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

Primary Human Osteoblast Cultures

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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 <0.2 g. 5. In early studies, osteoblastic phenotype was assessed using histochemical staining for alkaline phosphatise and secretion of osteocalcin. However, many researchers now prefer to undertake qPCR for ostoblastic genes (25). 6. We generally perform studies on cells at first passage (P1), but we have previously shown that cells isolated from replated explants cultures (E1P1, E2P1, and E3P1) are phenotypically stable with no reduction in the expression of osteoblastic markers (11). Furthermore, we have not detected any loss of phenotype at P2. Other investigators have studied the effects of repeated subculture on the phenotypic stability of HOBs and found that they eventually lose their osteoblast-like characteristics with successive passages. 7. Bone cells can be cultured from virtually any site, but an excellent source is the upper part of the femur of patients undergoing total hip replacement surgery for osteoarthritis. Trabecular bone is removed from this site prior to the insertion of the femoral prosthesis and would otherwise be discarded. The tissue obtained is remote from the hip joint itself, and thus from the site of pathology, and is free of contaminating soft tissue. 8. Bone can be stored for periods of up to 24 h at 4°C in PBS or serum-free DMEM prior to culture without any obvious deleterious effect on the ability of the tissue to give rise to populations of osteoblast-like cells. 9. Do not use disposable scalpel blades since these often shatter during this process. 10. We prefer Petri dishes to flasks because it is easier to remove the chips from them for subsequent culture and the scoring that is made on the plate surface when chopping the fragments appears to encourage cell spreading. 11. The first evidence of cellular outgrowth from the explants is usually observed within 5–7 days of plating and after 7–10 days

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cells can be observed migrating from the explants onto the surface of the culture dish (Fig. 4). If care is taken not to dislodge the explants when feeding, and they are left undisturbed between media changes, they rapidly become anchored to the substratum by the cellular outgrowths. Typical morphology of the cells is shown in Fig. 5, but cell shape varies between donors, from fibroblastic to cobblestone-like. 12. Cultures generally attain confluence 4–6 weeks postplating. 13. Using this technique, it is possible to obtain additional cell populations that continue to express osteoblast-like characteristics, including the ability to mineralise extracellular matrix, and maintain their cytokine expression profile (11). Presumably, these cultures are seeded by cells which are situated close to the bone surfaces, and which still retain the capacity for extensive proliferation and differentiation. The continued survival of these cells may be related to the gradual release over time in culture of the cytokines and growth factors that are present in the bone matrix. 14. As an alternative to using the cell filter, cells can be suspended in 2 ml serum-free DMEM containing 1 μg/ml DNAse I for each dish followed by aspiration with a narrow bore 2 ml pipette. However, we find that the act of passaging usually reduces clumping without the use of DNAse 1. 15. The yield of HOBs is typically 1−1.5 × 106 cells per 100 mm Petri dish of which ³75% are viable. In further experiments, we routinely subculture the HOBs at 5 × 103−104 cells/cm2 and achieve plating efficiencies of ³70% after 24 h. 16. The osteogenic potential of these cells can best be demonstrated by culturing the cells in diffusion chambers or on supports and transplanting them in vivo. Researchers wishing to study osteogenesis by transplanted HOBs should read the papers by Gundle et al. (19, 20) and the review by Abdallah et al. (26) and also to consult Chapter 7 in this volume. 17. Mineralising cultures can be generated from E1P1 or E2P1 cultures attesting to the phenotypic stability of E2 cells. Cells cultured in the continuous presence of ascorbate and treated with glucocorticoids at first passage show a patchy pattern of mineralisation, despite possessing similar amounts of extracellular matrix and alkaline phosphatase activity. Cells cultured without ascorbate secrete little extracellular matrix, and do not mineralise. The ability of the cells to mineralise their extracellular matrix is critically dependent on ascorbate being present continuously in primary culture. Moreover, the addition of ascorbate in secondary culture, even for extended periods, cannot compensate for its omission in primary culture. This indicates that maintenance of adequate levels of ascorbate during

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the early stages of explant culture is of critical importance for the survival of cells that retain the ability to proliferate extensively and give rise to precursors capable of undergoing osteogenic differentiation. Osteoblasts cultured continuously in the presence of ascorbate and glucocorticoids retain the ability to form bone when implanted in vivo within diffusion chambers in athymic mice (19). References 1. Bard, D. R., Dickens, M. J., Smith, A. U., and Zarek, J. M. (1972) Isolation of living cells from mature mammalian bone. Nature 236, 314–315. 2. Mills, B. G., Singer, F. R., Weiner, L. P., and Hoist, P. A. (1979) Long term culture of cells from bone affected with Paget’s disease. Calcif. Tissue Int. 29, 79–87. 3. Gallagher, J. A., Beresford, J. N., McGuire, M. K. B., Ebsworth, N. M., Meats, J. E., Gowen, M., Elford, P., Wright, P., Poser, J., Coulton, L. A., Sharrard, M., Imbimbo, B., Kanis, J. A., and Russell, R. G. G. (1984) Effects of glucocorticoids and anabolic steroids on cells derived from human skeletal and articular tissues in vitro. Adv. Exp. Med. Biol. 171, 279–292. 4. Beresford, J. N., Gallagher, J. A., Poser, J. W., and Russell, R. G. G. (1984a) Production of osteocalcin by human bone cells in vitro. Effects of 1,25(OH)2D3, parathyroid hormone and glucocorticoids. Metab. Bone Dis. Rel. Res. 5, 229–234. 5. MacDonald, B. R., Gallagher, J. A., AhnfeltRonne, I., Beresford, J. N., Gowen, M., and Russell, R. G. G. (1984) Effects of bovine parathyroid hormone and 1,25(OH)2D3 on the production of prostaglandins by cells derived from human bone. FEBS Lett. 169, 49–52. 6. MacDonald, B. R., Gallagher, J. A., and Russell, R. G. G. (1986) Parathyroid hormone stimulates the proliferation of cells derived from human bone. Endocrinology 118, 2245–2449. 7. Beresford, J. N., Gallagher, J. A., and Russell, R. G. G. (1986) 1,25-dihydroxy-vitamin D3 and human bone derived cells in vitro: effects on alkaline phosphatase, type I collagen and proliferation. Endocrinology 119, 1776–1785. 8. Gehron Robey, P., and Termine, J. D. (1985) Human bone cells in vitro. Calcif. Tissue Int. 37, 453–460. 9. Wergedal, J. E., and Baylink, D. J. (1984) Characterisation of cells isolated and cultured from human trabecular bone. Proc. Soc. Exp. Biol. Med. 176, 60–69.

