the increase in dendrite length induced by application of fluid flow shear stress. Although no effect was observed on dendrite lengthening in those transfected cells which remained under static culture conditions, the protein expression was significantly decreased in subsequent Western analysis for E11 (14). These experiments could be performed to determine if E11 or dendrites play a role in any osteocyte functions such as gap junction or hemichannel function, cell signaling, or osteocyte apoptosis. The same siRNA approach was used to show the role of β-catenin in osteocyte apoptosis and viability (15). 3.6. General Maintenance of MLO-A5 Late Osteoblast/Early Osteocyte-Like Cell-Line

All cell culture procedures should be performed in a Tissue Culture hood under aseptic conditions, and using sterile supplies. 1. Maintain cultures in α-MEM culture medium containing 5% FBS and 5% CS under sub-confluent conditions (85–90%) on collagen-coated plates. Grow in a humidified incubator at 37°C, at 5% CO2. 2. Passage at 1:15–1:20 every 3–4 days. 3. Freeze cells in 60% α-MEM, 30% FBS, 10% DMSO, at 1 × 106 cells/ml/cryovial.

3.7. Mineralization of MLO-A5

Please note that the pattern of mineralization obtained with MLOA5 is different from that described in other chapters (e.g., see Chapter 3, this volume). Here, mineralization is in sheets rather than in nodules. 1. For consistency, perform the experiment within 3–4 passages of defrosting a new batch of cells. 2. Plate cells into collagen-coated wells at ~3.5 × 104 cells/cm2, so the wells are 100% confluent in 2 days. Use α-MEM containing 5% FBS and 5% CS.

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3. At confluence (day 0), change the medium to mineralization medium: α-MEM/10% FBS containing 100 μg/ml ascorbic acid and 4–5 mM β-glycerophosphate (see Note 9). 4. Change the media in the wells every 2–3 days. Aspirate the spent medium using a sterile, beveled needle, taking care not to touch the cell layer. If leaving a small amount of the old medium (~100 μl is acceptable) make sure to be consistent between all wells. Work with four wells at a time so that the wells do not dry out. 5. On day 4–6, collagen fibrils start forming into a swirling, honeycomb pattern, which takes on a “shiny” and refractile appearance as the culture begins to mineralize. 6. Optimal mineralization is usually around day 10–12. 7. To harvest the cultures for Western blotting, you will need to use a 23 gauge needle attached to a syringe to vigorously mix the lysate (10–20 times), to break up the mineralized matrix. See Chapter 15, this volume, for details on Western blotting methods. 8. To harvest the cultures for mineralization, the cells are fixed in 10% buffered formalin for 10 min. and washed with H2O, before proceeding to Von Kossa or Alizarin Red staining. Alternatively the cultures are fixed in 95% ETOH for immunofluoresence staining.

4. Notes 1. Growing the cells in both types of serum is very important, as the CS maintains cell proliferation, while the FBS maintains cell differentiation. Switching to a new batch of serum, especially the FBS, can result in changes in both morphology and phenotypic expression. We suggest testing several new serum lots to make sure that the cells are performing as in the previous serum batch. We suggest you assay cell proliferation, cell morphology, dendrite length, gene and/or protein expression, and if applicable, the ability to mineralize. For heat inactivation, consult the Hyclone Web site. 2. For fluid flow, use positively charged glass slides (Thermo Shandon Colorfrost* Plus Slides cat#6776214) and collagen coat (C.C.) to insure optimal attachment of the cells onto the glass slides. Because the dimensions of the glass slides may be slightly different, you will need to pretest each slide in the Streamer chamber. If you encounter resistance, do not force it in, or it could break and obstruct the chamber. Sterilize the glass slides prior to C.C. by soaking in 70% ethanol or 100%

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isopropanol for at least 30 min. Air-dry or rinse in PBS before proceeding. (You can also sterilize the slides by wrapping and autoclaving.) To C.C., refer to the procedure given in Subheading 3.2. Place the slides in a sterile tissue culture dish, or other sterile container for coating. After coating, transfer the slides to sterile, square petri dishes using sterile forceps. Lean the slides on the side of the dish until they are dry before laying them flat inside the dish, otherwise, they will “stick” to the bottom, and will be difficult to remove later. 3. The MLOY4 osteocyte-like cells, which grow slower than osteoblast-like cell-lines, prefer to only make contact through their dendritic processes, not through their cell bodies. If grown too confluent, this can affect the dendritic characteristics, and the cells will start detaching. Since the cells do not produce their own extracellular matrix, the cells are maintained on collagen-coated plates to help maintain this dendritic morphology. 4. Try to limit the medium changes by providing enough medium at passage to last 3–4 days. Use 12 ml/100 mm dish or 30 ml/150 mm dish. Two reasons we recommend this are that (a) the cells love their own “conditioned media,” and actually become more dendritic, and (b) previously, we observed that frequent medium changes causes morphological changes in the cells. This is particularly important in experiments measuring cell body area or dendrite length. 5. We recommend letting the cells adhere to the glass slide for 48 h (but not longer than that), before performing fluid flow. Depending on the purpose of the experiment, and the endpoint, you may need to change the initial plating density. If the endpoint is to measure dendrite length, you want there to be adequate space between the cells so the dendrites have the space to elongate. 6. It is very important to be careful in filling the flow system. You need to fill the two plastic baffles (pulse dampeners) about 2/3 full. This is crucial to keep vibrations from the pump and bubbles from disturbing the cells. If you fail to adequately fill these baffles, the experiment is likely to fail, as the air bubbles dislodge cells off the slides. 7. For Western blotting, the general procedure given in Chapter 15, this volume, is used. To detect the osteocyte-specific marker E11, we use the hamster monoclonal antibody 8.1.1 from hybridoma conditioned media – a kind gift of Dr. Andrew Farr at the University of Washington. Dr. Farr has deposited 8.1.1 (anti-murine GP38; T1alpha; podoplanin) and the corresponding hybridoma with the Developmental Studies Hybridoma Bank (http://www.uiowa.edu/~dshbwww/) and

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it is available from there on request. This antibody works well in combination with peroxidase-conjugated goat anti-Syrian hamster IgG- (ImmunoResearch Laboratories Inc). 8. Oligofectamine is the preferred solution. Lipofectamine 2000, Lipofectamine Plus (Invitrogen,Carlsbad, CA), and TransITTKO Transfection reagent (Mirus, Madison, WI) affected cell morphology. 9. Ascorbic acid and β-glycerophosphate are added to the culture medium on the day of feeding. Make 100× stock solutions in medium without serum, filter-sterilize using a syringe filter, make small aliquots, wrap in foil, and freeze at −20°C. Aliquots are stable for several months. Since ascorbic acid is light sensitive, we tend to discard the thawed aliquot after use. The thawed aliquot of β-GP can be stored at 4°C and used for a week. 10. In this case, the initial dilution of the siRNA complex should be made as a 5X solution (at 125nM), so after the 50 μl complex is added to the 200 μl of growth medium, the final concentration of siRNA is 25nM. References 1. Mikuni-Takagaki, Y., Kakai, Y., Satoyoshi, M., Kawano, E., Suzuki, Y., Kawase, T., and Saito, S. (1995) Matrix mineralization and the differentiation of osteocyte-like cells in culture. J. Bone Miner. Res. 10, 231–242. 2. Bonewald, L. F. (2007) Osteocytes, In Osteoporosis (R. Marcus, D. Feldman, D. Nelson, C. Rosen, Ed.) 3 rd ed., pp 169–190, Elsevier. 3. Bonewald, L. F., Johnson, M. L. (2008) Osteocytes, Mechanosensing, and Wnt Signaling. Bone 42, 606–615. 4. Dallas, S. L., Bonewald, L. F. (2010) Dynamics of the Transition from Osteoblast to Osteocyte, Ann. N Y Acad. Sci 1192, 434–437. 5. Klein-Nulend, J., Bonewald, L. F. (2008) The Osteocyte, In Principles of Bone Biology (Bilezikian, J. P., Raisz, L.G., Ed.), Academic Press. 6. Kato, Y., Windle, J. J., Koop, B. A., Mundy, G. R., and Bonewald, L. F. (1997) Establishment of an osteocyte-like cell line, MLO-Y4, J. Bone Miner. Res. 12, 2014–2023. 7. Kato, Y., Boskey, A., Spevak, L., Dallas, M., Hori, M., and Bonewald, L. F. (2001) Establishment of an osteoid preosteocyte-like cell MLO-A5 that spontaneously mineralizes in culture. J. Bone Miner. Res. 16, 1622–1633. 8. Barragan-Adjemian, C., Nicolella, D., Dusevich, V., Dallas, M. R., Eick, J. D., and Bonewald, L. F. (2006) Mechanism by

9.

10.

11.

12.

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which MLO-A5 late osteoblasts/early osteocytes mineralize in culture: similarities with mineralization of lamellar bone. Calcif. Tissue In. 79, 340–353. Bellido, T., Ali, A. A., Gubrij, I., Plotkin, L. I., Fu, Q., O’Brien, C. A., Manolagas, S. C., and Jilka, R. L. (2005) Chronic elevation of parathyroid hormone in mice reduces expression of sclerostin by osteocytes: a novel mechanism for hormonal control of osteoblastogenesis. Endocrinology 146, 4577–4583. Papanicolaou, S. E., Phipps, R. J., Fyhrie, D. P., and Genetos, D. C. (2009) Modulation of sclerostin expression by mechanical loading and bone morphogenetic proteins in osteogenic cells. Biorheology 46, 389–399. Dallas, S. L., Veno, P. A., Rosser, J. L., Barragan-Adjemian, C., Rowe, D. W., Kalajzic, I., and Bonewald, L. F. (2009) Time lapse imaging techniques for comparison of mineralization dynamics in primary murine osteoblasts and the late osteoblast/early osteocyte-like cell line MLO-A5. Cells, tissues, organs 189, 6–11. Sittichockechaiwut, A., Scutt, A. M., Ryan, A. J., Bonewald, L. F., and Reilly, G. C. (2009) Use of rapidly mineralising osteoblasts and short periods of mechanical loading to accelerate matrix maturation in 3D scaffolds. Bone 44, 822–829. Schacht, V., Ramirez, M. I., Hong, Y. K., Hirakawa, S., Feng, D., Harvey, N., Williams, M., Dvorak, A. M., Dvorak, H. F., Oliver, G., and

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Detmar, M. (2003) T1alpha/podoplanin deficiency disrupts normal lymphatic vasculature formation and causes lymphedema. EMBO J. 22, 3546–3556. 14. Zhang, K., Barragan-Adjemian, C., Ye, L., Kotha, K., Dallas, M., Lu, Y., Zhao, S., Harris, M., Harris, S. E., Feng, J. Q., and Bonewald, L. F. (2006) E11/gp38 Selective Expression in osteocytes: regulation by mechanical strain and

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role in dendrite elongation. Mol. Biol Cell. 26, 4539–4552. 15. Kitase, Y., Barragan, L., Qing, H., Kondoh, S., Jiang, J., Johnson, M. L., Bonewald, L. F. (2010) Mechanical induction of PGE2 in osteocytes blocks glucocorticoid induced apoptosis through both the b-Catenin and PKA pathways. J. Bone Min. Res. 25, 2657–2668.

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Chapter 7 Isolation, Differentiation, and Characterisation of Skeletal Stem Cells from Human Bone Marrow In Vitro and In Vivo Rahul S. Tare, Peter D. Mitchell, Janos Kanczler, and Richard O.C. Oreffo Abstract In this chapter, we describe techniques for the isolation and characterization of skeletal stem cells from human bone marrow. The methods for enrichment of STRO-1 positive cells using magnetic activated cell sorting are described and we also cover techniques for establishing and characterising osteogenic, adipogenic, and chondrogenic cultures from these cells. Finally, we present methods for studying the ability of these cells to produce bone in vivo using diffusion chambers which have been implanted subcutaneously in mice. Key words: Stem cells, Osteoblast, Adipocyte, Chondrocyte, Bone marrow, Diffusion chamber

1. Introduction Adult human bone marrow stromal tissue contains multipotent progenitor cells referred to as skeletal stem cells (or more commonly mesenchymal stem cells), which are capable of differentiating into osteoblasts, chondrocytes, adipocytes, and myocytes. Mesenchymal stem cells can be identified by virtue of the fact that they express various cell surface markers which can be detected by specific antibodies. The most widely used of these is the monoclonal antibody STRO-1, which recognises a cell-surface trypsinresistant antigen expressed by a sub-population of bone marrow stromal cells that essentially includes all adherent high growthpotential clonogenic progenitors or fibroblastic colony-forming units CFU-F (1). This chapter provides an update on the methodologies for the isolation and enrichment of STRO-1 positive skeletal stem cells from bone marrow using immunolabelling followed by Miep H. Helfrich and Stuart H. Ralston (eds.), Bone Research Protocols, Methods in Molecular Biology, vol. 816, DOI 10.1007/978-1-61779-415-5_7, © Springer Science+Business Media, LLC 2012

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magnetic isolation. We also describe the techniques for differentiation of these cells down the osteogenic, chondrogenic and adipogenic lineages in vitro and the techniques that can be used to characterise the differentiated cells. Finally, we describe the methods for the in vivo analysis of stem cells by the use of diffusion chambers in nude mice.

2. Materials 2.1. Equipment for Tissue Culture and Stromal Cell Isolation

1. Class II tissue culture cabinet. 2. Microscope. 3. Incubator. 4. Centrifuge. 5. LS separation columns (Miltenyi Biotec). 6. VarioMACS assembly (Miltenyi Biotec).

2.2. Equipment for In Vivo Diffusion Chamber Experiments

1. Warming pad and sterile cover dressings. 2. Recovery incubator (temperature 28°C). 3. Michel clips & clip stapler. 4. Sterile forceps, scalpel blades No 10, scalpel blade holder, blunt dissection scissors, and artery forceps. 5. Diffusion chamber, outer diameter 14 mm, inner diameter of 10 × 2 mm (Millipore). 6. Mixed cellulose ester membranes, 13 mm diameter with 0.45 μm pore size (Millipore). 7. Cement fixative (Millipore).

2.3. Stromal Cell Isolation and Culture

1. α-MEM. 2. Phosphate-buffered saline (PBS). 3. Lymphoprep. 4. Foetal calf serum (FCS). 5. Trypsin/EDTA 0.05% Trypsin/0.02% EDTA. 6. 2% (w/v) Collagenase Type IV in α-MEM. 7. Normal human AB serum. 8. Hank’s buffered saline solution (HBSS) with 10 mM HEPES, pH 7.4. 9. Blocking buffer (HBSS with 10 mM HEPES, 10% normal human AB serum, 5% FCS, and 1% bovine serum albumin) (see Note 1).

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10. MACS buffer (2 mM ETDA in PBS with 0.5% (w/v) bovine serum albumin) (see Note 2). 11. Anti-human monoclonal STRO-1 antibody (R&D Systems). 12. Rat anti-mouse IgM-coated Microbeads (Miltenyi Biotec). 2.4. Stromal Cell Differentiation

1. Insulin, Transferrin, & Sodium selenite (ITS) medium supplement (Sigma). 2. 3-Isobutyl-1-1-methylxanthine (IBMX) (Sigma). 3. Human recombinant TGF-β3 (Calbiochem). 4. Osteogenic medium: α-MEM with 10% FCS, 100 μM Ascorbate-2-phosphate, 25 nM 1α, 25-dihydroxy Vitamin D3, and 1.8 mM Potassium dihydrogen phosphate). 5. Chondrogenic medium: α-MEM with 100 μM Ascorbate-2phosphate, 10 nM Dexamethasone, 1% ITS medium supplement, and 10 ng/ml TGF-β3. 6. Adipogenic medium: α-MEM with 3 g/l D+ Glucose, 10% FCS, 1% ITS, 1 μM Dexamethasone, 0.5 mM IBMX, and 100 μM Indomethacin. 7. Adipogenic insulin only medium: α-MEM with 3 g/l Glucose, 10% FCS, and 1% ITS.

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2.5. Alkaline Phosphatase Staining

1. 90% Ethanol.

2.6. Alkaline Phosphatase Activity

1. Assay buffer (33% 2-AMP alkaline buffer solution with 0.2% Igepal CA-630.

2. Alkaline Phosphatase staining solution (4% (v/v) Naphthol AS-MX solution with 0.024% (w/v) Fast Violet Salt) (see Note 3).

2. Assay substrate: 3.6 mM 4-Nitrophenyl phosphate disodium salt hexahydrate in 33% 2-AMP alkaline buffer solution. 3. 1 M NaOH. 4. 4-Nitrophenol. 5. Triton X-100 0.05% (v/v) in distilled water. 2.7. DNA Quantification

1. 10 mg/ml DNA stock (BDH laboratories). 2. 1× Tris/EDTA solution. 3. 0.5% PicoGreen® solution in 1× Tris/EDTA.

2.8. Von Kossa Staining

1. 1% (w/v) Silver nitrate in distilled water. 2. 2.5% (w/v) Sodium thiosulphate in distilled water. 3. Van Gieson’s stain: (0.09% (w/v) Acid Fuchsin in 50% (v/v) picric acid).

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2.9. Safranin O Staining

1. Weigert’s Iron Haematoxylin. Solution A: 1% (w/v) Haematoxylin in methanol. Solution B: 1.2% (w/v) Ferric chloride in 1% (v/v) concentrated HCl. Mix equal volumes of solutions A and B just before use. 2. Acid alcohol: 1% (v/v) concentrated HCl in methanol. 3. 0.001% (w/v) Fast green. 4. 1% (v/v) Acetic acid. 5. 0.1% (w/v) Safranin O.

2.10. Immunostaining for Type II Collagen and SOX-9

1. Rabbit anti-human SOX-9 primary antibody (Chemicon AB5535). 2. Rabbit anti-rodent/human Type II collagen primary antibody (Calbiochem 234187). 3. Biotinylated goat anti-rabbit secondary antibody (DakoCytomation E0432). 4. 0.01 M Sodium Citrate buffer, pH 6.0. 5. Hyaluronidase solution: 0.08% (w/v) hyaluronidase in PBS with 1% (w/v) BSA. 6. PBS with 1% (w/v) BSA. 7. 3% Hydrogen peroxide. 8. 30% Hydrogen peroxide. 9. Glacial acetic acid. 10. Histo-Clear. 11. Histology grade Methanol. 12. High salt solution: 400 mM NaCL, 50 mM Tris, 0.05% Tween20 (v/v) in dH2O. Adjust to pH 8.5. 13. Low salt solution: 150 mM NaCL, 50 mM Tris, 0.05% Tween20 (v/v) in dH2O. Adjust to pH 8.5. 14. Tris buffer solution: 100 mM Tris, 0.05% Tween-20 (v/v) in dH2O, pH 8.5. 15. ExtrAvidin Peroxidase solution: ExtrAvidin Peroxidase 2% (v/v) in PBS with 1% (w/v) BSA. 16. Substrate mixture: 50 mM AEC (3 amino-9-ethyl-carbazole) in dimethylformamide. Solution can be kept in fridge for up to 1 week. 17. Working substrate: dilute 0.5 ml substrate mixture in 9.5 ml 50 mM acetate buffer (pH 5.0) and add 5 μl of 30% H2O2. 18. Light Green stain: 2.5 mM Light Green in 2 mM acetic acid. Stir well and filter before use. 19. Alcian blue stain: 4 mM Alcian blue 8GX in 10 mM acetic acid. Stir well and filter before use. Staining solution can be used for approximately 1 week. 20. Crystal mount.

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1. 60% isopropanol. 2. 1% (v/v) Saturated Oil Red O in Isopropanol Stock solution. 3. Oil Red O working solution (see Note 4). 4. Formal Calcium Solution: (11% (v/v) formaldehyde in 1% Calcium Chloride.

2.12. Tissue Implant Models

1. Nude Mice (MF-1 nu/nu strain). 2. Class 1 flow cabinet. 3. Injectable anaesthetic (1:1 mixture of Hypnorm and Midazolam in sterile water).

3. Methods 3.1. Isolation of Bone Marrow Mononuclear Cells

The procedure is based upon immunomagnetic isolation of STRO-1 positive cells from a whole population of bone marrow mononuclear cells using a column that is placed in the magnetic field of a MACS separator as described by Stewart et al. (2). The magnetically labelled STRO-1 positive cells are held in the column under the influence of the magnetic field, while the unlabelled STRO-1 negative cells are eluted by repeated washes of the column. On removal of the column from the magnetic field, the magnetically labelled STRO-1 positive cells are eluted as a STRO1-enriched fraction. These cells can be cultured directly and characterised by immunostaining for stromal cell or stem cell markers (Fig. 1), or differentiated down the osteogenic (Subheading 3.4), chondrogenic (Subheading 3.5) or adipogenic (Subheading 3.6) lineages (Fig. 2). Whatever the researcher chooses to do, these procedures must be performed in a Class II laminar flow hood. 1. Obtain suitable human bone marrow tissue containing pieces of trabecular bone (see Note 5) and transfer into a 50-ml Falcon tube. Add 5 ml α-MEM to the sample and shake the sample vigorously for a couple of minutes. 2. Allow the sample to stand for a minute and pipette out the supernatant cell suspension into another 50-ml Falcon tube (see Note 6). 3. Repeat this process three to four times until most of the cells are released and the debris has been removed from the bone marrow sample. 4. Add α-MEM to make the volume of the cell suspension up to 50 ml and centrifuge at 250 × g for 5 min at 4°C. 5. Discard the supernatant taking care not to dislodge the cell pellet at the bottom of the Falcon tube (see Note 7).

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Fig. 1. Day 6 cultures of adult human bone marrow-derived STRO-1 positive skeletal stem cells in basal medium exhibiting robust immunoreactivity to the STRO-1, anti-Vimentin (primitive mesenchymal cell marker) and anti-SDF-1/Stromal cellDerived Factor-1 (stromal cell marker) antibodies. In addition to the STRO-1 antigen, cells of day 6 cultures co-express the CD105 antigen, a recognised marker of skeletal progenitor cells. Scale bar: 50 μm.

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Fig. 2. Multipotential ability of adult human bone marrow-derived STRO-1+ skeletal stem cells demonstrated by the expression of alkaline phosphatase in osteogenic culture conditions, formation of 3-D pellets, in chondrogenic medium, cells of which synthesise abundant proteoglycans (stained with Safranin O) and Type II collagen, and generation of adipocytes containing lipid droplets in adipogenic culture medium. Scale bar: 100 μm.

6. Resuspend the cell pellet in 10–15 ml α-MEM and filter the cell suspension into a fresh 50-ml Falcon tube through a 70–75-μm filter (see Note 8). 7. Add α-MEM to the filtered cell suspension to bring the volume to 25 ml and layer gently over 25 ml Lymphoprep which has been pre-warmed to room temperature (see Note 9). 8. Centrifuge at 800 × g for 40 min at 18°C with the centrifuge brake set to off. 9. Carefully remove the mononuclear cells from the interphase using a sterile pastette into another 50 ml Falcon tube and suspend the cells in 25 ml fresh α-MEM. Centrifuge at 250 × g, for 10 min at 4°C to remove any residual Lymphoprep. 10. Resuspend the cell pellet in 25 ml α-MEM and perform a cell count (see Note 10). 11. Pellet the cells by centrifugation at 250 × g for 10 min at 4°C. 3.2. Antibody Labelling of STRO-1 Positive Cells

1. Resuspend the cell pellet derived from Subheading 3.1, step 11 in 10 ml blocking solution. 2. Incubate for 30 min at 4°C, taking care to shake the tube every 10 min to avoid the cells from clumping and settling at the bottom of the tube (see Note 11).

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3. Centrifuge the cell suspension at 250 × g for 5 min at 4°C, and resuspend the cell pellet in 1 ml STRO-1 antibody at a concentration of 10 μg/ml (see Note 12). 4. Incubate the cell suspension/antibody solution for 30 min at 4°C, taking care to shake the tube every 10 min to avoid the cells from clumping and settling to the bottom of the tube (see Note 11). 5. Centrifuge the sample at 250 × g for 5 min at 4°C. 6. Remove the supernatant and resuspend the cell pellet in 50 ml MACS buffer which has been chilled to 4°C and centrifuge at 250 × g for 5 min at 4°C. 7. Repeat step 6. 8. Resuspend the cell pellet in 800 μl fresh MACS buffer and add 200 μl rat anti-mouse IgM Microbeads and mix thoroughly by gentle pipetting. 9. Incubate the cells in the secondary antibody solution for 30 min at 4°C, taking care to shake the tube every 10 min to avoid the cells from clumping and settling to the bottom of the tube (see Note 11). 10. Centrifuge the sample at 250 × g for 5 min at 4°C. 11. Remove the supernatant and wash the cells in chilled MACS buffer as described in step 6. 12. Resuspend the cell pellet in 3 ml MACS buffer and perform cell count. 3.3. Magnetic Separation of STRO-1 Positive Cells

1. Place the LS separation column in the magnetic field of the VarioMACS and place a 15 ml collection tube underneath. 2. Prepare the column by washing it with 3 ml MACS buffer. 3. Add 3 ml of immunolabelled cell suspension to the column and wait until the cell suspension passes through the column into the collection tube. 4. Wash the column three times with 3 ml MACS buffer. 5. Remove the column from the VarioMACS assembly and place in a fresh collection tube. Add 5 ml fresh MACS buffer and firmly flush out the STRO-1 positive cell fraction using the plunger supplied with the column. Suspend the eluted cells in 10 ml fresh MACS buffer. 6. Centrifuge the STRO-1 positive cell suspension at 250 × g for 5 min at 4°C and resuspend the resulting cell pellet in α-MEM supplemented with 10% FCS. 7. Place the cells in culture in α-MEM supplemented with 10% FCS for further characterisation and or differentiation as described in Subheadings 3.4–3.6.

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1. Seed 2 × 106 STRO-1 positive cells derived from Subheading 3.3 into a 25 cm3 tissue culture flask. 2. Maintain the cells in culture in α-MEM with 10% FCS for 9–12 days until they are between 50 and 80% confluent. 3. Aspirate the culture medium and replace with osteogenic medium (Subheading 2.4). 4. Continue the cultures for up to 28 days with changes of medium every 3 days. 5. Terminate the cultures and assay for evidence of osteoblast differentiation (alkaline phosphatase activity, bone nodule formation) as detailed in Subheadings 3.7–3.11.

3.5. Chondrogenic Differentiation of STRO-1 Positive Cells

1. Seed 2 × 106 STRO-1 positive cells derived from Subheading 3.3 into a 25 cm3 flask and culture in α-MEM supplemented with 10% FCS until the cells are about 90% confluent. 2. Remove the medium, wash with PBS, and add Trypsin/EDTA to detach the cell layer (see Note 13). 3. Pellet cells by centrifugation at 250 × g for 5 min at 4°C and resuspend the cell pellet in 5 ml α-MEM. Perform a cell count and adjust the volume so that the cell density is 5 × 105 cells/ml. 4. Add 1 ml of the cell suspension to a sterile 25 ml polycarbonate universal container. 5. Pellet the cells by centrifugation at 250 × g for 10 min at 4°C. 6. Carefully aspirate the supernatant, taking care not to disturb the cell pellet and gently add 1 ml of chondrogenic medium (Subheading 2.4) to each universal container. 7. Place the containers in a humidified CO2 incubator at 37°C with the lids slightly open for 21–28 days, with changes of medium every 2 days. 8. Terminate the experiment and analyse the cultures for evidence of chondrogenic differentiation as described in Subheadings 3.12 and 3.13.

3.6. Adipogenic Differentiation of STRO-1 Positive Cells

This method is used to generate adipocytes from STRO-1 positive cells by alternating periods of culture in adipogenic and insulin only medium. 1. Seed 2 × 106 STRO-1 positive cells derived from Subheading 3.3 into a 25 cm3 flask and culture in α-MEM in supplemented with 10% FCS until the cells are sub confluent. 2. Culture in adipogenic medium (Subheading 2.4) for 3 days and wash the cultures with PBS. 3. Culture in insulin only medium (described in Subheadings 2.4– 2.12) for 1 day and wash the cultures with PBS.

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4. Repeat steps 2 and 3 twice. 5. Continue the cultures in insulin only medium with medium changes every 3 days until adipocytes are observed in the cultures (see Note 14). 6. Terminate the experiment and analyse the cultures for evidence of adipogenic differentiation as described in Subheading 3.14. 3.7. Staining for Alkaline Phosphatase activity

1. Wash the tissue culture plates in PBS and fix in 90% ethanol for 15 min. 2. Remove ethanol and allow the plates to air-dry. 3. Add sufficient ALP staining solution to cover each sample (approximately 3 ml for a 25 cm3 flask). 4. Incubate at 37°C until red staining develops in the cells.

3.8. Analysis of Alkaline Phosphatase activity in cell lysates

1. Wash the tissue culture plates or flask in PBS and fix the cells by adding sufficient 90% ethanol to cover the cell layer and incubate for 15 min. 2. Remove ethanol and allow samples to air-dry. 3. Add sufficient 0.05% Triton-X-100 solution to cover each well in the culture plate. 4. Homogenise the cell layer using a cell scraper. 5. Freeze the plate at −20°C for 30 min and thaw at 37°C for 5 min. 6. Repeat step 5 three times, homogenising the cell layer using a cell scraper after each repeat. 7. Proceed to measure ALP activity and DNA content as described in Subheadings 3.9 and 3.10.

3.9. Measurement of Alkaline Phosphatase Activity

1. Prepare a standard curve by adding 100 μl 4-nitrophenol to the 96 well plate in triplicate over the concentration range 0–200 nmol/ml. 2. Add 10 μl lysate to each well of a 96 well plate in triplicate. 3. Add 90 μl 4-nitrophenol substrate to each well containing cell lysate (do not add to standards). 4. Note the time and incubate the plate at 37°C until the sample wells start to change colour to yellow. 5. Add 100 μl 1 M NaOH to all wells to stop the reaction and record the time elapsed. 6. Transfer the plate to a colorimetric plate reader and read at 410 nm absorbance. 7. Calculate the alkaline phosphatase activity as the amount of substrate converted per minute in relation to the standard curve. 8. Express the alkaline phosphatase activity as a function of DNA content as described in Subheading 3.10.

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1. Prepare a standard curve of DNA over the concentration range 0–1,000 ng and add 100 μl of each standard to a black 96-well cyto-fluor plate in triplicate. 2. Add 10 μl of cell lysate to each well in triplicate and add 90 μl of Tris/EDTA to each of the wells containing cell lysate. 3. Add 100 μl of 0.5% PicoGreen solution to all wells. 4. Read in a fluorimeter at 485 nm excitation and 530 nm emission. 5. Work out the DNA concentration in each sample well by comparing to the standard curve.

3.11. Von Kossa Staining for Bone Nodules and Osteoid

This method is used to visualise mineralised bone nodules and osteoid. The mineralised bone stains black and the osteoid red. 1. Add 3 ml silver nitrate per 25 cm3 flask. 2. Place under UV light for 20 min. 3. Wash thoroughly using ultra-pure water. 4. Add 3 ml sodium thiosulphate to each flask and leave for 8 min at room temperature. 5. Wash thoroughly using ultra-pure water. 6. Add 3 ml van Gieson stain and leave for 5 min at room temperature. 7. Remove any excess van Gieson stain from the flask by rinsing gently with ultra-pure water.

3.12. Safranin O Staining of Chondrocytes Cultures

This method is used to study production of cartilage glycosaminoglycans in sections of the chondrocyte cultures described in Subheading 3.5. The glycosaminoglycans stain orange whereas the cytoplasm of cells stain green and the nuclei black. 1. Fix the chondrogenic pellets by immersing in 90% ethanol or 4% paraformaldehyde overnight at 4°C. 2. Process the sections through graded methanol and chloroform solutions, embed in paraffin wax and prepare 7 μm sections for analysis using a microtome. 3. Deparaffinise the sections by immersing in Histoclear for 7 min and repeat once. 4. Immerse the sections on 100% methanol for 2 min and repeat once. 5. Repeat step 3 with 90% methanol. 6. Repeat step 4 with 50% methanol. 7. Repeat step 5 with water. 8. Add Weigert’s Iron Haematoxylin to the sections and incubate for 10 min.

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9. Wash the sections in running water for 10 min. 10. Clear the sections by dipping slides three-times in acid alcohol. 11. Wash in running water for 10 min. 12. Stain with Fast green for 5 min. 13. Rinse quickly in 1% acetic acid for no more than 10–15 s. 14. Stain with 0.1% Safranin O for 5 min. 15. Immerse in 100% methanol for 30 s and repeat once. 16. Immerse in 90% methanol for 30 s and repeat once. 17. Immerse in 50% methanol for 30 s and repeat once. 18. Immerse in Histoclear for 30 s and repeat once. 19. Mount by adding a drop of DPX to each section and gently drop a coverslip on top, taking care to avoid introducing any air bubbles. 3.13. Immunolocalisation of SOX-9 and Type II Collagen in Chondrocyte Cultures

This method is used to study production of the chondrocyte specific markers SOX-9 and type II collagen in sections of the chondrocyte cultures described in Subheading 3.5. For immunolocalisation of SOX-9, a microwave step is required for antigen retrieval just before addition of the primary antibody, whereas for type II collagen the sections should instead be incubated with hyaluronidase at this point. 1. Chondrogenic pellets are fixed in 90% ethanol or 4% paraformaldehyde overnight at 4°C, processed through graded methanol and chloroform solutions, embedded in paraffin wax and sequentially sectioned on the microtome at 7 μm. 2. Deparaffinise the sections by immersing in histoclear for 2 × 7 min. 3. Hydrate the sections by immersing in 100% methanol for 2 × 2 min, followed by 90% methanol 50% methanol, then dH2O, each for 2 min. 4. If the sections are to be stained for SOX9, follow step 4.1. If the sections are to be stained for type II collagen, follow step 4.2: 4.1. (Sox9 only) Immerse the slides in 0.01 M sodium citrate buffer in an appropriate container with a lid, put into a microwave and heat for 5 min at full power. Wash the slides in distilled water for 2 min and proceed to step 5. 4.2. (Type 2 collagen only) Add sufficient hyaluronidase solution to cover each section and incubate for 20 min at 37°C. Wash the slides by rinsing in running water and proceed to step 5. 5. Place slides flat on a staining tray, cover sections with 3% H2O2, incubate for 5 min, and wash briefly in running water.

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6. Add a drop of 1% BSA in PBS to each section and leave for 15 min. 7. Dilute the SOX9 (1:150) and collagen type II (1:500) primary antibody in PBS with 1% BSA and add enough antibody solution to cover each section (see Note 15). 8. Incubate the slides in a lidded staining tray for 1–3 h at room temperature or overnight at 4°C (see Note 16). 9. Rinse gently in running water and immerse slides in High Salt, then Low Salt, then Tris buffer for 5 min each. 10. Drain the slides and add sufficient diluted biotinylated secondary antibody (1:100) to cover each section. Incubate the slides in a lidded staining tray for 1–2 h at room temperature. 11. Repeat step 9. 12. Drain slides and cover the sections with ExtrAvidin Peroxidase solution and incubate for 30 min. 13. Repeat step 9. 14. Add sufficient AEC substrate buffer to cover each section and monitor the slides for the development of a colour change to brown (maximum incubation of 10 min). 15. Place the slides in a staining rack and place into the running water bath to terminate the reaction. 16. Add Light Green to the sections to counterstain and incubate for 1 min. 20 s (see Note 17). 17. Rinse in water bath and gently dab any excess droplets of water from the section using tissue paper. 18. Place a few drops of crystal mount directly onto the section and incubate at 37°C until set (approx. 1–2 h). Do not apply a coverslip. 3.14. Oil Red O Staining

The Oil red O method is used to visualise lipid droplets in adipocytes which stain bright red. 1. Fix cultures by adding sufficient Formal Calcium solution to cover the cell layer. 2. Rinse cells with 60% isopropanol. 3. Add 1 ml Oil Red O working solution per well or enough to cover flask base and leave to stain for 15 min. 4. Rinse three times with excess dH2O to remove lipid droplets released from cells, then add 1 ml dH2O or PBS to wells to visualise under light microscope, taking care not to force water from pipette to prevent damage to the cell monolayer. 5. Photograph within 1 h.

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3.15. Subcutaneous Implant Assay and Diffusion Chamber Assay

This method is used to generate skeletal tissue in vivo by subcutaneous implantation of a diffusion chamber containing skeletal stem cells into a nude mouse or by directly implanting samples of skeletal tissue into a nude mouse (see Note 18) (3, 4). The resulting tissue can then be analysed by various techniques such as imaging, histology and immunohistochemistry (5, 6). The whole procedure is carried out in a sterile operating theatre or in a class I tissue culture cabinet. 1. Sterilise the individual diffusion chamber ring sections overnight using UV exposure. 2. Assemble the diffusion chamber apparatus by attaching the ring to a membrane using the cement fixative (Fig. 3). 3. Add the cells, cell pellets, or bone constructs to the chamber and then add the other side membrane and fix using the cement (Fig. 3). 4. Submerge the constructs in tissue culture media and incubate at 37°C in 5% CO2/balanced air until ready for implantation. 5. Anaesthetise the mice by administering the anaesthetic at a dose of 10 ml/kg intraperitoneally using a 25 G needle and 1-ml syringe. 6. Place the mice on a warming pad, swab the back with an alcohol wipe and make a longitudinal 1–2 cm incision on one side of the vertebral column using a N°10 scalpel blade. 7. Create a subcutaneous pouch lateral to the vertebral column using blunt dissection large enough to contain the bone cell based implants or diffusion chambers required for the experiment. Scaffold Chamber rings

Cells Filter membrane Fig. 3. A schematic diagram of the diffusion chamber set up for in vivo subcutaneous implantation of skeletal scaffold constructs and cells.

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Fig. 4. A selective laser sintered scaffold (arrows) seeded with human foetal femur derived cells subcutaneously implanted into MF-1nu/nu mice for 28 days (a). The asterisk denotes the skin. Alcian blue (proteoglycans) and Sirius red (collagen) histological staining of a section of a selective laser sintered scaffold seeded with human foetal femur derived cells subcutaneously implanted into MF-1nu/nu mice for 28 days (b). Scale bar = 100 μm.

8. Using sterile forceps, gently insert the bone cell construct or the diffusion chamber (containing cells) into the subcutaneous pocket (see Note 19). 9. Close the incision using Michel staple clips. 10. Transfer mice to the recovery incubator until they recover from the anaesthetic and then place back in appropriate cages (see Note 20). 11. Continue the experiment for up to 28 days. 12. Sacrifice the mice using an approved method and extract the subcutaneous tissue implants or the diffusion chambers. 13. Proceed to molecular, histological, immunohistochemical, or microcomputed tomography analysis of the resulting tissue (Fig. 4).

4. Notes 1. The blocking buffer needs to be made up fresh for each experiment. 2. It is important to de-gas the MACS buffer and to minimise introduction of air bubbles while resuspending cells, since air bubbles in the sample may block the separation columns. 3. Add the fast violet salt to the Naphthol AS-MX solution up just before use. 4. Prepare from saturated Oil Red O solution by passing through filter paper twice. Add 3 ml of the filtered Oil Red O solution

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to 2 ml water and leave for at least 1 h. The working solution should be dark red but clear of sediment; should the solution remain cloudy, pass through a 0.22 μm filter. 5. We use bone tissue that would normally be discarded that is obtained during the course of routine orthopaedic surgical procedures. These samples should be obtained with the patients consent and with the approval of the local ethical committee. If foetal tissue is being used it is advisable to process the femur sample immediately after removal from the foetus. If this is not possible, the sample may be stored in culture media at 37°C, 5% CO2 for up to 48 h. 6. This process assists in releasing cells from the marrow sample and also gets rid of unwanted trabecular bone chips, and tissue debris. 7. The supernatant often contains fat. 8. Filtration is required to remove any residual bone chips and debris. A cell count can be performed at this stage, but is optional. 9. It is important that the Lymphoprep is at room temperature to separate the red cells from the bone marrow mononuclear cells. 10. The cell count should ideally be around 108 cells or higher. The greater the total number of cells, the better is the yield of STRO-1 positive cells. 11. Alternatively, place the tubes containing the cell suspension in the MACSmixTM tube rotator (Miltenyi Biotec). 12. If there are substantially more cells that 1 × 108, then the sample should be split so that 1 ml STRO-1 supernatant or 1 ml antibody is added for each 108 cells processed. 13. The cell layer can also be detached by first incubating with 2% collagenase type IV for 20 min. at 37°C, followed by a 5-min incubation with Trypsin-EDTA. This can help prevent the cell layer from lifting off the tissue culture plastic as a single sheet. 14. Adipocytes can be recognised by the accumulation of lipid droplets in the cytoplasm which appear yellow under phase contrast microscopy. 15. It is important not to let the sections dry out. 16. Each slide will typically require 60–75 μl antibody solution. 17. The sections can also be counterstained with Alcian blue. In this case, the sections should be covered with Alcian blue stain for 45 s before going onto the next step. 18. It can be problematic to differentiate between host and implanted tissue using the subcutaneous implant method, but the diffusion chamber model obviates this problem.

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19. Bilateral implants can be used to introduce diffusion chambers and tissue explants to either side of the vertebral column, thereby reducing the number of animals required for an individual experiment. 20. The duration of the anaesthesia is typically 30–40 min, and the recovery period is 120–240 min. References 1. Simmons, P., and Torok-Storb, B. (1991) Identification of stromal cell precursors in human bone marrow by a novel monoclonal antibody, STRO-1. Blood 78, 55–62. 2. Stewart, K., Walsh, S., Screen, J., Jefferiss, C. M., Chainey, J., Jordan, G. R., and Beresford, J. N. (1999) Further characterisation of cells expressing STRO-1in cultures of adult human bone marrow stromal cells. J. Bone Miner. Res. 14, 1345–1356. 3. Gundle, R., Joyner, C. J., and Triffitt, J. T. (1995) Human bone tissue formation in diffusion chamber culture in vivo by bone-derived cells and marrow stromal fibroblastic cells. Bone 16, 597–601.

4. Oreffo, R. O., and Triffitt, J. T. (1999) In vitro and in vivo methods to determine the interactions of osteogenic cells with biomaterials. J. Mater. Sci. Mater. Med. 10, 607–611. 5. Bolland, B. J., Kanczler, J. M., Dunlop, D. G., and Oreffo, R. O. (2008) Development of in vivo mCT evaluation of neovascularisation in tissue engineered bone constructs. Bone 43, 195–202. 6. Bolland, B. J., Kanczler, J. M., Ginty, P. J., Howdle, S. M., Shakesheff, K. M., Dunlop, D. G., and Oreffo, R. O. (2008) The application of human bone marrow stromal cells and poly(dl-lactic acid) as a biological bone graft extender in impaction bone grafting. Biomaterials 29, 3221–3227.

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Part II Culture of Osteoclasts

Chapter 8 Rodent Osteoclast Cultures Isabel R. Orriss and Timothy R. Arnett Abstract This chapter describes quantitative methods for isolating and culturing rodent osteoclasts on dentine, a bone-like, resorbable substrate. These techniques generate relatively large numbers of osteoclasts and allow the key processes of osteoclast formation and activation to be studied independently. A special focus will be on the role of extracellular pH, a critical factor in the control of osteoclast function. Key words: Osteoclast, Resorption, pH, Bone, Dentine

1. Introduction Osteoclasts are the cells responsible for resorbing bone and other mineralised tissues such as enamel, dentine, and cementum. They are derived from mononuclear progenitors of the monocyte/macrophage lineage and are usually large and multinucleated. Osteoclasts are the only cells known to be capable of removing both mineral and organic matrix, creating characteristic scalloped pits and trails with sharply defined edges. Compared to other bone cells, osteoclasts are relatively few in number, particularly in adult bone. The development of two types of in vitro model based on normal cells has helped to transform our understanding of osteoclast biology. Note that there are no immortalised cell lines currently available that can differentiate into authentic osteoclasts capable of forming true resorption pits. The development in 1984 of “disaggregated” osteoclast resorption cultures by Boyde et al. (1) and Chambers et al. (2) was a major step. The method relies on the relative abundance of mature osteoclasts in the bones of neonatal animals (reflecting the requirement for rapid remodelling during growth), which can be released by fragmenting and agitating the bones in a suitable liquid Miep H. Helfrich and Stuart H. Ralston (eds.), Bone Research Protocols, Methods in Molecular Biology, vol. 816, DOI 10.1007/978-1-61779-415-5_8, © Springer Science+Business Media, LLC 2012

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medium. The suspended osteoclasts, along with other cell types (including osteoblasts, fibroblasts, stromal, and other marrow cells) are then allowed to sediment on to bone or dentine slices, on which they excavate typical resorption pits. Variants of these simple assays were then adopted for quantifying the resorptive function of osteoclasts isolated from neonatal rat bones (3, 4) or chick bones (5). The second key step was the development of long-term osteoclast formation cultures, using haematopoietic precursor cells derived from bone marrow (6). When this assay system was first developed supplementation with factors such as 1α,25-dihydroxyvitamin D3, parathyroid hormone (PTH), and prostaglandin E2 (PGE2) (6, 7) was required. These factors act via the osteoblasts/ stromal cells present in the cultures to stimulate osteoclast differentiation (8). This method was superseded following the identification of macrophage colony stimulating factor (M-CSF) (9) and receptor activator for nuclear factor kB ligand (RANKL) (10, 11) as critical cytokines for osteoclastogenesis. M-CSF and RANKL are now commercially available (see Note 1) and are used to directly stimulate osteoclast formation in cultures from initially non-adherent mononuclear cells derived from marrow, spleen, or peripheral blood. These are now by far the most widely used methods for studying osteoclast function in vitro. There are several important advantages: (1) relatively large numbers of osteoclasts can be obtained (e.g. for biochemical analysis); (2) the key processes of osteoclast formation and activation can be studied independently; (3) osteoclastogenesis occurs in cultures that are relatively free of the confounding influence of stromal cells/pre-osteoblasts. This chapter describes the quantitative methods for studying the function and formation of osteoclasts from rodents. A special focus will be on the role of extracellular pH, a key factor in the control of osteoclast function (see Note 2).

2. Materials All solutions, instruments, and tissue culture plastics should be sterile. 1. Animals: The number of animals to be used depends on the number of treatment groups in the experiment. It should be borne in mind that variation within treatment groups is usually quite high in both the osteoclast formation assay and mature osteoclast resorption assay. Therefore, at least 7–8 replicate dentine discs should ideally be allowed for each treatment group. •

Mouse osteoclast formation assay: Generally, two 6–8-weekold mice will produce approximately nine treatment groups, each containing eight replicates.

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Rat mature osteoclast resorption assay: Four, 2–4-day-old rat pups are required for six treatment groups, and five animals for 7–8 groups. It is not recommended that more than five animals are used because the pooled, dissected bones need to be chopped very quickly (see Subheading 3.2).

2. Minimum essential medium (MEM): Add 10% foetal calf serum (FCS), 2 mM L-glutamine and 100 U/ml penicillin, 100 μg/ml streptomycin, 0.25 μg/ml amphotericin (mixture is known as antibiotic/antimycotic or AB/AM). 3. Step 1 MEM (S1MEM): To MEM add 10−7 M prostaglandin E2 (PGE2) and 2.5 ng/ml M-CSF (see Note 1). 4. Step 2 MEM (S2MEM): To MEM add 10−7 M prostaglandin E2 (PGE2), 10 ng/ml M-CSF (R&D Systems, Abingdon, UK; cat. no. 416-ML), and 3 ng/ml mouse RANKL (Escherichia coli expressed; R&D Systems; cat. no. 462-TEC) (see Notes 1 and 2). 5. Acidified MEM: To achieve a basal level of resorption the medium should be acidified by adding approximately 10 mEq/l of hydrogen ions. This can be achieved by adding 85 μl of concentrated hydrochloric acid (HCl) per 100 ml medium (12) (see Note 3). In the mouse osteoclast formation assay, acid is added to S2MEM to produce acidified medium for the final 48 h of the experiment; in the rat mature osteoclast culture basic MEM is used. 6. Phosphate-buffered saline (PBS): For storing tissues prior to use and for removing non-adherent cells from dentine discs. 7. Hydrochloric acid (HCl): Concentrated hydrochloric acid (11.5 M), to alter the pH of the culture medium. 8. Sodium hydroxide (NaOH): 6 M NaOH, to alter the pH of the culture medium. 9. Diamond saw: e.g. Buehler Isomet, to cut dentine slices. 10. Dentine slices: Substrate for osteoclast culture. •

Prepare the dentine slices by cutting 250-μm thick transverse wafers from a block of dentine (see Note 4) using a diamond saw operating at about 60% of maximum speed with a moderate blade weighting.

•

Soak the slices for 2 h in distilled water to reduce brittleness; cut 5 mm diameter discs from the wet wafers using a standard paper punch. These discs fit neatly into the wells of 96-multiwell plates.

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Wash the discs extensively by sonication in multiple changes of distilled water and store dry at room temperature.

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Before use, number the discs using a graphite pencil to aid identification and sterilise by immersing for 1 min in 100% ethanol.

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Allow the discs to air dry inside a tissue culture flow cabinet (>30 min) and rinse with sterile PBS.

11. Fixative: 2.5% (v/v) glutaraldehyde in PBS; prepare fresh before use. 12. Tartrate-resistant acid phosphatase (TRAP) staining: Leukocyte acid phosphatase kit (Sigma Kit 387-A). 13. Cell removal solution: 0.25 M ammonium hydroxide. 14. Resorption pit staining solution: 1% (w/v) toluidine blue in 1% (w/v) sodium borate solution. 15. Microscopes: A transmitted light microscope is used to count TRAP-positive osteoclasts and total number of cells. Number and/or area of resorption pits are determined using brightfield reflected light microscopy (13, 14). We use a Nikon Labophot 2A microscope, with 100 W epi-illumination and metallurgical objectives. 16. Tissue culture plastics: Large petri dishes (100 mm), 5 ml flatbottomed tubes, 15 and 50 ml centrifuge tubes, 75 cm2 tissue culture flasks and six-well tissue culture plates. Tissue culture plastics can be obtained from a number of different suppliers (e.g. BD Falcon, Nunc, Corning); we have found that the choice of supplier does not affect the formation of osteoclasts. 17. Dissection tools: Scalpels and blades (no. 20), tweezers, and scissors.

3. Methods To keep the cell cultures free from infection, normal sterile techniques be used (working in a flow cabinet, use of sterile media and instruments). 3.1. Mouse Osteoclast Formation Assay

1. Day 1: Kill two 6–8-week-old mice by cervical dislocation and sterilise with 70% ethanol. Place each cadaver in a large petri dish. 2. Remove the limbs by cutting with sharp scissors at the point closest to the body, preserving as much of the limb as possible. 3. Using a scalpel cut off the paws and cut the limb in half at the joint. 4. Remove the skin and scrape away the soft tissue from the limbs. 5. Cut off the epiphyses and flush the marrow out with PBS using a 25-gauge needle. 6. Collect the marrow in a 50-ml centrifuge tube and pellet the cells by spinning at 300 × g for 5 min at room temperature.

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7. Discard the supernatant and resuspend the cells in 2 ml of S1MEM. Put 14 ml of S1MEM into two 75 cm2 flasks; add 1 ml of cell suspension to each. 8. Incubate cells for 24 h at 37°C in a humidified 5% CO2 atmosphere to allow attachment of stromal cells. 9. Day 2: Place sterile dentine discs into the wells of a 96-well plate, numbered side facing down. Collect the non-adherent cells from each 75 cm2 flask; centrifuge at 300 × g for 5 min at room temperature. 10. Discard the supernatant and resuspend the cells at 5 × 106 cells/ml in S2MEM. 11. Add 200 μl of cell suspension (106 cells/disc) to the dentine slices and incubate overnight at 37°C in a humidified 5% CO2 atmosphere to allow attachment of the osteoclast precursors to discs (see Note 5). 12. Day 3: Transfer the dentine discs to six-well trays (4–5 discs/ well in 3 ml of S2MEM); add test substances as required. 13. Half the medium should be exchanged every 2–3 days. The pH should be monitored at every medium change (see Note 6) and maintained at ~pH 7.3 by the addition of 6 M NaOH (see Note 3). 14. Day 7: Acidify the culture to pH 7.0 by full medium change with S2MEM acidified by the addition of 11.5 M HCl (see Notes 3 and 7). This will “switch on” resorption pit formation. 15. Terminate the cultures 48 h after medium acidification (see Subheading 3.3 below). 3.2. Isolation of Mature Osteoclasts from Neonatal Rat Long Bones

1. Prior to commencing isolation of the cells, place sterile dentine discs into the wells of a 96-well plate, numbered side facing down, and add 50 μl culture medium to each well (MEM acidified to pH 7.0). Incubate for 30 min at 37°C. 2. Prepare 5 ml MEM (acidified to pH 7.0) containing test and control substances for each group and add to individual wells of a six-well plate. Place in a 37°C incubator containing a humidified 5% CO2 atmosphere at 5% CO2 for at least 30 min. 3. Sacrifice neonatal (2–4 days) rat pups by cervical dislocation or decapitation. Cut the limbs off and dissect the long bones dissected free of muscle, connective tissue, and cartilage. 4. Transfer the bones to a 35-mm diameter petri dish containing 3 ml of MEM. Chop the bones finely but rapidly with a scalpel blade, using fine forceps to hold the bones steady. 5. Create a suspension by aspirating the minced bones 10–20 times through a wide mouth polyethylene 3-ml transfer pipette with the tip cut back such that the opening is about 5 mm in diameter.

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6. Transfer the suspension (including the remaining small bone pieces) to a 15 ml centrifuge tube and vortex for 20–30 s. 7. Allow the mixture to settle for a few seconds and – avoiding the bone fragments – transfer the supernatant to a fresh 15 ml tube, using a 1-ml polyethylene pipette. 8. Wash the dish and remaining bone fragments with 2 ml culture medium, and vortex briefly. Aspirate the supernatant and combine with the cell suspension from step 7. 9. Quickly add 100 μl cell suspension to each well of the 96-well plate and allow to settle for 45 min at 37°C/5% CO2 (see Note 8). 10. Carefully remove the discs (containing adherent cells) from the 96-well plates using fine forceps or a 19-gauge needle and rinse by dipping in two changes of sterile PBS. 11. Transfer to pre-equilibrated MEM containing test substances or vehicle in a six-well plate (5–6 replicate discs in 5 ml/well). 12. Incubate for 24–28 h in a humidified atmosphere of 5% CO2/95% air at 37°C. 13. At the end of the experiment, measure the medium pH and pCO2 using a clinical blood gas analyser, with careful precautions to prevent CO2 loss (see Note 6). 3.3. Fixation and Staining

1. On termination of the experiment, wash the dentine discs twice with PBS. 2. Transfer to 2.5% glutaraldehyde for 5 min; wash twice more with PBS. 3. Perform light TRAP staining by following the directions in the kit. Count osteoclasts/osteoclast nuclei, as required (see Note 9).

3.4. Quantification of Resorption

1. Resorption pits are normally easy to see by reflected light microscopy (following TRAP staining), without the removal of cells. It is recommended that cells are not removed because discs can then be stored indefinitely (at room temperature) to provide a permanent record. Further analysis (e.g., of numbers of nuclei per cell) can be performed at a later date, as needed. We do not routinely stain nuclei; if the TRAP staining is kept light it is possible to clearly see the nuclei in the osteoclasts (see Note 9). 2. If it is necessary to remove cells from the discs, vigorous sonication for 5 min in 0.25 M ammonium hydroxide is reasonably effective (see Note 10). Discs may then need additional staining for 2 min in toluidine blue solution (followed by rinsing in water and air drying) to improve visualisation of pits (see Note 11). 3. Count the pits/measure pit surface area by scanning the entire surface of each disc using reflected light microscopy and a 10× objective (see Note 11).

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4. Express the results as number of pits/osteoclast or area resorbed/osteoclast and as pits/dentine disc or area resorbed/ dentine disc. It is usually preferable to normalise resorption to osteoclast numbers since the latter may vary within and between treatment groups (see Note 12). 3.5. Statistics

Depending on the data, we routinely use one-way analysis of variance (ANOVA) or non-parametric tests (Mann–Whitney) to analyse experiments. Although often neglected, adjustments for multiple comparisons between treatment groups (e.g., the Bonferroni correction) are frequently needed (see Note 13).

4. Notes 1. Batch testing of cytokines: For the murine osteoclast formation assay to work, good quality cytokines must be used. Failure to generate osteoclasts is usually caused by inactive M-CSF and/or RANKL. Both M-CSF and RANKL are commercially available from number of suppliers (such as Invitrogen, R&D systems, Peprotech); however, potency and quality are variable between both companies and batches. The concentrations provided here are optimised for our culture system, using M-CSF and the highly potent E. coli-expressed mouse RANKL from R&D Systems. The E. coli-expressed mouse RANKL from R&D Systems has the highest potency of any RANKL we have tested in both mouse and human osteoclastogenesis systems. When establishing this assay, we recommend that dose–response curves are generated to determine the optimal cytokine concentrations to use. This process should be repeated whenever cytokines are obtained from a new supplier or the batch received changes. 2. Hypoxia: Osteoclast formation is strongly increased in reduced oxygen (15), although permissive concentrations of RANKL and M-CSF are still required. If larger numbers of osteoclasts are required (e.g. for biochemical analyses), cultures may be incubated in an atmosphere of 2–5% O2/5% CO2/balance N2. This approach may also help to reduce cytokine costs. 3. Importance of pH: Extracellular pH is a critical factor in all osteoclast formation and bone resorption experiments. Rat osteoclasts are maximally activated to form resorption pits at pH ~6.9 and resorption is essentially “switched off” above pH ~7.2 (Fig. 1) (3, 14). Similar responses to extracellular acidification have been observed in all bone resorption systems examined to date, using cells or tissues derived from murine, avian, or human sources (Fig. 1) (16, 17). The action of many bone resorbing agents including RANKL (Fig. 2a) (18), parathyroid hormone (Fig. 2b) (3), 1,25-(OH)2-vitamin D3 (19),

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Fig. 1. Acid activation of osteoclasts cultured on dentine. Culture medium was adjusted by the addition of HCl or NaOH. Rat osteoclasts, isolated directly from the bone of neonates, are essentially “switched off” above pH 7.2; osteoclasts formed from mouse marrow behave similarly (data not shown). Osteoclasts from embryonic chick bones retain some resorptive activity at pH 7.4. Human osteoclasts, formed from human peripheral blood mononuclear cells, were cultured for 14 days at pH 7.4 with M-CSF and RANKL and then maintained for a further 48 h at the indicated pH; these cells also retain some resorptive activity at pH 7.4. Maximal acid-stimulation for all osteoclasts occurs at pH ~6.9. Values are means ± SEM (n = 5).

Fig. 2. Two-step stimulation of osteoclast resorption by low pH in combination with RANKL or PTH. (a) Mature osteoclasts, isolated directly from neonatal rat bones, cultured for 26 h. At “physiological” pH (~7.4), basal resorption was very low and RANKL exerts only a small stimulatory effect. Combined treatment with RANKL and low pH results in large, synergistic increases in resorption (courtesy of N Zanellato, UCL). Significantly different from control in the same pH group: **p < 0.01, ***p < 0.001. Significantly different from the same RANKL concentration at pH 7.4: ##p < 0.01. (b) Osteoclasts formed from human peripheral blood mononuclear cells, cultured for 14 days at pH 7.4 with MCSF and RANKL. Cells were cultured for a further 48 h in the presence or absence of PTH in medium maintained at pH ~7.4 or acidified to pH ~6.9. PTH increases resorption at both pH 7.4 and 6.9, with an additive stumulatory effect evident at the lower pH. Note that human osteoclasts are not completely “switched off” at pH 7.4 (see Fig. 1) (courtesy of A Brandao-Burch, UCL). Significantly different from control in the same pH group: *p < 0.05, **p < 0.01. Values are means ± SEM (n = 5–6).

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and extracellular nucleotides such as adenosine 5¢-triphosphate (ATP) (19) and adenosine 5¢-diphosphate (ADP) (20) is enhanced by acidification. These results indicate that a low pH is an essential requirement for the activation process; once this activation has occurred, further stimulation by a wide range of bone resorbing agents can take place. Most tissue culture media (including MEM) are buffered to pH ~7.20 when fully equilibrated with 5% CO2; this value corresponds to normal interstitial pH and is considerably more acidic than blood pH (7.35–7.40). The metabolic activity of cells cultured in the medium will act to lower the pH further. When cell numbers are high relative to the volume of medium (e.g., when dentine discs are cultured in 96 well plates), this effect can be sufficient to acidify the medium quite rapidly, with resultant activation of resorption pit formation. To activate resorption in a more controlled manner, relatively large volumes (³0.5 ml/dentine disc/24 h) of pre-acidified culture medium should be used. MEM may be acidified by the direct addition of small amounts of concentrated hydrochloric acid, which has the advantage of being self-sterilising (12, 14). This also has the effect of reducing HCO3− concentration (i.e., “metabolic acidosis”) and producing an operating pH close to 6.95 in a 5% CO2 environment (see Fig. 3), which is optimal for resorption pit formation (14). Further acidification in CO2/HCO3−-buffered media does not enhance resorption greatly and may ultimately

Fig. 3. Relationship between pH, pCO2, and HCO3− in tissue culture medium. Data obtained using a blood gas analyser.

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reduce cell survival. Addition of HCl does also have the effect of increasing medium chloride concentration slightly, but this does not appear to affect bone cell function. For the murine osteoclast formation assay (see Subheading 3.1), the best results are obtained when S2MEM is alkalinised to pH ~7.3 by the addition of ~7.5 mEq/L NaOH for the first 6–7 days; this enables osteoclast formation to occur with little or no resorption pit formation. For the last 2 days of culture, the medium is replaced with S2MEM, acidified to pH ~6.95, to activate resorption. Note, that if the S2MEM is maintained at pH 7.4 during the formation phase of the assay, the osteoclasts which form are very large (>30 nuclei) and sometimes vacuolated (Fig. 4). In contrast, if the culture medium is acidified (e.g. to pH 7.0) during the formation phase, osteoclastogenesis is reduced; however, resorption will still be relatively high because osteoclasts are activated as soon as they are formed. 4. Sources of dentine: Dentine can be obtained in the form of confiscated elephant ivory or sperm whale teeth from customs or fisheries and wildlife agencies (e.g., in the UK or USA). Dentine is a convenient osteoclast substrate because it is uniform, easy to cut, and lacks features such as Haversian systems and osteocytes which make quantification of resorption difficult. If bone slices are being used, they should be prepared from defatted and washed cortical bone from bovine femora; the slices should be transversely cut to reduce the likelihood of confusing in vitro resorption with endogenous features. We find that cortical bone is usually too brittle to permit the fabrication of uniform discs using a hole punch. 5. Other substrates: Mouse marrow cultures grown on plastic will form very large, multinucleate cells with a flat, rounded morphology and less intense TRAP staining than multinucleate osteoclasts formed on dentine or bone (Fig. 4). These cells formed on plastic also tend to exhibit less motile activity than typical mature osteoclasts isolated directly from rodent bones. A number of thin layer synthetic substrates are also available. Commercial preparations consisting of hydroxyapatite sintered at high temperature onto silica discs are available; these have the advantage of being translucent (e.g. for electrophysiology or cell fluorescence work), but in our experience may impair osteoclast survival. Alternatively, mineralised collagen films can be prepared more cheaply using the method of Lees et al. (21). The synthetic mineralised films also suffer from the disadvantage of disintegration and fragility in media acidified to pH 7.1 or below. 6. Measuring pH: Accurate measurement of the operating pH of mammalian osteoclast cultures is necessary for meaningful comparison of results from different laboratories. For

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Fig. 4. Dynamics of osteoclast formation and activation: comparison of dentine and plastic substrates. Osteoclasts formed from mouse marrow mononuclear cells cultured on dentine with RANKL and M-CSF, stained to demonstrate tartrateresistant acid phosphatase and viewed by transmitted light microscopy. (a) Very large, inactive multinucleated cells form in cultures maintained at “physiological” pH (7.4). (b) Acidification for the final 2 days of culture causes dramatic activation of resorption pit formation but tends to prevent further multinucleation of osteoclasts. Note that TRAP staining and transmitted light microscopy visualise osteoclasts (and their mononuclear precursors) as well as resorption pits (tan areas). (c) Osteoclasts cultured on plastic form large, inactive, multinucleate TRAP-positive cells. Scale bars = 10 μm (images on left ) and 50 μm (images on right ).

HCO3−/CO2-buffered media, accurate pH measurements can only be achieved by the use of a properly standardised blood gas analyser. We use a Radiometer ABL 705 blood gas analyser (Radiometer, Crawley, UK). The blood gas analyser uses a multi-electrode system to measure pH, pCO2, and pO2 in a 200-μl injected sample (cycle time ~2 min). The first medium measurement, taken immediately after removing the culture

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plates from the incubator, is assumed to provide a pCO2 value that is the same for all wells and that reflects the actual pCO2 during the 24-h incubation. (It is worth noting that opening the door of the incubator during experiments may cause perturbations in CO2 levels that affect measured pH and pCO2 values, and possibly osteoclast function.) Measured pCO2 typically drops for each subsequent reading from wells in a multiwell plate, causing pH values to rise accordingly. The pH readings for each well are then back-corrected to the pH value associated with the initially measured pCO2 value, using calibration curves previously measured for culture medium with different bicarbonate concentrations (Fig. 3). 7. Duration of cultures: The rate of osteoclastogenesis in murine cultures is dependent on the quality and potency of the M-CSF and RANKL used. Cytokines with a high biological activity will generate mature osteoclasts in 7 days or less (from the isolation of the bone marrow). To determine the optimum duration for these cultures, we initially recommend that discs are stopped and TRAP stained every other day (Fig. 5). It is important that these cultures are not maintained too long (12 days maximum); otherwise osteoclasts generated may start to die. Note that osteoclasts generated from human peripheral blood can normally survive for longer periods in culture (see Chapter 11, this volume). 8. Technique for plating out cell suspension: An automatic 2.5 ml multidispensing pipette (Gilson) is ideal for plating out the cell suspension. Delays at this stage can cause problems, because

Fig. 5. Time course of mouse osteoclast formation and activation on dentine. Primary osteoclasts were cultured on dentine discs for 9 days; tissue culture medium was acidified to ~pH 6.9 for the final 2 days to activate osteoclast resorption. These light microscopy images are representative of the cells present after 2, 5, 7, and 9 days of culture. At day 2, the cells present were all monocyte/macrophage precursors; by day 5 TRAP positive mononuclear cells were evident. At day 7, there were abundant mature but inactive osteoclasts present. Medium acidification resulted in widespread osteoclast resorption, as seen at day 9. Scale bar = 50 μm.

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the cells sediment rapidly from the suspension unless it is continuously agitated. Care should also be taken to ensure that dentine discs remain seated in the base of the wells and do not float up. To compensate for plating errors, the suspension should be dispensed sequentially across treatment groups, rather than dispensing to each treatment group in turn. 9. Identifying and counting osteoclasts: TRAP staining is a convenient and inexpensive method of visualising osteoclasts; however, it is not entirely specific because other cells of the haematopoetic lineage also express this enzyme (22). Furthermore, since mononuclear osteoclasts are observed in human cultures, osteoclasts are best defined as TRAP-positive cells that have two or more nuclei and/or excavate pits. Osteoclast number should be assessed “blind” to the treatment group using transmitted light microscopy and a 10× objective. During TRAP staining, the dentine discs should be carefully monitored to avoid permanent overstaining (which will obscure detail within the cells such as nuclei) (Figs. 4 and 5). The time needed for optimal, light staining may vary between 25 and 60 min. 10. Removal of cells: Sonication can remove the graphite pencil markings. In view of this, discs should be sonicated in a known sequence, so that identification numbers can be rewritten if necessary. 11. Visualising and quantifying resorption pits: Whilst reflected light microscopy often yields adequate images of resorption pits on unstained specimens, image quality is improved greatly by staining (because this has the effect of increasing reflectivity). Depending on the microscope system used, optimal reflected light images are obtained using either brightfield or darkfield modes. To assess resorption area, we normally use a simple dot counting morphometry system: output from the reflected light microscope via a standard colour video camera is displayed on a monitor, superimposed on which is an acetate sheet bearing a grid of dots. The dot grid is easily created by using a graph paper template that has been photocopied to the required magnification. It is possible to measure the volume of resorption pits using scanning EM or confocal microscopy (23), but this requires specialised and expensive equipment. Pit volume can also be estimated by measuring pit depth, and area, using reflected light microscopy (the fine focus control is usually calibrated in microns) and assuming that pits approximate to hemispheres; this method is not suited to determining the volume of individual pits to very high accuracy, but provides useful comparative data when multiple pits are measured. When very high levels of resorption occur (e.g. >25% of surface area), it may be possible to use automated image analysis of pits viewed by

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reflected light in a low-power dissecting microscope. For a method for automated quantification using low magnification reflected light microscopy see Chapter 12, this volume. 12. Assay variability: One of the most serious problems with the rat mature osteoclast assay (and to a lesser extent the murine osteoclast formation assay) is the high variability between experiments. First, osteoclast number in some culture preparations can be low, even if the procedure is followed accurately. Secondly, the basal level of resorption can vary from experiment to experiment, perhaps reflecting alterations in ambient concentrations of bone resorbing agents such as growth factors and nucleotides. Thus, culture conditions should be kept as identical as possible (i.e. freshness and pH of the medium; cytokine and serum batches; CO2 concentration of the incubator; origin, washing and sterilising of the discs). 13. Statistics: Because of interassay variability statistical comparisons should only be performed within one assay, and not between different assays. Variability within assays is likely to be high when cell numbers are low. The inherent “noise” in osteoclast resorption assays means that they are best suited for studying large, robust effects.

Acknowledgments The authors gratefully acknowledge the support of Arthritis Research UK and the European Union (Framework 7 Programme). References 1. Boyde, A., Ali, N. N., and Jones, S. J. (1984) Resorption of dentine by isolated osteoclasts in vitro. Br. Dent. J. 156, 216–220. 2. Chambers, T. J., Revell, P. A., Fuller, K., and Athanasou, N. A. (1984) Resorption of bone by isolated rabbit osteoclasts. J. Cell Sci. 66, 383–399. 3. Arnett, T. R., and Dempster, D. W. (1986) Effect of pH on bone resorption by rat osteoclasts in vitro. Endocrinology 119, 119–124. 4. McSheehy, P. M., and Chambers, T. J. (1986) Osteoblast-like cells in the presence of parathyroid hormone release soluble factor that stimulates osteoclastic bone resorption. Endocrinology 119, 1654–1659. 5. Arnett, T. R., and Dempster, D. W. (1987) A comparative study of disaggregated chick and rat osteoclasts in vitro: effects of calcitonin and prostaglandins. Endocrinology 120, 602–608.

6. Takahashi, N., Yamana, H., Yoshiki, S., Roodman, G. D., Mundy, G. R., Jones, S. J., Boyde, A., and Suda, T. (1988) Osteoclast-like cell formation and its regulation by osteotropic hormones in mouse bone marrow cultures. Endocrinology 122, 1373–1382. 7. Suda, T., Takahashi, N., and Martin, T. J. (1992) Modulation of osteoclast differentiation. Endocr. Rev. 13, 66–80. 8. Takahashi, N., Akatsu, T., Udagawa, N., Sasaki, T., Yamaguchi, A., Moseley, J. M., Martin, T. J., and Suda, T. (1988) Osteoblastic cells are involved in osteoclast formation. Endocrinology 123, 2600–2602. 9. Yoshida, H., Hayashi, S., Kunisada, T., Ogawa, M., Nishikawa, S., Okamura, H., Sudo, T., Shultz, L. D., and Nishikawa, S. (1990) The murine mutation osteopetrosis is in the coding region of the macrophage colony stimulating factor gene. Nature 345, 442–444.

8 10. Lacey, D. L., Timms, E., Tan, H. L., Kelley, M. J., Dunstan, C. R., Burgess, T., Elliott, R., Colombero, A., Elliott, G., Scully, S., Hsu, H., Sullivan, J., Hawkins, N., Davy, E., Capparelli, C., Eli, A., Qian, Y. X., Kaufman, S., Sarosi, I., Shalhoub, V., Senaldi, G., Guo, J., Delaney, J., and Boyle, W. J. (1998) Osteoprotegerin ligand is a cytokine that regulates osteoclast differentiation and activation. Cell 93, 165–176. 11. Yasuda, H., Shima, N., Nakagawa, N., Yamaguchi, K., Kinosaki, M., Mochizuki, S., Tomoyasu, A., Yano, K., Goto, M., Murakami, A., Tsuda, E., Morinaga, T., Higashio, K., Udagawa, N., Takahashi, N., and Suda, T. (1998) Osteoclast differentiation factor is a ligand for osteoprotegerin/osteoclastogenesisinhibitory factor and is identical to TRANCE/ RANKL. Proc. Natl. Acad. Sci. USA 95, 3597–3602. 12. Goldhaber, P. and Rabadjija, L. (1987) H+ stimulation of cell-mediated bone resorption in tissue culture. Am. J. Physiol 253, E90–E98. 13. Walsh, C. A., Beresford, J. N., Birch, M. A., Boothroyd, B., and Gallagher, J. A. (1991) Application of reflected light microscopy to identify and quantitate resorption by isolated osteoclasts. J. Bone Miner. Res. 6, 661–671. 14. Arnett, T. R., and Spowage, M. (1996) Modulation of the resorptive activity of rat osteoclasts by small changes in extracellular pH near the physiological range. Bone 18, 277–279. 15. Arnett, T. R., Gibbons, D. C., Utting, J. C., Orriss, I. R., Hoebertz, A., Rosendaal, M., and Meghji, S. (2003) Hypoxia is a major stimulator of osteoclast formation and bone resorption. J. Cell Physiol. 196, 2–8.


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16. Meghji, S., Morrison, M. S., Henderson, B., and Arnett, T. R. (2001) pH dependence of bone resorption: mouse calvarial osteoclasts are activated by acidosis. Am. J. Physiol. Endocrinol. Metab. 280, E112–E119. 17. Morrison, M. S. and Arnett, T. R. (1997) Effect of extracellular pH on resorption pit formation by chick osteoclasts. J. Bone Miner. Res. 12, S290–S290. 18. Bushinsky, D. A. (1987) Net calcium influx from live bone during chronic metabolic, but not respiratory, acidosis. Am. J. Physiol. 256, F836–F842. 19. Morrison, M. S., Turin, L., King, B. F., Burnstock, G., and Arnett, T. R. (1998) ATP is a potent stimulator of the activation and formation of rodent osteoclasts. J. Physiol. 511 (Pt 2), 495–500. 20. Hoebertz, A., Meghji, S., Burnstock, G., and Arnett, T. R. (2001) Extracellular ADP is a powerful osteolytic agent: evidence for signaling through the P2Y1 receptor on bone cells. FASEB J. 15, 1139–1148. 21. Lees, R. L., Sabharwal, V. K., and Heersche, J. N. (2001) Resorptive state and cell size influence intracellular pH regulation in rabbit osteoclasts cultured on collagen-hydroxyapatite films. Bone 28, 187–194. 22. Walsh, N. C., Cahill, M., Carninci, P., Kawai, J., Okazaki, Y., Hayashizaki, Y., Hume, D. A., and Cassady, A. I. (2003) Multiple tissue-specific promoters control expression of the murine tartrate-resistant acid phosphatase gene. Gene 307, 111–123. 23. Boyde, A., and Jones, S. J. (1991) Pitfalls in pit measurement. Calcif. Tissue Int. 49, 65–70.

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Chapter 9 Isolation and Culture of Primary Chicken Osteoclasts Patricia Collin-Osdoby and Philip Osdoby Abstract Osteoclasts originate from hematopoietic myeloid progenitors that differentiate into specialized multinucleated cells uniquely capable of resorbing bone in both physiological and pathological conditions. Osteoclast numbers and degradative activities increase in various inflammatory disorders of bone and certain bone oncologies, thereby causing bone loss that may weaken the skeleton, increase fracture incidence, and disturb marrow function. Many valuable insights have been obtained through the use of osteoclasts directly isolated from the bones of chickens fed a low calcium diet to enhance osteoclastogenesis and bone resorption. Particular advantages of this system include the abundance and highly resorptive nature of isolated chicken osteoclasts compared with those directly obtained from other species. After enzymatic release from the harvested bones, osteoclasts may be partially purified by density gradient sedimentation, bone substrate attachment, and/or immunomagnetic capture. Thereafter, osteoclast preparations may be analyzed, either directly or following some period of culture, to investigate their properties (biochemical, immunological, molecular, cell biological), resorptive function, and modulatory responses to various stimuli. Here, we present common procedures for the isolation, culture, and general study of chicken osteoclasts. Key words: Osteoclast isolation, Osteoclast culture, Osteoclast antigens, Bone resorption, Chicken, Avian

1. Introduction Bone is a dynamic tissue that is continually remodeled by the coordinated actions of bone-resorbing osteoclasts (OCs) and bone-forming osteoblasts. Whereas osteoblasts originate from mesenchymal cell precursors, OCs derive from hematopoietic precursors of the myeloid lineage present in the bone marrow and circulation. In response to specific hormonal or local signals, particularly the essential OC differentiation signal receptor activator of NF-κB ligand (RANKL), provided by osteoblasts, stromal cells,


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or other cells within the bone marrow, OC precursors fuse and differentiate into large multinucleated cells expressing characteristic morphological features, membrane polarization, adhesion molecules, ion pumps, enzyme activities, and antigenic profiles (1–3). Most importantly, they develop a capacity for bone pit resorption, the unique and defining functional attribute of OCs. Bone resorption and formation are normally carefully balanced processes in adults. However, in various pathologies, an imbalance arises such that the number of OCs, number of resorption sites initiated, and/ or rates of remodeling are altered, thereby resulting in either too much or too little bone turnover. Excessive bone loss occurs in many disorders including postmenopausal osteoporosis, rheumatoid arthritis, periodontal disease, tumor-associated osteolysis, and orthopedic implant loosening (4–7). It is therefore important to decipher the complex signals that control OC bone resorption to further our understanding and provide a rational basis for the design of novel therapeutic or preventative strategies to combat bone loss. In this regard, isolated primary cultures of in vivo formed OCs have proven to be an invaluable tool for investigating the characteristics, function, and regulation of OCs. Many valuable insights have been achieved through the use of avian OCs, cells that are highly active in their resorption of bone and are isolated in abundance from young chickens fed a low calcium diet (8). Following their enzymatic release from harvested bones, OCs can be partially purified by density gradient (Percoll) sedimentation owing to their large size, and further enriched by brief settling onto bone discs or rapid capture with magnetic beads pre-coupled with antibody that specifically recognizes OCs (9). OCs isolated via these procedures can be cultured and analyzed for biochemical, immunological, physiological, and functional properties, as well as modulator responses. Procedures for some of the most commonly used assays are presented.

2. Materials 2.1. Tissue Culture Medium, Solutions, and Supplies

All media and solutions should be prepared with glass distilled water and be sterilized. 1. Culture medium: Sterile α-minimum essential medium (α-MEM) supplemented with 5% fetal bovine serum (FBS, InvitrogenGibco) and 2.5% antibiotic/antimycotic (a/a, InvitrogenGibco); store at 4°C and prewarm to 37°C before use. 2. Hanks’ balanced salt solution, pH 7.2 (HBSS). 3. Moscona’s low bicarbonate (MLB): Add 8 g NaCl, 0.2 g KCl, 50 mg NaH2PO4, 0.2 g NaHCO3, 2 g dextrose, 10 ml a/a, 990 ml water, check pH is 7.2, and sterile-filter.

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4. Moscona’s low bicarbonate–EDTA (MLBE): Dissolve 1 g EDTA in 15 ml 1% KOH, add to 1 l of MLB, check pH is 7.2, and sterile-filter. 5. Phosphate-buffered saline, pH 7.2 (PBS). 6. Collagenase: Prepare 0.5 mg/ml stock solution in HBSS, store aliquots at −20°C, and dilute two parts of thawed stock solution with one part of MLB for use. 7. Trypsin: 1% Stock (1 g/100 ml) solution in MLB, store aliquots at –20°C, and dilute 11.25 ml of stock with 37.5 ml MLBE and 201.5 ml MLB for use. 8. Percoll for 35% Percoll, mix 65 ml of HBSS with 35 ml of Percoll (GE Healthcare Life Sciences); for 6% Percoll, mix 83 ml of the HBSS solution with 17 ml of 35% Percoll/HBSS. Adjust the pH of both solutions to 7.2, sterile-filter, and store at 4°C. 9. Heparin: 1,000 U/ml, GE Healthcare Life Sciences, store at 4°C. 10. Trypan blue: 0.4 g trypan blue dye in 100 ml water. 11. 1% Paraformaldehyde in HBSS (PF-HBSS): Preheat 100 ml HBSS on a hot plate to 60°C in a Pyrex beaker (monitor with a thermometer), move the beaker to a stir plate, add 1 g of PF, cover with foil to contain the vapors (keep the thermometer in place and briefly move the beaker back to the hot plate if the temperature falls below 50°C), slowly stir with a magnetic bar, and add 3–4 drops of 10 N NaOH just to dissolve. Let cool, filter through Whatman #1 paper into a brown glass bottle, and store at 4°C. 12. Protease inhibitor cocktail for cell pellet storage: Prepare an inhibitor stock solution A by dissolving 10 mg each of leupeptin, chymostatin, antipain, and pepstatin A in 1 ml of dimethyl sulfoxide, add 400 trypsin inhibitory units of aprotinin, and store this 1,000× cocktail in 0.1 ml aliquots at −20°C. Mix 10 μl of inhibitor stock solution A with 10 μl of a 1% stock solution of phenylmethylsulfonyl fluoride (PMSF) in ethanol (store at room temperature), 1.25 mg N-ethylmaleimide (NEM), 1.56 mg benzamidine, and 10 ml HBSS to yield inhibitor stock solution B. Store this at −80°C and overlay one drop (~50 μl) on top of each cell pellet to be stored frozen. 13. 350- and 110-μm Nitex filters: Sheets of Nitex (Tetko, Kansas City, MO) mesh are cut into squares larger (~50%) than the opening of a stackable plastic beaker, a filter square is stretched over the beaker, and the filter is tightly secured in place by fitting on a ring (~1-in. deep) made from a second plastic stackable beaker whose lower three-quarter portion has been cut off; filter squares can be washed well and reused.

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2.2. Preparation of Antibody-Conjugated Magnetic Beads

1. Magnetic polystyrene beads: 0.45 μm diameter, covalently conjugated with affinity purified sheep anti-mouse IgG (Invitrogen-Dynal Inc., store at 4°C). 2. Mouse Mab to OC-specific antigen: See Note 1. 3. Rotary mixer: To fit microcentrifuge tubes. 4. Magnet: Invitrogen-Dynal Inc. or other 20-lb pull magnet.

2.3. Fixation and TRAP Staining

Although not fully specific for OC, high tartrate resistant acid phosphatase (TRAP) activity plays a role in bone resorption, and it is upregulated and a convenient measure of OC development (10). 1. 1% Paraformaldehyde in HBSS: See Subheading 2.1, item 11. 2. TRAP staining: Prepare the following stock solutions and mix just before use (or purchase a staining kit from Sigma (cat. no. 386) and follow the manufacturer’s instructions, see Note 2). Solution A: Naphthol AS-BI phosphoric acid (12.5 mg/ml) in dimethyl formamide (store at −20°C). Solution B: 2.5 M acetate buffer, pH 5.2, store at 4°C. Solution C: 0.67 M tartrate solution, pH 5.2, store at 4°C. (a) Mix 0.4 ml of solution A, 0.4 ml of solution B, 0.4 ml of solution C, and 8.8 ml of deionized water (preheated to 37°C) in a 50 ml polypropylene tube, vortex well, and wrap the tube in foil. (b) Add 3 mg Fast Garnet GBC salt, vortex quickly to mix well, and filter the solution through Whatman #1 paper into a new foil-wrapped 50 ml polypropylene tube. Use immediately. 3. General stain: Use Diff-Quik (eosin Y, azure A, and methylene blue, Fisher Scientific) as recommended by the manufacturer.

2.4. Fixation and Immunostaining

For immunostaining, prepare the following solutions: 1. Blocking solution: 1% Bovine serum albumin (BSA) and 10% horse serum in PBS. 2. Monoclonal (MAb) or polyclonal (PAb) antibodies: Directed against OC antigens and appropriately diluted (typically 1:100 to 1:500 of 1 mg/ml stocks) in blocking solution just prior to use. 3. Biotinylated secondary antibodies: Directed against the primary antibody and appropriately diluted (typically 1:200 to 1:500) in blocking solution just prior to use (see Note 3). 4. Glycerol-buffered mounting medium: For example, EM Sciences, 80% glycerol in PBS, store at 4°C.