10. Marie, P. J., Sabbagh, A., De Vernejoul, M. C., and Lomri, A. (1988) Osteocalcin and deoxyribonucleic acid synthesis in vitro and histomorphometric indices of bone formation in postmenopausal osteoporosis. J. Clin. Invest. 69, 272–279. 11. Gallagher, J. A., Gundle, R., and Beresford, J. N. (1996) Isolation and culture of bone forming cells (osteoblasts) from human bone, in Human Cell Culture Protocols (Jones, G. E., ed.), Humana Press Totowa, New Jersey. 12. Clover, J. and Gowen, M. (1994) Are MG-63 and HOS TE85 human osteosarcoma cell lines representative models of the osteoblastic phenotype? Bone 15, 585–591. 13. Bilbe, G., Roberts, E., Birch, M., and Evans, D. B. (1996) PCR phenotyping of cytokines, growth factors and their receptors and bone matrix proteins in human osteoblast-like cell lines. Bone 19, 437–445. 14. Billiau, A., Edy, V. G., Heremans, H., Damme, J. V., Desmyter, J., Georgiades, J. A., et al. (1977) Human interferon: mass production in a newly established cell line, MG-63. Antimicrob. Agents Chemother. 12, 11–15. 15. Mulkins, M. A., Manolagas, S. C., Deftos, L. J., and Sussman, H. H. (1983) 1,25-Dihydroxyvitamin D3 increases bone alkaline phosphatase isoenzyme levels in human osteogenic sarcoma cells. J. Biol. Chem. 258, 6219–6225. 16. Rodan, S. B., Imai, Y., Thiede, M. A., Wesolowski, G., Thompson, D., Bar-Shavit, Z., et al. (1987) Characterization of a human osteosarcoma cell line (Saos-2) with osteoblastic properties. Cancer Res. 47, 4961–4966. 17. Bowler, W. B., Buckley, K. A., Gartland, A., Hipskind, R. A., Bilbe, G., and Gallagher, J. A. (2001) Extracellular nucleotide signaling: a mechanism for integrating local and systemic responses in the activation of bone remodelling. Bone 28, 507–512. 18. Buckley, K. A., Wagstaff, S. C., McKay, G., Gaw, A., Hipskind, R. A., Bilbe, G., Gallagher,

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J. A., and Bowler, W. B. (2001) Parathyroid hormone potentiates nucleotide-induced [Ca2+]i release in rat osteoblasts independently of Gq activation or cyclic monophosphate accumulation. A mechanism for localizing systemic responses in bone. J. Biol. Chem. 276, 9565–9571. 19. Gundle, R. G., 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 cells. Bone 16, 597. 20. Gundle, R., Joyner, C. J., and Triffitt, J. T. (1997) Interactions of human osteoprogenitors with porous ceramic following diffusion chamber implantation in a xenogeneic host. J. Mater. Sci. Mater. Med. 8, 519–523. 21. Walsh, S., Jefferiss, C., Stewart, K., Jordan, G. R., Screen, J., and Beresford, J. N. (2000) Expression of the developmental markers STRO-1 and alkaline phosphatase in cultures of human marrow stromal cells: regulation by fibroblast growth factor (FGF)-2 and relationship to the expression of FGF receptors 1–4. Bone 27, 185–195.

22. Eghbali-Fatourechi, G. Z., Lamsam, J., Fraser, D., Nagel, D., Riggs, B. L., and Khosla, S. (2005) Circulating osteoblast-lineage cells in humans. N. Engl. J. Med. 352, 1959–1966. 23. Rosada, C., Justesen, J., Melsvik, D., Ebbesen, P., and Kassem, M. (2003) The human umbilical cord blood, a potential source for osteoblast progenitor cells. Calcif. Tissue Int. 72, 135. 24. Liu, G., Li, Y., Sun, J., Zhou, H., Zhang, W., Cui, L., and Cao, Y. (2010) In vitro and in vivo evaluation of osteogenesis of human umbilical cord blood-derived mesenchymal stem cells on partially demineralized bone matrix. Tissue Eng. Part A. 16, 971–982. 25. Gartland, A., Buckley, K. A., Dillon, J. P., Curran, J. M., Hunt, J. A., and Gallagher, J. A. (2005) Isolation and culture of human osteoblasts. Methods Mol. Med. 107, 29–54. 26. Abdallah, B. M., Ditzel, N., and Kassem, M. (2008) Assessment of bone formation capacity using in vivo transplantation assays: procedure and tissue analysis. Methods Mol. Biol. 455, 89–100.

Chapter 2 Osteoblast Isolation from Murine Calvaria and Long Bones Astrid D. Bakker and Jenneke Klein-Nulend Abstract This chapter describes the isolation of primary mouse osteoblasts from adult mouse calvaria and long bones, as well as the process of isolation of bone cells from neonatal mouse calvaria. Osteoblasts from adult mouse bone are obtained as outgrowth from collagenase-treated bone pieces. Isolation of osteoblasts from neonatal calvaria is achieved by sequential enzymatic digestion of the bone matrix. Because of differences in origin and isolation method, each of the primary bone cell cultures described will have their own characteristics. Key words: Osteoblast, Osteocyte, Calvaria, Long bone, Cell culture, Mouse

1. Introduction When conducting in vitro research on bone, often a choice has to be made between the use of bone organ cultures or bone cell cultures. When using cell culture the choice is between primary cells or cell lines. The advantage of cell lines over freshly isolated cells lies in the immediate availability of large numbers of cells, the homogeneity of the cell cultures, and the expected stability of the phenotype. In the long run, however, many cell lines appear unstable to some extent, and subclones of cell lines tend to develop in different laboratories. In addition, the clonal selection generally favors rapidly growing cells, but these might not express all typical features of cells from a certain tissue, and the resulting cell line may thus not be entirely representative. In addition, although silencing RNA techniques are rapidly evolving, enabling the silencing of specific molecular targets in cell lines, transfection efficiency, and duration of silencing are still hard to control. An obvious advantage of using primary cells is that cells can be isolated from genetically

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_2, © Springer Science+Business Media, LLC 2012