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For fluorescence immunostaining: 5. Streptavidin conjugated with a fluorescent label [fluorescein isothiocyanate (FITC), Texas Red, or similar]: Appropriately diluted (typically 1:1,000 or more) in PBS (without serum) just prior to use. 6. 4¢,6-Diamidino-2-phenylindole (DAPI): (Molecular Probes), prepare 100 μg/ml of stock solution in water, store dark at 4°C, and dilute stock 1:300 in HBSS for use in fluorescent nuclear staining. For colorimetric immunostaining: 7. Streptavidin conjugated with β-galactosidase: Appropriately diluted (typically 1:100) in buffer A (see item 6). 8. Buffer A: 0.1 M Sodium phosphate, pH 7.2, containing 1.5 mM magnesium chloride, 2 mM β-mercaptoethanol and 0.05% sodium azide; store at 4°C and warm to room temperature before use. 9. Buffer B: 10 mM Sodium phosphate, pH 7.2, containing 150 mM sodium chloride, 3 mM potassium ferricyanide, 3 mM potassium ferrocyanide, and 1 mM magnesium chloride; store at 4°C and warm to room temperature before use. 10. Substrate solution (0.42 mg/ml X-gal in buffer B): Prepare a stock solution of X-gal (21 mg/ml) in dimethyl formamide, store at −20°C (e.g., in a Parafilm sealed, foil wrapped glass tube), and dilute this stock solution 1:50 in buffer B to prepare fresh substrate solution. 2.5. Preparation of Devitalized Bone or Ivory Discs for Bone Pit Resorption Studies

1. Ivory is obtained through donation from a local zoo or the Federal Department of Fish and Wildlife Services (in the USA; or a similar source in another country). Bovine cortical bone is obtained from a local slaughterhouse. Segments of ivory and bovine cortical bone are thoroughly cleaned and washed (multiple HBSS and 70% ethanol rinses), sliced into small chunks, and then reduced to rectangular 0.4-mm thick sheets using a low speed Isomet saw (Buehler, Lake Bluff, IL). 2. The sheets are rinsed three times with 70% ethanol, incubated in 70% ethanol overnight, and then washed for several hours in HBSS before circular discs are cut out using a 5-mm paper punch. 3. Punched discs are soaked repeatedly in 70% ethanol in sterile 50-ml tubes (alcohol changes can be gently poured off because the discs tend to stick to the side of the tube), and stored in 70% ethanol at −20°C. 4. For experimental use, remove the required number of discs from the tube using alcohol presoaked tweezers in a sterile hood, transfer the discs to a new sterile 50-ml polypropylene tube, rinse extensively by inversion and mild shaking at least three

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times with ~40 ml of sterile HBSS per wash, and transfer the discs using sterile tweezers into culture wells or dishes containing sterile HBSS for 3–24 h of pre-incubation in a tissue culture incubator prior to the plating of cells. Remove HBSS only immediately before seeding OC to prevent the discs from drying. 2.6. Preparation of Gold-Coated Glass Coverslips for Phagokinetic Motility Studies (see Note 4)

This procedure is a modification of the gold coverslip motility assay reported by Owens and Chambers (11). Glass coverslips are precoated with a thin layer of gelatin to enhance attachment and homogeneous coverage of the gold coating. All steps are performed in a sterile hood, using sterile reagents and supplies, and more coverslips (10–50%, depending on the skill you develop for this procedure) should be coated than you expect to need in the experiment. 1. Warm a solution of 2% gelatin in deionized water (stored at room temperature) to 37°C in a water bath and two 24-well tissue culture dishes in an incubator. 2. Assemble glass coverslips, sterile tweezers, and one or two 100-mm Petri dishes with fitted Whatman filter paper in a sterile hood. 3. Fill one well of a prewarmed 24-well dish with warm 2% gelatin solution, dip one coverslip at a time into the well using sterile tweezers, briefly drain against the side of the well, and place it gelatin side up on the filter paper in the open 100-mm dish to dry in the hood for at least 2 h. 4. Using tweezers, move each coverslip into one well of a sterile 24-well dish in the hood. 5. Prepare the gold coating solution (19.0 ml) in the hood by adding 11.0 ml of sterile deionized water, 4.44 ml of 0.2% gold chloride (in sterile water), and 3.36 ml of sterile 65.2 nM sodium carbonate into a 100-ml Pyrex beaker (foil covered and presterilized). Heat the solution on a hot plate (this can be performed outside the hood after the foil cover is replaced) just to boiling. 6. Place the covered hot beaker back into the sterile hood, add 1.8 ml of sterile 0.1% paraformaldehyde in water, and allow the gold solution to cool to 60°C. Monitor the temperature (this is critically important to achieve good gold coating) with an alcohol prewiped thermometer. 7. When the solution has cooled to 60°C, pipette 1 ml of the gold solution on top of each gelatin precoated coverslip and refrigerate the 24-well dish for 1 h. 8. Remove the excess solution, gently rinse the coverslips twice with HBSS, and place each coverslip onto filter paper in a 100-mm dish in the sterile hood to dry (several hours to overnight).

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9. Repeat items 5–8 to ensure adequate and even coverage of the gold particles on the coverslips. 10. After the second gold coating, store the coverslips on filter paper in a 100-mm dish in the hood for up to a few days prior to their use. Check one or two coverslips by placing into a 24-well dish with HBSS for at least 1 h to verify that the gold coating does not lift up, is sufficiently dark, and evenly coats the coverslip when viewed under the microscope (otherwise tracks produced by OCs will be hard to evaluate). If the coating lifts up, check a few other coverslips from that batch and discard all if they fail this test. If the gold coating is too sparse, repeat items 5–8 for a third time. 11. Before use, preincubate all coverslips for at least 1 h in HBSS early on the day of cell plating and plan to use only those that exhibit a firm, even, and dark gold coating.

3. Methods 3.1. Isolation of Osteoclasts from Calcium-Deficient Chicks (see Note 5)

After hatching, White Leghorn chicks are fed a normal diet for 4–6 days and then switched to a low calcium diet (0.15–0.25% calcium, analyzed before the feed is shipped from Purina) for at least 28 days (8). Typically, 15 chicks are used for each OC preparation and three people assist in the dissection and initial steps to minimize the time until the cells are isolated and plated (even 1 additional hour can affect the ultimate OC yield and viability, see Note 6). Animals are handled and euthanized in accordance with rules and procedures for the Institutional Animal Care and Use Committee and standards approved by the National Institutes of Health Guidelines for the Care and Use of Experimental Animals (or similar authority for countries other than the USA). 1. Just prior to dissection, multiple forceps, tweezers, and scissors are placed into beakers with 70% alcohol, all buffers are prechilled on ice, and the pH of MLB is readjusted if needed. 2. Fill several ice containers, and place two 100-mm Petri dishes with HBSS on ice. 3. Several people wearing alcohol-rinsed gloves each remove a group of birds immediately following their euthanization (do not delay), alcohol squirt the wings and legs of one bird just prior to its dissection, rapidly remove the tibiae and humeri using the alcohol-soaked scissors and forceps, clean off extraneous soft tissue without removing the bone ends (which are replete with OCs), and place the bones into one of the two HBSS-filled Petri dishes on ice. Repeat for each bird.

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4. When a number of bones have accumulated in the first HBSS dish on ice, this dish is given to one person to extract marrow from the bones in a sterile hood, while the other two people continue to dissect bones from the remaining birds and place them into the second HBSS dish on ice. 5. Marrow is removed from each bone by gripping it with alcohol-soaked tweezers over a 100-mm dish containing MHB, poking several small holes in each end of the bone using a 3-ml syringe with an 18-guage needle, and quickly flushing the marrow out by repeatedly inserting the tip of the syringe filled with MHB (from the dish) into one bone end and squirting fluid (not too hard) through the bone into the dish below. Hold the bone just above the dish, not in contact with the flushed marrow. 6. Carefully flip the bone held with tweezers over, and repeat step 5 to flush MHB several times through the other end of the bone. Place the flushed bone back into the original (non-marrow) dish of HBSS. Repeat for subsequent bones, each time flushing MHB through both ends of the bone before proceeding to the next bone. 7. After all of the marrow has been extruded, remove any bits of extra tissue carefully from the bones and then divide the bones into eight 50-ml tubes filled with 40 ml HBSS each. Shake gently by hand to wash the bones (~30 s). 8. Place the bones into two new dishes of HBSS on ice to split each bone lengthwise using sterile scissors while keeping the bones submersed in HBSS. 9. Transfer the split bones into eight 50-ml polypropylene tubes containing 40 ml of HBSS each, shake vigorously for 30 s, and pass the supernatants sequentially through 350- and 110-μm Nitex filters fitted over plastic beakers set on ice (be sure the filters are tightly fitted so that the cell suspensions do not leak into the bottom beaker unfiltered). 10. Refill each of the eight tubes containing bones with 40 ml MLB, shake, and filter the supernatants through the same 350and 110-μm filters into the beakers containing the first filtrates on ice. 11. Dispense the final filtered solutions into 50-ml centrifuge tubes on ice and centrifuge at 210 × g for 10 min at 4°C. This pellet represents a crude fraction containing the majority of the nonviable OC (which tend to be much larger in size on average than the surviving OC) and a minor proportion of viable OC. Because it provides a valuable source of OC material for enzyme-linked immunoassay (ELISA), SDS-PAGE, Western blotting, and other biochemical assays, it is routinely stored at −80°C as one or two cell pellets overlaid with a drop of protease inhibitor cocktail solution B.

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12. To obtain the viable OC fraction for cell culture, incubate the bones in eight 50-ml tubes, each containing 35 ml of 0.333 mg/ml collagenase in HBSS-MLB, for 30 min at 37°C in a water bath. 13. Gently shake the tubes, discard this solution, add 35 ml of MLB per tube to rinse the bones for 15 min, and then transfer the bones with tweezers to eight tubes containing 35 ml of 0.045% trypsin in MLB-MLBE. Incubate for 30 min at 37°C in a water bath to detach viable OCs from the bone surfaces. 14. Shake the bones vigorously for 3 min, and pass the cell suspensions through a 350-μm Nitex filter into a plastic beaker on ice containing 1 ml of heparin (1,000 U, to reduce clotting) and 5 ml of FBS (to inhibit further trypsin action). 15. Immediately refill the tubes with 20 ml of MLB, shake the bones vigorously for another 3 min, and filter this cell suspension through the same 350-μm Nitex filter into the beaker on ice containing the first shaken suspension from step 14. Repeat once more. 16. Split and pour each half of the combined filtrates through separate 110-μm Nitex filters into beakers on ice (two filters are recommended because they clog easily). Dispense the filtrates into twelve 50-ml centrifuge tubes on ice. 17. Centrifuge the cell suspensions at 300 × g for 10 min at 4°C. Gently pour out the lipid pad and supernatant, and wipe out lipid and matrix material clinging to the side of the tube with a clean tissue before righting each tube. Resuspend each pellet, using a 10-ml wide-bore pipette, in 2–5 ml of chilled MLB (see Note 7). 18. Transfer the OC suspension to six new 50-ml tubes, add 0.1 ml heparin to each, fill the tubes to 50 ml with chilled MLB, and invert to mix. Centrifuge as in step 17 to wash the cells. 19. Discard the supernatants and resuspend the cell pellets (see Subheading 3.1.1). If OCs will be cultured, sterile techniques and solutions should be initiated at this point for Percoll fractionations. 3.1.1. Percoll Purification of Osteoclasts

Use sterile solutions and techniques throughout. 1. Resuspend each of the six OC pellets in 5 ml of ice-cold 35% Percoll, combine into one new tube, add 0.6 ml heparin, vortex briefly at low speed, and divide the cell suspension into four 50-ml tubes. 2. Raise the volume of each tube to 10 ml with additional 35% Percoll, and then slowly overlay each tube with 3.0 ml ice-cold HBSS (do not deform the interface).

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3. Centrifuge the tubes in a swinging bucket rotor at 440 × g for 20 min at 4°C, and then carefully remove the tubes without disturbing the gradients. 4. Slowly withdraw the interface and top 5–8 ml with a pipette (see Note 7), transfer these solutions into four new 50-ml tubes on ice containing 25 ml HBSS, and add ice-cold HBSS up to 50 ml. Discard the residual pellets. 5. Centrifuge at 300 × g for 10 min at 4°C, retain the pellets, and discard the supernatants. 6. The 35% Percoll-fractionated OCs can be used at this point for immunomagnetic purification (see Subheading 3.1.2) or purified further by 6% Percoll fractionation as described in steps 7–12. 7. Set up four tubes containing 10 ml of 6% Percoll on ice. 8. Resuspend each of the four OC pellets from step 5 thoroughly in 3 ml ice-cold HBSS using a 10-ml pipette, combine, and briefly vortex (if clumping is a problem, add 0.12 ml of 1,000 U/ml heparin). 9. Slowly overlay 3–3.5 ml of this suspension on top of each of the four 6% Percoll gradient tubes, and let the tubes stand undisturbed on ice for 1 h to allow OCs to penetrate the Percoll layer. 10. Remove the top 4 ml from each tube and discard. Combine the bottom fractions pairwise and dilute with ice-cold HBSS to 50 ml each. Centrifuge the cell suspensions at 300 × g for 10 min at 4°C. 11. Resuspend each of the two OC pellets in 5 ml α-MEM medium, combine, mix gently, and microscopically evaluate by withdrawing 0.1 ml to mix with 0.1 ml of 0.4% trypan blue and immediately assess OC yield, viability (unstained cells) and purity in a hemocytometer. 12. Meanwhile, centrifuge the OC suspension again at 300 × g for 10 min at 4°C, and resuspend the OC pellet in prewarmed medium to disperse into sterile dishes or multiwell plates for culture. Typically, enrichments of at least 40% on a per cell basis (>80% on a per nucleus basis) are routinely achieved for OCs after 35% Percoll fractionation, and yields of 1–3 million OCs exhibiting >85% viability for OCs are obtained following 6% Percoll fractionation. Further enrichment of the 6% Percoll OC population can be readily accomplished by allowing these cells to settle and attach to bone or ivory in culture for 2.5–3 h, after which the unbound cells are removed and the adherent OCs are gently washed once or twice with fresh medium before further culture.

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OCs can be further enriched after 35 or 6% Percoll fractionation via immunomagnetic sorting (9). However, significantly higher OC yields are obtained if immunomagnetic capture is performed on 35%, rather than 6%, Percoll-separated populations. The last steps of bead preparation (steps 1–4 below) should be timed so that the beads are ready to add as soon as the OCs have been separated on the 35% Percoll gradients. Beads should be handled gently in all steps. 1. Just prior to use, magnetically sort the beads that have been coupled with an anti-OC MAb to remove the MAb coupling solution, wash the beads three times by gentle resuspension in PBS (~1 ml) and magnetic sorting, and incubate them for 30 min with 250 μl sterile 1% FBS in PBS to block nonspecific bead attachment to cells. 2. Wash the beads three times with PBS, and resuspend in 200 μl of PBS in preparation for addition to the 35% Percollfractionated OCs. 3. At this point, the four OC pellets from the 35% Percoll gradient (see Subheading 3.1.1, step 5) are resuspended in a 50-ml polypropylene tube in a total volume of 6 ml of ice-cold HBSS. 4. Add the MAb-coupled magnetic beads from step 2 (200 μl), swirl the tube gently to quickly mix cells and beads (see Note 9), place the tube into a bucket of ice at a ~45° angle (with the bead–cell mixture clearly visible from the top), and set the bucket on a rotary shaker to slowly mix the beads and cells for 30 min (see Note 9). 5. Remove from the shaker, stand the tube upright in the ice, and push a magnet down into the ice and tightly against the lower part of the tube. Let stand ~5 min to draw bead-bound OCs over to the magnet, and use a pipette to slowly remove the unbound (non-bead) cell supernatant for transfer to a new 50-ml tube on ice (this is resorted later to capture any lost beads bound with cells). 6. Move the tube away from the magnet, resuspend the beadbound cells in 40 ml ice-cold sterile HBSS to wash, invert several times to mix gently, and then place the tube back against the magnet in ice. Incubate undisturbed for 5 min to capture bead-bound OCs and transfer the wash using a pipette to another tube (to resort later for lost beads). 7. Repeat step 6 twice more to wash the bead-bound OCs a total of three times. 8. To recover any bead-bound OCs remaining in the original unbound cell supernatant (step 5) or lost during the wash steps (steps 6–7), incubate each of these tubes against the magnet for 5 min on ice, remove and discard the supernatants, and

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resuspend additional bead-bound OCs in a small volume of HBSS and add back to the main sample of bead-bound OCs. 9. Resuspend the final collection of bead-bound cells in 2–5 ml HBSS, mix 0.1 ml with 0.1 ml of 0.4% trypan blue, and immediately assess the sample for OC yield, viability (unstained cells), and purity in a hemocytometer. 10. Sort the remaining 2–5 ml of immunocaptured OCs with a magnet and either resuspend the cells in medium for culture, immediately extract RNA, prepare protein lysates, or use the cells as needed. Typically, five- to tenfold greater OC purity is achieved with MAb 121F immunomagnetic affinity capture in comparison with 6% Percoll density gradient fractionation, with immunomagnetic OC enrichments of up to 90% on a per cell basis and over 98% on a per nuclear basis. 3.2. Osteoclast Culture 3.2.1. Percoll-Purified OCs

1. Resuspend 6% Percoll-purified OCs in 5 ml of culture medium and immediately plate out at: (a) 0.5 ml per well (~100,000 OCs) in 10 wells of a 24-well dish, with or without a glass coverslip in the bottom of the well and/or 2–4 sterile discs of bone or ivory per well (see Subheading 2.5). (b) 0.2 ml per well (~50,000 OCs) in 20 wells of a 48-well dish, with or without one sterile disc of bone or ivory (see Note 10). 2. To enrich further for OCs on bone or ivory, change the medium after 2–3 h to selectively capture OCs onto the bone or ivory substrate. Modulators can be added at this point in fresh medium. Otherwise, change the medium after 16 h of incubation, and add modulators in fresh medium. 3. Culture for the designated period of time, typically 1–2 days (see Note 11).

3.2.2. Immunomagnetically Purified OC (see Note 8)

Because the yield of OCs is lower following immunomagnetic purification than Percoll density gradient separation, highly purified immunomagnetic OC populations are most useful for confirming in a limited fashion that biochemical or functional properties observed with Percoll-fractionated OC can be directly attributed to OCs and not rare contaminating cells. 1. Culture immunomagnetically purified OCs on bone, ivory, glass, or plastic in the presence or absence of modulators for up to several days as described in Subheading 3.2.1 above. 2. Although the temperature is kept at or below 4°C during immunomagnetic capture to prevent OCs from phagocytosing the beads bound to their outer surface, these are internalized within min once the cells are exposed to a higher temperature (see Note 12).

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3.3. Assay Techniques 3.3.1. Morphology and Ultrastructure


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Standard protocols are used to evaluate the morphological and ultrastructural characteristics of isolated chick OCs. When viewed by light microscopy, chick OCs appear as large multinucleated cells of varying sizes and shapes, with a grainy cast and often one or more pseudopodial extensions per cell. Immunomagnetically isolated chick OCs are typically decorated with multiple beads per cell and may be so thoroughly coated with MAb-conjugated beads that they resemble a ball of beads (Fig. 1a). On culture of immunomagnetically captured OCs on bone, ivory, glass or plastic, the cells spread out and internalize the beads, rather than shedding them as do nonphagocytic cells.

Fig. 1. Chick OC immunomagnetic purification and bone pit resorption. (a) Chick OCs from 35% Percoll preparations were affinity captured and purified via their binding to MAb 121F-coupled magnetic beads. Phase-contrast microscopy reveals numerous beads avidly attached to and covering OC cell surfaces. (b) SEM appearance of 6% Percoll-purified chick OC preparations cultured on plastic. Scale bar = 50 μm. (c) SEM analysis of MAb 121F immunomagnetically isolated chick OC and an associated resorption pit formed during culture on bone. Scale bar = 10 μm.

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OC morphology and ultrastructure can be analyzed using transmission (TEM) or scanning (SEM) electron microscopy as detailed in other chapters of this volume. Features characteristic of OC and evident by TEM include multiple nuclei often clustered within the cell and varying in number between cells, abundant mitochondria, numerous vesicles, extensive vacuolation, welldeveloped perinuclear Golgi complexes, prominent rough endoplasmic reticulum, free polysomes, and ruffled border membrane and clear zone domains. By SEM, chick OCs cultured on plastic typically appear as large cells having a complex morphology with many fine filopodial projections, microvilli and membrane blebs visible over the cell surface, and a peripheral cytoplasmic skirt (Fig. 1b). When cultured on bone or ivory, chick OCs appear by SEM either as large domed cells actively engaged in excavating a resorption cavity or as stretched inactive cells having a motile phenotype and characteristic leading and trailing membrane domains (Fig. 2f). OCs associated with resorption pits often exhibit membrane projections stretched back over a portion of the wellexcavated lacuna that has exposed collagen fibrils (Fig. 2f). Resorption pits formed by cultured chick OCs are typified by multilobulated excavations or long resorption tracks (which also may be multilobulated) or, less often, as a unilobular cavity adjacent to or underlying an OC actively involved in resorption (Fig. 2f). Immunomagnetically isolated OCs may exhibit less pit resorbing activity (Fig. 1c). The morphology of resorption pits can be examined more thoroughly by SEM following the removal of OCs from the bone or ivory substrate (either initially before a gold coating step or after viewing the sample and then recoating with gold to visualize the pits alone). 3.3.2. Cytochemical Staining

A quick and easy way to discriminate nuclear and cytoplasmic detail in cultured chick OCs is by use of the general differential stain Diff-Quik, which is simply incubated with the fixed chick OCs for several min and then rinsed off. However, by far the most commonly used stain to visualize OCs is based on their high level of TRAP activity which is upregulated early in OC development. Although not specific for OCs alone, this cytochemical stain readily identifies OCs in bone tissue sections and in isolated OC preparations (Fig. 3a, c). TRAP activity also can be quantified in cell extracts using a microplate enzymatic assay (and normalized for cell extract protein) (12). To stain for TRAP activity using freshly prepared solutions (see Subheading 2.3 and Note 2): 1. Remove the culture medium from the cells (and discard or save for other analyses). 2. Rinse the cells quickly three times with warm HBSS (tilt dish to add and remove solutions gently), add 1% PF-HBSS solution

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Fig. 2. Chick OCs form resorption pits on ivory and phagokinetic tracks on gold-coated coverslips. (a–c) 6% Percoll-purified chick OCs were cultured on ivory (2–3 days) and harvested for TRAP staining and resorption pit analysis. As viewed by light microscopy, OCs formed multilobulated resorption tracks that frequently were composed of connecting resorption lacunae. These represent periods of OC attachment and pit formation, followed by OC movement to an adjacent area of ivory for further resorption. (d) Resorption pits viewed by darkfield reflective light microscopy, as performed for quantifying the number and areas of resorption pits within the exact same fields evaluated for OC numbers. (e) 6% Percoll-purified chick OCs cultured on gold-coated coverslips for 16 h and subsequently stained for TRAP activity. OCs phagocytose the gold particles and thereby clear a path during their movement across the gold-coated coverslip. The numbers and areas of such phagokinetic tracks are measured, and expressed relative to the numbers of associated OCs. (f) SEM analysis of 6% Percoll-purified chick OCs engaged in bone pit resorption on ivory. Note the deep, well-excavated lacunae that are typically formed by chick OCs. Scale bar = 50 μm.

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(~300 μl per 48-well, 500 μl per 24-well-plate) to fix the cells for 15 min at room temperature, remove the fixative, rinse three times with HBSS and once with deionized water, and then either air dry the samples overnight or incubate them in −20°C methanol for several minutes followed by a water rinse to permeabilize the cells. 3. Add staining solution to cover the cells in the wells, and incubate the dish or plate at 37°C for 1 h in the dark. 4. Remove the stain, rinse the samples several times with water, and air dry the samples (on the dish or ivory) or mount coverslips by inverting coverslips onto a drop of glycerol-buffered mounting medium spotted on a glass slide (thereafter store at 4°C and rewarm before viewing in a microscope). 5. Alternatively, a commercially available staining kit (Sigma cat. no. 386) can be used as directed by the manufacturer (see Note 2). 3.3.3. Antigenic Profile

Together with specific morphological features and high TRAP activity levels, chick OCs exhibit various characteristic surface markers that are commonly monitored since they are not expressed (or only at lower levels) by related monocytes, macrophages, or macrophage polykaryons. These include expression of αvβ3 integrin (vitronectin receptor), H+-ATPase proton pump, carbonic anhydrase II, calcitonin receptor, galectin-3, and a series of antigens recognized by anti-OC MAbs, including 121F (Fig. 3b, d, f, g). Most or all of these OC markers play important roles in the bone resorption function and/or survival of OCs. Using specific antibodies, these markers can be detected on the surface of chick OCs cultured and fixed on bone, ivory, glass or plastic via immunostaining, alone or in combination with TRAP staining, F-actin cytoskeletal staining (with rhodamine-labeled phalloidin), and/or DAPI nuclear labeling (Fig. 3a–g). The relative surface level expression of these protein markers can also be measured for chick OCs following their fixation (as whole cells) in 96-well microtiter dishes and quantitative analysis by ELISA as detailed elsewhere (12). Alternatively, these markers may be monitored in total cell extracts or membrane lysates of chick OCs by ELISA, gel electrophoresis (with or without immunoprecipitation), or immunoblotting. OCs also express high intracellular levels of pp60c-src, a critical signal molecule required for OC bone resorption. This cytoskeletally associated protein can be detected by immunostaining in permeabilized cells (e.g., incubate fixed OCs in 0.1% Triton X-100 for 30 min prior to blocking) or by immunoblotting of electrophoresed cell extracts, with or without probing for phosphorylation status (see Chapter 15 this volume). Similar methods can be used to detect many other intracellular signaling molecules. A general protocol for OC

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Fig. 3. Cytochemical and immunostaining analysis of 6% Percoll-purified chick OCs. (a, c) Chick OCs cultured on plastic (1–2 days) and stained for TRAP activity. (b, d) Chick OCs cultured on plastic and immunostained using a MAb to the vitronectin receptor, integrin αvβ3 (LM 609), and a biotin-streptavidin β-galactosidase detection system. (e) Chick OCs cultured on plastic, fixed and permeabilized (Triton X-100), and double stained with rhodamine phalloidin to label cytoskeletal F-actin (red ) and DAPI to label multiple nuclei within OCs (blue). Note the peripheral actin ring formation characteristic of mature OCs. (f, g) Chick OCs cultured on plastic and immunostained with MAb 121F using a biotin-streptavidin FITC detection system. Both αvβ3 and the 121F antigen become highly expressed on the OC cell surface during their differentiation into bone-resorptive multinucleated cells, and each of these OC markers plays an important functional role in the resorption of bone by OCs.

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immunostaining is given here (use a minimum of 250 μl of reagent per well of a 24-well plate): 1. After culture of OCs on coverslips, bone or ivory, rinse the tissue culture wells gently and fix as for TRAP staining (see Subheading 3.3.2, steps 1–4). 2. Immediately process the samples for immunostaining and do not allow any part of the sample to dry during the following steps (including the edges of coverslips or bone/ivory slices since this will produce staining artifacts). 3. Block nonspecific binding sites by incubating with blocking solution for 1 h at room temperature. 4. Incubate with appropriate dilutions of anti-OC antibodies for 1 h at room temperature. Include sample(s) incubated in blocking solution alone or an irrelevant antibody to serve as negative controls for nonspecific staining. 5. Rinse three times briefly, and once for 10 min with PBS. 6. Incubate with a secondary biotin-conjugated antibody directed against the primary antibody for 1 h at room temperature. 7. Rinse three times briefly, and once for 10 min with PBS. 8. Incubate 30 min in the dark with streptavidin conjugated with FITC (or Texas Red). 9. Rinse three times briefly and once for 10 min with PBS. 10. Mount specimens onto glass slides with glycerol-buffered mounting medium (see Subheading 3.3.2, step 5), store dark at 4o C, and rewarm slides before viewing in a microscope. 11. If desired, OCs on coverslips can be briefly reacted with a membrane permeable fluorescent dye to label the nuclei (bright blue) by incubation in a 1:300 dilution of DAPI in HBSS for 1 min, followed by two rinses in HBSS before mounting. Immunostained samples can also be stained for TRAP activity (after step 9 above) before being mounted on glass slides. Antigen detection on OCs adherent to bone or ivory is difficult to measure using a fluorescent system unless confocal microscopy is used (see Chapter 25). A histochemical detection method may be better suited for this purpose, and also works well to immunostain OCs cultured on glass or plastic in place of the fluorescent system. We prefer to use antibodies coupled to β-galactosidase (which has negligible background problems and requires no specific blocking), but other enzymes (e.g. horseradish peroxidase) can also be used with good results if endogenous enzyme activities are quenched. For β-galactosidase-based immunostaining, perform steps 1–7 of the protocol given earlier in this subheading, then continue as follows: 1. Incubate for 30 min with streptavidin conjugated with β-galactosidase in buffer A.

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2. Rinse five times with buffer A over 30 min. 3. Incubate for 30 min (or longer, if necessary) in the dark with substrate solution. 4. Rinse five times over 30 min with PBS. 5. Store bone or ivory slices dry before viewing. Mount cells on coverslips onto glass slides as described in step 10 earlier in this subheading. 6. OCs immunostained by this method (on bone, ivory, glass or plastic) also can be double stained for TRAP activity (see Subheading 3.3.2). 3.3.4. Molecular Profile

Both Percoll-fractionated and immunomagnetically purified chick OCs are good RNA sources for analyzing the relative gene expression levels of various OC phenotypic and functional markers, either in freshly isolated cells or following OC culture on plastic, bone or ivory in the presence or absence of modulators. OC preparations provide sufficient RNA for ribonuclease protection assay (RPA), RT-PCR and other applications (13). Typically, one well of a 24-well dish seeded with 200,000 viable chick OCs (in 250 μl medium) yields 2–5 μg of total RNA, of which 100 ng–1 μg may be used per RT-PCR, or up to 5 μg for a single RPA assay. Chicken-specific primers for PCR amplification are available for some chick OC markers, and others can be generated based on interspecies sequence similarities reported for OC genes from human, mouse, or other species. Similarly, species homologous primers may be used to amplify chicken-specific genes and the PCR products cloned into appropriate vectors for the preparation of RPA probes (13). Methods for molecular analyses are given elsewhere in this volume (see Chapters 17 and 18).

3.3.5. Motility (see Note 4)

Because OCs are very large and vary considerably in size (and nuclear number), it is difficult to perform classical chemotaxis/ chemokinesis experiments through porous membranes to measure OC movement in response to various agents. However, OC movement can easily be monitored by culture on gold-coated coverslips, because OCs phagocytose the gold and thereby generate a cleared track in their wake (10). To perform this assay, 6% Percoll-purified OCs are used (see Subheading 3.1.1). 1. Resuspend the cells gently in 6–8 ml of culture medium, plate 0.5 ml (~100,000 OCs) per well of a 24-well dish containing rinsed and prewetted gold-coated coverslips (see Subheading 2.6), and culture for 2–3 h to allow OCs to attach. 2. Remove the nonadherent cells, add fresh medium with or without modulators, and culture the cells for 16–24 h. 3. Rinse gently three times with warm HBSS, fix the cells with 1% PF-HBSS for 15 min at room temperature, and rinse three times with HBSS.

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4. Stain the cells for TRAP activity (see Subheading 3.3.2 and Note 2). 5. Determine the number of TRAP-stained OCs, the number of phagokinetic tracks, and the cleared area of each track within a constant number of random adjacent fields using a microscope fitted with an ocular reticle and computer linked to an image analysis system (Fig. 2e). Calculate the mean track area and the total area of gold cleared, and normalize the data to determine the number of tracks per OC and the mean area cleared per OC. 3.3.6. Bone Resorption

OCs and other phagocytic cells can all resorb (ingest and degrade) very small particles of bone in vitro or secrete acid that dissolves hydroxyapatite [e.g., on calcium phosphate-coated dishes (Osteologic), BD Biosciences]. However, only OCs can create resorption pits on bone. This is, therefore, the key defining attribute and best assay for evaluating the bone resorptive function of fully developed OCs. Because the number of new sites initiated and the rate of resorption by OCs are major parameters controlling bone remodeling in both normal and pathological states, the in vitro bone pit resorption assay has become a very valuable investigational tool. Data obtained from this analysis reveals information about whether a modulator has altered the number of OCs on the bone or ivory (possibly reflecting effects on integrin-mediated attachment, cell survival or development), the number of pits formed (reflecting activation of OCs for initiating pit resorption), and the area of bone or ivory resorbed (overall or per pit, reflecting the amount and rate of resorption by OCs). 1. Culture chick OCs (6% Percoll purified) on bone or ivory in the presence or absence of modulators for 30–40 h (see Note 13). 2. Rinse, fix, and stain for TRAP activity as described in Subheading 3.3.2. 3. Evaluate resorption using a microscope fitted with an ocular reticle and computer-linked image analysis system (see Chapter 12 for description of a system) (Fig. 2a–c). 4. Count the number of TRAP-stained OCs within a constant number of random fields per bone slice, usually measured consecutively from an arbitrary starting location on the edge of the chip that is marked with a dot using a permanent ink marker. The number of fields chosen for analysis should encompass at least 100–300 OCs per bone chip (typically ~20 fields or half of the chip). To ensure that the exact same fields are subsequently analyzed for resorption pits, mark or draw the fields that have been evaluated for each chip on a grid log sheet.

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5. After all the bone chips have been analyzed for OC numbers (to ensure that no category has too few OCs), remove the OCs from the bone surface by soaking the chip for 1 min in 0.2 M NH4OH, rubbing the entire surface with gloved fingers, repeating this treatment a second time, and then rinsing the chip in deionized water. 6. Quantify the number and planar area of each resorption pit contained within the fields evaluated for OC numbers in step 4 using darkfield reflective light microscopy (Fig. 2d). 7. Express resorption measures as the mean number of OCs, number of pits, and total areas resorbed in this constant number of fields for each experimental condition. Also normalize the data to report the mean number of pits per OC, area resorbed per OC, and area (size) per individual pit. 8. In general, several trials, with 4–6 replicates each for control and treated groups, should be performed to achieve statistically significant results.

4. Notes 1. OC-specific antibodies that can be used for this purpose include MAb 121F (available from us upon request), antibodies to integrin αvβ3 (e.g., 23C6 or LM609) and other commercially available antibodies. 2. Cytochemical staining for TRAP activity is routinely performed using either freshly prepared reagents or a commercially available staining kit (Sigma cat. no. 386) as directed. Because TRAP staining intensity tends to be stronger with freshly prepared reagents, this protocol is preferred for tracking changes in OC TRAP activity (during development or in response to modulators), for discriminating OCs in quantitative resorption pit analyses, and for double staining of immunostained OCs. 3. Secondary antibodies may be directly conjugated with an enzymatic or fluorescent probe, but greater sensitivity is achieved if a biotinylated secondary antibody is used to amplify the primary antibody signal. 4. Gelatin pre-coating enables the gold particles to more reliably remain adherent to the glass coverslips. BSA cannot substitute for gelatin, as it interferes with chick OC movement on the gold-coated coverslips. Avoid plating too many OCs on the gold-coated coverslips because overlapping tracks become difficult to analyze. Similarly, incubation times should not exceed 16–24 h because phagokinetic tracks may become too long and convoluted (and overlap), migration differences in

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response to agents may diminish, and OCs may cease moving when overloaded with ingested gold particles. 5. Young posthatch growing chicks represent a highly abundant source of OCs, whose numbers are further increased by maintaining the chicks on a low calcium feed diet. However, the calcium level in this feed should not fall below 0.15% or the bones of the young chicks will become so soft and weak that the birds are unable to stand to walk, eat, or drink. The feed is prepared by special order (Purina) and can be stored at 4°C (kept dry to avoid mold) for up to 12 months. The birds should have free access to tap water (not deionized water which makes them too weak when they are on the low calcium feed). Whereas millions of OCs are obtained from such chicks, more than 1,000-fold fewer OCs are typically isolated from mouse, rat, or rabbit bone preparations, and only negligible OC numbers from most human bone tissue. Besides sharing morphological, phenotypic, and antigenic properties and most or all of the regulatory responses observed with OCs from other species, chick OCs are the most aggressively active species for bone pit resorption. Consequently, chick OCs provide a particularly sensitive assay system to measure the regulatory effects of various agents (many of which suppress, rather than stimulate, OC activity) on OC-mediated bone resorption. With the discovery of the RANKL-RANK-OPG regulatory pathway controlling OC development, resorption, and survival, many of the earlier restrictions on studies with OCs from other species have been alleviated since OCs may now be generated in vitro from precursors present in primary cell preparations (e.g., avian, mouse or human bone marrow or circulating monocytes) or cell lines (murine RAW 264.7 cells). Details of such procedures are given elsewhere in this volume. However, it may still be important in some cases to compare responses obtained with in vitro generated OCs against those of isolated OCs formed in vivo, especially if a particular agent cannot be directly investigated in vivo for its OC-related actions. 6. OC viability is dependent on the total length of time that it takes from the removal and processing of the bones until the 6% Percoll-fractionated or immunomagnetically isolated OC are placed into culture. This time should not exceed 6–7 h, as each additional hour will negatively impact the final OC viability. If the bone marrow is to be used (e.g., for OC precursor studies and RANKL development) from these same chicks, one person should blow out the bone marrow in a sterile hood from a group of bones while other individuals are harvesting or cleaning the remaining bones, and the dishes should be passed back and forth until they are all completed. One person then

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continues with the bone marrow cell preparation independently from those working on isolating OCs from the marrow-stripped bones (12). 7. Use only wide-bore pipettes or tips for any work in isolating or manipulating OC to avoid fragmenting these large multinucleated cells. Also, exercise care in resuspending, mixing, or vortexing OC preparations gently and for as little time as necessary. 8. Immunomagnetic isolation provides a rapid and highly efficient way to purify OCs from mixed cell populations. Magnetic beads coupled with 121F MAb are not species restricted and can effectively purify OCs from rat, rabbit, human and other sources. Other anti-OC MAbs also may be coupled to magnetic beads for use in purifying OCs via this procedure (see also Note 1). The procedure is performed on ice to prevent ingestion of the beads by OCs. However, because MAb 121F (bivalent or Fab fragments) partially inhibits OC bone pit resorption (14), and OC yields are lower in immunomagnetic compared to Percoll purifications, immunomagnetically isolated OCs are considered most useful for (1) obtaining highly purified OC samples for molecular or biochemical analysis and (2) confirming that responses observed in Percoll-purified OC preparations can be attributed to OC-specific effects. 9. For optimum OC immunomagnetic capture, avoid stirring the MAb coupled beads with the cells either too fast (which interferes with attachment) or too slow (which reduces binding due to poor mixing). Best results are achieved with 35% Percollseparated OC preparations used as the starting material as opposed to more crude preparations because the latter yields less pure OC populations and matrix reassembly is more problematic. In addition, 6% Percoll preparations typically yield fewer immunomagnetically isolated OCs than do 35% Percoll preparations. 10. As an alternative to seeding OC onto individual bone slices, OC may be seeded in 2.5 ml of medium onto ~24 bone or ivory discs spread out to fully cover the bottom of a 35-mm dish. After allowing OCs to selectively attach for 2.5–3 h, the nonadherent cells are removed, adherent OCs are gently washed with medium, and the bone or ivory discs are individually removed with sterile tweezers and placed into one well of a 48-well culture dish with fresh medium (250 μl). Modulators are typically administered (in 50 μl of medium, final volume of 300 μl per well) and the cells cultured for 30–40 h before harvest. 11. For resorption, OCs are routinely cultured for 30–40 h before harvest. For histochemical, enzymatic, or immunocytochemical

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analysis, OCs are cultured for 1–2 days before analysis. For molecular studies, RNA may be extracted directly from the Percoll-fractionated or immunomagnetically captured cells. Alternatively, RNA can be harvested from OCs cultured on bone, ivory or plastic for up to 3 days in the presence or absence of modulators. OC survival is enhanced if the cells are cultured on bone or ivory (due to integrin-mediated survival signals) as opposed to glass or plastic, so experiments under the latter conditions should be limited to a few days at most. Although some studies have indicated that OC resorb better under slightly acidic conditions, we find that chick (and human) OC performance is actually better in α-MEM supplemented with 5% FBS. 12. Immunomagnetically purified OCs can be cultured and will form resorption pits on bone or ivory, but because they rapidly ingest the bound beads at 37°C, it is conceivable that antibody and/or bead engagement of the OC cell surface may affect their physiology or resorptive function (Fig. 1a, c). It is possible to remove many, but not all, of the beads from the outer surface by physical (strong vortexing) or biochemical (low pH, protease digestion) methods, performed at 4°C, although some cell damage may occur during these procedures (9). 13. Because ethanol inhibits OC bone resorption, the bone/ivory slices must be well rinsed (and soaked >3 h) in HBSS before having cells plated onto them. Likewise, alcohol-soaked tweezers should be air-dried briefly before being used to move bone or ivory discs. In general, resorption pit analysis on bone is somewhat more complicated than on ivory due to the need to distinguish Haversian and Volkmann canals in the bone apart from the pits made by the cultured OCs. In our replicate studies using bone and ivory, no substrate-dependent differences have been noted to date in either basal or modulator evoked resorption parameters for isolated chicken OCs. Therefore, although ivory can be more difficult to obtain than bovine bone, it is preferable to use for quantitative resorption pit analysis.

Acknowledgments This work was supported by NIH Grants AR32927, AG 15435, and AR42715 to P.O.

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References 1. Hall, T. and Chambers, T. (1996) Molecular aspects of osteoclast function. Inflamm. Res. 45, 1–9. 2. Roodman, G. (1996) Advances in bone biology – the osteoclast. Endocr. Rev. 17, 308–332. 3. Suda, T., Udagawa, N., Nakamura, I., et al. (1995) Modulation of osteoclast differentiation by local factors. Bone 17, S87–S91. 4. Kanis, J. (1995) Bone and cancer: pathophysiology and treatment of metastases. Bone 17, 101S–105S. 5. Mundy, G. (1993) Cytokines and growth factors in the regulation of bone remodeling. J. Bone Miner. Res. 8, S505–S510. 6. Wiebe, S., Hafezi, M., Sandhu, H., et al. (1996) Osteoclast activation in inflammatory periodontal diseases. Oral Dis. 2, 167–180. 7. Manolagas, S., Bellido, T., and Jilka, R. (1995) New insights into the cellular, biochemical, and molecular basis of postmenopausal and senile osteoporosis: roles of IL-6 and gp130. Int. J. Immunopharmacol. 17, 109–116. 8. Oursler, M., Collin-Osdoby, P., Anderson, F., et al. (1991) Isolation of avian osteoclasts: improved techniques to preferentially purify viable cells. J. Bone Miner. Res. 6, 375–385. 9. Collin-Osdoby, P., Oursler, M., Webber, D., et al. (1991) Osteoclast-specific monoclonal antibodies coupled to magnetic beads provide a

10.

11.

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13.

14.

rapid and efficient method of purifying avian osteoclasts. J. Bone Miner. Res. 6, 1353–1365. Minkin, C. (1982) Bone acid phosphatase: tartrate-resistant acid phosphatase as a marker of osteoclast function. Calcif. Tissue Int. 34, 285–290. Owens, J. and Chambers, T. (1993) Macrophage colony-stimulating factor (M-CSF) induces migration in osteoclasts in vitro. Biochem. Biophys. Res. Commun. 195, 1401–1407. Collin-Osdoby, P., Oursler, M., Rothe, L., et al. (1995) Osteoclast 121F antigen expression during osteoblast conditioned medium induction of osteoclast-like cells in vitro: relationship to calcitonin responsiveness, tartrate resistant acid phosphatase levels, and bone resorptive activity. J. Bone Miner. Res. 10, 45–58. Sunyer, T., Rothe, L., Kirsch, D., et al. (1997) Ca2+ or phorbol ester but not inflammatory stimuli elevate inducible nitric oxide synthase messenger ribonucleic acid and nitric oxide (NO) release in avian osteoclasts: autocrine NO mediates Ca2+-inhibited bone resorption. Endocrinology 138, 2148–2162. Collin-Osdoby, P., Li, L., Rothe, L., et al. (1998) Inhibition of osteoclast bone resorption by monoclonal antibody 121F: a mechanism involving the osteoclast free radical system. J. Bone Miner. Res. 13, 67–78.

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Chapter 10 Isolation and Purification of Rabbit Osteoclasts Fraser P. Coxon, Michael J. Rogers, and Julie C. Crockett Abstract Newborn rabbits provide a useful and readily available source of authentic mature osteoclasts, which can be easily isolated directly from the long bones in relatively large numbers, compared to other rodents. Primary cultures of authentic rabbit osteoclasts on resorbable substrates in vitro are an ideal model of osteoclast behaviour in vivo, and for some studies may be preferable to osteoclast-like cells generated in vitro from bone marrow cultures or from human peripheral blood, for example in assessing osteoclastmediated bone resorption independently of effects on osteoclast formation. Rabbits also provide a particularly useful model for determining the effects of pharmacological agents on osteoclasts in vivo, by isolating osteoclasts using immunomagnetic bead separation (with an antibody to αVβ3) at the desired time following in vivo administration of the drug. Since osteoclasts are abundant in newborn rabbits, sufficient numbers of osteoclasts can be retrieved using this method for molecular and biochemical analyses. Key words: Rabbit, Osteoclast, Bone resorption, Polarisation, Vitronectin receptor

1. Introduction Osteoclasts are notoriously hard to study because of the difficulty in obtaining pure populations of cells in large numbers for biochemical and molecular analyses. Unlike with other rodents (e.g. mice and rats), mature osteoclasts can be obtained from rabbits in relatively large numbers and can be purified easily. For some studies, such primary cultures of authentic osteoclasts may be preferable to osteoclast-like cells generated in vitro from bone marrow cultures, or from human peripheral blood. Isolated rabbit osteoclasts are capable of resorbing mineralised substrates in vitro and are therefore useful for assessing the effect of pharmacologic agents on osteoclast-mediated bone resorption (1–3), independently of effects on osteoclast formation, which can complicate interpretation of studies using mouse or human osteoclasts. Furthermore, since they are isolated directly, rabbit osteoclasts cultured on a mineralized substrate in vitro are an ideal model of osteoclast Miep H. Helfrich and Stuart H. Ralston (eds.), Bone Research Protocols, Methods in Molecular Biology, vol. 816, DOI 10.1007/978-1-61779-415-5_10, © Springer Science+Business Media, LLC 2012

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behaviour in vivo (4, 5). We and others have also extracted protein or RNA from purified rabbit osteoclasts for studies on metabolic processes in osteoclasts, or molecular studies on osteoclast biology using western blotting, enzyme assays, or RT-PCR (3, 6–9). We routinely isolate osteoclasts from the long bones of neonatal rabbits using a method adapted from that initially described by Tezuka et al. (10) (see Subheading 3.1). Isolated rabbit osteoclasts can then be cultured on plastic, glass or mineralised substrates (such as cortical bone, elephant ivory, or whale dentine, or hydroxyapatite-coated surfaces). Culturing osteoclasts on glass coverslips (in multiwell plates) is useful for immunocytochemistry, since the coverslips can be mounted onto glass slides, enabling cells to be visualised using an upright microscope. For some applications, such as preparation of osteoclast lysates for western blot analysis, osteoclasts must be further purified from contaminating bone marrow cells. This can be done either by further washing culture dishes with PBS, or by removing contaminating adherent cells using a solution of pronase–EDTA (Subheading 3.2). This provides cultures of >95% pure, tartrate-resistant acid phosphatase (TRAcP)-positive, multinucleated osteoclasts and mononuclear, prefusion osteoclasts (Fig. 1a). When isolating mature osteoclasts we typically achieve a yield of approximately 5 × 104 purified osteoclasts from each rabbit. It is also possible to generate much larger numbers of osteoclast-like cells (up to 16 semi-confluent 10 cm petri dishes of TRAcPpositive, multinucleated cells per rabbit) (Fig. 2), by culturing bone marrow-derived cells in the presence of 1,25 dihydroxyvitamin D3 over a period of 10 days, using a method modified from David et al. (11) (see Subheading 3.3). For in vitro applications, purification of osteoclasts on culture dishes by pronase–EDTA digestion is sufficient to provide pure osteoclasts for biochemical studies. However, a pure population of

Fig. 1. (a) Phase-contrast photograph of multinucleated, rabbit osteoclasts cultured in a plastic petri dish following purification with PBS (bar = 25 μm). (b) A rabbit osteoclast showing vitronectin receptor expression by fluorescence immunostaining.

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Fig. 2. Phase-contrast photographs of a rabbit bone marrow culture (a) after 7 days, showing a developing multinucleated, osteoclast-like cell (arrow ) beneath the stromal cell layer; (b) following purification of osteoclast-like cells after 10 days (multinucleated cell shown in inset ). Bars = 20 μm.

Fig. 3. Scanning electron micrograph of an immunomagnetically isolated rabbit osteoclast cultured on dentine. The osteoclast is associated with a resorption lacuna and still has magnetic beads attached. Bar = 10 μm.

osteoclasts can also be isolated in relatively large numbers directly from rabbit bones, without prior cell culture in vitro. We have modified a technique (Subheading 3.4) developed by Collin-Osdoby et al. (12) that involves separation of osteoclasts from a mixed cell suspension using immunomagnetic beads and the 23C6 monoclonal antibody (Fig. 3). The latter specifically recognises the αVβ3 integrin (also known as the vitronectin receptor), which is highly expressed on osteoclasts (13) (Fig. 1b). This technique is particularly useful for determining the effects of pharmacological agents on osteoclasts in vivo, since the osteoclasts can be isolated and purified at the desired time following in vivo administration of the drug.

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The osteoclasts can then be examined by microscopy or lysed for analysis by western blotting or using other molecular and biochemical techniques. This approach has been used to examine the ability of bisphosphonates and phosphonocarboxylates (14–16), and statins (17, 18) to inhibit protein prenylation in osteoclasts in vivo, uptake of bisphosphonates in vivo (16) and to detect the accumulation of a toxic metabolite of clodronate (14) and a metabolite of the mevalonate pathway in osteoclasts in vivo (19). Importantly, the separation of osteoclasts from other cells within the bone marrow is sufficiently effective to determine whether the pharmacological agents under investigation act specifically on the osteoclasts. Rabbit osteoclasts in culture can be identified using markers that are highly abundant in these cells, for example by staining for TRAcP (20) (Subheading 3.5 step 1) and immunological detection of the vitronectin receptor and their (Fig. 1b) (see Subheading 3.5 step 2). Staining for the activity of TRAcP is useful for enabling the number of osteoclasts in culture to be counted (21), particularly when the cells are cultured on a substrate, such as dentine on which cells cannot easily be seen light microscopy. It should be noted, however, that this enzyme is not specific for osteoclasts and is present in other cell types, such as alveolar macrophages. Immunostaining for the vitronectin receptor is particularly useful for single cell studies using fluorescence microscopy, since it delineates the plasma membrane as well as selectively identifies the osteoclasts. When rabbit osteoclasts are cultured on a mineralised substrate such as dentine, actively resorbing osteoclasts can be identified by the presence of a characteristic ring of F-actin (see Subheading 3.5 step 3) and their ability to excavate resorption pits in the substrate (see Subheading 3.5 step 4) (Fig. 4).

Fig. 4. Resorptive activity of rabbit osteoclasts cultured on dentine discs. (a) Scanning electron micrograph showing a cultured rabbit osteoclast (arrow ) adjacent to a resorption pit (asterisk ). Bar = 10 micro m. (b) Rabbit osteoclasts purified through an FCS gradient and cultured on dentine, then stained with TRITC-phalloidin and analysed by fluorescence microscopy. Characteristic F-actin rings are evident, denoted by arrows. (c) Reflected light microscopic image of resorption pits excavated by rabbit osteoclasts (original magnification ×100).

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2. Materials 2.1. General Reagents

1. α-Minimum essential medium (α-MEM) supplemented with 100 U/ml penicillin, 100 μg/ml streptomycin, and 1 mM glutamine. 2. Foetal calf serum (FCS). 3. Phosphate-buffered saline (PBS). 4. 4% Formaldehyde in PBS. 5. Hank’s buffered salt solution (HBSS).

2.2. Isolation and Purification of Rabbit Osteoclasts

1. Sharp scissors. 2. Blunt-ended forceps. 3. Disposable scalpel (for removing tissue). 4. Autoclaved scalpel handle and disposable scalpel blade (for mincing bones). 5. 10 cm diameter glass petri dishes. 6. PBS containing 0.001% (w/v) pronase and 0.002% (w/v) EDTA. Filter-sterilise (0.2 μm filter) before use. Pronase can be prepared as a concentrated stock solution in PBS and stored frozen as aliquots at −20°C, and then be diluted in PBS/EDTA before use. 7. HBSS (for serum gradient enrichment of osteoclasts).

2.3. Generation of Rabbit OsteoclastLike Cells

1. 1,25 Dihydroxyvitamin D3 (Sigma, Poole, UK).

2.4. Isolation of Rabbit Osteoclasts Using Immunomagnetic Beads

1. Anti-αVβ3 mAb (clone 23C6; Serotec, Oxford, UK). 2. 0.1% (w/v) Bovine serum albumin (BSA) in PBS. 3. Magnetic beads conjugated to rat anti-mouse IgG (e.g. Dynabeads from Invitrogen, Oregon, USA). 4. Magnet or magnetic particle concentrator.

2.5. Staining for Tartrate Resistant Acid Phosphatase

1. 10 mg/ml Naphthol-AS-BI-phosphate substrate in dimethylformamide (stable at 4°C for about 2 weeks). 2. 4% (w/v) Sodium nitrite. 3. Pararosanilin: add 1 g of pararosanilin to 20 ml dH2O and then add 5 ml concentrated HCl. In a fume hood, heat the solution carefully with constant stirring in a water bath for 30 min and then filter after cooling (stable at 4°C for several months).

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4. Veronal buffer: 11.7 g/l anhydrous sodium acetate, 29.4 g/l veronal (barbital) in dH2O (toxic solution) (stable at 4°C for several months). 5. 0.1 N acetate buffer, pH 5.2: dissolve 0.82 g anhydrous sodium acetate in 100 ml dH2O. Adjust the pH of this solution to 5.2 using a solution of 0.6 ml glacial acetic acid made up to 100 ml with dH2O (stable at 4°C for several months). 6. Acetate buffer plus tartrate: add 2.3 g sodium tartrate to 100 ml acetate buffer, to give a stock solution of 100 mM tartrate (stable at 4°C for several months). 2.6. Immunostaining for av b3 Integrin (Vitronectin Receptor)

1. Anti vitronectin receptor mAb (clone 23C6; Serotec, Oxford, UK). 2. Fluorescently conjugated secondary antibody (e.g. Alexa Fluor 555 goat anti-mouse IgG; Invitrogen, Oregon, USA). 3. Nuclear stain, such as 0.5 μg/ml 4,6-diamidino-2-phenylindole (DAPI) or Sytox Green (Invitrogen).

2.7. Detection of F-actin Rings

1. 0.5% (v/v) Triton X-100 in PBS.

2.8. Resorption Pit Assay

1. Dentine discs: cut 200–400 μm sections of 1 cm2 blocks of elephant ivory using a Buehler low speed saw with wafering blade. The surface of the slices can be polished by rubbing vigorously with abrasive tissue paper, then the slices can be punched into discs simply using a paper hole punch. Mark the discs with a non-symmetrical identifier in pencil on one side, to enable the disc to be orientated correctly when handling, which is useful when removing from 96-well plates to mount on slides. Ensure that the mark is on the lower surface when plated so that the cells are seeded onto the unmarked surface. The discs should be sterilised and stored in 70% ethanol prior to use.

2. Tetramethylrhodamine isothiocyanate (TRITC)- or fluorescein isothiocyanate (FITC)-phalloidin (Sigma, Poole, UK).

2. 20% (w/v) Sodium hypochlorite solution.

3. Methods 3.1. Isolation and Culture of Rabbit Osteoclasts

1. Euthanise 2- to 4-day-old rabbits under halothane. 2. Remove all four limbs entirely, skin, and keep in PBS on ice. 3. In a glass petri dish, remove all tissue from the femora, tibiae, ulnae, radii with a disposable scalpel. Transfer the bones to PBS as soon as they are dissected. 4. Create a mixed cell suspension in a glass petri dish by mincing all isolated bones from one rabbit in α-MEM (approx. 20 ml) using a scalpel. At this point, osteoclasts can be partially purified

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through a serum density gradient (see Subheading 3.2), in which case it is better to mince the cells in HBSS rather than α-MEM. For the larger bones, it is best to first cut longitudinally, then scrape out the marrow and the inside of the bones before mincing the remaining bone. It is important to perform this part of the procedure as quickly as possible, since the osteoclasts settle and adhere to the dish. 5. Transfer the cell suspension and bone fragments to a 50 ml conical tube and vortex vigorously for three 10 s bursts. Allow the bone fragments to settle for 1 min and then decant the cell suspension to a fresh tube. If enriching the osteoclasts through a serum density gradient see Subheading 3.2 below; otherwise, add α-MEM and supplement with FCS to a final concentration of 10% (v/v) in a final volume of 25 or 50 ml (see Table 1). This suspension should contain approximately 1 × 108 total cells. 6. Plate out the mixed cell suspension into petri dishes (see Notes 1 and 2) or multi-well plates using the guidelines in Table 1. 7. For cultures on dentine discs, allow seeded osteoclasts to adhere for 2 h then gently rinse the discs in PBS to remove the non-adherent cells (see Note 3). At this stage, add any agents further agents (e.g. drugs or cytokines) into fresh α-MEM containing 10% (v/v) FCS. 8. Incubate cultures on plastic overnight in 5% CO2 at 37°C, then remove non-adherent cells by washing gently in PBS using a sterile, wide-bore Pasteur pipette. Three washes are usually sufficient to remove most of the non-osteoclastic cells (see Notes 4 and 5). The remaining adherent cells are mainly osteoclasts, prefusion mononuclear cells, and stromal cells.

Table 1 Recommended density of bone marrow cells for preparing cultures of rabbit osteoclasts (see Note 2) Culture vessel

Total volume of cell suspension (ml)

Volume per well, ml (approx. cell number)

10-cm Petri dish

50

16 (3.2 × 107)

6-Well plate

25

2 (8 × 106)

24-Well plate

25

0.5 (2 × 106)

48-Well plate

25

0.3 (1.2 × 106)

96-Well plate

25

0.125 (0.5 × 106)

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3.2. Enrichment of Osteoclasts Using Foetal Calf Serum Gradients

Osteoclast preparations can be crudely purified prior to seeding by serum density gradient fractionation, using a simplified version of that described by Collin-Osdoby et al. (22). This procedure can improve cultures on resorbing surfaces, by removing the bulk of the contaminating cells that interfere with single cell analysis by fluorescence microscopy, and by increasing the number of resorbing osteoclasts on each disc of dentine. The following volumes are suitable for cells isolated from a maximum of five rabbits. 1. Prepare 30 ml of 70% FCS in HBSS (21 ml FCS plus 9 ml HBSS) and 30 ml 40% FCS in HBSS (12 ml FCS plus 19 ml HBSS) in separate 50-ml tubes. 2. Mix vigorously then transfer 15 ml of 70% FCS/HBSS into a fresh 50-ml tube. 3. Very carefully layer 15 ml of 40% FCS/HBSS on to the 70% FCS/HBSS in each tube, holding the tube at 45° to avoid mixing. 4. Centrifuge isolated bone marrow cells at 100 × g for 5 min at 4°C. 5. Resuspend cells in HBSS, using 15 ml for each gradient to be prepared. 6. Layer the cell suspension extremely carefully on to the FCS gradient, again holding the tube at 45°, then place in a rack and allow cells to settle for 30 min (see Note 4). 7. Remove the upper fraction and discard; this contains primarily red blood cells and mononuclear cells. 8. The middle fraction should contain mainly mononuclear cells plus some multinucleated cells, while the lower fraction should contain a much higher proportion of multinucleated osteoclasts. Therefore, to achieve the highest purity, only the lower fraction should be retained. To achieve the greatest yield, both the middle and lower fractions should be retained. Desired fractions should be transferred to fresh tubes and centrifuge at 100 × g for 5 min. 9. Resuspend pellets in 3 ml supplemented α-MEM and check purity/count in haemocytometer. 10. Seed osteoclasts on to dentine discs at up to 500 per well. Since counting low cell numbers is not particularly reliable, we usually dilute the cell fractions with supplemented α-MEM to approximately 5 ml medium for each rabbit used (up to 10 ml if middle and lower fractions have been combined) then seed into tissue culture plates using the usual volumes (e.g. 100 μl for 96 well plates).

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1. If significant numbers of contaminating cells remain even after rinsing with PBS, incubate the adherent cells for 5–10 min (or until the non-osteoclastic cells are released) in pre-warmed 0.001% (w/v) pronase, 0.002% EDTA in PBS, at 37°C. 2. Wash the plates four times in PBS and culture the remaining purified (typically >95% pure) osteoclasts and prefusion mononuclear cells in α-MEM supplemented with 10% (v/v) FCS. The purified osteoclast cultures are typically about 10–20% confluent (higher if purified through an FCS gradient), and each 10 cm petri-dish usually yields 100–200 μg of cellular protein following lysis (see Note 6).

3.4. Generating Large Numbers of Rabbit Osteoclast-Like Cells in Vitro

This simple culture can be set up using whole marrow cell preparations, the non-adherent cells from isolated osteoclast cultures on plastic dishes, or cells from the middle fraction of FCS gradient separations, and is useful for biochemical studies requiring large numbers of osteoclasts, 1. Seed the cells into 10 cm diameter petri dishes at 3 × 107 cells per dish (or other plates at an equivalent density) in α-MEM supplemented with 10% (v/v) FCS and containing 1 × 10−8 M 1,25(OH)2 vitamin D3. 2. Replace half of the medium (containing 1,25(OH)2 vitamin D3) every 2 days. 3. After 10 days, remove the stromal layer by washing extensively with PBS. This usually yields >95% pure multinucleated, TRAcP-positive osteoclast-like cells (capable of resorbing bone mineral when the marrow cells are cultured on dentine discs); therefore, further purification using pronase–EDTA is not usually required.

3.5. Isolation of Rabbit Osteoclasts Using Immunomagnetic Beads

This approach has proved particularly useful for studying the effects of pharmacological agents on osteoclasts in vivo (14–19), since it enables effective purification of the osteoclasts directly from bone cell preparations. 1. Prepare a mixed cell suspension from the long bones of a neonatal rabbit as described in Subheading 3.1. 2. Centrifuge the mixed cell suspension at 300 × g (10 min) and resuspend the cell pellet in 1.0 ml αMEM containing 3.3 μg/ml 23C6 antibody for 30 min at 37°C. 3. Centrifuge the cells at 300 × g (5 min) then wash in 0.1% (w/v) BSA in PBS and resuspend in 0.1% (w/v) BSA in PBS containing 2 × 107 magnetic Dynal beads conjugated to rat anti-mouse IgG. Incubate at 4°C on a rotating mixer for 20 min. 4. Separate the vitronectin receptor-positive from vitronectin receptor-negative cells by placing in a Dynal magnetic particle

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concentrator for 5 min. Wash the vitronectin receptor-positive cells (osteoclasts) four times in 0.1% (w/v) BSA in PBS, separating the bead-coated cells in the magnet for 1 min after each wash. Finally, place the vitronectin receptor-negative fraction into the magnet to retrieve any lost beads, wash these beads and add to the vitronectin receptor-positive fraction. 5. Count the number of purified osteoclasts using a haemocytometer. This technique typically yields approximately 2 × 104 TRAcP-positive multinucleated cells that are capable of resorption when then cultured on a mineralised substrate (Fig. 3) (see Note 7). 3.6. Characterisation of Rabbit Osteoclasts

1. Rinse cells in PBS then fix in 4% (v/v) formaldehyde for 10 min.

3.6.1. Fluorescence Immunostaining for av b3 Integrin (Vitronectin Receptor)

3. Incubate cells in 2 μg/ml 23C6 monoclonal antibody in PBS/5% FCS for 30 min at room temperature.

2. Incubate cells in 10% (v/v) FCS in PBS for 20 min.

4. Wash four times in PBS containing 0.1% (v/v) FCS. 5. Incubate with fluorescently labelled, secondary antibody according to the manufacturer’s guidelines. We routinely use goat anti-mouse IgG [conjugated to either Alexa Fluor 488 (green emission) or Alexa Fluor 555 (red emission)] from Invitrogen at 1:200 (25 μg/ml) in PBS/5% goat serum, for 30 min. 6. To identify multinucleated cells, nuclei can be fluorescently stained by incubating cells for 10 min with a fluorescent nuclear dye of an appropriate wavelength such as DAPI or Sytox Green. 7. Rinse cells several times with PBS and then visualise using a fluorescence microscope equipped with a ×20 or ×40 objective and appropriate filters (Fig. 1b).

3.6.2. Detection of F-actin Rings in Osteoclasts Cultured on Dentine Discs (see Note 8)

1. Prepare osteoclasts as described in Subheading 3.1 or 3.3, and seed onto dentine discs in 96-well plates. 2. At the end of the culture period (see Note 9), fix the cells on the discs in 4% (v/v) formaldehyde. 3. Rinse cells in PBS then permeabilise with 0.5% (v/v) Triton X-100 in PBS for 15 min. 4. Incubate with 0.5 μg/ml TRITC-phalloidin or FITCphalloidin in PBS for 30 min (see Note 10). 5. Rinse twice in PBS and then store in PBS at 4°C, protected from light. 6. Visualise actin rings using a fluorescence microscope with appropriate filters (Fig. 4b).

3.6.3. Staining for Tartrate Resistant Acid Phosphatase (see Note 11)

1. Osteoclast cultures fixed in 4% (v/v) formaldehyde can be stored at 4°C in PBS for up to 2 weeks before staining for TRAcP.

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2. Prepare staining solution by mixing solutions A and B described below (see note 12). Once prepared, this staining solution should be used the same day. Solution A: 150 μl naphthol-ASBI-phosphate stock, 750 μl veronal buffer (pH 10.1), 0.45 ml acetate buffer, and 1.35 ml acetate buffer/100 mM tartrate. Solution B: 120 μl pararosanilin and 120 μl sodium nitrite (4% w/v). Filter staining solution through a 0.2-μm filter before use. 3. Incubate osteoclasts in filtered staining solution at 37°C for 30–60 min. TRAcP-positive cells metabolise the substrate to a red-coloured product which appears as granular staining throughout the cytoplasm of osteoclasts. The cells should be rinsed in PBS and can then be stored in 70% (v/v) ethanol at 4°C for several weeks. 4. Count the number of osteoclasts (TRAcP-positive, multinucleated cells with >2 nuclei) under bright field illumination at ×20 magnification. Since mononuclear prefusion osteoclasts are also TRAcP positive, it is important to verify the number of nuclei (easily distinguished by negative contrast as unstained areas) within each cell when counting. 3.6.4. Resorption Pit Assay

1. After analysis of cells (e.g. counting actin ring number and osteoclast number as described above), immerse the discs in 20% (w/v) sodium hypochlorite solution, followed by vigorous rubbing with paper towel to remove cells. 2. Visualise resorption pits using a reflected light microscope. Areas of resorption appear dark since the uneven surface of the disc scatters the light, whereas unresorbed, flat surfaces appear bright since they reflect light (Fig. 4c). Alternatively, pits can be visualised using a conventional light microscope after staining the discs with 0.5% (v/v) toluidine blue, or by fluorescence microscopy following staining with a fluorescent bisphosphonate (see Chapter 25 on confocal microscopy, this volume). 3. The area of resorption pits can be quantitated using image analysis software. We use a Leitz Q500MC image analysis system (Leitz, Milton Keynes, UK) with Aphelion-based software developed in-house. The cultures prepared as described above usually result in 0.5–1 mm2 of resorption pit area per disc.

4. Notes 1. The source of petri dishes appears to influence the yield of osteoclasts. In our hands, tissue-culture grade Falcon 10 cm diameter petri dishes produce the best results.

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2. It is difficult to obtain confluent monolayers of purified rabbit osteoclasts, since seeding the cells at higher densities than those indicated in Table 1 prevents attachment of the osteoclasts to tissue culture dishes. 3. When seeding osteoclasts onto dentine discs to assess resorptive activity, it is important to gently wash off the non-adherent cells after seeding, since subsequent resorption appears to be greatly reduced in the presence of the non-adherent cells. The non-adherent cells can be removed as little as 1 h after seeding cells onto dentine discs. This will result in a purer population, but lower yield, of osteoclasts. For this reason, partial purification of the osteoclasts using the FCS gradient is often beneficial for resorption assays. 4. Sedimentation time will affect the yield of osteoclasts: shorter sedimentation times will produce purer osteoclast preparations but with lower overall yield, while longer sedimentation will improve yield at the cost of purity. 5. Extensive washing with PBS is often sufficient to remove contaminating stromal cells, which may lift off as a single layer. In these cases digestion with pronase–EDTA is unnecessary. However, cultures should be only gently rinsed with PBS using a wide-bore, plastic Pasteur pipette, to avoid damaging the osteoclasts. 6. Rabbit osteoclasts have a relatively long life-span when cultured in vitro, even following purification. Although cell number gradually declines, some rabbit osteoclasts remain viable after more than 1 week in culture, without the addition of exogenous growth factors or supplements other than FCS. 7. Rabbit osteoclasts are extremely adherent to tissue culture plastic and difficult to remove enzymatically. Therefore, when preparing osteoclast lysates for western blot analysis, we lyse the cells directly in the petri dish. These lysates can be concentrated if necessary by centrifuging through a microconcentrator (we use 12 kDa molecular weight cut-off). 8. We use immunomagnetic separation specifically to isolate osteoclasts for the preparation of cell lysates (e.g. for western blot analysis) or for single cell analysis following in vivo administration of pharmacologic agents (12–17). Although the vitronectin receptor-positive osteoclasts with magnetic beads attached can be cultured on dentine discs, the resorptive function of these cells is typically 20% of that of osteoclasts that have not been separated using magnetic beads; therefore, this is not recommended for routine in vitro cultures. 9. F-actin-rich podosomes can also be observed around the periphery of osteoclasts cultured on plastic or glass following

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staining with TRITC-phalloidin, but on these surfaces osteoclasts do not polarise and form a genuine “F-actin ring”. 10. Rabbit osteoclasts adhere rapidly to dentine discs but, in our hands, do not begin to resorb until about 12 h after seeding. We routinely incubate cultures of rabbit osteoclasts on dentine discs for 48 h to assess resorptive activity. Only around 25–50% of the TRAcP-positive, multinucleated osteoclasts seeded onto dentine discs are active (i.e. exhibit actin rings) at any one time. 11. Staining for TRITC-phalloidin is useful in combination with staining for the vitronectin receptor, in which case the phalloidin can be added to the secondary antibody solution. We prefer to use TRITC-phalloidin with Alexa Fluor 488-conjugated secondary antibody, when staining in this way. However, it is important to note that staining with phalloidin-conjugates is usually better if the cells have previously been permeabilised. 12. If analysing actin rings as well as the number of TRAcP-positive osteoclasts, staining and analysis of the actin rings must be done first, since this relies on fluorescence microscopy, and will not be able to be seen following histochemical staining for TRAcP. 13. A final tartrate concentration of 50 mM is used when staining for TRAcP in rabbit osteoclast cultures, which is higher than that used for staining for TRAcP in osteoclasts from other species, e.g. mouse (where we use a final tartrate concentration of 30 mM). References 1. Shakespeare, W., Yang, M., Bohacek, R., Cerasoli, F., Stebbins, K., Sundaramoorthi, R., Azimioara, M., Vu, C., Pradeepan, S., Metcalf, C., III, Haraldson, C., Merry, T., Dalgarno, D., Narula, S., Hatada, M., Lu, X., van Schravendijk, M. R., Adams, S., Violette, S., Smith, J., Guan, W., Bartlett, C., Herson, J., Iuliucci, J., Weigele, M., and Sawyer, T. (2000) Structure-based design of an osteoclast-selective, nonpeptide src homology 2 inhibitor with in vivo antiresorptive activity. Proc. Natl. Acad. Sci. USA 97, 9373–9378. 2. Fisher, J. E., Rogers, M. J., Halasy, J. M., Luckman, S. P., Hughes, D. E., Masarachia, P. J., Wesolowski, G., Russell, R. G. G., Rodan, G. A., and Reszka, A. A. (1999) Alendronate mechanism of action: geranylgeraniol, an intermediate in the mevalonate pathway, prevents inhibition of osteoclast formation, bone resorption and kinase activation in vitro. Proc. Natl. Acad. Sci. USA 96, 133–138.

3. Coxon, F. P., Helfrich, M. H., van’t Hof, R. J., Sebti, S. M., Ralston, S. H., Hamilton, A. D., and Rogers, M. J. (2000) Protein geranylgeranylation is required for osteoclast formation, function, and survival: inhibition by bisphosphonates and GGTI-298. J. Bone Miner. Res. 15, 1467–1476. 4. Stenbeck, G., and Horton, M. A. (2004) Endocytic trafficking in actively resorbing osteoclasts. J. Cell Sci. 117, 827–836. 5. Coxon, F. P., Thompson, K., Roelofs, A. J., Ebetino, F. H., and Rogers, M. J. (2008). Visualizing mineral binding and uptake of bisphosphonate by osteoclasts and non-resorbing cells. Bone 42, 848–860. 6. Benford, H. L., McGowan, N. W., Helfrich, M. H., Nuttall, M. E., and Rogers, M. J. (2001) Visualization of bisphosphonateinduced caspase-3 activity in apoptotic osteoclasts in vitro. Bone 28, 465–473. 7. Weidema, A. F., Dixon, S. J., and Sims, S. M. (2001) Activation of P2Y but not P2X(4)

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F.P. Coxon et al. nucleotide receptors causes elevation of [Ca2+]i in mammalian osteoclasts. Am. J. Physiol. Cell Physiol. 280, C1531–C1539. Lees, R. L., Sabharwal, V. K., and Heersche, J. N. (2001) Resorptive state and cell size influence intracellular pH regulation in rabbit osteoclasts cultured on collagen-hydroxyapatite films. Bone 28, 187–194. Chikazu, D., Hakeda, Y., Ogata, N., Nemoto, K., Itabashi, A., Takato, T., Kumegawa, M., Nakamura, K., and Kawaguchi, H. (2000) Fibroblast growth factor (FGF)-2 directly stimulates mature osteoclast function through activation of FGF receptor 1 and p42/p44 MAP kinase. J. Biol. Chem. 275, 31444–31450. Tezuka, K., Sato, T., Kamioka, H., Nijweide, P. J., Tanaka, K., Matsuo, T., Ohta, M., Kurihara, N., Hakeda, Y., and Kumegawa, M. (1992) Identification of osteopontin in isolated rabbit osteoclasts. Biochem. Biophys. Res. Commun. 186, 911–917. David, J. P., Neff, L., Chen, Y., Rincon, M., Horne, W. C., and Baron, R. (1998) A new method to isolate large numbers of rabbit osteoclasts and osteoclast-like cells: application to the characterization of serum response element binding proteins during osteoclast differentiation. J. Bone Miner. Res. 13, 1730–1738. Collin-Osdoby, P., Oursler, M. J., Webber, D., and Osdoby, P. (1991) Osteoclast-specific monoclonal antibodies coupled to magnetic beads provide a rapid and efficient method of purifying avian osteoclasts. J. Bone Miner. Res. 6, 1353–1365. Nesbitt, S., Nesbit, A., Helfrich, M., and Horton, M. (1993) Biochemical characterization of human osteoclast integrins. Osteoclasts express alpha v beta 3, alpha 2 beta 1, and alpha v beta 1 integrins. J. Biol. Chem. 268, 16737–16745. Frith, J. C., Mönkkönen, J., Auriola, S., Mönkkönen, H., and Rogers, M. J. (2001) The molecular mechanism of action of the antiresorptive and anti-inflammatory drug clodronate. Evidence for the formation in vivo of a metabolite that inhibits bone resorption and causes osteoclast and macrophage apoptosis. Arthritis Rheum. 44, 2201–2210.

15. Coxon, F. P., Ebetino, F. H., Mules, E. H., Seabra, M. C., McKenna, C. E., and Rogers, M. J. (2005) Phosphonocarboxylate inhibitors of Rab geranylgeranyl transferase disrupt the prenylation and membrane localization of Rab proteins in osteoclasts in vitro and in vivo. Bone 37, 349–358. 16. Roelofs, A. J., Coxon, F. P., Ebetino, F. H., Lundy, M. W., Henneman, Z. J., Nancollas, G. H., Sun, S., Blazewska, K. M., Lynn, F. B., Kashemirov, B. A., Khalid, A. B., McKenna, C. E., and Rogers, M. J. (2010) Fluorescent risedronate analogs reveal bisphosphonate uptake by bone marrow monocytes and localization around osteocytes in vivo. J. Bone Miner. Res. 25, 606–616. 17. Staal, A., Frith, J. C., French, M. H., Swartz, J., Gungor, T., Harrity, T. W., Tamasi, J., Rogers, M. J., and Feyen, J. H. M. (2003) The ability of statins to inhibit bone resorption is directly related to their inhibitory effect on HMG-CoA reductase activity. J. Bone Miner. Res. 18, 88–96. 18. Hughes, A., Idris, A., Rogers, M. J., and Crockett, J. C. (2007) Rosuvastatin inhibits osteoclast function in vitro and prevents in vivo bone loss via an anti-resorptive mechanism. Calcif. Tissue Int. 81, 403–413. 19. Räikkönen, J., Crockett, J. C., Rogers, M. J., Mönkkönen, H., Auriola, S., and Mönkkönen, J. (2009) Zoledronic acid induces the formation of a pro-apoptotic ATP analogue and isopentenyl pyrophosphate in osteoclasts in vivo and in MCF-7 cells in vitro. Br. J. Pharmacol. 157, 427–435. 20. Minkin, C. (1982) Bone acid phosphatase: tartrate-resistant acid phosphatase as a marker of osteoclast function. Calcif. Tissue Int. 34, 285–290. 21. Van’t Hof, R. J., Tuinenburg-Bol, R. A., and Nijweide, P. J. (1995) Induction of osteoclast characteristics in cultured avian blood monocytes; modulation by osteoblasts and 1,25(OH)2 vitamin D3. Int. J. Exp. Pathol. 76, 205–214. 22. Collin-Osdoby, P., Yu, X., Zheng, H., and Osdoby, P. (2003) RANKL-mediated osteoclast formation from murine RAW 264.7 cells. Methods Mol. Med. 80, 153–166.

Chapter 11 Generation of Human Osteoclasts from Peripheral Blood Kim Henriksen, Morten A. Karsdal, Adam Taylor, Denise Tosh, and Fraser P. Coxon Abstract Osteoclasts are multi-nucleated cells that have the unique ability to resorb calcified bone matrix. They derive from haematopoietic precursor cells, and can be generated in vitro by stimulation of peripheral blood mononuclear cells with the cytokines M-CSF and RANKL. In this chapter, we describe the method for generating human osteoclast from peripheral blood or buffy coats, as well as methods for studying both the differentiation and resorbing activity of these cells. Key words: Osteoclast, Resorption, Cytoskeleton, Monocytes, Assays

1. Introduction Osteoclasts are large, multi-nucleated cells, which arise from haematopoietic stem cells found in the bone marrow, spleen, and peripheral blood (1–6). The formation of resorbing osteoclasts is a complex multi-step process involving the commitment of cells from the monocyte lineage to differentiate into osteoclast precursors, the fusion of these cells to form mature multi-nuclear osteoclasts, and finally the activation of the osteoclasts to resorb bone (3, 5, 7–9). Osteoclastogenesis is dependent on a series of molecules with four major functions: (1) the survival and proliferation of the cells (M-CSF, c-fms, c-fos, PU.1); (2) differentiation into osteoclasts (NFATc1, NFkB, DAP12, FcRg, RANK, RANKL); (3) fusion into multi-nuclear osteoclasts (DC-STAMP, d2 subunit of the V-ATPase); and (4) cytoskeletal reorganization (PYK2, c-src, avb3, MitF) (5, 7, 10–12). The cytokines M-CSF and RANKL are both essential and sufficient for differentiation and activation of osteoclasts (4, 5).


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Osteoclasts are polarized cells that, through the formation of an actin ring, form a sealing zone, where the osteoclast attaches to the bone surface, effectively isolating the resorption lacuna (5). Within the sealing zone, bone resorption occurs through the activity of hydrochloric acid, which is actively secreted into the resorption lacuna via the V-ATPase protein pump and the ClC-7 chloride antiporter, and the cysteine protease cathepsin K, which is secreted from lysosomal vesicles (13, 14). Cultures of human osteoclasts from osteopetrosis patients with mutations in these proteins have helped to indicate their crucial roles in resorption; osteoclasts form normally, but do not resorb effectively due to failure to acidify the extracellular space (V-ATPase or ClC-7 mutations) or to degrade the collagen matrix of bone (cathepsin K mutations) (15, 16). Hallmarks of osteoclasts are the expression of TRAcP, the calcitonin receptor, αvβ3 (also known as the vitronectin receptor; VNR), and MMP-9, which serve as useful tools for identification of human osteoclasts (5). Many studies have used mature human osteoclasts isolated from human osteoclastomas (giant cell tumours of bone) to obtain basic information about osteoclasts (12). Alternatively, other studies used osteoclasts generated by culture of human peripheral blood mononuclear cells (PBMCs) with an osteoblastic cell line, acting as a feeder layer for the osteoclast precursors (17). However, since recombinant RANKL became commercially available, in vitro methods for human osteoclast formation using peripheral blood as the starting material have become widely accessible. To generate human osteoclasts, PBMCs are first isolated by density centrifugation. Optionally, osteoclast precursors can be purified further by isolating CD14+ cells using magnetic sorting (18). PBMCs or CD14+ cells are cultured in the presence of M-CSF to expand the precursor population, and then with both RANKL and M-CSF to generate osteoclasts (Figs. 1 and 2). The resulting mature, multi-nucleated cells have the ability to degrade dentine and cortical bone (18–25), and display all the classical osteoclast markers, such as CTR, TRAcP, Cathepsin, ClC-7, V-ATPase a3 subunit, and MMP-9 (Fig. 3) (20). Finally, and importantly, they respond to known bone resorption inhibitors, such as bafilomycin, bisphosphonates, calcitonin, and the cysteine protease inhibitor E64 (20). Using human osteoclasts generated in cultures with synthetic growth factors is advantageous in many experimental protocols for the following reasons. 1. It enables human osteoclasts to be studied without the necessity for invasive techniques (unlike osteoclastoma-derived cells). 2. Osteoclast precursors can be obtained readily and in high numbers, enabling cultures to be set up on relatively large scales for biochemical analyses.

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Fig. 1. Schematic showing our general protocol for generating osteoclasts from peripheral blood or buffy coats and culturing these cells for analysis of differentiation or resorptive activity.

3. It enables assessment of osteoclastogenesis, osteoclast function, and osteoclast lifespan in a number of clinical conditions, where osteoclast function is affected, such as in osteoporosis, osteopetrosis, pycnodysostosis, Paget’s disease, and multiple myeloma (16, 19, 23, 26, 27). 4. Osteoclasts generated in vitro are extremely robust and longlived, and are amenable to transfection at an early step in their differentiation (24), as described in Chapter 14, this volume.

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Fig. 2. Phase-contrast images showing human osteoclasts generated from cells isolated from buffy coats and cultured with M-CSF and RANKL for the indicated number of days. After 6 days of treatment with RANKL, the culture contains predominantly multi-nucleated osteoclasts.

Fig. 3. Human osteoclasts generated from buffy coats express αvβ3 integrin (left panel, immunostained using 23C6 antibody), form F-actin rings (middle panel, stained using TRITC-phalloidin), and resorb dentine (right panel, reflected light microscopy).

5. Osteoclast cultures are essentially free from other cells types, unlike co-cultures with osteoblasts or osteoclasts isolated from osteoclastomas.

2. Materials All reagents and materials to be used for collecting the blood sample, performing the cell purification, and culturing the cells should be sterile. General materials: 1. Fresh sample of peripheral blood or buffy coat, collected into an anticoagulent vessel (e.g. EDTA or heparin). 2. Phosphate-buffered saline (PBS). 3. Ficoll-Paque (GE Healthcare) or lymphoprep (Axis-Shield) (see Note 2).

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4. 50-ml tubes. 5. Trypsin. 6. Cell culture flasks (T75) and multi-well plates. 7. Cortical bovine bone slices (Nordic Bioscience A/S) or dentine slices. 8. Dynabeads® CD14 (Cat#111-49D, Invitrogen). 9. Magnets compatible with 15-ml tubes (Invitrogen). 2.1. Tissue Culture Reagents

1. Culture medium consisting of alpha minimal essential medium (α-MEM) supplemented with 100 U/ml penicillin–streptomycin and 10% foetal calf serum (FCS). Of importance is the need for testing serum batches, and it is recommended to obtain five different sera from different suppliers and test their ability to support osteoclastogenesis. 2. hM-CSF (R&D Systems , cat#216-MC), use at working concentration of 20 ng/ml. 3. hRANKL (R&D Systems, cat# 390-TN), use at working concentration of 25 ng/ml. 4. mRANKL(R&D Systems, cat#462-TR), use at working concentration of 2 ng/ml. 5. Bone slices: We use cortical bone slices from the femurs of cows/bulls or dentine slices from elephant ivory. The former are prepared from femoral shafts, which are drilled into sticks (diameter fitting into a 96-well plate) along the length of the shaft. These are then cut into slices (0.2-mm thickness) using a slow-speed saw equipped with a diamond blade. The slices are then irradiated in a UV oven for 5 min, and stored in 70% ethanol until further use. Preparation of dentine slices is similar.