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modified animals, expressing a stable phenotype. This means that in certain experiments, the use of primary bone cells is preferred above the use of cell lines. Peck and coworkers initiated the use of primary bone cell cultures in 1964 (1). They isolated cells from frontal and parietal bones of fetal and neonatal rat calvaria by collagenase digestion of the uncalcified bone matrix. The isolated cells were viable, proliferated during culture, and exhibited high activity of the osteoblast marker alkaline phosphatase (ALP). The real nature of the cells, however, especially the amount of contamination with connective tissue fibroblasts, could not be unambiguously defined (1). Wong and Con (1974) tried to isolate a better defined and more homogeneous cell population by removing the outer layers of the periosteum with successive collagenase treatments (2). While this method led to cell cultures that were more osteoblastic in nature, these were still not completely free from other cell types, such as osteoclast precursors (3). Other investigators have tried to improve the osteoblastic character of the isolated bone cell populations by removing the fibroblastic outer periosteum before using enzymatic digestion to isolate the cells from the calvarium (4, 5). This method resulted in two cell populations, one of which was still osteogenic after prolonged culture time (osteoblastic cells), and another one which was not osteogenic after prolonged culture (periosteal fibroblasts) (6). Nowadays, a broad range of methods is available for obtaining well-defined osteoblast-like cells in vitro (also see Chapters 1 and 3), which are widely used as tools in bone biology (7–9). This chapter describes the isolation of primary mouse bone cells from adult mouse calvaria and long bones, as well as the process of isolation of bone cells from neonatal mouse calvaria. Due to their difference in origin and method of isolation, one might expect that each of the primary bone cell cultures described will have their own characteristics. For instance, it is likely that neonatal cell cultures contain more immature rapidly growing cells than cultures from adult bone. Indeed, it has been shown that neonatal cells show a higher basal release of nitric oxide and a higher response to 1,25-dihydroxyvitamin D3 treatment than bone cells obtained from adult bone (10). Thus, for in vitro studies investigating the cellular behavior of adult bone, it seems advisable to use cells from adult bone fragments to best reproduce the inherent cellular properties of the adult tissue. There are also profound differences between bones from the skull and the axial and appendicular skeleton, with regard to matrix composition (11), osteoclast functionality (12), and osteocyte morphology (13). Therefore, one might expect that differences exist between cell cultures derived from calvaria and long bones, and care should be taken which cells are used for in vitro research. We found no difference between cultured osteoblasts from adult mouse calvaria or adult mouse long bones with regard to the nitric

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oxide response to mechanical loading (10), suggesting that either cell culture can be used for in vitro mechanotransduction experiments. Interestingly, it has been reported that the differential mechanosensitivity of bones from C57BL/6J and C3H/HeJ mice in vivo is also reflected in the in vitro mechanical loading response of osteoblasts derived from these mouse strains (14). All these factors should be taken into consideration when planning experiments with murine osteoblasts.

2. Materials 2.1. Tissues

Cells are obtained from the long bones and the calvaria of adult (age 9 weeks or older) mice, or from the calvaria from neonatal mouse pups (age 3–4 days).

2.2. Instruments

All of the following materials are supposed to be sterile. 1. Polystyrene plate and needles for fixing the mice. 2. Scalpels (no. 10 and 11), scissors, tweezers, and curved forceps. 3. 5 ml and 10 ml syringes, 27G½ needles and 0.2 μm disposable filter units (Schleicher & Schuell GmbH, Dassel, Germany). 4. 25 and 75 cm2 tissue culture flasks (Nunc, Roskilde, Denmark), 94/16 mm cellstar petri dishes (Greiner), and 145/20 mm cellstar (large) petri dishes. 5. 100 × 16 mm (10 ml) conical base test tubes with screw cap (Bibby Sterilin ltd, Staffordshire, UK) and conical base, 15 and 25 ml polypropylene centrifuge tubes (Greiner).

2.3. Media and Solutions

1. Sterile phosphate-buffered saline (PBS), pH 7.4. 2. Dulbecco’s modified Eagle’s medium, 1 g/l glucose + L-glutamine + pyruvate (DMEM). 3. Complete culture medium (cCM):DMEM supplemented with 100 U/ml penicillin, 50 μg/ml streptomycin sulfate, 50 μg/ ml gentamycin, 1.25 μg/ml fungizone, 100 μg/ml ascorbate, and 10% FBS (e.g., Hyclone, Logan, UT, USA; see Note 1). Make fresh before each medium change and filter sterilize. 4. Collagenase II solution. 2 mg collagenase II (260 U/mg; Worthington, Lakewood, NJ, USA) per ml DMEM. Make fresh and filter sterilize. 5. Trypsin solution. 0.25% trypsin 1:250 (Gibco) plus 0.1% EDTA in PBS, filter sterilize.

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6. Digestion solution: Add 1 ml trypsin solution plus 3.2 mg collagenase II (Worthington) to 4 ml PBS. Make up fresh just before use. 7. Stock solution of collagenase type I: 10 mg/ml collagenase type I in Hank’s balanced salt solution (HBSS). Filter sterilize and freeze aliquots for single use at −20°C. 8. Collagenase I work solution: Dilute stock solution of collagenase type I in HBSS to 1 mg/ml just before use. 9. 4 μM EDTA in PBS. Filter sterilize and store at 4°C.

3. Methods Normal techniques for working under sterile conditions (use of sterile media and instruments and working in a flow cabinet) should be adhered to. 3.1. Isolation and Culture of Primary Bone Cells from Adult Mouse Long Bones

1. Euthanise one or two adult mice. 2. Fixate the mouse in a supine position on a polystyrene plate or in a large petri dish, and rinse the abdomen and extremities using a small amount of 70% ethanol. 3. Make a single incision through the skin, starting at the top of the sternum and ending a few millimeters above the genitals, using a number 10 scalpel. Make a second incision starting from the top of the first incision and ending at the wrist of the upper left extremity. Repeat this procedure with the other paws. Carefully remove the skin from the abdomen with a blade. 4. Change your blade for a sterile number 11 scalpel. Carefully remove all the muscles from one of the long bones (femur, tibia and fibula, or humerus, radius and ulna), and thoroughly scrape the bone with a scalpel until it is clean (see Note 2). Excise the long bone and place it in a petri dish with PBS. Continue until all the bones are excised. 5. When all the long bones have been removed, cut off the epiphyses. 6. Thoroughly flush out the bone marrow with PBS, using a 5 ml syringe and a 27 gauge needle. 7. Cut the clean diaphyses into little pieces of approximately 1–2 mm2 using scissors. 8. The bone pieces are washed several times with PBS, and incubated in 4 ml collagenase II solution at 37°C in a shaking water bath in order to remove all remaining soft tissue and adherent cells.

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9. Vigorously shake the collagenase II solution containing the bone pieces, by hand, every 30 min. 10. After 2 h, rinse the bone pieces three times with cCM, shaking the solution containing the pieces for a few seconds during every wash step. 11. Transfer the bone pieces to 25 cm2 flasks, containing 5 ml cCM, at a density of about 20–30 fragments per flask. Replace culture medium three times per week. After each medium replacement, make sure that the bone chips are evenly distributed over the bottom of the culture flask, by gently swirling the culture flask. 12. Adult mouse bone cells will start to migrate from the bone chips after 3–5 days. On average the cells growing from the bone fragments will be ready for use after 11–15 days. Do not allow the cells to become over-confluent around the bone chips. 13. To obtain more cells, cCM is removed from the flask, cells, and bone pieces are gently washed three times with PBS, and incubated in 1 ml trypsin solution at 37°C for 10 min. Swirl and tap the flask on a level surface a few times during the 10 min incubation period. Carefully check whether the cells are released from the tissue culture plastic using a microscope. Remove the trypsin solution containing the cells using a small pipette. Leave the bone pieces in the flask, and discard. 14. The cells are plated at 2.5−5 × 103 cells per cm2 in T25 or T75 flasks in cCM. 15. Medium is changed three times per week, and after approximately 7–10 days cells will reach subconfluency, upon which they can be used for experiments (see Note 3). The average number of cells thus obtained is between 4 × 106 and 6 × 106 cells. 3.2. Isolation and Culture of Primary Bone Cells from Adult Mouse Calvaria

1. Euthanise two adult mice and fixate them on a polystyrene plate, or in a large petri dish. 2. Clean the head using 70% ethanol, and make a cut through the skin at the base of the skull, using scissors. 3. Make an incision starting at the nose bridge, and ending at the base of the skull. Remove the skin from the top of the head. 4. Use scissors to cut through the bone at the base of the neck. Cut the calvaria loose while holding the head with curved forceps placed in the orbita (see Fig. 1). 5. Transfer the calvaria to a petri dish with PBS and remove the soft tissues using tweezers or by scraping with a knife (see Note 2). 6. Remove the sutures using scissors, and chop the remaining bone into small fragments of approximately 1–2 mm2.

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Fig. 1. Mouse calvaria. Make a cut through the skin at the base of the skull, using scissors. For adult tissue, make an incision starting at the nose bridge, ending at the base of the skull, and remove the skin from the top of the head. Use scissors to cut away the skin from the top of the neonatal mouse heads. Dissect calvaria as indicated by the dotted line and remove as much soft tissue as possible. Do not include the area near the neck, since this will result in heavy fibroblast contamination of the cultures. Sf skin flap, C calvaria.

7. Incubate the fragments for 30 min in 4 ml collagenase II solution at 37°C in a shaking waterbath. 8. Discard the collagenase II solution and replace with fresh collagenase II solution. Incubate another 30 min and then replace the collagenase II solution for trypsin solution. 9. Incubate in trypsin for 30 min. Replace by 4 ml collagenase II solution for the fourth and final incubation step of 30 min.

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10. Rinse the bone pieces three times with cCM and transfer the bone pieces to 25 cm2 flasks, containing 5 ml cCM, at a density of about 20–30 fragments per flask. 11. Medium is changed three times per week. After each medium replacement, make sure that the bone chips are evenly distributed over the bottom of the culture flask, by gently swirling the culture flask. Adult mouse bone cells will start to migrate from the bone chips after 3–5 days. 12. On average the cells growing from the bone fragments will reach subconfluency after 11–15 days, upon which the cells are incubated with 1 ml trypsin solution at 37°C for 10 min. Swirl and tap the flask on a level surface a few times during the 10 min incubation period. Remove the trypsin solution containing the cells using a small pipet. Leave the bone pieces in the flask and discard. 13. The cells are plated at 2.5 to 5 × 103 cells per cm2 in T25 or T75 flasks in cCM. 14. Medium is changed three times per week, and after approximately 7–10 days cells will reach subconfluency, upon which they can be used for experiments. The average number of cells thus obtained lies between 4 × 106 and 6 × 106 cells. 3.3. Isolation and Culture of Bone Cells from Neonatal Mouse Calvaria

1. Euthanise 20–30 neonatal mice pups (2 nests) by means of decapitation, or inhalation of halothane, and place the heads in a petri dish with PBS (see Note 4). 2. Grab a head in the nape of the neck, and cut the skin away using scissors. 3. Hold the head with curved forceps placed through the orbita, cut the calvaria loose along the edge, and place it in a petri dish with PBS (see Fig. 1). 4. Pin the calvaria down with tweezers and cut away the edges and sutures with a small scalpel. Transfer the calvaria halves to a 25 ml tube with PBS and wash twice with PBS. 5. Incubate the calvaria in 4 ml digestion solution at 37°C in shaking water bath. After 10 min shake the calvaria by hand for a few seconds. 6. Incubate for a total of 20 min, and then transfer the supernatant containing cells to a 10 ml tube. Add 700 μl FBS to the cell suspension to inhibit trypsin activity. 7. Wash the calvaria with 3 ml DMEM (without FBS!), shake well, and add the supernatant to the tube containing the cell suspension. This is population number 1. 8. Add new digestion solution to the calvaria, and repeat the previous three steps to obtain population number 2. During the 20 min that the calvaria have to incubate in the water bath,

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centrifuge cell population number 1 at 300 × g for 5 min. Discard supernatant, resuspend cell pellet in 1 ml cCM, and add to 17 ml cCM. Pipette in a 6-well plate at 3 ml cell suspension per well. 9. Repeat the entire procedure for a total of four times to obtain population 1–4. 10. Culture medium is changed 1 day after isolation of the bone cells. 11. Within approximately 5 days cells will reach subconfluency, upon which they are trypsinized by incubation with 200 μl trypsin solution per well, at 37°C for 10 min. 12. To enhance the number of cells available for experiments, population 1 and 2, resembling osteoblast progenitor cells, are usually pooled, as well as population 3 and 4. This latter pooled cell population is enriched with cells exhibiting biochemical characteristics of differentiated osteoblasts, such as high ALP activity and osteopontin expression. Both pooled and unpooled populations can be directly used for experiments. 13. The number of cells obtained using this method varies between 6 × 106 and 10 × 106 cells. 3.4. Isolation and Culture of Bone Cells from Neonatal Mouse Calvaria (Alternative)