2.2. TRAcP Activity Reagents

1. 8.8 mg/ml L-Ascorbic acid (Sigma A0278), in distilled H2O, freshly made. 2. 46 mg/ml Di-sodium tartrate (Merck 106663), in distilled H2O, freshly made. 3. 18 mg/ml 4-Nitrophenylphosphate (Sigma N4645), in distilled H2O, freshly made. 4. Reaction buffer (1 M acetate, 0,5% Triton X-100, 1 M NaCl, 10 mM EDTA, pH 5.5): 59 ml acetic acid 100%, 5 ml Triton X-100, 58 g NaCl, 3.72 g EDTA, 700 ml distilled H2O, adjust pH to 5.5 with 6 M NaOH and add distilled H2O to 1,000 ml. This buffer needs to be cooled down to dissolve completely. 5. STOP buffer (0.3 M NaOH in distilled H2O).

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2.3. Staining

1. Mouse anti-VNR antibody, clone 23C6, Serotec, Oxford, UK. 2. Anti-Mouse IgG-488; Invitrogen. 3. Synergy HT BioTek or similar fluorescence place reader.

3. Methods All steps from obtaining the blood sample to the end point of the culture should be carried out under sterile conditions and using sterile tubes, media, and instrumentation. Be careful when working with human blood and adhere to any local rules pertaining to the use of unscreened blood. Wear gloves and perform the cell isolation and culture procedures in a sterile flow hood. 3.1. Isolation of PBMCs from Peripheral Blood or Buffy Coats

PBMCs for generating human osteoclasts in vitro can be isolated from venous blood samples or from buffy coats, which are usually available from the local blood bank. Ethical approval should be in place for all studies with human tissues. Venous blood samples are a source of fresh cells available on demand and enable the study of osteoclasts from patients with bone diseases (see Note 1). The major advantage of using buffy coats is that a single sample (~50 ml) contains the white cells of 450 ml of whole blood of one donor, and therefore enables isolation of far greater number of osteoclast precursors. The procedure for isolating osteoclast precursors from both samples is very similar (Fig. 1). 1. Dilute the blood sample or buffy coat with an equal volume of PBS. 2. Prepare 50-ml tubes containing 15-ml Ficoll or lymphoprep per tube (see Note 2). 3. Carefully layer 30 ml of the diluted blood/buffy coat on to 15-ml Ficoll/lymphoprep (see Note 3). 4. Centrifuge at 800 × g for 20 min, with brake set to zero. 5. Collect the mononuclear cell layer from each tube and transfer to two new 50-ml tubes. 6. Add PBS to a volume of 45 ml per tube to wash the cells. 7. Centrifuge at 300 × g for 10 min (brake can be used from this stage onwards). 8. Discard the supernatants and resuspend the cell pellet in each tube in 40 ml PBS (second wash) (see Note 4). 9. Centrifuge at 300 × g for 10 min and discard the supernatants from each tube.

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10. The cells can then be further purified using magnetic bead separation (see Subheading 3.2), put straight into culture to expand the M-CSF-dependent macrophages (see Subheading 3.3), or frozen down for future use (see Subheading 3.4). 3.2. Purification of CD14+ Osteoclast Precursor Cells

An optional step following the isolation of PBMCs is to purify CD14+ cells, thereby obtaining a pure osteoclast precursor population. Since other cells capable of affecting osteoclastogenesis (such as stromal cells and lymphocytes) are absent, this type of culture is more suitable for studies investigating direct effects of growth factors, cytokines, and glucocorticoids, for example, on osteoclast formation or function. Figure 4 shows that TRAcP-positive osteoclast-like cells are observed in unpurified PBMC cultures even when cultured without RANKL, suggesting the presence of nonosteoclastic cells that express osteoclastogenic factors. Such osteoclast-like cells are not seen in cultures of CD14+ cells without RANKL. The disadvantages of cell selection are the extra time

Fig. 4. Images of TRAcP-stained cells after 10 days of culture. In the PBMC cultures, several multi-nuclear TRAcP-positive cells are seen even after culture in the absence of RANKL (black arrows) while in the CD14+ sorted preparation, multinuclear TRAcP-positive cells are seen only following culture in the presence of RANKL.

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required and the high cost of the CD14+ beads. In addition, the volume of blood needed is increased, as CD14+ cells constitute only ~10% of the PBMCs. Please note that the beads are autofluorescent, especially towards the red end of the visible spectrum, and this may interfere with fluorescent staining at the end of the culture period. 3.2.1. Preparation of the Magnetic Beads

Please note that the numbers and volumes in this protocol are adjusted for the purification of one buffy coat, which corresponds to 450 ml of whole blood. 1. Shake the vial containing the beads vigorously (without vortexing). 2. Add 5 ml cold PBS into 2× 15-ml tubes. 3. Add 125 μl beads (4 × 108 beads/ml) to each tube and mix gently. 4. Place the tubes in the magnetic device for 1 min. 5. Remove the supernatant while the tube is in the magnetic device. 6. Wash three times with 5 ml/tube cold PBS and discard supernatant.

3.2.2. CD14+ Cell Purification

1. Resuspend the cell pellets (from Subheading 3.1, step 10) in cold PBS containing 2% FCS and combine to a final volume of 10 ml. 2. Add 5 ml cell suspension/tube to the beads (in both tubes) (see Note 5). 3. Incubate the cells/beads suspension at 4°C for 20 min with end-over-end rotation. 4. Place the tubes in the magnetic devices for 2 min. 5. Discard the supernatants while the tube is in the magnetic devices. 6. Wash the cells with 5 ml cold PBS + 2% FCS/tube. Pipette gently up and down to resuspend the beads. 7. Place the tubes in the magnetic devices for 2 min. 8. Discard the supernatants while the tubes are in the magnetic devices. 9. Repeat the washing step five times. Resuspend in 20 ml medium and count the cells (see Note 6).

3.3. Expansion of Macrophages/ Osteoclast Precursors

By expanding the macrophages prior to differentiating the osteoclasts, differences between donors can be largely eliminated as identical numbers of osteoclast precursors can be seeded prior to the addition of RANKL. In addition, expansion of precursors in bulk prior to seeding for osteoclast generation also prevents the

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variability that arises if the osteoclast precursors are expanded directly in the culture plate to be used for analysis. We normally expand precursors in 75-cm2 flasks. 1. Resuspend cells in supplemented α-MEM containing 20 ng/ ml M-CSF (see Note 7). Seed at a density of 250,000 cells/ cm2 if using non-purified PBMCs or seed at 150,000 cells/cm2 if using CD14+ cells. If using PBMCs from venous blood samples, it is not necessary to count the cells; seed the cells isolated from 15 to 20 ml blood in one 75-cm2 flask (typically, 1.5–2 × 107 cells). 2. Culture the proliferating cells, feeding with fresh media containing 20 ng/ml M-CSF every 3 days, until they reach a suitable confluency. For osteoclast generation in situ (in the same flask), culture until cells are 80% confluent and continue the protocol as under Subheading 3.6. For osteoclast generation in other culture vessels or on glass or dentine, precursors can be cultured until 95% confluent. Typically, the expansion period is 6–7 days, but each culture behaves differently (see Note 8). 3. To harvest the precursors, wash twice in PBS, digest with trypsin for 15–30 min, and remove cells with a cell scraper, see Subheading 3.6. Typical yields are 0.5–2 × 106 cells/flask. 3.4. Cryopreservation of Osteoclast Precursors

PBMCs (isolated as described in Subheading 3.1) can be cryopreserved for future use. An alternative and the preferred option is, however, to cryopreserve the expanded macrophages, which can be cultured directly with RANKL upon thawing, significantly shortening the time required to generate osteoclasts (see Note 9). These approaches are particularly useful for valuable samples, such as patient cells, in which all the necessary analyses may not be able to be carried out immediately. 1. The freezing solution consists of 45% α-MEM, 45% FCS, and 10% DMSO. To prepare, add an equal volume of ice-cold 20% DMSO in FCS to cells at twice the required freezing density, suspended in α-MEM containing 10% FCS. 2. Freeze PBMCs at 60–100 × 106 cells per vial to be able to seed two 75-cm2 flasks with one vial. 3. Freeze macrophages at 0.5–2 × 106 cells/vial.

3.5. Generation of Osteoclasts for Assessment of Osteoclastogenesis

1. Adjust concentration of macrophages to 2 × 105/ml in supplemented α-MEM containing 20 ng/ml MCSF plus RANKL (20 ng/ml human RANKL or 2 ng/ml murine RANKL; see Note 10), and seed 100 μl (i.e. 2 × 104 cells) into wells of a 96-well plate. Use replicates of at least four for each separate treatment, and include cells incubated without RANKL (i.e. medium with M-CSF alone) as a control.

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2. The following day, feed the cells with fresh α-MEM containing M-CSF and RANKL as appropriate, plus the experimental treatments (see Note 11). 3. Feed cells as above every 3 days until osteoclasts are formed. This is typically 4–6 days from first addition of RANKL. 4. Remove the medium, rinse off the non-adherent cells with PBS, and then fix the cells in 4% paraformaldehyde in PBS. 5. Osteoclasts can now be stained as necessary. Staining for VNR and nuclear DNA, described in Subheading 3.7.1 below, is ideal for determining osteoclastogenesis. Furthermore, by counting VNR-positive cells with one to two nuclei and those with three or more nuclei separately, effects on differentiation of osteoclast precursors can be distinguished from effects on precursor fusion. 3.6. Generation of Osteoclasts for Functional Studies

While it is possible to differentiate osteoclasts directly on mineralized surfaces, using the conditions described in Subheading 3.5, the precursors often adhere erratically, leading to large variation even between replicas. Variability is greatly reduced by using mature osteoclasts in resorption assays. In addition, this removes further osteoclast formation as a confounder from resorption studies. To perform functional studies with mature osteoclasts, proceed as below. 1. After expansion of macrophages to ~80% confluency (see Subheading 3.3, step 2), consider how many osteoclasts are required and transfer the appropriate amount of expanded macrophages to an appropriately sized culture vessel. You may also proceed with the T75 flask and generate osteoclasts in situ (see Note 11). 2. Feed the cells with fresh medium containing M-CSF and RANKL to drive osteoclastogenesis. 3. Change the medium every 3 days by removing all medium and replacing it with fresh medium containing M-CSF and RANKL. 4. After app. 6–7 days, multi-nucleated osteoclasts have formed in high numbers and they are ready to be reseeded on a resorbable substrate. 5. Wash cells 3× in PBS. 6. Add 4 ml trypsin per 75 cm2 flask (or equivalent if using different size culture vessel) and incubate for 15 min or until most cells become shrunken in appearance with spiky edges. Note that if the osteoclasts are large and well-spread, this process may take up to 1 h. In our experience, this does no lasting damage to the majority of cells and they are able to adhere as normal even after 1 h in trypsin (see Note 12).

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7. Neutralize trypsin by adding α-MEM containing 10% FCS. 8. Detach the remaining adherent cells by scraping very carefully using a cell scraper while keeping the tissue culture vessel flat, and transfer the cells and the medium to a 50-ml tube. 9. Pellet the osteoclasts by centrifuging for 5 min at 200 × g. 10. Resuspend the pellet in α-MEM containing MCSF and RANKL cells. 11. Count osteoclasts and seed on cortical bone, dentine, or other mineralized substrate (see Note 13) at a density of up to 30,000 cells/well of a 96-well plate (see Note 14). 12. Allow osteoclasts to attach for at least 3 h before changing medium and adding treatments. 13. Culture osteoclasts for the desired period. Normally, sufficient resorption has been achieved after 48 h to measure the effect of resorption inhibitors; however, the cultures can be continued for 10–15 days if required, changing the medium and other additives every 3rd day. If quantifying CTX-I or calcium release (see Subheading 3.8), culture supernatants should be collected when changing medium. 14. At the end of the experiment, fix cells in 4% paraformaldehyde in PBS for cellular analysis or alternatively place the slices of bone (or dentine or other mineralized substrate) slices in distilled water to remove the cells for resorption pit analysis (see Subheadings 3.7 and 3.8). 3.7. Analysis of Osteoclast Formation 3.7.1. Assessment of Vitronectin ReceptorPositive Cells

Osteoclast formation on plastic, e.g. in 96-well plates, can be quantified by staining for VNR and a nuclear stain, such as DAPI. Using fluorescent microscopy, the number of multi-nucleated, VNRpositive cells can be manually counted and distinguished from VNR-positive cells containing only one or two nuclei differentiating effects on osteoclast formation from those on osteoclast fusion. Alternatively, the fluorescence intensity in each well can be measured using a fluorescence plate reader to provide an objective measure of osteoclast formation. To do so, proceed from Subheading 3.6, step 13, as follows. 1. Fix cells in 4% paraformaldehyde in PBS for 10 min. 2. Wash cells three times in PBS. 3. Block non-specific antibody binding by incubating in PBS containing 10% FCS for 30 min at room temperature. 4. Incubate with anti-VNR antibody at 1 μg/ml in PBS containing 5% FCS for 1 h at room temperature. Make sure to earmark a couple of wells as “negative control” to assess non-specific staining. These wells are incubated with PBS containing 5% PBS only.

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5. Remove the antibody and wash the cells with three changes of fresh PBS. 6. Incubate the cells with an Alexofluor fluorescent secondary antibody at 1:150 dilution in PBS containing 5% FCS for 1 h at room temperature. 7. Remove the secondary antibody and wash the wells with three changes of fresh PBS. 8. Finally, add 100 μl PBS to each well and count the cells using an inverted fluorescence microscope using an appropriate filterset. 9. Alternatively, read the fluorescence from the bottom of each well (excitation 488 nm; emission 528 nm) using an appropriate plate reader (we currently use a Synergy HT BioTek). To calculate fluorescence intensity per well, deduct the average value obtained from the negative-control wells. 3.7.2. Measurement of TRAcP Activity in the Supernatant

Osteoclast numbers can be assessed by measuring TRAcP activity in supernatants using a colorimetric assay (19). Supernatants from the appropriate matrix, i.e. bone slices incubated without osteoclasts, should be used to provide the background values. Supernatants collected for TRAcP activity measurements can be stored in −20°C until analysis. 1. Use 8 ml of TRAcP solution buffer (see Subheading 2.2) for each 96-well plate. Transfer 20 μl (see Note 15) sample to a 96-well plate in duplicates. 2. Add 80 μl TRAcP solution buffer to each well. 3. Cover the plate with tin foil to protect from light and prevent evaporation. 4. Incubate the plate on a shaker at 3–400 rpm at 37°C for 1 h. 5. Stop the reaction by adding 100 μL 0.3 N NaOH per well. 6. Measure the absorbance at 650 nm while subtracting OD 405 nm as background values. 7. Subtract the background values from each experimental well and plot the mean absorbance of each duplicate sample; the higher the absorbance value, the greater the activity of TRAcP and therefore the greater the number of osteoclasts (18).

3.7.3. Alternative Measurements of Osteoclast Formation

To assess overall human osteoclast formation, western blotting of lysates can be used to analyze the expression of osteoclast-specific proteins, such as cathepsin K, TRAcP, and MMP9, all proteins for which useful reagents are commercially available. In addition, qPCR can be used to analyze osteoclast-specific gene expression at the mRNA level.

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3.8.1. Microscopic Analysis of Resorption Pits


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The most common ways to assess osteoclast activity are to measure the area of the resorbed pits on the dentine or to measure the amount of calcium or collagen fragments released into the medium. To allow for possible differences in the number of osteoclasts, it is recommended to express as a function of osteoclast number. We routinely assess osteoclast number by immunostaining for VNR (see Subheading 3.7.1 above) and counting the number of polarized, active cells according to the presence of an F-actin ring after staining with fluorescent phalloidin conjugates. Expressing resorptive activity as a function of osteoclasts containing F-actin rings can help to elucidate mechanistic effects. For example, normal numbers of osteoclasts expressing actin rings, together with decreased resorption, indicate a defect in the effector pathways of resorption, such as vesicular trafficking to the ruffled border. This is seen in osteoclasts from osteopetrosis patients carrying mutations in ClC-7 or Plekhm1 (19, 23, 28) or in osteoclasts treated with phosphonocarboxylate inhibitors of Rab prenylation. 1. Remove the cells from the bone slices using a cotton swab. 2. Wash away cell debris in MilliQ. 3. Incubate the bone slices for 8 min in Mayer’s hematoxylin. 4. Wash the bone slices in MilliQ water. 5. The resorbed area (see Note 16) can be measured under conventional light microscopy using CAST-Grid (Olympus). Alternatively, resorption pit area can be measured using reflected light microscopy, without the need for the staining with hematoxylin (see, for example, Chapter 10, this volume, on rabbit osteoclasts).

3.8.2. Measurement of Calcium or Collagen Fragments in the Supernatant

The concentration of calcium or degraded collagen fragments in the supernatant are other quantitative measures of resorptive activity, which have the potential advantage that they take account of the overall volume of resorption, rather than just the area of the resorption pits. Calcium can be measured in culture supernatants by colorimetric assay using a Hitachi 912 Automatic Analyzer (Roche Diagnostics), using supernatants from bone slices incubated without osteoclasts as background values (29). The release of the c-terminal type I collagen fragments (CTX-I) from bone or dentine slices can be determined using the CrossLaps for Culture kit (IDS Nordic), according to the manufacturer’s instructions. Once again, supernatants from slices incubated without osteoclasts should be used to provide background values.

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4. Notes 1. PBMCs can still be isolated successfully 48 or 72 h after blood collection, provided the samples are stored at 4°C. This allows cultures to be set up for diagnostic assessment from fresh patient samples. If no reliable courier can be found to guarantee shipment within this time, it is safer to isolate PBMCs at the site of sample collection, freeze the cells as per the protocol in Subheading 3.4), and send the cells on dry ice. 2. Ficoll and lymphoprep are equally effective for isolating PBMCs (30). 3. Layering 25-ml diluted buffy coat sample onto 25-ml Ficoll/ lymphoprep (i.e. 1:1 ratio) is also effective. 4. If the pellets contain a lot of red blood cells, they can be resuspended and incubated in Pharm Lyse (BD Biosciences) for 15 min to eliminate this. Centrifuge at 200 × g for 10 min after the incubation and continue with the PBS wash step, but reduce centrifuge steps to 200 × g. This is particularly useful for patient blood samples that may have been in transit for 24–48 h. 5. Titration of Dynabeads® numbers is very important, and this needs to be done for individual types of blood preparations. 6. We always work with a fixed protocol for a fixed blood volume. The yield is usually around 6 × 108 cells from one buffy coat, of which around 6 × 107 are CD14+ cells. 7. M-CSF concentrations used vary widely between different laboratories, and we recommend a titration step if setting up the procedure for continuous use. Otherwise, 20 ng/ml is the recommended concentration. 8. Osteoclast precursors tend to stop proliferating after around 8 days in culture with M-CSF; therefore, there is little benefit to culturing precursors beyond 8 days. Furthermore, the osteoclastogenic potential also appears to be lost with prolonged expansion. 9. We usually allow the thawed macrophages to recover overnight in medium containing M-CSF, before stimulating osteoclastogenesis with RANKL. 10. Although the M-CSF used must be of human origin, either human or mouse RANKL can be used. Mouse RANKL is the more potent of the two, effective at a concentration of 2 ng/ ml. We recommend performing a titration experiment to establish the optimal concentration to give the best osteoclast yield. 11. Osteoclasts can be formed in different sizes of culture vessel. Small-scale cultures may be appropriate, where only relatively

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small numbers of cells are required, for example if pretreating with compounds or transducing with a virus prior to analysis of resorptive activity. 12. During trypsinisation and scraping, many of the larger osteoclasts do not survive. It is, therefore, important to transfer osteoclasts before the culture has progressed to the point at which all the osteoclasts are too large; in our experience, they should be no bigger than seven to eight nuclei. 13. Cortical bovine bone slices and dentine slices are the best-characterized substrate for human osteoclasts. 14. Thirty thousand cells is a very high cell density, but is often required if calcium release is used as a measure of resorption. For analysis of pit area or CTX-I evaluation, the number of cells can be reduced. We routinely seed cells derived from one well of a 6-well plate onto 20 dentine slices. 15. The dilution of the supernatant needed for the measurement of TRAcP cannot be pre-specified as it not only depends on the number of osteoclasts, but also on the sensitivity of the plate reader. Therefore, it is recommended to start out with 20 μL, and then test lower volumes if the signal is saturated. 16. Resorption pits generated by human osteoclasts are generally resorption trails as a result of the high motility of the osteoclasts. It is, therefore, difficult to score the number of individual pits. The preferred method is, therefore, to measure the resorption area. References 1. Roodman, G. D. (1999) Cell biology of the osteoclast. Exp. Hematol. 27, 1229–1241. 2. Baron, R., Neff, L., Louvard, D., and Courtoy, P. J. (1985) Cell-mediated extracellular acidification and bone resorption: evidence for a low pH in resorbing lacunae and localization of a 100-kD lysosomal membrane protein at the osteoclast ruffled border. J. Cell Biol. 101, 2210–2222. 3. Osdoby, P., Martini, M. C., and Caplan, A. I. (1982) Isolated osteoclasts and their presumed progenitor cells, the monocyte, in culture. J. Exp. Zool. 224, 331–344. 4. Boyle, W. J., Simonet, W. S., and Lacey, D. L. (2003) Osteoclast differentiation and activation. Nature 423, 337–342. 5. Segovia-Silvestre, T., Neutzsky-Wulff, A. V., Sorensen, M. G., Christiansen, C., Bollerslev, J., Karsdal, M. A., and Henriksen, K. (2009) Advances in osteoclast biology resulting from the study of osteopetrotic mutations. Hum. Genet. 124, 561–577.

6. Fujikawa, Y., Quinn, J. M., Sabokbar, A., McGee, J. O., and Athanasou, N. A. (1996) The human osteoclast precursor circulates in the monocyte fraction. Endocrinology 137, 4058–4060. 7. Vaananen, H. K., and Horton, M. (1995) The osteoclast clear zone is a specialized cell-extracellular matrix adhesion structure. J. Cell Sci. 108(Pt 8), 2729–2732. 8. Teitelbaum, S. L. (2000) Bone resorption by osteoclasts. Science 289, 1504–1508. 9. Baron, R. (2003) Anatomy and biology of bone matrix and cellular elements, in Primer on the Metabolic Bone Diseases and Disorders of Mineral Metabolism, American Society for Bone and Mineral Research, Washington, pp. 1–8. 10. Lacey, D. L., Timms, E., Tan, H. L., Kelley, M. J., Dunstan, C. R., Burgess, T., Elliott, R., Colombero, A., Elliott, G., Scully, S., Hsu, H., Sullivan, J., Hawkins, N., Davy, E., Capparelli, C., Eli, A., Qian, Y. X., Kaufman, S., Sarosi, I.,

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K. Henriksen et al. Shalhoub, V., Senaldi, G., Guo, J., Delaney, J., and Boyle, W. J. (1998) Osteoprotegerin ligand is a cytokine that regulates osteoclast differentiation and activation. Cell 93, 165–176. Yasuda, H., Shima, N., Nakagawa, N., Mochizuki, S. I., Yano, K., Fujise, N., Sato, Y., Goto, M., Yamaguchi, K., Kuriyama, M., Kanno, T., Murakami, A., Tsuda, E., Morinaga, T., and Higashio, K. (1998) Identity of osteoclastogenesis inhibitory factor (OCIF) and osteoprotegerin (OPG): a mechanism by which OPG/OCIF inhibits osteoclastogenesis in vitro. Endocrinology 139, 1329–1337. Horton, M. A., Rimmer, E. F., Lewis, D., Pringle, J. A., Fuller, K., and Chambers, T. J. (1984) Cell surface characterization of the human osteoclast: phenotypic relationship to other bone marrow-derived cell types. J. Pathol. 144, 281–294. Coxon, F. P. and Taylor, A. (2008) Vesicular trafficking in osteoclasts. Semin. Cell Dev. Biol. 19, 424–433. Blair, H. C., Teitelbaum, S. L., Ghiselli, R., and Gluck, S. (1989) Osteoclastic bone resorption by a polarized vacuolar proton pump. Science 245, 855–857. Helfrich, M., Crockett, J. C., Hocking, L. J., and Coxon, F. P. (2007) The Pathogenesis of osteoclast diseases: some knowns, but still many unknowns. BoneKey-Osteovision 4, 61–77. Chavassieux, P., Karsdal, M. A., SegoviaSilvestre, T., Neutzsky-Wulff, A. V., Chapurlat, R., Boivin, G., and Delmas, P. D. (2008) Mechanisms of the anabolic effects of teriparatide on bone: insight from the treatment of a patient with pycnodysostosis. J. Bone Miner. Res. 23, 1076–1083. Fujikawa, Y., Sabokbar, A., Neale, S., and Athanasou, N. A. (1996) Human osteoclast formation and bone resorption by monocytes and synovial macrophages in rheumatoid arthritis. Ann. Rheum. Dis. 55, 816–822. Karsdal, M. A., Hjorth, P., Henriksen, K., Kirkegaard, T., Nielsen, K. L., Lou, H., Delaisse, J. M., and Foged, N. T. (2003) Transforming growth factor-beta controls human osteoclastogenesis through the p38 MAPK and regulation of RANK expression. J. Biol. Chem. 278, 44975–44987. Henriksen, K., Gram, J., Schaller, S., Dahl, B. H., Dziegiel, M. H., Bollerslev, J., and Karsdal, M. A. (2004) Characterization of osteoclasts from patients harboring a G215R mutation in ClC-7 causing autosomal dominant osteopetrosis type II. Am. J. Pathol. 164, 1537–1545.

20. Sorensen, M. G., Henriksen, K., Schaller, S., Henriksen, D. B., Nielsen, F. C., Dziegiel, M. H., and Karsdal, M. A. (2007) Characterization of osteoclasts derived from CD14+ monocytes isolated from peripheral blood. J. Bone Miner. Metab. 25, 36–45. 21. Henriksen, K., Sorensen, M. G., Nielsen, R. H., Gram, J., Schaller, S., Dziegiel, M. H., Everts, V., Bollerslev, J., and Karsdal, M. A. (2006) Degradation of the organic phase of bone by osteoclasts: a secondary role for lysosomal acidification. J. Bone Miner. Res. 21, 58–66. 22. Sorensen, M. G., Henriksen, K., NeutzskyWulff, A. V., Dziegiel, M. H., and Karsdal, M. A. (2007) Diphyllin, a novel and naturally potent V-ATPase inhibitor, abrogates acidification of the osteoclastic resorption lacunae and bone resorption. J. Bone Miner. Res. 22, 1640–1648. 23. Van Wesenbeeck, L., Odgren, P. R., Coxon, F. P., Frattini, A., Moens, P., Perdu, B., MacKay, C. A., Van Hul, E., Timmermans, J. P., Vanhoenacker, F., Jacobs, R., Peruzzi, B., Teti, A., Helfrich, M. H., Rogers, M. J., Villa, A., and Van Hul, W. (2007) Involvement of PLEKHM1 in osteoclastic vesicular transport and osteopetrosis in incisors absent rats and humans. J. Clin. Invest. 117, 919–930. 24. Taylor, A., Rogers, M. J., Tosh, D., and Coxon, F. P. (2007) A novel method for efficient generation of transfected human osteoclasts. Calcif. Tissue Int. 80, 132–136. 25. Coxon, F. P., Helfrich, M. H., Larijani, B., Muzylak, M., Dunford, J. E., Marshall, D., McKinnon, A. D., Nesbitt, S. A., Horton, M. A., Seabra, M. C., Ebetino, F. H., and Rogers, M. J. (2001) Identification of a novel phosphonocarboxylate inhibitor of Rab geranylgeranyl transferase that specifically prevents Rab prenylation in osteoclasts and macrophages. J. Biol. Chem. 276, 48213–48222. 26. Henriksen, K., Gram, J., Hoegh-Andersen, P., Jemtland, R., Ueland, T., Dziegiel, M. H., Schaller, S., Bollerslev, J., and Karsdal, M. A. (2005) Osteoclasts from patients with Autosomal Dominant Osteopetrosis type I (ADOI) caused by a T253I mutation in LRP5 are normal in vitro, but have decreased resorption capacity in vivo. Am. J. Pathol. 167, 1341–1348. 27. Del Fattore, A., Peruzzi, B., Rucci, N., Recchia, I., Cappariello, A., Longo, M., Fortunati, D., Ballanti, P., Iacobini, M., Luciani, M., Devito, R., Pinto, R., Caniglia, M., Lanino, E., Messina, C., Cesaro, S., Letizia, C., Bianchini, G., Fryssira, H., Grabowski, P., Shaw, N., Bishop, N., Hughes, D., Kapur, R. P., Datta, H. K.,

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Taranta, A., Fornari, R., Migliaccio, S., and Teti, A. (2006) Clinical, genetic, and cellular analysis of 49 osteopetrotic patients: implications for diagnosis and treatment. J. Med. Genet. 43, 315–325. 28. Karsdal, M. A., Henriksen, K., Sorensen, M. G., Gram, J., Schaller, S., Dziegiel, M. H., Heegaard, A. M., Christophersen, P., Martin, T. J., Christiansen, C., and Bollerslev, J. (2005) Acidification of the osteoclastic resorption compartment provides insight into the coupling of bone formation to bone resorption. Am. J. Pathol. 166, 467–476.

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Chapter 12 Osteoclast Formation in Mouse Co-cultures Cecile Itzstein and Robert J. van ’t Hof Abstract The murine co-culture assay is used to generate mature osteoclasts from bone marrow precursors by culturing them with osteoblasts that are stimulated with 1,25-dihydroxy vitamin D3 and prostaglandin E2. This assay is used particularly to analyse osteoblast–osteoclast interactions and to determine the cell type affected in knock-out or transgenic mice. This chapter describes also the isolation of bone marrow cells from mice and the methods to purify and replate mature osteoclasts. Key words: Co-culture, Osteoclasts, Osteoblasts

1. Introduction The murine co-culture assay originally described by Takahashi et al. (1), was the first culture system developed that generated genuine, bone-resorbing osteoclasts. In this assay, osteoblasts are stimulated with 1,25-dihydroxy vitamin D3 (1,25-(OH)2D3) and prostaglandin E2 (PGE2) to promote RANKL and M-CSF expression. These factors then stimulate early osteoclast precursors present in the spleen or bone marrow cell populations to differentiate into mature osteoclasts. At the end of the culture, osteoclasts can be identified by TRAcP staining, and, when the cultures are performed on dentine slices, resorption activity can be measured as well. Even though nowadays it is possible to generate osteoclasts from bone marrow cells alone by treating the cultures with RANKL and M-CSF, the co-culture system is still a useful model for studying osteoblast–osteoclast interactions and to determine the cell type affected in knock-out or transgenic mice. It has been widely used to study the origin of the osteoclast (2) and the effects of growth factors and drugs on osteoclast formation (3, 4). In studies with osteopetrotic mice, the co-culture assay has been used to Miep H. Helfrich and Stuart H. Ralston (eds.), Bone Research Protocols, Methods in Molecular Biology, vol. 816, DOI 10.1007/978-1-61779-415-5_12, © Springer Science+Business Media, LLC 2012

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determine whether the underlying mechanism was due to a defect in the osteoblasts or a defect in the osteoclast precursors (5). We also describe here the methods for detaching and replating mature osteoclasts using collagen gel-coated plates and for purifying osteoclasts formed in co-culture.

2. Materials 2.1. General Reagents/ Materials

1. Sterile instruments (scissors, forceps, scalpels). 2. Sterile syringes and needles (19 and 25 G). 3. Ficoll or Lymphoprep (Axis-Shield). 4. Sterile Petri dishes. 5. Conical polypropylene centrifuge tubes. 6. Falcon tissue culture plates (see Note 1).

2.2. Tissue Culture Reagents

1. Culture medium: α-MEM supplemented with 10% FCS and antibiotics. 2. Hank’s balanced salt solution (HBSS). 3. HBSS supplemented with 10% FCS. 4. Collagen gel (cellmatrix type 1A) from Nitta Gelatin Inc. (Japan) (available from Wako Chemicals in Europe and USA). To the 100-ml bottle of collagen gel, add 20 ml of 1 mM HCl pH 3.0 and store the bottle at 4°C. 5. 5× concentrarted α-MEM. 6. 200 mM hydroxyethylpiperazine-N¢-2-ethanesulfonic acid (HEPES) buffer, pH 7.4 containing 2.2% NaHCO3. 7. 1,000× concentrated stock of 1,25(OH)2D3 (10−5 M in ethanol, Sigma), further referred to as D3. 8. 1,000× concentrated stock of Prostaglandin E2 (10−3 M in ethanol, Sigma), further referred to as PGE2. 9. Collagenase solution: α-MEM containing 0.1% collagenase type IA (Sigma) and sterilised using a 0.22-μm filter. Make fresh before use. 10. Pronase-EDTA solution: PBS containing 0.001% pronase (Sigma) and 0.02% EDTA (Sigma). Make fresh before use. 11. Dentine slices: we use elephant ivory, cut into slices of approximately 200 μm thickness using a Buehler Isomet low-speed saw with a diamond wafering blade (series 15 HC). Out of these slices we punch discs that fit the wells of a 96-well plate, using a paper puncher (see Notes 2 and 3).

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1. Naphtol-AS-BI-phosphate stock: 10 mg/ml Naphtol-AS-BIphosphate in dimethylformamide (Sigma). Stable ±1 week at 4°C. 2. Veronal buffer: 1.17 g sodium acetate anhydrous, 2.94 g veronal (sodium barbiturate). Dissolve in 100 ml distilled water. 3. Acetate buffer 0.1 N, pH 5.2: (a) Dissolve 0.82 g sodium acetate anhydrous in 100 ml distilled water. (b) 0.6 ml Glacial acetic acid, make up to 100 ml with distilled water. (c) Adjust the pH of solution (a) to pH 5.2 with solution (b). 4. 100 mM Tartrate: Dissolve 2.3 g of sodium tartrate in 100 ml of acetate buffer. 5. Pararosanilin, acridinfrei: add 1 g Pararosanilin to 20 ml distilled water and add 5 ml concentrated hydrochloric acid, heat carefully for 15 min in a 95°C water bath while stirring, and filter once the solution has cooled down. Solutions 2–5 are stable for months if kept protected from light in a refrigerator.

3. Methods 3.1. Osteoblasts

The assay starts with the isolation of the cell populations needed. Although some groups have reported good results with osteoblastlike cell lines, such as ST2 cells (6), we have not been very successful with these and use primary osteoblasts isolated from the calvaria of 2–3 day old neonatal mice (see Chapter 2, this volume). The osteoblasts are plated on plastic, collagen gel or dentine 1 day before the addition of the bone marrow cells.

3.2. Isolation of Bone Marrow Cells

Although the assay was originally described using spleen cells ((1), see Note 4), we generally use bone marrow cells as the source of osteoclast progenitors. Furthermore, others have successfully used certain haemopoietic stem cell lines, such as C2GM cells (7). 1. Dissect the femurs and tibia out of a mouse (3–8 weeks old). 2. Flush out the marrow using a 25 G needle and HBSS + 10% FBS. 3. The cells are collected by centrifugation at 300 × g for 3 min and resuspended in 1 ml of culture medium. Alternatively: ●

Get a single cell suspension by squeezing the cell suspension through needles of decreasing size (start with 19 G, end with 25 G).

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Remove red blood cells by Ficoll density centrifugation (600 × g, 25 min, brake off).

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Harvest the bone marrow cells from interface and wash once in HBSS.

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Resuspend in 1 ml of culture medium.

4. Keep the cells on ice until use (but try and get the cells in culture as soon as possible). To perform mix-match experiments using osteoblasts and bone marrow cells from mice of different genotypes, it is necessary to isolate the haematopoietic precursors from the total population of bone marrow cells. It is possible to use Ficoll gradients or to simply culture the cells overnight in culture medium allowing the mesenchymal stromal cells to adhere to the tissue culture plate. Harvest the non-adherent bone marrow cells by centrifuging the medium the next day and proceed as described in Subheading 3.3 using similar cell numbers. 3.3. Setting Up the Co-culture

The optimal number of bone marrow cells and osteoblasts per well may vary and is dependent on the mouse strain used. For the C57Bl/6 mouse strain, we use routinely the following seeding densities give optimal numbers of osteoclasts. In a 6-well plate: 1. On day 0, plate 1 × 105 osteoblasts in 2 ml medium per well. 2. On day 1, remove the medium and add 2 × 106 freshly isolated bone marrow cells or non-adherent bone marrow cells in 2 ml medium/well. The medium should contain 10 nM D3 + 1 μM PGE2. 3. On day 3, gently remove 1.5–1.7 ml medium from each well and add 2 ml of fresh medium containing 10 nM D3 + 1 μM PGE2 (see Note 5). 4. Proceed with the co-culture for up to 4–6 more days by changing the media every other day or until multinucleated osteoclasts are observed. The medium changes need to be done very carefully because the confluent layer of osteoblasts can be quite easily disturbed, and come off (see Note 5). This would result in a total absence of osteoclasts. Usually, the first osteoclasts appear on day 4 or 5 (see Note 6). Reasonable numbers of osteoclasts (see Note 7) are present between day 7 and 9.

3.4. Collagen Gel Culture

Detaching mouse osteoclasts formed on plastic is very difficult even when using trypsin-EDTA or collagenase treatment. Therefore performing the co-culture assay on collagen gel-coated plates allows us to easily obtain mature functioning mouse osteoclasts (8) that can then be replated on various substrates such as dentine, plastic culture plates, or glass coverslips.

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1. To coat a 6-well plate, prepare the collagen gel-coating solution in a 50-ml tube by mixing: ●

7 parts of collagen gel.

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2 parts of 5× concentrated αMEM.

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1 part of 200 mM Hepes pH 7.4, 2.2% NaHCO3 buffer.

2. Mix without creating air bubbles by inverting gently the tube several times. 3. Add 1.5 ml of this solution to each well. 4. Incubate 30 min at 37°C until polymerisation. 5. Plate osteoblasts and bone marrow cells as described in Subheading 3.3. 6. Once osteoclasts are formed, wash the cells with α-MEM without serum. 7. Add 1 ml of 0.1% collagenase solution per well. 8. Incubate 15–20 min at 37°C with gentle shaking (30 cycles/ min in a shaking incubator or give a gentle swirl every 5 min). 9. The cells are collected by centrifugation at 300 × g for 3 min and resuspended in 5–8 ml of culture medium. 10. Plate 100 μl onto dentine slices to quantify the bone resorbing activity or on 96-well plate to evaluate osteoclast numbers. 3.5. Purification of Osteoclasts Obtained by Co-culture

In a co-culture, osteoclasts represent only 2–5% of the cells and further purification is essential for biochemical studies (9, 10). Osteoblasts and stromal cells can easily be removed either by peeling away the osteoblast monolayer or by treatment with pronaseEDTA solution. ●

If osteoblasts and stromal cells are forming a monolayer, rinse with PBS and gently squirt PBS on the wall of the well using a plastic Pasteur pipette to detach a corner of the layer. Once detached, rinse the cells with PBS.

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If the cells are not confluent, rinse once with PBS and add 1 ml/well of pronase-EDTA solution. Incubate 5–10 min at 37°C and gently wash three times with PBS.

Using these techniques, the purity of osteoclasts is increased to more than 80% (9, 10). Purified osteoclasts need to be used rapidly because no growth factors, cytokines, or osteoblasts are present to maintain them alive. 3.6. Tartrate Resistant Acid Phosphatase Staining

Osteoclasts express very high levels of the enzyme tartrate resistant acid phosphatase (TRAcP) and can therefore be easily visualised by staining for this enzyme as follows ((11), see Notes 8–11). As an alternative to the protocol described here, a staining kit from Sigma (387-A, Leukocyte Acid Phosphatase staining kit) can be used.

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This kit uses fast Garnet as the dye, and this leads to a very dark purple stain (see Chapter 8, this volume). 1. Rinse the culture with PBS. 2. Fix the cells for 5 min with 4% formaldehyde. 3. Rinse with PBS. 4. Prepare staining solution: Solution 1 In a glass container, add 150 μl Naphtol-AS-BI-phosphate stock to 750 μl Veronal buffer (pH 10.1). Then, add 0.9 ml Acetate buffer. Add 0.9 ml Acetate buffer with 100 mM Tartrate. Solution 2 120 μl Pararosanilin. 120 μl 4% NaNO2. Mix solutions 1 and 2, filter through a 0.45-μm filter and use immediately. 5. Incubate the cells for 30–60 min at 37°C with staining solution (50–100 μl/well of a 96-well plate). 6. Rinse with distilled water. 7. Store in 70% ethanol. Osteoclasts and mononuclear osteoclast precursors should be visible as bright red stained cells (see Fig. 1a). 3.7. Quantification of the Resorption Area

After the osteoclasts have been stained and counted, the slices are cleaned and the resorption pits can be visualised either by staining with dyes such as Toluidine blue or Coomassie blue, by scanning electron microscopy or by reflected light microscopy. We routinely use reflected light microscopy because it is easy to perform, the slices only need thorough cleaning and no staining, and the image obtained can be fairly easily quantified using image analysis. We use a Zeiss Axiolab reflected light microscope, fitted with a 2.5× lens, wide field c-mount adapter, and Diagnostics Instruments Insight B/W large chip digital camera. This set-up allows us to capture an entire bone slice in one image at sufficient resolution to identify and measure the resorption pits (see Fig. 1b). We developed our own image analysis software package using the Aphelion ActiveX image analysis toolkit from ADCIS (ADCIS SA, Hérouville-SaintClair, France). The program prompts the user to focus the slice to be measured (see Fig. 2a), captures the image, and identifies the dentine slice (see Fig. 2b) so that any dark objects outside the dentine slice can be automatically removed. Next, the resorption pits are identified using grey level thresholding and selection by shape (see Fig. 2c), and the resorption area is calculated. The entire process takes 2–3 min per slice. A non-automated system for quantification that works well is described in Chapter 8, this volume.

Fig. 1. End result of a co-culture. (a) Multinucleated osteoclast identified by TRAcP staining. (b) Resorption pits visualised by reflected light microscopy. The resorption pits stand out as dark objects. (c) Actin rings in the osteoclasts, visualised by phalloidin staining.

Fig. 2. Measurement of resorption pits. Clockwise from the top left: An image of the dentine slice is captured using a digital camera; The dentine slice is identified by the software; The resorption pits are detected using grey level thresholding.

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4. Notes 1. Osteoblasts and bone marrow cells appeared to adhere better on Falcon tissue culture plates leading to better osteoclast differentiation. 2. Ivory is the preferred to cortical bone for this assay, since it is very even and does not contain osteocyte lacunae, which interfere with the identification of the resorption pits. 3. As we use reflected light microscopy and automated image capture to visualise the resorption pits at the end of the culture, it is important that the surface of the slices is as smooth and shiny as possible without compromising the cell adhesion to the dentine. Therefore, the slices are polished using a paper towel until the surface of the disc appears even under light reflected microscopy. Any remaining polish particles are removed by sonicating the dentine slices for ±15 min in 70% ethanol. The slices are stored in 70% ethanol until use. 4. Spleen cells can be used as an alternative to bone marrow cells in the co-culture. The advantage is, that they are easier to isolate than bone marrow cells. However, we generally get more consistent results using bone marrow. To use spleen cells, dissect the spleens out of two young adult mice. Use a bent 19 G needle to press the cells out of the spleen. Thoroughly resuspend the cells, load onto Ficoll, and proceed as described in the alternative method in Subheading 3.2, step 3 using similar cell numbers. 5. The most common problem with the co-culture is the osteoblast layer contracting and coming off the slice. This is usually due either to the plating density of the osteoblasts being too high or an uncareful medium refresh. Because of the last point, we do not remove completely the medium when changing it and we leave around 250–300 μl in each well. Make sure that the tip of the pipette does not touch the osteoblast layer. 6. Any drugs or factors to be tested can be present during different parts of the co-culture. To test effects on mature osteoclasts, drugs can be added during the last 2–3 days of the culture, whereas having the drugs present during the first 3–4 days gives an indication of effects on osteoclast precursors. 7. The usual yield of osteoclasts should be between 150 and 300 osteoclasts per slice. If the numbers are substantially lower, this may be due to non-optimal seeding densities. Although the seeding densities mentioned above work well for cells from C57Bl/6 mice, other mouse strains may need different densities. It should be noted that numbers of osteoblasts and bone marrow cells that are either too high or too low will both lead

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to a reduction of osteoclast numbers and a series of densities should be tested. 8. In this murine assay, the most convenient procedure is the TRAcP stain. However, it should be noted that in long term cultures, macrophage polykaryons become TRAcP positive as well (12). These macrophage polykaryons can be distinguished from osteoclasts by their staining for the macrophage antigen F4/80. Furthermore, the TRAcP stain as described here works fine for murine osteoclasts. However, when staining osteoclasts from different species, the concentration of tartrate may need to be changed. For human osteoclasts for example, we use a final concentration of 100 mM tartrate. 9. In some species, osteoclasts can easily be identified by immunohistochemical or immunofluorescent staining for the vitronectin receptor (13). However, reagents for detection of vitronectin receptor in the mouse are not easily available. 10. Osteoclasts can also be identified by the presence of calcitonin receptors (14). However, this procedure is too involved and time consuming for routine analysis as it involves using radiolabeled calcitonin and autoradiography. 11. Osteoclasts that are actively resorbing display an actin ring, and this can be visualised by staining the actin with phalloidin, coupled to rhodamine (Molecular Probes, or Sigma). Comparing total number of TRAcP positive osteoclasts with the number of cells displaying the actin ring gives an indication of the fraction of osteoclasts actively resorbing bone. References 1. Takahashi, N., Akatsu, T., Udagawa, N., Sasaki, T., Yamaguchi, A., Moseley, J. M., Martin, T. J., and Suda, T. (1988) Osteoblastic cells are involved in osteoclast formation. Endocrinology 123, 2600–2602. 2. Hagenaars, C. E., Kawilarang-De Haas, E. W., van der Kraan, A. A., Spooncer, E., Dexter, T. M., and Nijweide, P. J. (1991) Interleukin-3dependent hematopoietic stem cell lines capable of osteoclast formation in vitro. J. Bone. Min. Res. 6, 947–954. 3. van ’t Hof, R. J. and Ralston, S. H. (1997) Cytokine-induced nitric oxide inhibits bone resorption by inducing apoptosis of osteoclast progenitors and suppressing osteoclast activity. J. Bone. Min. Res 12, 1797–1804. 4. van ’t Hof, R. J., Armour, K. J., Smith, L. M., Armour, K. E., Wei, X., Liew, F. Y., and Ralston, S. H. (2000) Requirement of the inducible nitric oxide synthase pathway for IL-1- induced osteoclastic bone resorption. Proc. Natl. Acad. Sci. USA. 97, 7993–7998.

5. Lowe, C., Yoneda, T., Boyce, B. F., Chen, H., Mundy, G. R., and Soriano, P. (1993) Osteopetrosis in Src-deficient mice is due to an autonomous defect of osteoclasts. Proc. Natl. Acad. Sci. USA 90, 4485–4489. 6. Udagawa, N., Takahashi, N., Akatsu, T., Tanaka, H., Sasaki, T., Nishihara, T., Koga, T, Martin, T. J., and Suda, T. (1990) Origin of osteoclasts: mature monocytes and macrophages are capable of differentiating into osteoclasts under a suitable microenvironment prepared by bone marrow-derived stromal cells. Proc. Natl. Acad. Sci. USA 87, 7260–7264. 7. De Grooth, R., Mieremet, R. H., Kawilarang-De Haas, E. W., and Nijweide, P. J. (1994) Murine macrophage precursor cell lines are unable to differentiate into osteoclasts: a possible implication for osteoclast ontogeny. Int. J. Exp. Pathol. 75, 265–275. 8. Akatsu, T., Tamura, T., Takahashi, N., Udagawa, N., Tanaka, S., Sasaki, T., Yamaguchi, A., Nagata, N., and Suda, T. (1992) Preparation

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and characterization of a mouse osteoclast-like multinucleated cell population. J. Bone Miner. Res. 7, 1297–1306. 9. Nakamura, I., Takahashi, N., Sasaki, T., Tanaka, S., Udagawa, N., Murakami, H., Kimura, K., Kabuyama, Y., Kurokawa, T., Suda, T., and Fukui, Y. (1995) Wortmannin, a specific inhibitor of phosphatidylinositol-3 kinase, blocks osteoclastic bone resorption. FEBS Lett. 361, 79–84. 10. Jimi, E., Ikebe, T., Takahashi, N., Hirata, M., Suda, T., and Koga, T. (1996) Interleukin-1 alpha activates an NF-kappaB-like factor in osteoclast-like cells. J. Biol. Chem. 271, 4605–4608. 11. Barka, T. and Anderson, P. J. (1962) Histochemical method for acid phosphatase

using hexazonium pararosanilin as coupler. J. Histochem. Cytochem. 10, 741–753. 12. Modderman, W. E., Tuinenburg-Bol Raap, A. C., and Nijweide, P. J. (1991) Tartrate-resistant acid phosphatase is not an exclusive marker for mouse osteoclasts in cell culture. Bone 12, 81–87. 13. Horton, M. A., Taylor, M. L., Arnett, T. R., and Helfrich, M. H. (1991) Arg-Gly-Asp (RGD) peptides and the anti-vitronectin receptor antibody 23 C6 inhibit dentine resorption and cell spreading by osteoclasts. Exp. Cell Res. 195, 368–375. 14. Nicholson, G. C., Horton, M. A., Sexton, P. M., D’Santos, C. S., Moseley, J. M., Kemp, B. E., Pringle, J. A., and Martin, T. J. (1987) Calcitonin receptors of human osteoclastoma. Horm. Metab Res. 19, 585–589.

Chapter 13 RANKL-Mediated Osteoclast Formation from Murine RAW 264.7 cells Patricia Collin-Osdoby and Philip Osdoby Abstract Extensive research efforts over the years have provided us with great insights into how bone-resorbing osteoclasts (OCs) develop and function and, based on such work, valuable antiresorptive therapies have been developed to help combat the excessive bone loss that occurs in numerous skeletal disorders. The RAW 264.7 murine cell line has proven to be an important tool for in vitro studies of OC formation and function, having particular advantages over the use of OCs generated from primary bone marrow cell populations or directly isolated from murine bones. These include their ready access and availability, simple culture for this pure macrophage/pre-OC population, sensitive and rapid development into highly boneresorptive OCs expressing hallmark OC characteristics following their RANKL stimulation, abundance of RAW cell-derived OCs that can be generated to provide large amounts of study material, relative ease of transfection for genetic and regulatory manipulation, and close correlation in characteristics, gene expression, signaling, and developmental or functional processes between RAW cell-derived OCs and OCs either directly isolated from murine bones or formed in vitro from primary bone marrow precursor cells. Here, we describe methods for the culture and RANKL-mediated differentiation of RAW cells into boneresorptive OCs as well as procedures for their enrichment, characterization, and general use in diverse analytical assays. Key words: Osteoclast, Osteoclast development, Bone resorption, RANKL, Mouse macrophage, RAW 264.7 cells

1. Introduction Osteoclasts (OCs) are cells uniquely responsible for dissolving the organic and inorganic components of bone during bone development and remodeling throughout life. They originate from hematopoietic precursors of the monocyte/macrophage lineage present in the bone marrow and peripheral circulation, and their numbers and/or activity frequently increase in a wide array of


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clinical disorders associated with excessive bone loss (1). For many years, investigations into how OCs develop and function were hampered by considerable difficulties associated with isolating and culturing these normally rare cells. Whereas cell lines have frequently provided an invaluable research tool and are widely used to decipher mechanisms involved in osteoblast (OB) differentiation and bone formation, no immortalized cell lines for mature OCs exist and the few pre-OC cell lines that were reported either did not undergo full OC differentiation (2, 3) or involved coculture systems and cells that were not readily available to all researchers (4–6). To further compound the problem, it was difficult to reliably generate bone-resorptive OCs expressing mature OC characteristics from primary bone marrow or circulating precursor cells in vitro. This all changed with the breakthrough discovery of the key OC differentiation signal, receptor activator of nuclear factor κB ligand (RANKL), that triggers the full development and activation of OCs both in vitro and in vivo (7–9). During OB development or in response to specific hormonal or local signals, RANKL becomes highly expressed on the surface of OB/stromal cells and interacts with a receptor, RANK, upregulated by macrophage colony-stimulating factor (M-CSF) on the surface of pre-OCs to stimulate their fusion, differentiation, and resorptive function. Many researchers now routinely form OCs in vitro through the exogenous addition of soluble recombinant RANKL (in combination with M-CSF to stimulate pre-OC proliferation, survival, and RANK expression) to cultures of primary bone marrow cells or peripheral blood monocytes derived from various species (e.g., human, mouse, rat, rabbit, or chicken, as discussed in other chapters in this volume). However, such procedures still require the isolation of primary precursor populations, and in sufficient numbers, to provide enough in vitro generated OCs for experimentation or characterization. In addition to primary cells, at least one pre-OC cell line, murine macrophage RAW 264.7 cells, responds to RANKL stimulation in vitro to generate bone resorbing multinucleated OCs (RAW-OCs) with the hallmark characteristics expected for fully differentiated OCs (10–12). RAW cells have been extensively employed in macrophage studies for >30 years and were originally established from the ascites of a tumor induced in a male mouse by intraperitoneal injection of Abelson leukemia virus (although RAW cells do not secrete detectable virus particles) (13). RAW cells express the c-fms receptor for M-CSF (14) as well as M-CSF, perhaps explaining why they also express high levels of RANK (10) and do not require M-CSF as a permissive factor in their RANKL-induced formation into RAW-OCs. RAW cells are often used in studies of OC differentiation and function, in parallel or as a prelude to studies with OCs formed from primary cells. There are many advantages of this system over the

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generation of OCs from primary cell populations, including the following: (1) ready access (making it unnecessary to schedule experiments around when primary cells may become available) and widespread availability of this cell line to most researchers, (2) easy culture and homogeneous nature of the pre-OC population (devoid of OBs, stromal, lymphocytes, or other cell types), (3) sensitive and very rapid RANKL-mediated formation of RAWOCs (within days), (4) very large number of RAW-OCs that can be generated (and, thus, RNA or protein for study), (5) high bone pit resorptive capability and expression of OC characteristics exhibited by RAW-OCs, (6) relative ease of transfection for genetic and regulatory manipulation, and (7) close correlation in characteristics, gene expression, signaling, and developmental or functional processes between RAW-OCs, OCs formed from primary precursor cells in vitro, and isolated in vivo formed OCs. In this chapter, we describe methods for the culture and RANKLmediated differentiation of RAW cells into bone-resorptive RAWOCs, the preparation of RAW-OC enriched populations by serum density gradient fractionation, and the culture and characterization of RAW-OCs. Such in vitro generated OCs can be analyzed using biochemical, immunological, physiological, molecular, functional, or other assays according to commonly employed procedures; see also various other chapters on osteoclasts in this volume.

2. Materials 2.1. Tissue Culture Medium, Solutions, and Supplies

All media and solutions are prepared with glass distilled water. 1. Culture medium: mix 90 ml of sterile Dulbecco’s modified Eagle medium (DMEM) supplemented with 4 mM L-glutamine, 1.5 g/l sodium bicarbonate, 4.5 g/l glucose, and 1.0 mM sodium pyruvate with 10 ml of fetal bovine serum (FBS, Invitrogen-Gibco) and 1 ml of a 100× stock of antibiotic/antimycotic (a/a, Invitrogen-Gibco); store at 4°C and prewarm to 37°C for use with cells. 2. Phosphate buffered saline, pH 7.2 (PBS). 3. RANKL (Enzo Life Sciences, PeproTech, EMD4Biosciences, R&D Systems, or homemade): reconstitute and store as a concentrated stock solution (typically 100 μg/ml in PBS) in aliquots (~10–50 μl) at −80°C as recommended by the manufacturer, briefly thaw and dilute into culture medium (to 35 ng/ml final concentration for murine recombinant soluble RANKL) immediately before use with RAW cells, and refreeze remaining RANKL (and aim to thaw individual vials no more than three times to retain optimal bioactivity).

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4. Moscona’s high bicarbonate (MHB): add 8 g of NaCl, 0.2 g of KCl, 50 mg of NaH2PO4, 1.0 g of NaHCO3, 2 g of dextrose, 10 ml of a/a, and 990 ml of water; check pH is 7.2 and sterilefilter. 5. Hanks’ balanced salt solution (HBSS, Invitrogen-Gibco), pH 7.2. 6. Collagenase (Type 3): prepare a 3% stock (3 g in 100 ml) solution in HBSS; store in aliquots (0.5–1.0 ml) at −20°C. 7. Trypsin: 1% stock (1 g in 100 ml) solution in HBSS; store in aliquots (1.0 ml) at −20°C. 8. Collagenase–trypsin digestion solution: briefly thaw and add 71 μl of 3% collagenase solution and 141 μl of 1% trypsin solution to 3 ml of MHB (per dish) immediately before use with cells. 9. Protease (EC 3.4.24.31, Sigma P-8811): 0.1% (100 mg in 100 ml) stock solution in PBS; store at 4°C for up to several months or in aliquots (0.5 ml) at −20°C for long-term storage. 10. EDTA: 2% (2 g in 100 ml) stock solution (using EDTA sodium salt) in PBS; store at 4°C. 11. Protease–EDTA digestion solution: briefly thaw and add 50 μl of 0.1% protease solution and 50 μl of 2% EDTA solution to 5 ml of PBS (per dish) immediately before use with cells. 12. Supplies: sterile bottles, flasks, and tissue culture dishes; rubber cell scrapers (Fisher); hemocytometer. 13. Devitalised bone or dentine slices, prepared as described (see Section 3.1). Ivory is obtained through donation from a local zoo or, in the USA, the Federal Department of Fish and Wildlife Services (or similar Department in other countries). Bovine cortical bone is obtained from a local slaughterhouse.

3. Methods 3.1. Preparation of Devitalized Bone or Dentine Slices

1. Segments of ivory and bovine cortical bone are thoroughly cleaned and washed (multiple HBSS and 70% ethanol rinses), sliced into small chunks and then reduced to rectangular 0.4mm thick sheets using a low-speed Isomet saw (Buehler, Lake Bluff, IL). 2. The sheets are rinsed three times with 70% ethanol, incubated in 70% ethanol overnight, and then washed for several hours in HBSS before circular disks are cut using a 5-mm paper punch.

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3. The disks are soaked repeatedly in 70% ethanol in sterile 50-ml tubes (alcohol changes can be gently poured off because the disks tend to stick to the side of the tube), and stored in 70% ethanol at −20°C. 4. For experimental use, the required number of disks are removed from the tube using alcohol-presoaked tweezers (to maintain sterility) in a tissue culture hood, transferred to a new sterile 50-ml polypropylene tube, rinsed extensively by inversion and mild shaking at least three times with ~40 ml sterile HBSS per wash, and the disks transferred using sterile tweezers into culture wells or dishes containing sterile HBSS for 3–24 h of preincubation in a tissue culture incubator prior to the plating of cells. HBSS is removed only immediately before the disks are used so that they do not dry prior to RAW cell or RAW-OC seeding. 3.2. RAW 264.7 Cell Culture

RAW 264.7 cells are obtained from the ATCC or similar cell repository. They represent a murine macrophage cell line that has the capability to be grown indefinitely as an OC precursor population or can be differentiated by treatment with RANKL into multinucleated bone resorptive OCs expressing the hallmark characteristics of in vivo formed OCs (see Subheading 3.3). All work should be performed in a sterile hood using sterile solutions and supplies. 1. If starting from a frozen (liquid nitrogen) vial of RAW cells, quickly (6 months of age. 3.20. Cortical Bone Histomorphometry

The usual sites for cortical bone histomorphometry in rats and mice are the femoral or tibial midshaft. In mice, we mostly use the femoral diaphysis, since mouse femurs are less fragile and less curved than the tibiae, and it is easier to define the femoral midshaft than the tibial midshaft. We measure structural parameters on 15- to 20-mm-thick microground sections stained with toluidine blue (Fig. 8). We quantify cross-sectional area (Tt.Ar), cortical area (Ct.Ar), marrow area (Ma.Ar), cortical thickness (Ct.Th), and

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number and area of intracortical pores (N.Po and Po.Ar) using Zeiss AxioVision 4.7 image analysis software. We perform image acquisition using a Zeiss Axioskop microscope with a ×2.5 objective for the mouse samples, or a Zeiss SV11 stereomicroscope for the rat samples, together with a Diagnostic Instruments Insight digital camera. We measure cortical thickness directly as a primary parameter, using 90 radii originating from the center of gravity of the bone section (Fig. 8). In addition to the absolute values for Ct.Ar, Ma.Ar, and Po.Ar, we always calculate the relative values for cortical and marrow areas (Ct.Ar/Tt.Ar and Ma.Ar/Tt.Ar, %), and intracortical pore area (Po.Ar/Ct.Ar, %). Relative values are helpful when samples of different size need to be compared. To evaluate the bone resorptive activity at endocortical bone surfaces, the endocortical eroded perimeter (Ec.E.Pm/B.Pm, %) can be measured (11). The percentage of endocortical eroded perimeter is traced with a ×10 objective, using an interactive image analysis system. Similar to cancellous bone, it is useful to use a cutoff value of at least 3 mm to exclude very shallow erosions. We use the Osteomeasure interactive system for this measurement. 3.21. Periosteal and Endocortical Bone Formation

In order to quantify periosteal and endocortical bone formation rates, it is necessary to measure the mineralizing perimeter and the mineral apposition rate at both surfaces. Similar to cancellous bone histomorphometry, we use only the double-labeled perimeter for the calculation of the mineralizing perimeter (M.Pm/B.Pm = Db. Lb.Pm/B.Pm). For most purposes, it is enough to measure one section. We measure the periosteal and endocortical mineralizing perimeter with the Osteomeasure interactive system, using ×10, ×20, or ×40 objectives, depending on the interlabel distance. Rats and mice lack true Haversian remodeling. However, intracortical remodeling can be induced in rats by drugs or microdamage accumulation (12–16). Therefore, in rare cases, the intracortical bone formation rate can be of interest. It is measured in analogy to the endocortical and periosteal envelopes by assessing MAR and M.Pm/B.Pm within cortical bone. When periosteal BFR is an important end point of a study employing aged mice and rats, the marker intervals used for cancellous bone double labeling may result in inseparable double labels at periosteal surfaces because MAR may be much lower there compared with cancellous and endocortical bone. To solve this problem, a second pair of markers with longer marker interval can be used, such as administration of alizarine complexone at Days 11 and 1 before necropsy for assessment of periosteal surfaces, and calcein at Days 5 and 2 before necropsy for assessment of cancellous bone BFR in a mouse study. In young mice and rats, this is usually not a problem due to the high periosteal MAR.

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4. Notes 1. Dry the benzoyl peroxide using a desiccator or by incubating at 40°C using a drying oven. Dried benzoyl peroxide should be handled with caution, since it may explode! 2. Prepare the MMA solution freshly before use and stir for at least 1 h before adding N,N-dimethyl-p-toluidine. 3. Make up fresh before each use by adding 50 ml 38% formaldehyde to 150 ml 0.138 M sodium carbonate. 4. Make up fresh by adding 10 ml potassium ferrocyanide to 200 ml 10% thiosulphate; use within 1 h of preparation. 5. Combine the reagents and heat in water bath or oven for 12 h at 50°C. Incubate for a further three days at 37°C and filter into a brown flask. 6. Filter prior to use. The solution can be reused for staining multiple sections. 7. The TRAcP reagent needs to be made up fresh and used within 1 h of preparation. For negative controls, the sections can be stained with TRAcP reagent that has to be prepared without the addition of napthol AS-MX. 8. Filter before use. The solution can be reused for staining multiple sections. 9. Filter prior to use. The solution can be reused for staining multiple sections. 10. For larger bones such as the femurs and tibiae, it is advantageous to open the marrow cavity for better fixation and infiltration. We, therefore, suggest that these bones should be cut in half at the diaphysis to facilitate fixation. 11. Alkaline phosphatase enzyme activity is very sensitive to aldehyde-containing fixatives (16, 17). If histochemical analysis of alkaline phosphatase activity in bone is desired, we recommend that 40% ethanol fixation be used instead of PFA. 12. The wash step is only required if the samples have been fixed in PFA. 13. The incubation times for the dehydration and infiltration steps depend on sample size and age of the animals. Longer durations should be used for larger samples, samples from older animals or from animals treated with antiresorptives. Shorter durations can be used for bones from young animals or small bones. 14. Smaller glass vials can also be used for smaller bone samples. In this case, adjust the amounts of the MMA solution proportionately depending on the size of the vial.

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15. To prevent premature polymerization or the MMA, it is critical that all steps subsequent to addition of N,N-dimethyl-ptoluidine be carried out at 4°C (on water ice) and that the vials are prechilled on ice and kept on ice until they are transferred to a cooling unit for polymerization. If you have many samples to process, we recommend that you prepare fresh MMA/N,Ndimethyl-p-toluidine solution every 2 h. 16. Accurate temperature control during polymerization is critical. We use a Binder environmental test chamber for the complex temperature profiles that are required. 17. For many antigens it is possible to store the samples at room temperature, but this needs to be determined empirically on a case by case basis. 18. Older solutions of silver nitrate may require longer incubation times. 19. The TRAcP activity can be enhanced by incubating the sections in 0.2 M Tris–HCl buffer, pH 9.0, for 1 h at 37°C after the embedding material has been removed following completion of step 3 of the protocol. 20. It is important that you do not allow the sections to dry out. 21. Cancellous bone osteopenia in the appendicular skeleton is a typical feature of aging in mice. Accordingly, in aged mice or osteopenic rats, there may be only very little cancellous bone left in tibiae or femurs which precludes a meaningful analysis of cancellous bone turnover. To avoid this problem, the lumbar vertebrae should be harvested in aged mice or severely osteopenic rats. In osteopenic animals, the amount of cancellous bone available for analysis of bone turnover may be minimal. In order to arrive at meaningful results, we measure at least 20 mm of cancellous bone perimeter in rats, and 5 mm of cancellous bone perimeter in mice. If several sections need to be analyzed to arrive at these minimum requirements, it is important to always analyze the whole measuring area in individual sections (the same measuring area in all sections), and not just subregions. Preferential analysis of subregions may bias the measurement. 22. In experiments where growing animals are treated with potent antiresorptive drugs, the newly formed bone deposited during endochondral bone growth may form an area of densely packed bone spicules underneath the growth plate. This area of bone which is deposited under the influence of the drug is normally excluded from cancellous bone analyses. 23. Normally, the amount of unmineralized bone (osteoid) in rat and murine cancellous bone is very low, in the range of 1–2%. The method to measure structural histomorphometric data by image analysis of von Kossa-stained sections described in

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Subheading 3.11 does not pick up osteoid. In the absence of increased amounts of osteoid, the error is very small and thus acceptable. However, image analysis of von Kossa-stained sections will yield erroneous values when bone mineralization is disturbed, resulting in increased amounts of osteoid. In the latter case, structural histomorphometric data need to be assessed by the interactive measurements described under Subheading 3.15 for mice and Subheading 3.17 for rats. 24. In humans, the adjusted apposition rate (Aj.AR) is often used to calculate the formation period. The Aj.AR is given by multiplying MAR with the M.Pm/O.Pm ratio, and includes so called OFF periods during the formation period, when osteoblasts temporarily stop synthesizing collagen. This is not applicable to mice and rats because M.Pm/O.Pm is ³1.0 in many cases (17). Thus, we use only the active formation period in rodents. References 1. Parfitt, A. M., Drezner, M. K., Glorieux, F. H., Kanis, J. A., Malluche, H., Meunier, P. J., Ott, S. M., and Recker, R. R. (1987) Bone histomorphometry: standardization of nomenclature, symbols, and units. Report of the ASBMR Histomorphometry Nomenclature Committee. J. Bone Miner. Res. 2, 595–610. 2. Erben, R. G. (1997) Embedding of bone samples in methylmethacrylate: An improved method suitable for bone histomorphometry, histochemistry, and immunohistochemistry. J. Histochem. Cytochem. 45, 307–313. 3. Schenk, R. K., Olah, A. J., and Herrmann, W. (1984) Preparation of calcified tissues for light microscopy. In Methods of calcified tissue preparation (Dickson, G.R. Ed.), pp. 1–56. Elsevier, Amsterdam. 4. Reim, N. S., Breig, B., Stahr, K., Eberle, J., Hoeflich, A., Wolf, E., and Erben, R. G. (2008) Cortical bone loss in androgen-deficient aged male rats is mainly caused by increased endocortical bone remodeling. J. Bone Miner. Res. 23, 694–704. 5. Baron, R., Vignery, A., Neff, L., Silverglate, A., and Santa Maria, A. (1983) Processing of undecalcified bone specimens for bone histomorphometry. In Bone Histomorphometry: Techniques and Interpretation (Recker, R.R. Ed.), pp. 13–35. CRC Press, Boca Raton, FL. 6. Erben, R. G. (2003) Bone Labeling Techniques. In Handbook of Histology Methods for Bone and Cartilage (An, Y.H. and Martin, K.L. Eds.), pp. 99–117. Humana Press Inc, Totowa, NJ, USA. 7. Erben, R. G., Scutt, A. M., Miao, D. S., Kollenkirchen, U., and Haberey, M. (1997)

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12.

13.

Short-term treatment of rats with high dose 1,25-dihydroxyvitamin D3 stimulates bone formation and increases the number of osteoblast precursor cells in bone marrow. Endocrinology 138, 4629–4635. Frost, H. M. (1983) Bone histomorphometry: choice of marking agent and labeling schedule. In Bone Histomorphometry: Techniques and Interpretation (Recker, R.R. Ed.), pp. 37–52. CRC Press, Boca Raton, FL. Frost, H. M. (1983) Bone histomorphometry: analysis of trabecular bone dynamics. In Bone Histomorphometry: Techniques and Interpretation (Recker, R.R. Ed.), pp. 109– 131. CRC Press, Boca Raton, FL. Erben, R. G. (1996) Trabecular and endocortical bone surfaces in the rat: Modeling or remodeling? Anat. Rec. 246, 39–46. Reim, N. S., Breig, B., Stahr, K., Eberle, J., Hoeflich, A., Wolf, E., and Erben, R. G. (2008) Cortical bone loss in androgen-deficient aged male rats is mainly caused by increased endocortical bone remodeling. J. Bone Miner. Res. 23, 694–704. Ibbotson, K. J., Orcutt, C. M., D’Souza, S. M., Paddock, C. L., Arthur, J. A., Jankowsky, M. L., and Boyce, R. W. (1992) Contrasting effects of parathyroid hormone and insulin-like growth factor I in an aged ovariectomized rat model of postmenopausal osteoporosis. J. Bone. Miner. Res., 7, 425–432. Lauritzen, D. B., Balena, R., Shea, M., Seedor, J. G., Markatos, A., Le, H. M., Toolan, B. C., Myers, E. R., Rodan, G. A., and Hayes, W. C. (1993) Effects of combined prostaglandin and alendronate treatment on the histomorphometry

19 and biomechanical properties of bone in ovariectomized rats. J. Bone Miner. Res. 8, 871–879. 14. Uzawa, T., Hori, M., Ejiri, S. and Ozawa, H. (1995) Comparison of the effects of intermittent and continuous administration of human parathyroid hormone(1–34) on rat bone. Bone 16, 477–484. 15. Bentolila, V., Boyce, T. M., Fyhrie, D. P., Drumb, R., Skerry, T. M., and Schaffler, M. B. (1998) Intracortical remodeling in adult rat long bones after fatigue loading. Bone 23, 275–281.


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16. Weber, K., Kaschig, C., and Erben, R. G. (2004) 1alpha-Hydroxyvitamin D2 and 1alphahydroxyvitamin D3 have anabolic effects on cortical bone, but induce intracortical remodeling at toxic doses in ovariectomized rats. Bone 35, 704–710. 17. Erben, R. G., Eberle, J., Stahr, K., and Goldberg, M. (2000) Androgen deficiency induces high turnover osteopenia in aged male rats: a sequential histomorphometric study. J. Bone Miner. Res. 15, 1085–1098.

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Chapter 20 Studying Gene Expression in Bone by In Situ Hybridization Ina Kramer, Rishard Salie, Mira Susa, and Michaela Kneissel Abstract Here, we described a method for carrying out nonradioactive in situ hybridization to detect mRNA transcripts in cryostat sections of mouse bone using the CryoJane® Tape-Transfer System and digoxigenin (DIG)-labeled riboprobes. Key words: In situ hybridization, ISH, Bone, Histology, Gene expression, mRNA, Transcript

1. Introduction The technique of in situ hybridization (ISH) (1) is a valuable method for assessing the temporal and spatial patterns of gene expression in various organs and tissues including bone (2, 3). It involves hybridizing a labeled nucleic acid probe to cells, tissue sections or whole mount tissues, and embryos. The difference between ISH and other methods of assessing gene expression such as quantitative real-time polymerase chain reaction (PCR) is that it allows the researcher to visualize expression of a particular gene of interest in tissue samples where the morphology has been preserved. Therefore, it is possible to determine which cell types express the gene of interest and to assess in a semiquantitative manner, if transcript abundance varies in different cell types. Moreover, ISH is frequently used for assessing temporal changes in gene expression by performing time course analyses of cellular differentiation and tissue maturation during embryonic and postnatal development (4). Disadvantages of ISH include the fact that it is relatively time-consuming and technically demanding and less suitable than real-time PCR for the detection of low abundance transcripts and quantitative gene expression analyses. Here, we


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described a method for ISH based on nonradioactive digoxigenin (DIG)-labeled riboprobes performed on cryo-sections of mouse bone generated with the CryoJane® Tape-Transfer System (5).

2. Materials 2.1. Tissue Collection, Fixation, Decalcification, and Embedding

1. 0.22-μm pore sized filter cups. 2. Plastic peel-away embedding molds. 3. Ketamine and xylazine. 4. Phosphate buffer: 1 M sodium phosphate in distilled water, pH 7.4. Autoclave and store at room temperature. 5. Fixative: 4% (w/v) paraformaldehyde (PFA) in 0.1 M sodium phosphate buffer, pH 7.4. Sterile filter and store at 4°C for a maximum of 2 days or freeze in aliquots at −20°C for longterm storage (see Note 1). 6. Decalcification solution (see Note 2). 0.48 M ethylenediaminetetraacetic acid (EDTA) in distilled diethylpyrocarbonate (DEPC) treated water, pH 7.4. Sterile filter and store at 4°C for up to 4 weeks. 7. 30% (w/v) sucrose in PBS without calcium and magnesium. Sterile filter and store at 4°C.

2.2. Tissue Sectioning

1. Cryostat for tissue sectioning equipped with the CryoJane® Tape-Transfer System (Instrumedics, St. Louis, MO). 2. Disposable or permanent metal knife for cryo-sectioning. 3. Razor blades to trim the frozen tissue block by hand prior to cryo-sectioning. 4. Small artist’s paintbrush for cleaning the knife. 5. Fine metal tweezers. 6. CryoJane® ultraviolet (UV) light-sensitive adhesive-coated slides (Instrumedics). 7. CryoJane® tissue tape and hand roller (Instrumedics). 8. Microscope Cover Slips, 24 × 60 mm.

2.3. Generation of DIG-Labeled Riboprobes

1. 5 μg template DNA (see Note 3). 2. Restriction enzymes (see Note 4). 3. DIG RNA labeling Mix (Roche). 4. T3, T7, and SP6 RNA polymerases (Roche). 5. RNAse inhibitor (40 U/μl). 6. DEPC-treated water.

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7. RNase-free DNase I (Roche). 8. 8 M LiCl. 9. 0.2 M EDTA in DEPC water, pH 8.0. 10. DIG-labeled control RNA (Roche). 11. Agarose. 12. TBE buffer (50 mM Tris–HCl, pH 7.8, 50 mM orthoboric acid, 2 mM NaEDTA). 13. Ethidium bromide. 14. Petri dish. 15. 100% ethanol. 16. 70% ethanol. 17. 20% (w/v) Glycogen in distilled water. 2.4. Hybridization, Washes, Probe Detection, and Signal Development

1. Metal slide racks, glass slide troughs, and staining containers. 2. Small plastic slide box. 3. Hybridization oven. 4. 1 M Tris, pH 7.5 and pH 9.5. 5. Proteinase K stock: 10 mg/ml proteinase K in distilled water. Aliquot and freeze at −20°C. 6. Proteinase K buffer: 1 μg/ml proteinase K, 6.25 mM EDTA in 0.05 M Tris, pH 7.5. Add proteinase K fresh from the stock solution just before use. 7. Acetylation buffer: 1.16% (v/v) triethanolamine in distilled water. 8. 20× SSC buffer: 3 M NaCl and 0.3 M tri-sodium citrate (Na3C6H5O7) in DEPC water, pH 7.0. Autoclave to sterilize. 9. 50× Denhardt’s solution: 1% (w/v) Ficoll (Type 400), 1% (w/v) polyvinylpyrrolidone, and 1% (w/v) bovine serum albumin in distilled water. Filter sterilize and store at −20°C in 1–5 ml aliquots. 10. Baker’s yeast total RNA: 25 mg/ml in distilled water. Aliquot and freeze at −20°C. 11. Hybridization buffer: 50% (v/v) formamide, 5× SSC, 5× Denhardt’s solution, 0.25 mg/ml baker’s yeast RNA, 0.5 mg/ml single-stranded (ss) DNA from fish sperm in distilled water. Prepare 12-ml aliquots and store at −20°C. 12. 50% Formamide in distilled water. 13. B1 buffer: 0.1 M maleic acid, 0.15 M NaCl in distilled water, pH 7.5. Autoclave and store at RT. 14. Blocking reagent for nucleic acid hybridization and detection (Roche).

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15. 10% (w/v) blocking reagent in B1 buffer (see Note 5). Autoclave and store in aliquots at −20°C. 16. B2 blocking buffer: 2% (v/v) blocking reagent in B1 buffer. Prepare fresh just before use from 10% stock. 17. Anti-DIG sheep IgG conjugated to alkaline phosphatase (antiDIG-AP antibody; Roche). Prepare 0.05% (v/v) working dilution in B2 buffer fresh and keep on ice until use. 18. B3 detection buffer: 0.1 M NaCl, 5 mM MgCl2 in 0.1 M Tris, pH 9.5. Sterile filter. 19. 10% (v/v) Tween-20 stock solution in distilled water. 20. 0.1% (v/v) Tween-20 in B3 detection buffer. Prepare fresh before use. 21. 5-Bromo-4-chloro-3-indolylphosphate/4-nitro blue tetrazolium chloride (BCIP/NBT) Alkaline Phosphatase substrate kit. 22. Levamisole. 23. B4 developing solution (see Note 6). 24. TE buffer: 0.01 M Tris, 1 mM EDTA in distilled water, pH 8.0.

3. Methods 3.1. Tissue Collection, Fixation, Decalcification, and Embedding

1. Anesthetize the animal by intraperitoneal injection of a cocktail of 120 mg ketamine per kg body weight and 25 mg xylazine per kg. 2. Verify that the animal is fully anesthetized, as reflected by absence of toe and eye blink reflexes. 3. Spray the animal’s skin and fur at the ventral abdomen with 70% ethanol. 4. Carefully cut the skin, abdominal muscle wall, and ribcage with sharp scissors from the midabdomen to about the throat level. 5. Remove the diaphragm and ventral ribs from the thoracic cavity, taking care not to rupture heart, lungs, or major blood vessels. 6. Puncture the right atrium of the heart with sharp scissors taking care not to damage the remainder of the heart. 7. Slowly inject 10 ml of ice-cold PBS, pH 7.4 into the left ventricle of the beating heart at a rate of approximately 3 ml/min to exsanguinate the animal (see Note 7). 8. When exsanguination is complete, perfuse the animal with icecold 4% paraformaldehyde fixative (see Note 8).

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9. Carefully dissect out the bones and any other tissues that you wish to examine as quickly possible (see Note 9). 10. Immediately following dissection, fix the samples by immersing in ice-cold 4% paraformaldehyde and incubate on a shaking platform at 4°C. Fixation times vary from 6 h up to 3 days depending on the age of the animal and type of bone being analyzed (Table 1). 11. When fixation is complete, wash the tissue by immersing in PBS at 4°C for 30 min. 12. Transfer the samples to decalcification solution (see Note 10) and incubate on a shaking platform at 4°C with daily changes of decalcification solution for up to 5 days, depending on the age of the animal and type of bone being analyzed (Table 2). 13. Rinse the samples in ice-cold PBS for 5–10 min. 14. Transfer to 30% sucrose in PBS for 8–24 h at 4°C on a shaking platform. 15. When the sucrose infiltration is complete (see Note 11) remove excess sucrose solution with a paper towel.

Table 1 Suggested duration of fixation step at 4°C Mouse age (days)

Mouse calvaria

Mouse long bone, vertebrae

0–2.5

6h

8–12 h (overnight)

3–7.5


1 day

8–14.5

1 day

1–2 days

15–21.5

1–2 days

3 days

Juvenile and adult (>21.5)

3 days

3 days

Table 2 Suggested duration of decalcification step at 4°C Mouse age (days)

Mouse calvaria

Mouse long bone, vertebrae

0–2.5


1–2 days

3–7.5

1 day

3–5 days

8–14.5

3 days

3–5 days

Juvenile and adult (>14.5)

3–5 days

3–5 days

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16. Carefully orientate the samples in plastic peel-away embedding molds filled with OCT compound and freeze on dry ice (see Note 12). 3.2. Cryo-Sectioning

1. Place the CryoJane® tissue tape, adhesive-coated slides, hand roller, and the frozen tissue block in the cryostat set at −24°C and leave for about 10 min (see Note 13). 2. If necessary, trim the sides of the block not containing any embedded tissue by hand with a razor blade. 3. Place the trimmed tissue block in the cryostat block holder in the desired position. 4. Section the tissue in increments of 10–20 μm thickness until just before the level of interest has been reached. 5. Adjust the section thickness to 5–7 μm and adhere a cold adhesive tape segment to the block surface by slowly peeling off the removable backing. 6. Place the adhesive side of the tape on the embedded frozen sample and ensure that it is firmly attached to the tissue block by applying pressure with the hand roller. 7. Raise the lower end of the tape above the level of the knifeedge and slowly cut a section in the usual manner. 8. Using fine tweezers, transfer the tape containing the freshly cut section to the adhesive-coated slide. 9. Using the hand roller, apply pressure on the tape to ensure that the section is firmly attached to the slide surface. 10. Remove the slide from the cryostat chamber, place it briefly on the back of your hand for 1–3 s, return to the cryostat chamber and apply pressure with the hand roller once again to ensure that the tissue is firmly attached to the slide (see Note 14). 11. Insert the slide into the CryoJane® flash tray and treat with a single UV flash to polymerize the adhesive coating of the slide. 12. Leave the slide in the coldest part of the cryostat for about 2–3 min. 13. With the slide remaining in the cryostat, carefully remove the adhesive backing of the tape from the slide using cold forceps, keeping the delaminated backing adhered to the slide to prevent the section from coming off with the tape. 14. Air dry the slides at room temperature for a minimum of 20 min and store the slides at −20°C until further use.

3.3. Linearization of the Plasmid Containing the Probe of Interest

1. Linearize 5 μg of plasmid DNA containing the probe of interest in a reaction volume of 100 μl in a sterile microcentrifuge tube using an appropriate restriction enzyme (see Notes 16 and 17).

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2. Confirm that the plasmid is completely linearized by running about 5 μl of the digested plasmid on a 1% agarose gel alongside an uncut plasmid sample. 3. Purify the linearized DNA by adding 100 μl of phenol/chloroform/IAA to the plasmid and vortex to mix. 4. Place the tube in a microcentrifuge and spin at 10,000 × g for 3 min. 5. Carefully aspirate about 90 μl of the upper aqueous phase into a new sterile microcentrifuge tube. 6. Add 1 μl glycogen, 45 μl 7.5 M ammonium acetate, and 225 μl 100% ethanol and mix by vortexing. 7. Place the sample in a −70°C or −80°C freezer for 15–30 min. 8. Place the tube in a microcentrifuge and spin at 10,000 × g for 10 min at room temperature. 9. Carefully aspirate the supernatant taking care not to dislodge the DNA pellet. 10. Wash the pellet by adding 1 ml 70% ethanol to the tube, place in a microcentrifuge, and spin at 10,000 × g for 10 min at room temperature. 11. Air dry the DNA pellet at room temperature for about 5 min. 12. Resuspend the plasmid in 10 μl of RNAse-free 1× transcription buffer provided with the DIG RNA labeling mix kit. 13. Estimate the concentration of plasmid by running 1 μl of the resuspended DNA on a 1% agarose gel in TBE buffer alongside a DNA marker of known concentration. 14. Store the linearized plasmid at −20°C until further use. 3.4. Generation of DIG-Labeled Riboprobes

1. Add 1 μg of linearized plasmid (Subheading 3.3, step 14), 2 μl 10× transcription buffer, 2 μl DIG-NTP labeling mix, 0.5 μl RNAse inhibitor, 1.5 μl of the appropriate RNA polymerase (T7, T3 or SP6) to a sterile microcentrifuge tube, and bring the reaction volume up to 20 μl with DEPC water. 2. Collect the reaction mix at the bottom by pulsing in a microcentrifuge for 10 s. 3. Incubate the reaction mixture at 37°C for 2–3 h (see Note 18). 4. Add 1 μl of RNAse-free DNAse I to the reaction and incubate for 15 min at 37°C. 5. Stop the reaction by adding 2 μl RNAse-free 0.2 M EDTA. 6. Add 1.25 μl 8 M LiCl and 75 μl 100% ethanol. Mix the sample by vortexing gently and transfer to a −70°C or −80°C freezer for 15–30 min to precipitate the RNA. 7. Place the sample in a microcentrifuge and spin at 10,000 × g for 15 min.