Although we prefer the use of collagenase type II, other groups have reported use of crude collagenase type I, which is cheaper and intrinsically contains trypsin as a contaminant. Therefore, an alternative protocol for isolation of osteoblasts from neonatal mouse calvaria, which uses alternate collagenase I and EDTA incubations to remove as much mineralized matrix as possible, and increases cellular yield, is given below: 1. Dissect calvaria as described in Subheading 3.3 steps 1–3 and collect them in 3 ml HBSS in a 25 ml centrifuge tube in a shaking waterbath at 37°C. 2. Incubate calvaria in 3 ml collagenase I work solution for 10 min in a shaking waterbath at 37°C. 3. Replace the collagenase I work solution (fraction 1) with fresh solution and discard fraction 1. 4. Incubate for 30 min in collagenase I work solution. Collect the solution containing cells (fraction 2) and place in a conical centrifuge tube. Wash calvaria in 7 ml of PBS and add wash to fraction 2. 5. Add EDTA solution to the calvaria and incubate for 10 min at 37°C. Collect EDTA solution (fraction 3). Wash calvaria in 7 ml HBSS and add wash to fraction 3. 6. Add collagenase I work solution to the calvaria and incubate for 30 min at 37°C. Collect the solution containing cells (fraction 4)

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and place in a conical centrifuge tube. Wash in HBSS and add wash to fraction 4. 7. Further fractions, presumably containing more mature osteocyte-like cells, can be collected by repeating steps 5 and 6, but cell yields will be increasingly lower. 8. Centrifuge all fractions immediately after collection (250 × g for 5 min) and resuspend pellets in cCM (see Subheading 2.3.3). 9. Plate out pooled or single fractions in 75 cm2 culture flasks using cells derived from 2 to 3 animals/flask. 10. Cultures will be confluent in 3–4 days. To minimize contamination by other adherent cell types, replace medium once cells have adhered (2–3 h after plating). Use the more differentiated fractions (3 and 4) for osteoclast cocultures (see Chapter 12, this volume).

4. Notes 1. Addition of serum to the medium is necessary for the survival and stimulation of proliferation of the primary mouse bone cells. However, “serum” is not a constant and homogeneous product, and the growth rate of primary bone cells can vary considerably between several batches of serum. It is therefore recommended to test several batches of serum on their cell proliferative ability and continue to use the one that produces the best results. 2. Sometimes, the primary bone cultures can contain fibroblasts, which grow faster than the bone cells and can quickly overgrow the primary bone cell cultures. If this problem occurs, care should be taken to remove all soft tissues by scraping the bones with a knife before starting the collagenase treatment. Also make sure the collagenase is not expired, and that the collagenase solution is made fresh every time. 3. Primary bone cell cultures are not 100% pure and may contain some fibroblasts and other cell types that are not from the osteoblast lineage. The osteoblastic phenotype of the primary mouse bone cell cultures can be determined by stimulating the cells with 1,25dihydroxyvitamin D3, which should lead to enhanced ALP activity (11). The exact nature of the bone cells that are isolated from adult long bones and adult calvaria has not been fully determined. Since the cell isolation protocols involve mechanically removing the soft tissues first and then all adhering cells by means of incubation with collagenase, the cells that are isolated from the bones might represent osteocytes that reverted to proliferation after several days of exposure to serum.

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Fig. 2. Phase contrast microscopy of primary mouse bone cell cultures. (a) Adult mouse bone cells growing out of the bone chips, day 6 of culture. (b) Subconfluent layer of adult mouse bone cells, first passage. (c) Neonatal mouse calvarial cells, population number 1 and 2, day 2 of culture. (d) Neonatal mouse calvarial cells, population number 3 and 4, day 2 of culture. Note the oblong trapezoid shaped morphology of the osteoblastsl.

The microscopical appearance of the isolated bone cells is mostly osteoblastic (Fig. 2), but mRNA expression of several markers for (pre)osteocytes, such as MEPE, Phex, and DMP1 can be detected by means of PCR. Absence of staining for von Willebrand factor (factor VIII) shows that the bone cell cultures do not contain endothelial cells. 4. Smaller numbers of calvaria can be used successfully, leading to a proportionately lower yield of osteoblasts. References 1. Peck, W. A., Birge, S. J., and Fedak, S. A. (1964) Bone cells: biochemical and biological studies after enzymatic isolation. Science 146, 1476–1477. 2. Wong, G. L., and Cohn, D. V. (1974) Separation of parathyroid hormone and calcitonin-sensitive cells from non-responsive cells. Nature 252, 713–715. 3. Burger, E. H., Boonekamp, P. M., and Nijweide, P. J. (1986) Osteoblast and osteoclast precursors in primary cultures of calvarial bone cells. Anat. Rec. 214, 32–40. 4. Yagiela, J. A. and Woodbury, D. M. (1977) Enzymatic isolation of osteoblasts from fetal rat calvaria. Anat. Rec. 188, 287–306.

5. Nijweide, P. J., van der Plas, A., and Scherft, P. J. (1981) Biochemical and histological studies on various bone cell preparations. Calcif. Tissue Int. 33, 529–540. 6. Nijweide, P. J., van Iperen-van Gent, A. S., Kawilarang-de Haas, E. W. M., van der Plas, A., and Wassenaar, A. M. (1982) Bone formation and calcification by isolated osteoblast-like cells. J. Cell Biol. 93, 318–323. 7. Klein-Nulend, J., Semeins, C. M., Ajubi, N. E., Nijweide, P. J., and Burger, E. H. (1995) Pulsating fluid flow increases nitric oxide (NO) synthesis by osteocytes but not periosteal fibroblasts: correlation with prostaglandin upregulation. Biochem. Biophys. Res. Commun. 217, 640–648.