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8. Carefully remove the supernatant taking care not to lose the RNA pellet. 9. Add 1 ml 70% ethanol to the tube to wash the pellet. 10. Place the sample in a microcentrifuge and spin at 10,000 × g for 5 min. 11. Carefully aspirate the supernatant and discard. 12. Allow the RNA pellet to air dry for about 5 min at room temperature. 13. Dissolve the RNA in 50 μl distilled water containing 0.4 U/μl RNAse inhibitor. 14. Prepare a 1% agarose gel containing a few drops of ethidium bromide in a Petri dish and dot 1 μl of the riboprobe solution directly onto the gel surface alongside with serial dilutions of DIG-labeled control RNA at known concentrations. 15. Estimate the amount of DIG-labeled riboprobe present in the sample by comparing the intensity of the dot with the known amounts of DIG-labeled control RNA. 16. Store the probe frozen at −20°C until further use or for up to 12 months. 3.5. Hybridization

A summary of the sequential steps involving immersion of the slides in staining containers or horizontal incubation in humidified chambers is outlined schematically in Fig. 1. Unless otherwise indicated all steps are carried out at room temperature. Generally, both negative and positive controls should be included when performing ISH (see Note 19). 1. Take an aliquot of hybridization solution from the freezer and prewarm at 37°C. 2. Place the slides in a rack and fix the sections by immersing the slides in 4% paraformaldehyde fixative for 10 min. 3. Wash the slides by immersing three times in PBS for 3 min at a time. 4. Immerse the slide in proteinase K buffer containing 1 μg/ml proteinase K for 5 min. 5. Transfer the slides back to 4% paraformaldehyde fixative and incubate for 5 min. 6. Wash the slides by immersing three times in PBS for 3 min at a time. 7. Transfer the slides to a staining dish placed on top of a magnetic stirrer containing acetylation buffer. 8. Keeping the slides just below the solution surface add 1.3 ml acetic anhydride per 500 ml acetylation buffer, while simultaneously stirring using a sterile magnetic bar. Incubate for 10 min (see Note 20).

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a TISSUE PREPARATION Tissue Dissection

Decalcification 0.5 –5 d, 4°C

Fixation 0.5 –3 d, 4°C Embedding & Tissue storage -70 to -80°C

Cryo-protection 8 -24 h, 4°C

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b DIG-LABELED RIBOPROBE GENERATION DNA Purification & Quantification

Linearization 1 –2 h, 37°C

Cloned DNA with Probe of Interest In Vitro Transcription, DIG-RNA Labeling 2 –3 h, 37°C

Riboprobe Dot Quantification

RNA Purification

c DAY 1: HYBRIDIZATION Fix 10‘

PBS

PBS

PBS

3‘

3‘

3‘

ProtK 5‘

Fix

PBS

PBS

PBS

y Acetyl

5‘

3‘

3‘

3‘

10‘

PBS

PBS

PBS

Prehyb

Hyb

3‘

3‘

3‘

1h

o/n 50-72°C

d DAY 2: WASHES, PROBE DETECTION AND SIGNAL DEVELOPMENT 5SSC

SSC

SSC

B1

B2

5‘

1h

5‘

5‘

1h

α-DIG

B1

B1

B1

B3

3 h or o/n, 4°C

5‘

5‘

5‘

5‘

T20/B3

B4

TE

dH2O

5‘

up to o/n in the Dark

5‘

10‘

70°C

Air Dry, Mount & Store @ 4°C

Fig. 1. Schematic outline of ISH. Schematic representation of the sequential processing steps for ISH with incubation times indicated starting with the tissue (a) and riboprobe (b) preparation, followed by the actual hybridization on the first day (c) and the washes, probe detection, and signal development on the second day (d). Immersion steps using slide racks immersed in staining containers are indicated by trapezoids, horizontal incubation of slides in humidified chambers is depicted by rectangles. If not indicated otherwise, all steps are carried out at RT. Abbreviations: α-DIG: anti-DIG-AP antibody, Acetyl: TAE acetylation buffer with freshly added acetic anhydride, B1: B1 buffer, B2: B2 blocking buffer, B3: B3 detection buffer, B4: B4 developing solution, dH2O: distilled water, Fix: 4% PFA fixative, Hyb: hybridization solution with probe added, o/n: overnight, PBS: phosphate-buffered saline, Prehyb: hybridization solution without probe, ProtK: Proteinase K buffer with proteinase K added just before use, 5SSC: 5× SSC buffer, SSC: 0.2× SSC, T20/B3: 0.1% Tween20 in B3 buffer, TE: TE buffer.

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9. Wash the slides in PBS for 3 min. 10. Remove the slides from the rack, and lay horizontally in a tray humidified with water-soaked paper towels. Add 800 μl hybridization solution to each slide and incubate for between 1 and 3 h. 11. Add sufficient DIG-labeled riboprobe to 1 ml of hybridization solution to give a final RNA concentration of 1.0 μg/ml. 12. Heat the diluted riboprobe/hybridization solution to 80°C for 5 min and transfer immediately to ice for 1 min. Remove from the ice and maintain at room temperature until adding to the slides. 13. Place a folded paper towel soaked in 50% formamide solution at the bottom of a small plastic slide box to serve as a humidified hybridization cassette (see Note 21). 14. Remove the prehybridization solution and add 250 μl of the riboprobe/hybridization solution over the entire surface of the slide. Carefully place a glass coverslip on top of the slide covering all tissue sections, avoiding entrapment of air bubbles. 15. Place the slides horizontally in the prepared humidified plastic box. Tightly seal the cassette with tape to avoid evaporation overnight, while keeping the box upright to prevent leakage of the hybridization solution. Place the box standing, i.e., with the slides orientated horizontally and parallel to the ground in a hybridization oven preset to an appropriate temperature (see Note 22) and incubate overnight. 3.6. Washes, Probe Detection, and Signal Development

1. Heat 5× SSC and 0.2× SSC solutions to 70°C in covered-glass staining dishes with empty slide racks in a water bath (see Note 23). 2. Transfer the slides with coverslips to the 5× SSC solution and incubate for 5 min at 70°C. 3. Transfer the slides one by one from the 5× SSC to the 0.2× SSC solution. (The coverslips should automatically come off at this point and remain in the 5× SSC solution). 4. Incubate slides at 70°C in 0.2× SSC for 1 h. 5. Remove the entire 0.2× SSC staining dish with the slides from the water bath and allow to cool for 5 min at room temperature. 6. Transfer the slides to B1 buffer and incubate for 5 min at room temperature. 7. Lay the slides horizontally in a humidified chamber, cover with 800 μl 2% blocking solution, and incubate for 1 h at room temperature.

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8. Aspirate the blocking solution from the slides, replace with 700 μl of diluted anti-DIG-AP antibody solution, and incubate for 3 h at room temperature or overnight at 4°C in a humidified tray. 9. Aspirate the antibody solution from each slide and transfer all slides to a slide rack. 10. Wash three times for 5 min in B1 buffer in a staining container at room temperature. 11. Transfer to B3 buffer and incubate for 5 min at room temperature. 12. Lay slides horizontally in a humidified chamber and add 0.9 ml 0.1% Tween-20/B3 solution each slide. 13. Aspirate the Tween-20/B3 buffer and replace with 600 μl B4 developing solution. 14. Cover the humidified chamber with aluminum foil to block out the light and incubate at room temperature for up to 12 h, monitoring progression of signal development periodically under a dissecting microscopy (see Note 24). 15. When the signal has developed to the desired level, stop the reaction by transferring the slide to a slide rack in a staining container filled with TE buffer. Incubate at room temperature for 5 min. 16. Wash the slides in distilled water for 10 min at room temperature and air dry for 10–20 min. 17. Mount slides by adding about 350 μl Kaiser’s glycerin gelatin in a drop-by-drop fashion to each slide and store at 4°C.

4. Notes 1. PFA is toxic and should always be handled under a fume hood while wearing personal protective equipment. 2. Dissolve the EDTA by adjusting the pH to 7.4 using 14.8 M ammonium hydroxide rather than sodium hydroxide solution, as EDTA dissolved in ammonium hydroxide solution has been shown to result in more efficient decalcification of bone samples (6). 3. Plasmid with cloned cDNA or part of a genomic coding sequence. Suitable plasmids with recognition sites for T7, T3, or SP6 polymerase for generating riboprobes are commercially available via different sources. Expressed sequence tags (ESTs, complete cDNAs and cDNA fragments) can be obtained from the I.M.A.G.E. (Integrated Molecular Analysis of Genomes

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and their Expression) Consortium (http://image.hudsonalpha.org/). 4. A selection of restriction enzymes is required to linearize the plasmid DNA at the 5¢ or 3¢ end of the probe sequence to prepare antisense or sense riboprobes, respectively. 5. To prepare a blocking reagent stock solution, dissolve the blocking reagent in B1 buffer with shaking and heating either on a heating block or in a microwave oven. 6. To prepare the B4 developing solution first mix all reagents in the BCIP/NBT Alkaline Phosphatase substrate kit by vortexing to dissolve potential precipitates that may have formed upon storage. Next add six drops of the NBT solution (Reagent 1) to 15 ml B3 buffer and mix well by vortexing. Then add six drops of the BCIP solution (Reagent 2) to the solution and mix well again by vortexing. Next add three drops of levamisole (inhibitor of endogenous alkaline phosphatases) to the solution and mix well by vortexing. Finally, add 0.05 ml of 10% Tween-20 stock to the solution. The antibody-conjugated alkaline phosphatase will oxidize BCIP to indigo and reduce NBT to diformazan. The reaction products then form a waterinsoluble dark blue to magenta precipitate whose production rate is proportional to the amount of anti-DIG-AP antibody bound to the hybridized riboprobe. 7. For smaller or larger animals use less or more PBS according to the respective body weight. As the blood is washed out from the body, the liver turns from a deep red to a yellowish color indicating successful perfusion. 8. Following successful perfusion fixation the animal should be rigid. 9. We usually isolate femora, tibiae, lumbar vertebrae, and the calvaria and sometimes harvest the humerii, ulnae, and radii. Care should be taken not to remove all soft tissue surrounding the bone in order to preserve cellular structures at the periosteal surface. 10. For optimal cryo-sectioning results with the CryoJane® TapeTransfer System partial decalcification is recommended, as can be accomplished in 5 days or less. The volume of decalcification solution should be at least ten times higher than the actual tissue volume and the solution should be daily changed to ensure successful decalcification. 11. The tissue samples should sink when they are completely infiltrated with the sucrose solution. 12. Avoid formation of air bubbles when pouring the viscous OCT compound. Frozen tissue blocks should be wrapped in aluminum foil and stored in air-tight plastic bags at −80°C until sectioning to prevent dehydration of the sample.

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13. This is to ensure that the tissue block and other materials are at the same temperature as the cryostat. 14. From experience we have determined that the final sections have superior morphology and adhere better, if the frozen slide with the laminated tape and tissue section on it is gently heated on the back of the hand and then re-rolled immediately thereafter. Appropriate lamination is achieved when cortical bone tissue no longer appears bright white on the tape. The CryoJane® Tape-Transfer System requires some practice to cut high-quality sections of adult mouse long bone. 15. The slides can be left for up to 8 h at room temperature after air drying if necessary. 16. The optimum riboprobe length for ISH is between 200 and 500 nucleotides, but this needs to be determined empirically for each application. 17. If using a restriction enzyme such as ApaI or KpnI, which generates a 3 overhang, proceed with a Klenow/T4 DNA polymerase reaction to remove the overhang in order to prevent false priming of the RNA polymerase. 18. If using less than 1 μg linearized template DNA, extend the incubation time to 4 h. 19. It is important to include controls to assess probe specificity. False-positive signal may arise from target sequence independent binding of the probe to nucleic acid and/or nonspecific binding to other tissue components. Nonspecific binding to nucleic acid can be determined by using the probe in a Northern or Southern blot to determine, if it hybridizes specifically with the target sequence of the expected molecular size. Nonspecific binding to tissue components can be uncovered by using sense riboprobe having an identical GC content (see examples in Figs. 2 and 3) or by pretreatment of the tissue with RNase, which should abolish all hybridization to RNA and thus reveal unspecific binding to other tissue components. Nonspecific staining can be reduced by increasing the hybridization temperature and/or salt concentration. Positive controls can be included by using sections of tissue known to express the respective gene of interest. 20. Acetylation blocks reactive amine groups present in the tissue and reduces nonspecific binding. As the half-life of acetic anhydride is very short in aqueous solution, care should be taken to rapidly disperse it across all slides, while simultaneously stirring the solution. 21. For a 25-slide-plastic box use 2 ml 50% formamide. Thoroughly clean the box after each usage with water. When analyzing expression of several marker genes in parallel, we recommend

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Fig. 2. ISH staining for the osteoblastic lineage marker gene type 1 collagen alpha 1. 10-μm thin cryosections of partially decalcified late stage skeletally growing (3-month-old) wild-type (C57BL/6) mouse femur (a–d) and tibia (e–f ) were hybridized with DIG-labeled control sense (a, b) or antisense (c-f ) riboprobe of the osteoblastic lineage marker gene type 1 collagen alpha 1. Boxes in (a, c, e) denote corresponding higher magnifications depicted in (b, d, f ), respectively. Note strong type 1 collagen alpha 1 expression in bone surface attached endocortical and trabecular osteoblasts (arrowheads; d, f) compared to weaker expression in cortical (c) and trabecular (t) osteocytes (arrows; d, f ). No expression signal is detected on sections hybridized with sense control probe (a, b). Scale bar: 0.4 mm (a, c, e); 50 μm (b, d, f ).

that separate boxes be used for each probe and that new gloves are used for each probe to avoid cross-contamination. 22. Hybridization temperature usually needs to be determined empirically for an optimized signal-to-noise ratio. In practice, most probes can be hybridized at 55–58°C, but some require higher temperatures. In theory (7), the hybridization temperature is based on the melting temperature (Tm) of the respective double-stranded RNA–RNA molecule, which depends on the length and G/C content of the exact-match riboprobe. The Tm

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Fig. 3. ISH staining for the hypertrophic chondrocyte marker gene type 10 collagen. 5-μm thin cryosections of decalcified juvenile skeletally growing (3-week-old) wild-type (C57BL/6) mouse femur (a–d) hybridized with DIG-labeled control sense (a, b) or antisense (c, d) riboprobe of the hypertrophic chondrocyte marker gene type 10 collagen. Note strong specific type 10 collagen expression in terminally differentiated, hypertrophic chondrocytes in the growth plate (c, d). No expression signal is detected on sections hybridized with sense control probe (dashed lines demark zone of hypertrophy in a, b). Scale bar: 0.1 mm (a, c); 50 μm (b, d).

is calculated according to the following formula: Tm = 79.8°C + 18.5(log[Na+]) + 0.584(%GC) + 0.0012(%GC)2 − 820/n − 0.35 (%F), with %F: formamide concentration, %GC = percentage of guanine and cytosine in the entire probe sequence, log[Na+]: log molar sodium concentration, and n: probe length. A 1% mismatch in the probe sequence will reduce the melting temperature by 1°C. As sequence polymorphisms are usually not known, but should be taken into account, choosing a hybridization temperature that is 1–5°C lower than the calculated Tm for the exact-match probe sequence is advisable. 23. Never place glass staining dishes containing cold solution into the preheated water bath, as they may crack! 24. It is advisable to check for the development of signal every 5–10 min initially and then every 30–60 min for several hours up to overnight incubation.

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Acknowledgments We thank Dr. M. John (Novartis Institutes for BioMedical Research, Basel, Switzerland) and Prof. E. Schipani (Massachusetts General Hospital and Harvard Medical School, Boston, MA, USA) for providing the DNA constructs for generation of type 1 collagen alpha 1 and type 10 collagen riboprobes. We are grateful to Profs. D. W. Rowe and I. Kalajzic (University of Connecticut Health Center, Farmington, CT, USA) as well as Prof. S. E. Harris (University of Texas Health Science Center at San Antonio, San Antonio, TX, USA) for advice on fixation and decalcification times, as well as cryo-sectioning with the CryoJane® Tape-Transfer System and ISH on skeletal tissue. References 1. Darby, I. A., and Hewitson, T. D. (2006) In Situ Hybridization Protocols (Series: Methods in Molecular Biology), 3 rd edn. Humana Press, Totowa. 2. Nomura, S., Hirakawa, K., Nagoshi, J., Hirota, S., Kim, H., Takemura, T., Nakase, T., Takaoka, K., Matsumoto, S., Nakajima, Y., Takebayashi, K., Takano-Yamamoto, T., Ikeda, T., and Kitamura, Y. (1993) Method for Detecting the Expression of Bone Matrix Protein by In Situ Hybridization Using Decalcified Mineralized Tissue. Acta Histochem. Cytochem. 26, 303–309. 3. Salie, R., Li, H., Jiang, X., Rowe, D. W., Kalajzic, I., and Susa, M. (2008) A Rapid, Nonradioactive In Situ Hybridization Technique for Use on Cryosectioned Adult Mouse Bone. Calcif. Tissue Int. 83, 212–221.

4. Witte, F., Dokas, J., Neuendorf, F., Mundlos, and S., Stricker, S. (2009) Comprehensive expression analysis of all Wnt genes and their major secreted antagonists during mouse limb development and cartilage differentiation. Gene Expr. Patterns. 9, 215–223. 5. Jiang, X., Kalajzic, Z., Maye, P., Braut, A., Bellizzi, J., Mina, M., and Rowe, D. W. (2005) Histological analysis of GFP expression in murine bone. J. Histochem. Cytochem. 53, 593–602. 6. Sanderson, C., Radley, K., and Mayton, L. (1995) Ethylenediaminetetraacetic acid in ammonium hydroxide for reducing decalcification time. Biotech. Histochem. 70, 12–18. 7. Farrell, R. E, (2009) RNA Methodologies: A Laboratory Guide for Isolation and Characterization. 4th edn. Elsevier, Oxford.

Chapter 21 Immunostaining of Skeletal Tissues Tobias B. Kurth and Cosimo De Bari Abstract Immunohistochemistry (IHC) is a routinely used technique in clinical diagnosis of pathological conditions and in basic research. It combines anatomical, immunological, and biochemical methods and relies on the specific binding of an antibody to an antigen. Using the technique with mineralised tissues is more complicated than with soft tissues. This can in most cases be overcome by demineralising the samples, which allows embedding in paraffin wax and a simpler work-up than for resin-embedded, or for frozen samples. This chapter describes methods for IHC on paraffin-embedded formaldehyde fixed sections to detect antigens in the musculoskeletal tissues. Key words: Immunohistochemistry, Antigen retrieval, Mouse knee joint, Immunofluorescence, Paraffin section

1. Introduction The use of immunohistochemistry (IHC) dates back to the early 1940s when Coons et al. (1) used FITC-labelled antibodies to detect Pneumococcal antigens in infected tissues (1). Since then, IHC has become one of the most powerful routine methods in diagnostics and basic research. It combines histological, immunological, and biochemical techniques and is based on the principle that antigens can be detected in cells or tissues using specific antibodies. In this chapter, IHC on paraffin-embedded paraformaldehyde-fixed tissue sections using either enzyme-based or fluorescence-based methods (see Fig. 1) are described. We prefer the use of paraffin over frozen sections because the overall tissue morphology is better preserved allowing best identification of the labelled tissue components in relation to other structures. However, a disadvantage of using fixed and embedded material is the fact that many commercially available antibodies work better in frozen sections. Increasingly though, companies screen their primary


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Fig. 1. (a) Immunohistochemical staining for Collagen type II in the articular cartilage of a 3-month-old mouse. In this protocol, we used hyaluronidase treatment (4,000 U/ml for 60 min at 37°C) as antigen retrieval step. A peroxidase-based staining was performed using DAB which results in a brown signal of the antigen. Nuclei were counterstained with haematoxylin. Scale bar: 10 mm. (b) Immunofluorescence staining for the pan-haematopoietic marker CD45 in a bone marrow cavity of a 3-month-old mouse. Positive cells are demonstrated by a red membrane associated signal. Nuclei are counterstained with DAPI (blue). Scale bar: 10 mm.

antibodies on fixed embedded material and the use of a variety of antigen retrieval methods can rescue antigenicity in such tissues. So in short, our method of choice is formaldehyde fixation to contribute to better tissue preservation, followed by antigen retrieval to unblock the cross-linked amino groups of fixed proteins. We then apply a primary antibody that is tagged with either a fluorochrome or an enzyme (direct IHC), or we detect binding of an un-conjugated primary antibody with a secondary conjugated antibody (indirect IHC). To enhance intensity, sensitivity, and specificity of the signal, additional systems such as use of avidin–biotin complexes, or use of tyramide-based signal amplification are possible. When using enzyme-conjugated antibodies, a suitable enzyme substrate that precipitates at the site it is formed needs to be available. Many companies now produce kits for such histochemical reactions that are well tested and include appropriate blockers for endogenous enzyme activity in the tissue. IHC methods to detect expressed antigens can be usefully combined with demonstration of artificial labels deliberately integrated in the tissue, such as nucleoside analogues (e.g. BrdU) for pulse chase to detect proliferating cells. We recently used a double nucleoside analogue labelling strategy in a mouse model of articular cartilage knee joint injury to identify and characterise functional mesenchymal stem cells within the synovium in vivo (2). Of note, this study was performed using mainly IHC methods.

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Overall, IHC is an extremely useful method to combine anatomical and biochemical information and the methods described here can be adapted for use with tissues prepared in different ways, such as tissue embedded in methyl-methacrylate, or in acrylic resins such as Lowicryl HM20 after appropriate optimisation of each step. In general, optimisation of all steps is required for each tissue type and for each primary antibody. Use of automated robotic stainers is increasingly common in diagnostic settings and in larger research units and can help standardise staining between slides by minimising inter-slide variability, and allow quantification of staining intensity. Such methods are not described here in detail as they are largely dictated by the equipment and kits used for detection. While we have found such equipment very useful, especially as the detection kits are highly optimised to give superb sensitivity and therefore save on primary antibody, autostainers remain expensive to run in a standard laboratory setting. We therefore concentrate on manual protocols here, but encourage the reader to amend and adapt the principles to their specific experimental condition.

2. Materials Unless stated otherwise, materials can be obtained from Sigma or similar chemical suppliers. 1. Phosphate-buffered saline (PBS): use tablets and dilute with the required amount of distilled water. 2. Fixation solution, 2% paraformaldehyde (PFA) + 0.05% glutaraldehyde in PBS: to prepare 2% PFA solution place 450 ml of distilled water in a glass beaker. Heat to 60°C using a hot plate with stirring facility. While stirring, add 10 g of paraformaldehyde powder to the heated water. Cover and maintain at 60°C. Add five drops of 2N NaOH (one drop per 100 ml). The solution should clear within a short time (there will be some fine particles that will not disappear). Do not heat solution above 70°C, PFA will break down at temperatures above 70°C. Remove from heat and add 50 ml of 10× PBS. Adjust pH to 7.2; you may have to add some HCl. Final volume will be 500 ml. Filter and add 0.05% glutaraldehyde (50 ml in 100 ml). Place on ice when using it immediately or freeze aliquots at −20°C and thaw when needed (see Note 1). 3. Decalcifying Solution-Lite. 4. 4% EDTA in PBS, adjust pH to 7.2–7.4 using NaOH. 5. Superfrost+ slides (Menzer), 25 mm × 75 mm. 6. Citrate buffer: 10 mM citric acid, 0.05% Tween 20, pH 6.0. The solution can be stored at RT for up to 3 months or even longer when stored at 4°C.

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7. Tris–EDTA buffer: 10 mM Tris base, 1 mM EDTA solution, 0.05% Tween 20, pH 9.0. The solution can be stored at RT for up to 3 months or even longer when stored at 4°C. 8. Hydrochloric acid (0.2 N): add 3 ml of fuming hydrochloric acid (37%) to 497 ml of distilled water. 9. Pepsin solution: use porcine pepsin at a concentration of 0.5–3 mg/ml diluted in 0.2N hydrochloric acid. Shake gently and keep at 37°C until the crystals have dissolved. 10. 3% H2O2 in distilled water. 11. Tris-buffered saline (TBS) 20× stock: add 122 g Trizma base and 180 g NaCl to 900 ml of distilled water. Stir until dissolved and adjust pH to 7.6 using concentrated HCl. Fill aliquots in 50 ml falcon tubes and freeze at −20°C. Prepare 1× TBS using one falcon tube and fill up to 1,000 ml with distilled water. 12. Washing buffer: 0.2% Triton X-100 in TBS. 13. Avidin blocking solution: ready-to-use solution (VECTOR, www.vectorlabs.com). 14. Humidified chamber: these are commercially available (e.g. staining tray from VWR) or can be made by yourself: use a box that is large enough to put in two 5 or 10 ml plastic pipettes to keep the slides raised up. Cover the bottom with some washing buffer and close the box with a lid. For immunofluorescence (IF) staining these humidified chambers must be impervious to light. 15. Biotin blocking solution: ready-to-use solution (VECTOR). 16. Blocking solution: 1% bovine serum albumin (BSA) in washing buffer. 17. Mouse-On-Mouse (MOM) Ig Blocking reagent (VECTOR): add two drops of stock solution to 2.5 ml of washing buffer. 18. MOM Diluent (VECTOR): add 600 ml of protein concentrate stock solution to 7.5 ml of washing buffer. 19. DNase solution: dilute Desoxyribonuclease I from bovine pancreas to a concentration of 1,000 U/ml with 0.15 M NaCl. Store stocks of 250 ml at −20°C and dilute with 250 ml of TBS Triton X-100 to achieve working solution. 20. Parafilm: cut small pieces (depends on the size of your section; we are using approximately 20 mm × 40 mm in size for mouse knee joint samples) of parafilm and fold approximately 5 mm from one shorter end that the parafilm side forms a 90° angle. Strip of paper and cover with the parafilm side that was protected by the paper to cover the section. 21. Avidin–biotin complex (ABC) reagent (VECTOR): add two drops of reagent A to 5 ml TBS Triton X-100 and mix gently.

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Add two drops of reagent B to this solution and mix immediately. Allow to stand for 30 min. 22. DAB solution (VECTOR): add two drops of buffer stock solution to 5 ml of distilled water and mix well. Add four drops of DAB stock solution and mix well. Add two drops of hydrogen peroxide solution and mix well. Alternatively, you can add two drops of Nickel stock solution and mix well to receive a black reaction product. 23. Haematoxylin QS: ready-to-use solution (VECTOR). 24. DePex mounting medium. 25. Ammonium chloride solution: add 0.5 g of NH4CI to 200 ml of TBS and stir until dissolved. 26. Mowiol: mix 6.0 g glycerol with 2.4 g Mowiol 4–88 and dissolve with frequent agitation for 1 h at RT. Add 6.0 ml distilled water and stir for one more hour at RT. Add 12.0 ml 0.2 M Tris–HCl (pH 8.5) and incubate for 2 h at 50°C under periodical stirring (every 20 min for 2 min). Note: in many cases Mowiol does not dissolve completely. We recommend centrifugation for 15 min at 5,000 × g. Continue with the supernatant. Add 25 mg/ml 1,4-diazabicyclo[2.2.2]octan (DABCO) and stir until complete dissolution. Aliquot 1 ml into 1.5 ml Eppendorf tubes. Add 1 ml DAPI stock solution (see below) and mix well. Store for long term at −20°C. Before using thaw at RT. 27. DAPI stock solution: dilute 4¢,6-diamidino-2-phenylindole (DAPI) at a concentration of 0.5 mg/ml in distilled water. Freeze 10 ml aliquots at −20°C.

3. Methods 3.1. Preparation of Mouse Knee Joint Paraffin Blocks

1. Dissect knee joints from the mouse, remove skin, and strip muscle as far as possible (do not cut into the joint!). 2. Wash 3 × 10 min in PBS by gently shaking at room temperature (RT). 3. Fix the samples in 2% PFA and 0.05% glutaraldehyde in PBS at RT for 1 h. 4. Wash 3 × 10 min in PBS by gently shaking at RT. 5. Decalcify samples: (a) In Decalcifying Solution-Lite (20:1 ratio for solution:tissue) at RT for 1 h; rinse extensively in tap water (this method is rapid, but destructive for a number of antigens). (b) In 4% EDTA in PBS at 4°C by gently shaking for 2 weeks (change solution every 2–3 days).

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6. Wash 3 × 10 min in PBS by gently shaking at RT. 7. Put the samples in 70% ethanol at 4°C and embed in paraffin wax using a tissue processor and standard wax protocol (see Note 2). 8. Cut 5-mm thick sections on a rotary microtome (Leica), float to stretch in a warmed water bath (45°C) and collect on Superfrost+ slides; allow the sections to dry overnight (this is important!) before starting with the staining protocol. 3.2. Immunohistochemical Staining Using EnzymeConjugated Antibodies

1. De-wax and rehydrate 5-mm thick paraffin sections using the following protocol: 2. 2 × 5 min xylene, 2 × 2 min ethanol 100%, 2 min ethanol 95%, 2 min ethanol 70%, and 5 min H2O. 3. Perform antigen retrieval using heat-mediated epitope retrieval (HIER) and/or proteolytic-induced epitope retrieval (PIER) (see Note 3). 4. Rinse 2 × 5 min in H2O. 5. Quench endogenous peroxidase with 3% H2O2 in H2O for 10 min (see Note 4). 6. Rinse 2 × 5 min in H2O. 7. Rinse in TBS for 5 min. 8. Rinse in washing buffer for 5 min. 9. Put one drop of avidin blocking solution on the section and incubate in a humidified chamber for 15 min (see Note 5). 10. Rinse in washing buffer for 5 min. 11. Block with one drop of biotin blocking solution in a humidified chamber for 15 min. 12. Block with blocking solution for 45 min (see Note 6). 13. Blot the excess blocking solution off, but do not allow to dry and do not wash. 14. Incubate with primary antibody diluted in blocking solution at RT for 1 h or at 4°C overnight (or in case of IdU staining in DNase solution at RT for 1 h); cover section with a small piece of parafilm to prevent evaporation (see Note 7). 15. Rinse 3 × 5 min in washing buffer. 16. Incubate with biotinylated secondary antibody at RT for 30 min (see Note 8). 17. Rinse 3 × 5 min in washing buffer. 18. Incubate with ABC reagent for 30 min (see Note 9). 19. Rinse 3 × 5 min in washing buffer.

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20. Incubate with peroxidase substrate solution for 2–12 min; monitor development of the staining under a microscope (see Note 10). 21. Rinse in tap water for 5 min. 22. Counterstain with Haematoxylin QS for 5 s. 23. Rinse in tap water until water is colourless. 24. Dehydrate using the following protocol: 3 min ethanol 70%, 2 × 3 min ethanol 100%, and 2 × 3 min xylene. 25. Mount with DePex and apply coverslip. 26. Sections can be analysed after polymerisation of the mounting medium (usually overnight) using a brightfield microscope and can be stored long term at room temperature. 3.3. Immunofluorescence Staining Using FluorochromeConjugated Antibodies

1. De-wax and rehydrate 5-mm thick paraffin sections as described in Subheading 3.2 step 1. 2. Perform antigen retrieval as described in Subheading 3.2 step 2. 3. Rinse in H2O for 5 min. 4. Rinse in TBS for 5 min. 5. Quench autofluorescence 2 × 5 min with TBS containing 50 mM NH4CI (see Note 11). 6. Wash 2 × 5 min in washing buffer. 7. Block with blocking solution at room temperature for 45 min. 8. Blot the excess blocking solution off but do not allow to dry and do not wash. 9. Incubate with primary antibody diluted in blocking solution for 1 h at RT or at 4°C overnight (or in case of IdU staining in DNase solution for 1 h at RT). Apply a small piece of parafilm over the section to prevent evaporation. 10. Wash 3 × 5 min in washing buffer. 11. Incubate with fluorochrome-conjugated secondary antibody at RT for 30 min. From here on incubation should be performed in the dark; avoid exposing the sections too long to light sources as this might affect the fluorescence intensity of the secondary antibody. 12. Wash 3 × 5 min in washing buffer. 13. Mount with Mowiol containing DAPI. 14. Sections can be analysed after polymerisation of the mounting medium (usually overnight) under a fluorescent microscope and can be stored long term at −20°C. A series of website that list useful information about reagents and procedures for IHC and IF is given in Note 12.

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4. Notes 1. The process of fixation prevents the decay of the tissue that might happen through intrinsic components like endogenous proteolytic enzymes or extrinsic factors like bacteria. Frozen samples are usually not fixed. However, a post-fixation step on the sections is performed by using rather short times of incubation in formaldehyde solutions, acetone, or methanol. Whereas formaldehyde preserves and strengthens structure within the tissue, it can result in tissue shrinkage and alcoholic fixatives may destroy morphological details like nuclei. Among the many fixation agents that are described in detail in histology textbooks (3), the mostly commonly used is aldehyde-based crosslinking. Usually samples are immersed in a 3.7% formaldehyde solution (also called 10% Neutral Buffered Formalin) for at least 1 h. The often stated risk of potential over-fixation of tissue leading to compromised immunohistochemical staining was recently tested and found to be small when tissue showed good immunoreactivity even after 7 weeks of fixation (4). The advantages of formaldehyde are its fast penetration and the possibility to store samples for long term. By contrast glutaraldehyde penetrates more slowly, but preserves cellular morphology better. For immunostaining formaldehyde generally gives better results, but for both reagents antigen retrieval has to be considered (see Note 3). We prefer a mixture of both aldehyde fixation agents as a compromise between preservation of tissue and cellular details, penetration speed, and the quality of immunohistochemical signals. 2. Embedding samples in paraffin and sectioning paraffin blocks is best done using tissue processors which eliminate exposure to solvents. This equipment can be found in all Pathology Departments or in most Histology facilities. You may of course use manual methods, but beware of solvents and use a chemical fume hood. Methods for embedding can be found in all Histology text books, for example in “Theory and Practice of Histological Techniques” (3). 3. Antigen retrieval: a detailed overview of antigen retrieval methods was recently published by D’Amico et al. (5). Two general principles are HIER and PIER. The choice between these methods depends on the fixation status of the antigen, the primary antibody to be used and the tissue of interest. For example, detection of antigens in synovial tissue might be facilitated by PIER as this tissue is rich in fibrous extracellular matrix which has to be opened by the enzyme. The same antigen in bone marrow, however, might be destroyed by PIER and HIER should be the method of choice in this case. In HIER,

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sections are boiled in a specific buffer using various devices such as microwave oven, steamer, pressure cooker, autoclave, or water bath. The length and the temperature of the boiling step are crucial and should be evaluated for each antibody used. Also the choice of buffer is important and may vary between antibodies. Two mostly used buffers are citrate buffer of pH 6 and Tris–EDTA buffer of pH 9 (these buffers are available in consistent quality from a broad range of suppliers like DAKO or Vector, but can also be prepared fresh as detailed in Subheading 2). In our laboratory, we used an autostainer (Bone Max, Leica) to perform HIER. This staining robot performs automated de-waxing and rehydration steps and can be programmed to heat sections at 99°C for 10–30 min using either the buffer of pH 6 or 9. The advantage of this staining robot is that the temperature is consistent over the whole section and that the section stays in a horizontal position which prevents it from floating off the slide. However, remember that the process of boiling is destructive to joint sections. Especially, articular cartilage experiences a high degree of shrinkage and sections tend to detach from the slide. Even with all these artefacts, HIER is less destructive to the antigens than PIER and results in good signal quality. Enzymes used in PIER include pepsin, trypsin, Proteinase K or pronase, or hyaluronidase (see Fig. 1a). As mentioned above the destructive nature of the enzyme can affect antigens and even tissue morphology. A careful evaluation of incubation time and concentration of the protease is therefore crucial. Also the solution in which the enzyme is diluted is of importance. Pepsin, for example works only in a high acidic environment that might irreversibly denature antigens. However, in our laboratory we used 15- and 45-min incubation with porcine pepsin (0.5–3 mg/ml) successfully to detect a range of antigens. Antigen retrieval for nucleoside analogues such as IdU requires harsh antigen retrieval. We obtained very good signals when we used a pepsin solution; the low pH and the proteolytic activity combined, helps to denature the DNA and to remove nuclear proteins, to result in single-stranded DNA. Additionally, we diluted the primary antibody in a DNase solution (see Subheading 2, item 19) and incubated for 1 h at RT. The enzyme cuts the DNA strands into small pieces and the access of the primary antibody to the nucleoside is greatly facilitated resulting in excellent staining with very good signal to noise ratio. 4. If the enzyme peroxidase is used in the detection system, the tissue of interest should be tested for endogenous peroxidase expression. This is done by de-waxing and re-hydrating your section and then applying a drop of peroxidase substrate solution like DAB (see Note 10). If endogenous peroxidase activity

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is detected this can be blocked by incubating the section in 3% hydrogen peroxide in water for 10–30 min. Some people use 0.3% hydrogen peroxide in methanol, but methanol affects nuclear morphology and therefore we would not recommend this method. We use this blocking step after de-waxing and rehydration and wash 2 × 5 min with water to remove hydrogen peroxide residues. Blocking of endogenous peroxide can be done later in the protocol as long as it is done before the peroxidase-conjugated reagent is introduced. 5. The use of the avidin–biotin system greatly enhances signals, but endogenous biotin can cause problems with false-positive signals. VECTOR offers a special kit to prevent such problems by blocking the endogenous biotin (www.vector.com). 6. Blocking solutions are used to inhibit specific or non-specific background staining. These false-positive signals occur generally through binding of the antibody as a consequence of electrostatic forces within the tissue. To prevent this, a blocking solution with high protein content can be applied to the section to cover non-specific binding sites for antibodies. Frequently used is a blocking solution that contains up to 20% of serum from the species in which the secondary antibody is produced. In our laboratory, we successfully block with a 1% BSA solution. A special case arises when you use an antibody that is raised in the species as the test tissue. This happens most frequently when using mouse monoclonal antibodies on mouse tissues. The use of the Mouse-On-Mouse-Kit (VECTOR) can be helpful in this case. In the first step, sections are incubated with a mouse Ig blocking reagent for 1 h and then, after a quick wash, sections are additionally incubated in a mouse protein cocktail for 5 min. This treatment, in our experience, results in highly specific staining. 7. The choice of a good primary antibody is sometimes a challenge. We usually start by checking published data on antibodies used for the antigen we wish to detect. Another source of information are the datasheets (helpfully available online for most suppliers) in which suppliers may state whether the antibody can be used on frozen or paraffin embedded, formalinfixed sections. Comparison websites can help source the companies that make antibodies to the antigen of your choice (e.g. www.biocompare.com). Monoclonal antibodies give highly specific signals and are low in background staining, but are often sensitive to fixation and paraffin embedding which might change conformation of the antigen. By contrast, polyclonal antibodies are more robust, but can give false-positive signals due to presence of irrelevant antibodies in the immunised animal. An appropriate control in this case would be preimmune serum, but this is not always available. The antibody

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concentration has to be determined empirically using a range of concentrations spanning the optimal dilution recommended by the supplier. This is necessary because the antigen of interest may, in your test situation, be located in different tissue compartments, or you may wish to use different antigen retrieval methods, or the abundance of antigen is quite different than in the supplier’s test situation. Although antibodies can be used for years if stored correctly (do read the instruction from the supplier and abide by them), their affinity may decline over time. If you expect this is the case, do not throw out the antibody just yet, but re-optimise the optimal dilution. For each staining a positive and a negative control should be used. The positive control should be a section similarly prepared as the experimental sample, i.e. identical fixation and embedding protocol. This could be a different tissue, or even better, a different organ/tissue compartment inside the same section which is known to express the antigen of interest (internal positive control). However, remember that different tissues may require different antigen retrieval methods or primary antibody dilutions to reveal the antigen so as to optimise the conditions for the positive control properly. The negative control sections of your experimental sample should ideally be incubated with an Ig control antibody raised in the same species and containing the same Ig subclass as your primary antibody (isotype negative control). For polyclonal antibodies, pre-immune serum should be used as negative control. Concentration of Ig (NOT dilution of antibody) should be the same as for the optimised primary antibody. If the concentration of the primary antibody is unknown, an option is to omit the first antibody entirely and replace with incubation in wash buffer alone. Non-specific binding of an antibody to proteins other than the antigen can sometimes occur. To determine whether the staining is specific, a blocking experiment with an immunising peptide (usually available from the commercial supplier of the antibody) can be performed. Prior to staining, the primary antibody is neutralised by incubation with an excess of peptide that corresponds to the epitope recognised by the antibody. The antibody that is bound to the blocking peptide is no longer available to bind to its epitope. Therefore, specific staining will be absent in the immunostaining performed with the neutralised antibody. 8. The secondary antibody is raised against the species from which the primary antibody is generated. Secondary antibodies are usually tagged by conjugation to a fluorochrome or an enzyme. Secondary antibodies can also be tagged with biotin and then be used with signal enhancement systems such as the

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ABC-complex. If double staining is performed, a mix of the secondary antibodies can only be used if none of the secondaries is raised in the same species than one of the primary antibodies. For example, if one of the primary antibodies is raised in goat, a mix of two secondary antibodies that includes one that is raised in goat would lead to cross-reactivity between the secondaries. In this case, secondary antibodies should be applied sequentially with an intermediate blocking step using goat serum: first antibodies → wash → anti-goat secondary antibody → wash → blocking with goat serum → secondary antibody raised in goat. In the same way as for the primary antibody, optimal concentrations for secondary antibodies should be determined empirically using datasheets for guidance. 9. Amplification systems greatly enhance signal strength and can be used in enzyme-based IHC. Avidin–biotin conjugates that are coupled to peroxidase molecules exist as large polymers and can bind to biotinylated secondary antibodies. As a consequence multiple enzyme reaction sites are offered in contrast to only single sites when the enzyme is directly conjugated to the secondary antibody. A similar signal amplification can be obtained in IF staining, when the tyramide amplification system from Perkin Elmer is used (6). Here, a secondary antibody conjugated with peroxidase is used to react with a substrate that deposits fluorophore-labelled tyramide. This method has the potential problem of giving a high specific background if the method is not carefully optimised. Therefore, always include a negative control (by omitting primary antibody) and a control without TSA. The tyramide reaction itself is very quick (less than 10 min). 10. If peroxidase is used as a detection enzyme in IHC, 3,3¢-diaminobenzidine (DAB) is the most widely used substrate which gives a brown or by adding a nickel solution a black reaction product. It is very sensitive, the reaction is very fast (sometimes within seconds; recommended incubation times are 2–12 min; monitor the staining under a light microscope to prevent overstaining), gives good contrast and easy to use as many suppliers offer the solution in kit format. However, handling of DAB must be done with caution as it is known to be a carcinogen. VECTOR now offers other substrates that are also very good in contrast like NovaRed (red reaction product) or VIP (violet reaction product) or can be used like AEC (red reaction product) with aqueous mounting media. Whereas the counterstain with haematoxylin gives a good contrast to DAB, these other substrates might benefit from being used with different nuclear counterstains such as Methyl Green. Vector provides a useful chart which substrate is best combined with which counterstain (www.vector.com).