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8. Bakker, A. D., Soejima, K., Klein-Nulend, J., and Burger, E. H. (2001) The production of nitric oxide and prostaglandin E2 by primary bone cells is shear stress dependent. J. Biomech. 34, 671–677. 9. Armstrong, V. J., Muzylak, M., Sunters, A., Zaman, G., Saxon, L. K., Price, J. S., and Lanyon, L. E. (2007) Wnt/beta-catenin signaling is a component of osteoblastic bone cell early responses to load-bearing and requires estrogen receptor alpha. J. Biol. Chem. 282, 20715–20727. 10. Soejima, K., Klein-Nulend, J., Semeins, C. M., and Burger, E. H. (2001) Different responsiveness of cells from adult and neonatal mouse bone to mechanical and biochemical challenge. J. Cell Phys. 186, 366–370. 11. van den Bos, T., Speijer, D., Bank, R. A., Brömme, D., and Everts, V. (2008) Differences in matrix composition between calvaria and long bone in mice suggest differences in biomechanical properties and resorption: special emphasis on collagen. Bone 43, 459–468.

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12. Jansen, I. D., Mardones, P., Lecanda, F., de Vries, T. J., Recalde, S., Hoeben, K. A., Schoenmaker, T., Ravesloot, J. H., van Borren, M. M., van Eijden, T. M., Bronckers, A. L., Kellokumpu, S., Medina, J. F., Everts, V., and Oude Elferink, R. P. (2009) Ae2(a,b)deficient mice exhibit osteopetrosis of long bones but not of calvaria. FASEB J. 23, 3470–3481. 13. Vatsa, A., Breuls, R. G., Semeins, C. M., Salmon, P. L., Smit, T. H., and Klein-Nulend, J. (2008) Osteocyte morphology in fibula and calvaria – is there a role for mechanosensing? Bone 43, 452–458. 14. Lau, K. H., Kapur, S., Kesavan, C., and Baylink, D. J. (2006) Up-regulation of the Wnt, estrogen receptor, insulin-like growth factor-I, and bone morphogenetic protein pathways in C57BL/6J osteoblasts as opposed to C3H/ HeJ osteoblasts in part contributes to the differential anabolic response to fluid shear. J. Biol. Chem. 281, 9576–9588.

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Chapter 3 Rat Osteoblast Cultures Isabel R. Orriss, Sarah E.B. Taylor, and Timothy R. Arnett Abstract This chapter describes the isolation, culture and staining of primary osteoblasts from the calvaria and long bones of neonatal rats. The key advantages of this assay are that it allows direct measurement of bone matrix deposition and mineralisation, as well as yielding good quantities of osteoblasts at defined stages of differentiation for molecular and histological analysis. A special focus of this chapter is on the role of β-glycerophosphate in cell-mediated mineralisation in these cultures. Key words: Osteoblast, Bone formation, Mineralisation, pH

1. Introduction Osteoblasts are derived from mesenchymal stem cells and are responsible for bone formation. A number of different approaches have been developed to study osteoblasts in vitro, including bone organ cultures, primary cell cultures, and immortalised osteoblastlike cell lines. Combined these methods have provided abundant information about the regulation of osteoblast proliferation, differentiation and function. Osteoblastic cell lines are widely used as a convenient source of large numbers of cells with a relatively stable phenotype. However, many of these osteoblast-like cells do not express the whole range of bone-specific genes and/or form bone in vitro. Furthermore, repeated passaging has been shown to cause the loss of the osteoblastic phenotype in some cell lines (1). Primary bone cell cultures were first described in 1964 by Peck and colleagues (2), who isolated cells from the parietal and frontal bones of foetal and neonatal rat calvaria using collagenase digestion.

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_3, © Springer Science+Business Media, LLC 2012

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The isolated cells proliferated in vitro and exhibited high alkaline phosphatase (ALP) activity, although the cultures were contaminated with other cell types such as fibroblasts. In 1974, Wong and Cohn (3) used sequential collagenase digestion to obtain a more homogenous population of osteoblastic cells. The first description of the formation of bone nodules by differentiating osteoblasts released enzymically from calvarial bones and cultured with β-glycerophosphate (β-GP) ascorbate and dexamethasone was by Bellows and colleagues in 1986 (4). This chapter describes the isolation, culture, and staining of primary osteoblasts from the calvaria and long bones of neonatal rats (see Note 1). The osteoblast bone formation assay has a number of advantages. Firstly, it allows the key function of osteoblasts, namely bone formation, to be studied quantitatively (5). Bone formation in this essentially 2-dimensional culture system appears to recapitulate the intramembranous (woven) bone formation seen in histological sections from developing animals – i.e., large-scale structures resembling trabeculae appear. Secondly, it offers the opportunity to study separately the processes of bone matrix deposition and mineralisation (6). Thirdly, it allows the extracellular environment to be tightly regulated (e.g., pH, pO2) in a manner which is not possible using bone organ cultures or in vivo (7, 8). Fourthly, osteoblast activity can be studied in an environment that is relatively free from the influence of other cell types normally found in bone, such as endothelial cells, nerves, and haematopoietic cells. Lastly, osteoblasts can be studied at clearly identified stages of differentiation from the immature, proliferating cells present early in the cultures through to the mature bone-forming osteoblasts in late stage cultures. Compared to mice of the same age, neonatal rats yield higher numbers of cells per animal. The rat model is therefore more convenient to use, and is appropriate for studies requiring larger numbers of osteoblasts. Two important pitfalls should be highlighted here. The first concerns the use of inappropriately high concentrations of the ALP substrate, β-GP in many published descriptions of osteoblast cultures. At high concentrations, β-GP causes generalised mineral deposition and impaired cell viability (the osteoblasts effectively fossilise), with the result that the cultures fail to progress beyond the formation of small nodules. The second, related pitfall concerns the frequent confusion between mineral deposition (which also occurs on teeth, in caves and in kettles) and true bone formation, which involves selective mineralisation of a collagenous matrix deposited in characteristic patterns. Osteoblast cell lines (and many other cell types) express ALP and can mineralise in the presence of sufficient β-GP – but this is not bone formation.

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2. Materials All solutions, instruments, and tissue culture plastics should be sterile. 1. Animals: The cells are obtained from the calvaria and long bones of neonatal (2–3 day old) rats. The number of animals to use depends on the experiments to be performed; typically, one animal will yield 107 and 5 × 106 cells from the calvaria and long bones, respectively (see Notes 1 and 2). 2. Phosphate-buffered saline (PBS): For storing tissues prior to use and washing cells. 3. Supplemented Dulbecco’s modified essential medium (sDMEM): 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/ antimitotic or AB/AM). This sDMEM should be stable at 4°C for at least 4 weeks. 4. Osteogenesis DMEM (osDMEM): To the sDMEM described above add 2 mM β-GP (see Note 3), 10 nM dexamethasone, and 50 μg/ml ascorbate. Always make fresh on day of use. 5. Sodium hydroxide (NaOH): 6 M NaOH to alter the pH of the culture medium if required (see Note 4). 6. Trypsin-EDTA: 0.25% (w/v) trypsin with 1 mM EDTA; free of calcium and magnesium, which inhibit trypsin activity (Invitrogen 25200056). 7. Collagenase: 0.2% (w/v) collagenase solution (Type II collagenase from Clostridium histolyticum, Sigma C6885) made up in Hank’s balanced salt solution (HBSS); filter sterilise. Note that HBSS contains calcium, which is required for collagenase activity. 8. Fixative: 2.5% glutaraldehyde in dH2O or 70% ethanol; prepare fresh before use. 9. Alizarin red stain for mineralised bone nodules: 1% (w/v) alizarin red in dH2O; prepare fresh before use. 10. ALP staining: Alkaline phosphatase kit (Sigma kit 86C-1KT). 11. Sirius red stain for deposited collagen: Sircol™ dye reagent (Biocolor kit S1005). 12. Tissue culture plastics: Large petri dishes (100 mm), 5 ml flat bottomed tubes, 15 and 50 ml centrifuge tubes, 75 cm2 tissue culture flasks, and 24-well tissue culture plates. 13. Dissection tools: Scalpels and blades (no. 20), tweezers, and scissors.

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3. Methods To keep the cell cultures sterile, normal techniques for working under sterile conditions should be employed (working in a flow cabinet, use of sterile media and instruments, etc.). 3.1. Isolation and Culture of Primary Osteoblasts from Neonatal Rat Calvaria

1. Sacrifice two neonatal (2–3 days old) rats and sterilise with 70% ethanol. Place each cadaver in a large petri dish. 2. Remove the head using large scissors; transfer the body to a separate petri dish for long bone isolation (see Subheading 3.2). 3. Grasp the head at the nape of the neck and make a small incision along the base of the skull (small scissors will make the cleanest cut). Carefully remove the skin and the brain tissue from the skull using a scapel and tweezers. 4. Cut away the jaw and scrape off any excess tissue and cartilage from around the edge of the calvaria. 5. Cut the calvaria in half and place in a flat-bottomed 5 ml tube; wash with PBS. 6. Repeat steps 2–5 for the second animal. 7. Incubate the calvariae in 1% trypsin (1 ml/calvarial bone) for 10 min at 37°C. Remove and discard the trypsin solution; wash in sDMEM (serum and calcium will inactivate any residual trypsin). 8. Incubate in 0.2% collagenase solution (800 μl/calvarial bone) for 30 min at 37°C. 9. Remove the collagenase digest, discard, and replace with fresh solution for a further 60 min at 37°C. 10. Keep the final digest and transfer to a 15 ml conical base centrifuge tube. Wash calvaria with sDMEM (5 ml), transfer the solution to the 15 ml tube containing the final digest. 11. Spin the cell solution at 1,500 × g for 5 min at room temperature. Discard the supernatant and resuspend the cell pellet in sDMEM (1 ml/calvarial bone). Pool the cell suspensions. 12. Add 20 ml of sDMEM to 2 × 75 cm2 flasks; add 1 ml of cell suspension to each flask. 13. Incubate the flask at 37°C/5% CO2 until the cells reach confluency (~3 days).

3.2. Isolation and Culture of Primary Osteoblasts from the Long Bones of Neonatal Rats

1. Take the bodies of the animals used for the calvarial osteoblast isolation (see Subheading 3.1). 2. Remove the limbs from the body by cutting with sharp scissors at the point closest to the body, preserving as much of the limb as possible.

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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 place the bone fragments into a flatbottomed 5 ml tube. Wash and vortex in PBS. 6. Incubate the bone fragments in 1% trypsin (1 ml/animal) for 10 min at 37°C. Remove and discard the trypsin solution; wash in sDMEM. 7. Incubate in 0.2% collagenase solution (1 ml) for 30 min at 37°C. 8. Remove the collagenase digest, discard, and replace with fresh solution for a further 60 min at 37°C. 9. Keep the final digest and transfer to a 15 ml conical base centrifuge tube. Wash remaining bone fragments with sDMEM (5 ml) and transfer the solution to the 15 ml tube containing the final digest. 10. Spin the cell solution at 1,500 × g for 5 min at room temperature. Discard the supernatant and resuspend the cell pellet in sDMEM (1 ml). The intitial cell population may be slightly more heterogeneous than a calvarial bone isolate but behaves similarly on subsequent culture in osDMEM. 11. Add 20 ml of sDMEM to a 75 cm2 flasks; add cell suspension. 12. Incubate the flask at 37°C/5% CO2 until the cells reach confluency (~3 days). 3.3. Bone Formation Assay

This protocol is used for osteoblasts from both calvarial and long bone isolations. 1. Once the cells in the 75 cm2 have reached confluency (~3 days), remove the sDMEM and wash with PBS. Add 1% trypsin (2 ml/flask) and incubate for 10 min at 37°C. 2. Inactivate the trypsin by adding 10 ml of sDMEM; transfer the cell suspension to a 15 ml centrifuge tube. 3. Spin the cell solution at 1,500 × g for 5 min at room temperature. Discard the supernatant and resuspend the cell pellet in sDMEM (1 ml/flask). Pool the cell suspensions. 4. Perform a cell count using a haemocytometer; seed the cells in 24-well tissue culture trays at a density of 2.5 × 104 cells/well in osDMEM. 5. Half the medium should be exchanged every 2–3 days. Cultures will typically be fully confluent by day 4; they will begin to form discrete bone nodules from about day 10, with extensive networks of mineralising “trabeculae” developing from day 14 (Figs. 1 and 2).

Fig. 1. Phase contrast microscopy of primary rat osteoblast cultures. Representative images are of unstained cell layers. By day 4 of culture, a confluent monolayer of cells is evident. At day 7, the cells are more compacted and organic matrix is beginning to be deposited (as shown by the arrow ). By day 10, there is abundant deposition of unmineralised collagenous matrix and mineralisation (dark area at lower left of image) is commencing. After 14 days of culture, there is widespread formation of mineralised matrix networks. Scale bar = 250 μm.