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11. Autofluorescence is an artefact that is not easy to deal with. If the experiment consists of an immunohistochemical staining for a single antigen we recommend non-fluorescent methods as an option. However, some experiments require double or triple staining and co-localisation of these signals is important to show that two antigens are simultaneously expressed by the same cell or in the same compartment of that cell. Some autofluorescent signals can be spotted readily in the microscope and distinguished from real signals. For example, using a long-pass green emission filter, green fluorescent tags like GFP or Alexa Fluor 488 appear in bright green colour and not yellowish like autofluorescent structures. It is, however, challenging when it comes to take pictures. Autofluorescence might be caused by fixation agents which contain aldehydes that react with amines and proteins and therefore create autofluorescent structures. Proper antigen retrieval can reduce these artefacts, but cannot avoid them. Other sources of autofluorescene are biochemical molecules like lipofuscin, a break-down product of red blood cells, collagen, or elastin. A strong source of autofluorescence which is seen in bone marrow or sites of bleeding, are red blood cells due to the porphyrin structure of haemoglobin. They give strong signals in the excitation spectrum of blue and green lasers. Red blood cells can be easily spotted after nuclear counterstain (e.g. DAPI) as they are devoid of a nucleus. We observed that erythrocyte autofluorescence disappeared after pepsin treatment for antigen retrieval while it persisted after HIER. Other methods that have been used to block autofluorescence are treating sections with agents like Sudan Black in 70% ethanol or copper sulphate in ammonium acetate buffer (7). However, it has to be noted that such treatments may also reduce the fluorescent signal of the staining. Other approaches to reduce autofluorescence are the incubation in ammonium chloride or pretreatment of sections for some hours by irradiation with UV light to photobleach autofluorescent structures (8). 12. The following websites provide useful information about IHC and IF and reagents: http://jhc.sagepub.com/ – Official Journal of the Histochemical Society. www.ihcworld.com – Webpage around IHC with background information, technical support, and also a store that offers a range of products including antibodies. www.proteinatlas.org/ – Database showing staining for a wide range of antibodies directed against human antigens in different tissues. http://www.biocompare.com/ProductCategories/ 2045/Antibodies.html – Good webpage that lists antibodies of different suppliers.

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http://dshb.biology.uiowa.edu/ – Developmental Studies Hybridioma Bank (University of Iowa); offers monoclonal antibodies for use in research at reasonable prices. www.vector.com – Supplier specialised in IHC. www.dako.com – Supplier with a range of IHC products including antibodies used in diagnostics

Acknowledgements The authors are grateful for the support from Arthritis Research UK (grants 19271 and 19429). Professor De Bari is a Fellow of the Medical Research Council, UK (grant G108/620). References 1. Coons, A. H., Creech, H. J., Jones, R. N., and Berliner, E. (1942) The demonstration of pneumococcal antigen in tissues by the use of fluorescent antibody. J. Immunol. 45, 159–170. 2. Kurth, T. B., Dell’Accio, F., Crouch, V., Augello, A., Sharpe, P. T., and De Bari, C. (2011) Functional mesenchymal stem cell niches in adult mouse knee joint synovium in vivo. Arthritis Rheum. 63, 1289–1300. 3. Bancroft, J. D., and Gamble, M. (2001) Theory and Practice of Histological Techniques. Churchill Livingstone, Philadelphia. 4. Webster, J. D., Miller, M. A., DuSold, D., Ramos-Vara, J. (2009) Effects of Prolonged Formalin Fixation on Diagnostic Immunohistochemistry in Domestic Animals. J. Histochem. Cytochem. 57, 753–61.

5. D’Amico, F., Skarmoutsou, E., and Stivala, F. (2009) State of the art in antigen retrieval for immunohistochemistry. J. Immunol. Meth. 341, 1–18. 6. Liu, G., Amin, S., Okuhama, N. N., Liao, G., and Mingle, L. A. (2006) A quantitative evaluation of peroxidase inhibitors for tyramide signal amplification mediated cytochemistry and histochemistry. Histochem. Cell Biol. 126, 283–291. 7. Schnell, S. A., Staines, W. A., and Wessendorf, M. W. (1999) Reduction of lipofuscin-like autofluorescence in fluorescently labelled tissue. J. Histochem. Cytochem. 47, 719–730. 8. Neumann, M., and Gabel, D. (2002) Simple method for reduction of autofluorescence in fluorescence microscopy. J. Histochem. Cytochem. 50, 437–9.

Chapter 22 Techniques for the Study of Apoptosis in Bone Sudeh Riahi and Brendon Noble Abstract There has been great interest in the identifying the mechanisms by which apoptosis is regulated in bone over recent years and in the biological role that this process plays in bone metabolism and bone disease. Here, we describe several methods for the detection of apoptosis in bone sections and in bone cell cultures. Key words: Apoptosis, Osteocyte, Bone, Caspase, Nick translation

1. Introduction Apoptosis or programmed cell death plays a key role in normal physiology and in various pathological processes, such as cancer and inflammation. The first descriptions of apoptosis were based on morphological changes in cells, such as shrinkage, condensation and margination of chromatin nuclear fragmentation, and production of membrane-bound apoptotic bodies. While these criteria are still regarded as the “gold standard” for identifying apoptotic cells, additional markers of apoptosis have been identified more recently, such as activation of caspase enzymes, and expression of phosphatidylserine on the external aspect of the cell membrane (1). The loss of cells through apoptosis has wide-ranging effects on all body tissues and bone is no exception. Apoptosis is known to play a role in endochondral ossification (2), in regulating new bone formation (3), and osteoclastic bone resorption (4). Osteocyte apoptosis has been suggested to play a role in targeting of bone remodelling (5) by release of osteocyte apoptotic bodies which promote osteoclastic bone resorption (6). There is also evidence to suggest that apoptosis is perturbed or deregulated in a number of bone diseases (7–9).


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Studies of apoptosis in bone are technically demanding due to the mineralised nature of the tissue. Under normal circumstances, bone needs to be embedded in material, such as methyl methacrylate prior to sectioning but this greatly hinder penetration of the tissue by the enzymes, antibodies, and other reagents that are required to detect apoptosis. Although it is possible to remove the embedding material with a deplasticising step, this can result in the loss of small fragments of DNA associated with apoptosis, thereby interfering with the detection of apoptosis. In view of this, apoptosis is best studied in freshly cut cryosections of bone which can be prepared using a tungsten carbide edged knife and a heavy-duty cryostat. Here, we describe several methods for assessing apoptosis in bone, including morphological detection of apoptotic cells by toluidine blue staining; assessment of cell viability using the lactate dehydrogenase (LDH) assay, assessment of DNA fragmentation by the nick translation assay and gel electrophoresis, and by the assessment of caspase activity in bone sections in situ. 1.1. Toluidine Blue Staining

Apoptotic cells can be identified in sections of bone by toluidine blue staining. This relatively straightforward procedure stains the nuclei blue allowing the researcher to visualise features of apoptosis by light microscopy, such as nuclear condensation, blebbing, and fragmentation.

1.2. LDH Assay

Cells maintain intact cell membranes and active metabolic processes throughout most of the apoptotic process. This is in distinct contrast to necrotic death, where the cell membrane ruptures and metabolic activity rapidly declines. Hence, detection of the DNA fragmentation in cells with intact membranes and active metabolic enzymes will indicate apoptosis rather than necrosis. Loss of cell viability also represents the final “outcome” of the apoptotic process and in the case of the osteocytes, which are entombed within lacunae in bone, this is indicative of cell death, although it does not discriminate between apoptosis and necrosis. In order to assess cell viability in cryosections of bone, we have developed techniques to detect LDH enzyme activity histochemically. The technique is highly sensitive to ensure that any active enzyme present is identified. Because apoptosis occurs only in living cells, the LDH assay can be used alongside other techniques, such as DNA laddering or nick translation to determine which cells were alive when the tissue was prepared for sectioning.

1.3. Nick Translation

The technique of nick translation uses DNA polymerase I to incorporate DIG-conjugated nucleotides into DNA strand breaks to identify cells in culture or in tissue sections that contain large amounts of fragmented DNA (Fig. 1). The technique has been purposely designed to be relatively insensitive to small amounts of DNA fragmentation that might be present in normal cells or necrotic cells,

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Fig. 1. Cells containing large amounts of highly fragmented DNA are labelled using a nick translation technique. Fragmented DNA in osteocytes resident in bone is identified after incorporation of labelled nucleotides using a nick translation reaction. (a) Propidium iodide staining of osteocyte nuclei. (b) Apoptotic osteocytes labelled positive for fragmented DNA (FITC). Arrows denote two example cells positive for fragmented DNA. (c) PI staining of osteocyte nuclei in the negative control (no polymerase enzyme). (d) Negative control (no polymerase enzyme) showing lack of FITC-labelled cells.

and hence has high specificity for apoptosis (7). This technique provides a more consistent method of identification of apoptosis than the more commonly used Terminal deoxynucleotidyl transferase dUTP Nick End Labelling (TUNEL) staining, which employs terminal deoxynucleotide transferase (TdT). This might be due to the fact that TUNEL greatly amplifies the fragmentation signal due to the addition of multiple labelled nucleotides at 3¢ termini of a break, whereas nick translation adds only a single nucleotide. The TUNEL method also includes a proteinase K digestion step which might cause positive results due to small levels of DNAse contamination in the proteinase K. 1.4. DNA Laddering

Fragmentation of DNA into nucleosomal sized fragments of 180– 200 base pairs is a hallmark of apoptosis and leads to the production of a “DNA ladder” when the DNA is analysed on an agarose gel (10). Some cells produce much larger DNA fragments during apoptosis (200–300 and 30–50 kbp) (11). It is thought that these are produced as a prelude to the production of nucleosomal fragments and that apoptotic cells not showing DNA ladders have stopped DNA fragmentation at this earlier stage in the process. It is possible to identify these larger fragments using pulse field electrophoresis, but this is not covered in this chapter.

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1.5. Caspase Activation

Caspases or cysteine-aspartic proteases are activated in response to proapoptotic signals and cause the cleavage of protein substrates that eventually lead to disassembly of the cells (12, 13). Based on their actual or predicted roles, caspases are divided into two subgroups, initiator caspases (caspase-2, -8, -9, and -10) and effector caspases (caspase-3, -6, and -7) (14). Effector caspases are thought to be responsible for demolition of the cell during apoptosis. Caspase-9 simultaneously activates caspase-3 and -7 (15). Caspase-3 is required for the activation of four other caspases in this pathway (-2, -6, -8, and -10) and is also engaged in a feedback loop involving caspase-9 (15). Here, we describe the used of the Image-iT™ LIVE Green Caspase-3 and -7 Detection Kit (Invitrogen) to detect active caspase 3-7 in bone cells.

2. Materials 2.1. General Materials

1. Phosphate-buffered saline (PBS). 2. Paraformaldehyde: 4% (w/v) paraformaldehyde in PBS (see Note 1). 3. DAPI: 4¢, 6-diamidino-2-phenylindole dehydrochloride 4 μg/ ml in PBS. 4. Propidium Iodide (PI): Propidium iodide 1 μg/ml in PBS (see Note 2). 5. Fixogum adhesive or clear nail varnish. 6. Fluorescent mounting medium (DAKO). 7. Cover slips.

2.2. Toluidine Blue Staining

1. Picric formalin: 6% (v/v) Formalin (40% aqueous formaldehyde), 50% (v/v) ethanol 95%, and 4% (v/v) glacial acetic acid in distilled water (see Note 3). 2. Toluidine blue solution: 0.1% (w/v) toluidine blue in distilled water. 3. n-butyl alcohol. 4. Citifluor (Agar Scientific). 5. Light green: 1% (w/v) light green in distilled water.

2.3. LDH Assay

1. Polypep solution: 40% Polypep (w/v), in 0.1 M diglycine, and 17 mM NaOH (pH 8.0) (see Note 4). 2. Reaction mix: 0.175% (w/v) nicotinamide adenine dinucleotide and 0.3% (w/v) nitroblue tetrazolium in 60 mM lactic acid (pH 8.0) (see Note 5).

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3. 10 M NaOH. 4. Polymethylmethacrylate plastic rings, 10–15 mm in diameter (see Note 6). 5. Vaseline. 2.4. Nick Translation

1. Decalcification solution: 0.25 M EDTA in 50 mM Tris–HCl, pH 7.4. 2. Digoxigenin-11-dUTP, alkali-labile (DIG-11 dUTP). 3. Nick translation buffer without DNA polymerase: 3 μM dATP, 3 μM dCTP and 3 μM dGTP and 0.08 nM DIG-11 dUTP, 50 mM Tris–HCl, 5 mM MgCl2, 0.1 mM dithiothreitol, (pH 7.5). 4. Nick translation buffer with DNA polymerase: 3 μM dATP, 3 μM dCTP and 3 μM dGTP and 0.08 nM DIG-11 dUTP, 50 mM Tris–HCl, 5 mM MgCl2, 0.1 mM dithiothreitol (pH 7.5) containing 0.5% (v/v) DNA polymerase 1. 5. Anti-DIG FITC fab mix: 11% (v/v) sheep anti-digoxigeninfluorescein (FITC), fab fragments, and 4% sheep serum in PBS. 6. DNAse I solution: 0.2% DNase 1 in PBS.

2.5. DNA Laddering

1. Nucleon tissue DNA extraction kit (Nucleon Biosciences, Glasgow, UK). 2. Ribonuclease A. 3. Chloroform. 4. 100% ethanol. 5. 3 M sodium acetate. 6. Tris–Borate EDTA (TBE): 89 mM Tris base, 89 mM boric acid, and 2 mM EDTA, in distilled water (pH 8.0). 7. Agarose. 8. Loading buffer: 0.25% (w/v) bromophenol blue, 0.25% (w/v) xylene cayanol FF, 30% (v/v) glycerol in distilled water (see Note 7). 9. 100 and 1,000 (bp) DNA ladder (Gibco). 10. Horizontal gel electrophoresis tank and power supply delivering up to 150 V. 11. UV transilluminator.

2.6. Caspase 3-7 Detection

1. Image -iT™ Live Green caspase-3 and -7 detection kit (Invitrogen). 2. 30% (v/v) hydrogen peroxide.

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3. Methods 3.1. Detection of Apoptosis by Toluidine Blue Staining

The procedure stains the nuclei blue and enables visualisation of condensation, blebbing, or fragmentation of the nucleus prior to packaging of nuclear and cytoplasmic contents into apoptotic bodies. 1. Fix cryostat sections in picric-formalin for 10 min at room temperature. 2. Add toluidine blue to the sections and incubate for 30 min. 3. Blot dry and place in PBS for resin sections or n-butyl alcohol for frozen sections for 2 min (see Note 8). 4. If a counterstain is required, add 1% light green to the section, incubate for 2 min and rinse with distilled water. 5. Mount in Citifluor (see Note 9).

3.2. Assessment of Cell Viability Using the LDH Assay

The method described is a modification of the methods of Wong et al. (8) and Farquharson et al. (16). Purple staining indicates viable cells; the absence of staining implies a dead cell or the presence of an empty lacuna (Fig. 2). 1. Prepare cryostat sections 10 μm thick from the tissue to be analysed, mount on microscope slides and store at −20°C or −80°C. 2. Defrost the slides at the room temperature for few minutes. 3. Place plastic rings onto the slides using Vaseline and add about 400 μl reaction mix to each the ring, making sure that are bubbles are excluded. 4. Smear some Vaseline on cover slips and place on top of the rings to prevent the reaction mix evaporating. 5. Incubate for 3 h at 37°C in a humidified chamber. 6. Carefully remove the rings and Vaseline and then rinse in warm water at approximately 50°C. 7. Rinse in acetone for 30 s. 8. Fix sections in 4% paraformaldehyde for 10 min. 9. Wash the sections three times in PBS. 10. Add DAPI to each section and incubate for 10 min at room temperature. 11. Wash the sections three times in PBS. 12. Mount slides in DAKO fluorescent mounting medium. 13. Add cover slips and seal with clear nail varnish or Fixogum adhesive.

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Fig. 2. Cell viability determined in situ using LDH activity as a marker. Cells in frozen sections are stained for lactate dehydrogenase activity and examined using light microscopy. (a) Live osteocytes stained dark for active lactate dehydrogenase. Arrows illustrate two examples of live cells. (b) Region of bone containing dead osteocytes showing no staining for LDH staining. Cells on bone surface stain positive. (c) Diagram illustrates the use of plastic rings for LDH staining. The reaction mix is placed in a plastic ring sealed at the base and top with Vaseline to allow prolonged incubation at 37°C.

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3.3. Detection of Fragmented DNA Using Nick Translation 3.3.1. Preparation of Cells and Sections

This method can be used for detection of fragmented DNA in cryostat sections, cultured bone cells or cytospin preparations. 1. Prepare fresh 7–10 μm cryostat sections of bone (see Note 10). 2. Fix the sections or cells in 4% paraformaldehyde in PBS for 10 min at room temperature (see Note 11). 3. If sections are being studied, immerse the slides in decalcification buffer for 10 min. 4. Wash the slides three times in PBS. 5. Allow the sections or cells to dry thoroughly and store at 4°C until ready to use.

3.3.2. Nick Translation Assay

1. Add 50 μl DNase I solution to one section and incubate for 1 h at 37°C as a positive control. 2. Add 50 μl nick translation buffer without DNA polymerase to one section as a negative control and incubate for 1 h at 37°C in a humidified chamber. 3. Incubate all other sections (including the positive control from step 1) with nick translation buffer containing DNA polymerase, for 1 h at 37°C in a humidified chamber. 4. Wash three times in PBS, taking care to keep the sections moist at all times. 5. Add sufficient anti-DIG FITC fab mix to cover each section and incubate for 1 h at room temperature. 6. Wash three times in PBS. 7. Counterstain the nuclei by adding propidium iodide to each section for 2 min (or 30 s if cells are being studied). 8. Wash thoroughly in distilled water. 9. Mount in DAKO fluorescent mounting medium and keep in the dark. 10. Analyse the sections or cells with a fluorescent microscope to distinguish FITC-stained apoptotic cells (green) from all cells (stained red with PI) (see Note 12).

3.4. Detecting of DNA Laddering in Cells and Tissue Sections

All bone cells that we have tested so far (from various species) yield DNA ladders when undergoing apoptosis, but it is wise to always include a positive control in the assay described below in which apoptosis has been induced by heating cultured cells to 44°C for 30 min, or by using sections of materials which have been shown to contain apoptotic cells by another technique, such as nick translation (Fig. 3).

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Fig. 3. DNA ladders indicative of apoptosis. DNA from apoptotic cells produces multiple bands of approximately 180-bp increments when run on an agarose gel. Lanes A, B: DNA from apoptotic cells producing characteristic “ladder” pattern. Individual bands are highlighted with arrows. Lane C: 1,000-bp markers.

3.4.1. Preparation of Cells for Analysis

1. Using a confluent T75 flask of bone cells (approximately 4 × 106 cells), aspirate medium and wash the monolayer gently in PBS. 2. Aspirate to dryness, and place in a −80°C freezer immediately, for a minimum of 3 h. 3. Remove the flask from the freezer, add 1 ml of PBS, and scrape the cells into an Eppendorf tube. 4. Centrifuge at 600 × g, at 4°C for 5 min. 5. Remove the supernatant and proceed to extract DNA from the cell pellet as described in Subheading 3.4.3.

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3.4.2. Preparation of Sections for Analysis

1. Cut 15–20 separate bone sections of 10 μm thickness from the tissue of interest using a cryostat. 2. Transfer the sections directly into a Bijou and place immediately into a −80°C freezer until ready to analyse.

3.4.3. Isolation of DNA

The DNA isolation method described here is based on the use of a Nucleon DNA extraction kit. 1. Add 340 μl of reagent B to the cell pellet or sections. Vortex and incubate at room temperature for 40 min. 2. Centrifuge at 600 × g for 5 min. Decant the supernatant to another tube. 3. Add ribonuclease A to the supernatant to give a final concentration of 50 μg/ml and incubate for 30 min at room temperature. 4. Add 100 μl sodium perchlorate to each tube and transfer to a rotary mixer. Incubate at 37°C for 20 min followed by 65°C for 20 min. 5. Add 580 μl chloroform (stored at −20°C) to each tube and transfer to a rotary mixer for 20 min at room temperature. 6. Transfer the reaction to a 2-ml Nucleon tube. 7. Centrifuge at 1,300 × g for 1 min. 8. Add 45 μl of Nucleon silica suspension to each tube and mix well. 9. Centrifuge at 1,300 × g for 4 min. 10. Pour off the upper aqueous phase containing the DNA into a fresh tube. 11. Centrifuge at 1,300 × g for 30 s to pellet any remaining silica and transfer the supernatant to a fresh tube. 12. Add 880 μl of 100% ethanol to the supernatant and invert the tube to mix. 13. Centrifuge at 4,000 × g for 5 min to pellet the DNA and discard the supernatant (see Note 13). 14. Wash the DNA pellet by adding 1 ml of 70% ethanol to the tube and transfer to a rotary mixer at room temperature for 20 min. 15. Centrifuge at 4,000 × g for 5 min to collect the DNA pellet at the bottom of the tube. 16. Carefully aspirate the ethanol and leave the pellet to air dry. 17. Dissolve the DNA sample in TBE (see Note 14). 18. Run the samples through a 1% agarose gel in TBE with a 100 and 1,000 bp molecular weight markers in one lane and stain with ethidium bromide. 19. Analyse the gel under UV light for evidence of DNA fragments of the expected size (see Note 15).

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3.5. Detection of Caspase Activation

The method is based on a fluorescent inhibitor of caspase (FLICA™) (17) which consists of a fluoromethyl ketone (FMK) moiety that can react covalently with a cysteine and a carboxy fluorescein group (FAM) that acts as a fluorescent reporter (17). The FLICA reagent is thought to interact with the enzymatic reactive centre of activated caspase via the recognition sequence, aspartic acid-glutamic acid-valine-aspartic acid (DEVD) for the caspase-3 and -7 reagent, and then to attach covalently through the FMK moiety. The FLICA inhibitor is cell permeable and nontoxic and is an excellent reagent for visualising caspase activity in situ (17).

3.5.1. Preparation of Cells

1. Seed the cells into a 24-well tissue culture plate at a density of 5×103 cells and culture overnight. 2. Prepare several positive control wells by adding H2O2 concentration at concentrations ranging between 20 and 600 μM to individual wells, keeping the cells in culture for between 8 and 16 h with periodic monitoring for evidence of apoptosis (see Note 16). 3. Wash the positive control wells and test wells gently with culture medium three times (see Note 17). 4. Prepare 150 × FLICA reagent stock according to the manufacturer’s instructions and store in aliquots of 5 μl, store them at −20°C protected from light.

3.5.2. Preparing the Labelling Reagent

1. Prepare the FLICA reagent solution 150× concentrate as described in the manufacturer’s protocol. 2. Dilute the 150 × FLICA concentrate 1 in 5 in PBS to generate a 30 × stock solution. 3. Prepare about 5 ml of 1 × FLICA reagent in tissue culture medium by making a 1 in 30 dilution of the 30 × FLICA stock solution. 4. Add ~200 μl FLICA reagent to each well and incubate for 1 h at 37°C with the tissue culture plate wrapped in foil. 5. Remove the reagent using a syringe and wash the cells gently three times with culture medium. 6. Add propidium iodide solution to the medium to give a final concentration of 1 μg/ml. 7. Incubate the cells for 30 min at 37°C with the tissue culture plate wrapped in foil. 8. Wash cells three times with PBS. 9. Fix the cells in 4% paraformaldehyde for 15 min (see Note 18). 10. Wash the cells three times with PBS. 11. Counterstain the nuclei by adding DAPI 4 μg/ml to each well (see Note 19). 12. Wash the cells three times with PBS.

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Fig. 4. (a) Light-microscopic morphology of apoptotic and necrotic cells. Cells are treated with H2O2 as an apoptotic stimulus. Apoptotic cells are highlighted with arrows. Swollen necrotic cells are shown with triangles (scale bar 50 μm). (b) DAPI staining of treated cell. (c) Apoptotic cells determined using caspase 3-7 kit. Apoptotic cells stained positive for caspase 3-7 activity and showed both intracellular and nucleus staining (arrows). (d) Phase image of cells (scale bar 50 μm). (e) Untreated cells with H2O2 had no staining for caspase 3-7. ( f ) Phase and DAPI image of untreated cells. (g and h) negative control (no staining with caspase reagent).

13. Add a drop of DAKO fluorescent mounting media to each well and add a 13 mm cover slip. Keep the plates in the dark until analysis. 14. Analyse cells for evidence of caspase activation using fluorescent microscopy (Fig. 4).

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4. Notes 1. The solution can be gently heated to assist with dissolution, but do not exceed a temperature of 60°C. This should be made fresh on the day of assay or prepared in advance in aliquots and stored frozen at −20°C until use. 2. The stock solution is stable for 6 months at 4°C, if stored in the dark. Make a working solution by adding 10 μl of PI stock to 10 ml distilled water just before use (final conc. 1 μg/ml). 3. The solution keeps for about 4 months at room temperature. Make a fresh batch as if a cloudy precipitate forms. 4. Prepare by adding NaOH to the 0.1 M diglycine solution. Then, add the Polypep and heat to approximately 37°C with stirring so that the Polypep goes into solution. Store the stock solution at 4°C. Diglycine is also known as Gly–Gly and Glycyl–Glycine. 5. To make 10 ml of reaction mix, melt 40% Polypep in a jar surrounded by hot water and to 10 ml of Polypep add 44 μl of lactic acid, 17.5 mg of NAD, and then adjust pH to 8.0 with 10 M NaOH (~75 μl NaOH for 10 ml mixture gives a pH ~ 8.12) before adding 30 mg NBT. 6. The plastic rings are made by cutting thin cross-sectional slices of polymethylmethacrylate tubing with a scalpel. 7. The loading buffer is stable for 6 months at 4°C. 8. Resin embedded sections should be washed in buffer rather than n-butyl alcohol to avoid shrinkage and crinkling of the section. 9. The standard technique involves dehydrating the sections and mounting in DePeX which may or may not suit the material being stained, but we find that bone sections do not require this dehydration step. 10. Freshly cut sections and freshly prepared cells should always be used for nick translation. Defrosting frozen sections can induce damage of DNA yielding false-positive results. 11. If cultured cells are being studied, aspirate the culture medium from the tissue culture wells and add enough 4% paraformaldehyde to cover the cell layer. 12. Positive controls should show a large number of cells with fragmented DNA (FITC positive) and negative controls should not show any FITC-positive cells, with low background fluorescence in the bone. The nuclei can also be counterstained by adding DAPI to the sections or cells and incubating at room temperature for 10 min. In this case, the cell nuclei are stained blue rather than red as with PI.

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13. It is possible to add 100 μl 3 M sodium acetate to the ethanol waste at this point and leave at −20°C overnight to precipitate further DNA. 14. The DNA can be frozen stored at −20°C at this point and analysed by gel electrophoresis later. 15. The smallest amount of DNA in a single band that can reliably be detected with ethidium bromide is approximately 10 ng and about 60 pg with SYBR® Green stain. The maximum amount of DNA that can be run as a sharp, clean band is about 100 ng. Overloaded DNA results in trailing and smearing, a problem that will become more severe as the size of DNA increases. 16. Monitor the induction of apoptosis by checking cells under the light microscope periodically for morphological evidence of apoptosis (18) (Fig. 4). 17. Washing steps should be undertaken very gently as the cells are loose and can easily detach from the plate. We suggest using a syringe and blunt ended needle to take out the medium. 18. We have also used the fixative used in the Image –iT kit with good results. 19. Hoechst dye in PBS at a concentration of 1 μM can also be used to stain the nuclei. References 1. Wyllie, A. H., Kerr, J. F., and Currie, A. R. (1980) Cell death: the significance of apoptosis. Int. Rev. Cytol. 68, 251–306. 2. Stevens, H. Y., Reeve, J., and. Noble, B. S. (2000) Bcl-2, tissue transglutaminase and p53 protein expression in the apoptotic cascade in ribs of premature infants. J. Anat. 196, 181–191. 3. Jilka, R. L., Weinstein, R. S., Bellido, T., Roberson, P., Parfitt, A. M., and Manolagas, S. C. (1999) Increased bone formation by prevention of osteoblast apoptosis with parathyroid hormone. J. Clin. Invest. 104, 439–446. 4. Kameda, T., Ishikawa, H., and Tsutsui, T. (1995) Detection and characterization of apoptosis in osteoclasts in vitro. Biochem. Biophys. Res. Commun. 207, 753–760. 5. Verborgt, O., Gibson, G. J., and Schaffler, M. B. (2000) Loss of Osteocyte Integrity in Association with Microdamage and Bone Remodeling After Fatigue In Vivo. J. Bone Miner. Res. 15, 60–67. 6. Kogianni, G., Mann, V., and Noble, B. S. (2008) Apoptotic bodies convey activity capable of initiating osteoclastogenesis and local-

7.

8.

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ized bone destruction. J. Bone Miner. Res. 23, 915–927. Noble, B. S., Stevens, H., Loveridge, N., and Reeve, J. (1997) Identification of apoptotic changes in osteocytes in normal and pathological human bone. Bone 20, 273–282. Wong, S. Y. P., Evans, R.A., Needs, C., Dunstan, C., Hills, E., and Garvan, J. (1987) The pathogenesis of osteoarthritis of the hip: evidence for primary osteocyte death. Clin. Orthop. Rel. Res. 214, 305–312. Canalis, E., Mazziotii, G., Giustina, A., and Bilezikian, J. (2007) Glucocorticoid-induced osteoporosis: pathophysiology and therapy. Osteoporosis Int. 18, 1319–1328. Wyllie, A. H., (1980) Glucocorticoid-induced thymocyte apoptosis is associated with endogenous endonuclease activation. Nature 284, 555–556. Oberhammer, F., Wilson, J. W., Dive, C., et al. (1993) Apoptotic death in epithelial cells: cleavage of DNA to 300 and/or 50 kb fragments prior to or in the absence of internucleosomal fragmentation. EMBO J. 12, 3679–3684.

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12. Slee, E. A., Adrain, C., and Martin, S. J. (1999) Serial killers: ordering caspase activation events in apoptosis. Cell Death Diff. 6, 1067–1074. 13. Yuan, J., Shaham, S., Ledoux, S., Ellis, H. M., and Horvitz, H. R. (1993) The C. elegans cell death gene ced-3 encodes a protein similar to mammalian interleukin-1 [beta]-converting enzyme. Cell 75, 641–652. 14. Creagh, E., and Martin, S. (2001) Caspases: cellular demolition experts. Biochem. Soc. Trans. 29, 696–701. 15. Slee, E. A., Harte, M. T., Kluck, R. M. et al. (1999) Ordering the cytochrome c-initiated Caspase cascade: hierarchical activation of Caspases-2, -3, -6, -7, -8, and -10 in a

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Caspase-9-dependent manner. J. Cell Biol. 144, 281–292. 16. Farquharson, C., Whitehead, C. Rennie, S., Thorp, B., and Loveridge, N. (1992) Cell proliferation and enzyme activities associated with the development of avian tibial dyschondroplasia: an in situ biochemical study. Bone 13, 59–67. 17. Ekert, P.G., Silke, J., and Vaux, D.L. (1999) Caspase inhibitors. Cell Death Diff. 6, 1081–1086. 18. Mann, V., Huber, C., Kogianni, G., Collins, F., and Noble, B. (2007) The antioxidant effect of estrogen and Selective Estrogen Receptor Modulators in the inhibition of osteocyte apoptosis in vitro. Bone 40, 674–684.

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Chapter 23 Transmission Electron Microscopy of Bone Vincent Everts, Anneke Niehof, Wikky Tigchelaar-Gutter, and Wouter Beertsen Abstract This chapter describes procedures to process mineralized tissues obtained from different sources for transmission electron microscopy (TEM). Methods for fixation, resin embedding, staining of semi-thin sections and ultrathin sections are presented. In addition, attention will be paid to processing of cultured bone explants for TEM analysis. Key words: TEM, Ultrastructure, Bone, Mineral, Decalcification

1. Introduction Ultrastructural analysis of bone and other mineralized tissues like calcified cartilage and dentin is essential for the understanding of the cell–cell/cell–matrix interaction, composition and threedimensional organization of these tissues. A wide variety of techniques have been introduced to process such tissues. This chapter describes a few methods to process mineralized tissues obtained from different sources for ultrastructural analysis. In addition, attention will be paid to processing of cultured bone explants for electron microscopic analysis.

2. Materials Prepare all solutions containing fixative in a fume hood and use gloves. All compounds are very toxic and most are volatile.


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2.1. Fixative

4% formaldehyde and 1% glutaraldehyde in 0.1 M sodium cacodylate buffer (pH 7.4) (see Notes 1 and 2). 1. Heat 200 ml distilled water to 70°C. 2. Dissolve 40 g paraformaldehyde and add approximately 2.5 g sodium hydroxide pellets. Allow the solution to cool (the solution should be clear). 3. Add 21.4 g sodium cacodylate. 4. Add 40 ml 25% glutaraldehyde and fill up to 800 ml with distilled water. 5. Adjust the pH to 7.4 with 1 N HCl and fill up to 1,000 ml. 6. This solution should be stored at 4°C and new fixative should be prepared each week (see Note 1).

2.2. Osmium and Ferrocyanide Postfixative

1% osmium tetroxide and 1.5% potassium ferrocyanide (K4Fe(CN)6)⋅3H2O) in 0.1 M sodium cacodylate buffer (pH 7.4). Stock solutions should be stored at 4°C (see Note 3). 1. 2% OsO4 stock solution: Add 1 g OsO4 crystals (EMS, crystalline, highest purity 99.95%) to 50 ml double distilled water in a stoppered dark glass vial. Gently (!) shake the solution till the crystals are dissolved. Store the solution in the tightly closed vial at 4°C. To avoid blackening of the solution prior to its use, the vial has to be thoroughly cleaned with acetone to remove lipids (osmium is an excellent fixative for lipids!), washed in double distilled water, and dried. Use gloves and avoid any contact with the skin. 2. 0.2 M sodium cacodylate buffer: Dissolve 42.8 g sodium cacodylate in 900 ml distilled water. Adjust the pH to 7.4 and add distilled water to a volume of 1,000 ml. 3. 3% ferrocyanide stock solution: Dissolve 3 g potassium ferrocyanide in 0.2 M sodium cacodylate buffer. 4. Prior to use, mix one volume of 2% OsO4 solution with one volume of 3% ferrocyanide solution.

2.3. Osmium and Cacodylate Postfixative

1% osmium tetroxide in 0.075 M sodium cacodylate buffer (see Note 3). 1. 4% osmium tetroxide stock solution: Dissolve 1 g OsO4 crystals in 25 ml distilled water according to the method described above (Subheading 2.2, step 1). 2. 0.1 M sodium cacodylate buffer: Dissolve 21.4 g sodium cacodylate in 900 ml distilled water, adjust the pH to 7.4 and add distilled water to a volume of 1,000 ml. 3. Mix prior to fixation one volume of the 4% OsO4 solution with three volumes of 0.1 M sodium cacodylate buffer.

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2.4. Decalcification Solution

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1.9% glutaraldehyde and 0.15 M EDTA (Titriplex III, ethylenedinitrilo tetraacetic acid disodium salt dihydrate) in 0.06 M sodium cacodylate buffer. 1. Dissolve 38.53 g sodium cacodylate and 167.52 g Titriplex III in 2,000 ml distilled water. 2. Stir the solution and as soon as all Titriplex is dissolved (the solution should be clear) add 232 ml 25% glutaraldehyde. 3. Adjust the pH to 7.4, first by adding approximately 10 g sodium hydroxide pellets followed by adding 2 N sodium hydroxide. Add distilled water to a volume of 3,000 ml (see Note 1). This solution is stable for several months at 4°C.

2.5. Goldner’s Masson Trichrome

1. Dissolve 1.25 g hematoxilin in 100 ml 25% ethanol. 2. Dissolve 0.15 g light green SF yellowish and 0.2 ml glacial acetic acid in 100 ml distilled water. 3. Ponceau de xylidine stock: Dissolve 1 g ponceau de xylidine and 1 ml glacial acetic acid in 100 ml distilled water. 4. Acid fuchsin stock: Dissolve 1 g acid fuchsin and 1 ml glacial acetic acid in 100 ml distilled water. 5. Ponceau-acid fuchsin stock: Two parts of ponceau de xylidine stock (see step 3) with one part of acid fuchsin stock (see step 4). 6. Orange G stock: Dissolve 1 g Orange G in 100 ml distilled water. 7. Ponceau-acid fuchsin staining solution: One part of ponceauacid fuchsin stock (see step 5), one part of Orange G stock (see step 6) and eight parts of distilled water. 8. Dissolve 1.0 g phosphomolybdic acid hydrate in 100 ml distilled water. 9. Dissolve 2.5 g ferric chloride and 1 ml concentrated HCl in 99 ml distilled water. 10. Rinsing solution: 5.2 ml 96% acetic acid in 1,000 ml distilled water. 11. All staining solutions are stable for months and are stored at ambient temperature.

2.6. Methylene Blue

1. Dissolve 2 g methylene blue in 100 ml distilled water (solution a) 2. Dissolve 0.5 g Azure II in 50 ml distilled water (solution b). 3. Dissolve 2 g Borax (di-sodiumtetraborat-10-hydrat) in 100 ml distilled water (Solution c). 4. Mix solutions a:b:c = 2:1:1, and store at 4°C. Staining solution is stable for months. 5. Filter just before use.

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2.7. Von Kossa

1. Dissolve 0.5 g silver lactate in 100 ml distilled water. 2. Dissolve 0.5 g hydrochinon in 100 ml distilled water. 3. Dissolve 5 g sodium thiosulfate pentahydrate in 100 ml distilled water. All solutions are made fresh just before use.

2.8. Uranyl Acetate

1. Dissolve 0.35 g uranyl acetate in 10 ml double distilled water. Store at 4°C.

2.9. Lead Nitrate

1. Boil and cool 50 ml double distilled water. 2. Dissolve 1.33 g lead nitrate and 1.76 g trisodium citrate dihydrate in 30 ml water. 3. Shake for 1 min vigorously and shake it a few times during the next 30 min. 4. Add 8 ml 1 N NaOH and add cooled boiled water to a volume of 50 ml. Store at 4°C.

2.10. Epoxy Resin

Use gloves and a fume hood when preparing the stock of epoxy resin. 1. Mix under continuous stirring the resin components (Ladd Res. Industries, Burlington, Vermont), adding the next component when the previous one is completely dissolved. The components should be added in the following order: 100 g LX-112, 72.4 g DDSA, 40.4 g NMA, and 3.9 g DMP-30. 2. Stir very well for another 30 min and collect the mixed resin in small plastic vials (10 ml) with cap. 3. Store these vials at –80°C. The frozen vials can be kept at this temperature for a very long time (at least for 1 year). 4. Prior to embedding, warm an appropriate number of vials at ambient temperature. Open the vial only when the resin is at room temperature.

3. Methods 3.1. Perfusion Fixation of Animal Bones

1. Use a perfusion fixation system consisting of a perfusion pump or a bottle with a rubber cap hanging upside down at a height of approximately 50 cm above the working place. If a pump is used, the tube is inserted in a bottle with fixative. If a hanging bottle is used, a needle (0.8 × 40 mm) is fixed to a tube and inserted into the rubber cap of the bottle. In addition, inserting a second tube with a needle into the cap makes an air inlet. The other end of this tube is fixed to the side of the bottle, with its opening above the fluid level in the bottle. To the tube used for

23

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fixation a hypodermic needle (0.6 × 30 mm for small animals [e.g., young mice], 0.8 × 40 mm for larger animals) is fixed for insertion into the heart. A valve should be placed somewhere along the length of the tube that is used for fixation. 2. Anesthetize the animal and fix it on his back on a plateau. Open the belly and cut the thorax left and right from the sternum. Reflect the skin and open the thorax, expose the heart, and carefully cut the pericardium. 3. Fix the heart with two fingers (use well-fitting gloves) and insert the needle through the wall of the left ventricle. Open the tube valve (hanging bottle system) or switch on the perfusion pump (set to 2.5 ml/min). Wait a few seconds and cut the right atrium with a fine pair of scissors to let the perfusate escape. 4. The quality of fixation is checked by testing the stiffness of soft tissues like the lip, the bleaching of the liver and rigidness of the paws. After 5–10 min, the perfusion is stopped and the tissue samples are collected and stored in fixative. 3.2. Immersion Fixation of Animal Bones

1. After killing the animal and exposure of the bones of interest, dissect the bones and immerse them as quickly as possible in the fixative (freshly prepared 4% formaldehyde and 1% glutaraldehyde in 0.1 M sodium cacodylate buffer, pH 7.4). If bones are collected from larger animals, the bones should be cut into smaller pieces. Cutting is preferably done in fixative. (Bones of young mice can be fixed without further cutting). 2. Fix at ambient temperature for at least 4 h. After this, the tissue samples can be left in fixative overnight at 4°C. 3. Wash the sample in 0.1 M sodium cacodylate buffer. 4. Transfer to postfixative for 1 h (see Note 3). 5. Wash the sample in 0.1 M sodium cacodylate buffer. 6. Proceed with the embedding protocol (see Subheading 3.6 and following).

3.3. Immersion Fixation of Human Bone Samples

3.4. Immersion Fixation of Cultured Mineralized Tissues

Immersion fixation and processing of bone samples obtained from humans is similar to protocol 3.2. It is essential that the samples are immersed into the fixative as quickly as possible and that the size of the fragments is small. Try to keep a maximal thickness of approximately 3–5 mm. Cutting of the bones into smaller fragments has to be performed in fixative. 1. Collect the bone explants after the preferred culture period and place in fixative (see Note 4). 2. Leave at ambient temperature for at least 4 h (after this period, the fixed bones can be kept in the fixative or in buffer at 4°C for another day).

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3. Process the bones further with or without decalcification (see Subheading 3.5). Bones obtained from (very) young animals (e.g., mice

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