Fig. 2. Different staining methods in primary rat osteoblast cultures. Images show 14-day osteoblast cultures either left unstained, or stained with alizarin red (to demonstrate calcium), alkaline phosphatase (ALP) or sirius red (to demonstrate collagen). Bone formed in these cultures exhibits a typical “trabecular” morphology, with discrete mineralisation confined to matrix nodules, as shown in the unstained and alizarin red-stained images. In the unstained image at lower left, unmineralised matrix appears brown and mineralised matrix black. Intense ALP staining (also “trabecular”) is evident in mature, bone forming osteoblasts. Sirius red staining shows the presence of collagen fibres in these cultures. Scale bars = 0.1 cm (cell well scans, upper row) and 250 μm (phase contrast images, lower row ).

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6. The pH should be monitored at every medium change and maintained at ~ pH 7.4 by the addition of 6 M NaOH if required (see Notes 4 and 5). 7. ALP activity (ALP assay kit, Biotron) and soluble collagen production (Sircol Red assay, Biocolor) can be assayed throughout the 14-day culture period using commercially available kits. 3.4. Fixation and Staining

1. On termination of the experiment, carefully wash the cell layers with PBS. 2. Transfer to fixative: 2.5% glutaraldehyde for 5 min for alizarin red and ALP staining, 70% ethanol for 1 h for sirius red staining. 3. Wash twice with PBS and leave to air dry (sirius red staining only). 4. For cell layers to be stained with alizarin red or for ALP, wash three times with 70% ethanol and leave to air dry. 5. Stain as required (Fig. 2): (a) Alizarin red for 5 min followed by three washes with 50% ethanol; allow to air dry. (b) ALP for ~30 min in the dark; wash with dH2O and allow to air dry. (c) Sirius red for 1 h; wash with dH2O and allow to air dry.

3.5. Quantification of Mineralised Bone Formation

1. Once fully dry, scan plates at 2,000 dots/cm2 on a highresolution flat-bed scanner. We use an Epson Perfection 4990 photo/slide scanner. Transmitted light scanning of alizarin red-stained plates is commonly used but unstained cell layers also yield excellent high-contrast images in both transmitted and reflected light modes (the latter showing the white bone trabeculae against a dark background) (Fig. 2). 2. Apply a circular mask to the image of each cell well and convert to a binary image (e.g., using “Adobe Photoshop”). 3. Use an image analysis program (e.g., “Image J” – available free from http:www.rsbweb.nih.gov/ij/) to determine the number and surface area of mineralised bone nodules in the binary images of each individual well, using constant threshold and minimum particle size levels.

3.6. Statistics

We routinely use one-way analysis of variance (ANOVA) to analyse experiments. Although often neglected, adjustments for multiple comparisons between treatment groups (e.g., the Bonferroni correction) are frequently needed (see Note 6).

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4. Notes 1. Calvarial cells vs. long bone cells : Although the flat bones of the skull and the limb bones are formed by different developmental processes, osteoblasts derived from either source appear to behave similarly in culture with regard to proliferation and mineralization. 2. Young cells vs. old cells : An important reason why the rat osteoblast culture system described here is able to form bone is that the initial cell population is obtained from neonates – and thus has considerable growth potential. Cells derived from adult animals (including human donors) may have progressed too far toward their “Hayflick limits” to be capable of the rapid expansion to high cell density required for true osteogenic differentiation. 3. Concentration of β-GP : Bone mineralisation requires a supply of calcium and orthophosphate (Pi), both of which are required for hydroxyapatite crystal formation. A key source of phosphate in bone is alkaline phosphatase, which hydrolyzes a range of phosphate-containing substrates, including adenosine triphosphate (ATP) and inorganic pyrophosphate (6). An additional important mechanism by which ALP promotes mineralisation, however, is by hydrolysing pyrophosphate, a ubiquitous physicochemical inhibitor of calcium phosphate deposition. To form mineralised bone nodules in vitro, an additional source of phosphate is required. The most widely used phosphate source is β-GP, although phosphate itself can also be used. Figure 3 illustrates the critical importance of β-GP concentration in this culture system (see also ref. 6). In osteoblast cultures lacking β-GP, organic matrix is deposited but mineralisation fails to occur. Osteoblasts cultured with 2 mM β-GP reproducibly form abundant bony structures with characteristic “trabecular” morphology; alizarin red staining shows that mineralisation is confined to these matrix structures. In contrast, culture with 5–10 mM β-GP causes a widespread, dystrophic deposition of mineral as soon as the differentiating osteoblasts begin to express significant amounts of ALP, leading to mineralisation of the cells themselves (Fig. 3a). This impairs cell viability (Fig. 3b), with the result that only small mineralised nodules are able to form (rather than the striking trabecular structures evident with lower concentrations of β-GP). Thus, it is critical that a low concentration of β-GP is used in this assay. 4. The importance of pH : Extracellular pH is an important factor in the regulation of bone mineralisation (7). Acidosis causes a selective, inhibitory action on matrix mineralisation by

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Fig. 3. The effect of the β-glycerophosphate concentration on bone mineralisation and osteoblast viability in vitro. (a) Rat calvarial osteoblasts were cultured for 14 days before staining with alizarin red to demonstrate mineral deposition. Rat long bone osteoblasts were cultured for 28 days before fixation and transmission electron microscopy (TEM). In the absence of β-glycerophosphate (β-GP), bone mineralisation fails to occur due to insufficient inorganic phosphate; widespread, unmineralised collagenous matrix is evident in the phase contrast and TEM images. Bone formed in the presence of 2 mM β-GP exhibits a typical “trabecular” morphology; mineralisation is confined to these structures. In the presence of 5–10 mM βGP, widespread, nonspecific (dystrophic) deposition of bone mineral occurs across the cell monolayer, with inhibition of normal matrix deposition. Intracellular deposition of mineral that causes damage to cell membranes and organelles is also evident. (b) Increased β-GP concentration is associated with decreased osteoblast viability, assessed as lactate dehydrogenase (LDH) release after 14 days in culture. Values are means ± standard error of the mean (SEM) (n = 6); significantly different from control ***p < 0.001, **p < 0.01. Scale bars = 0.1 cm (cell well scans), 250 μm (phase contrast images), and 1 μm (TEM).

inhibiting ALP expression and activity, while increasing mineral solubility (7). For example, even a reduction in pH from pH 7.43 to pH 7.32 will reduce bone nodule formation in vitro by ~60% (Fig. 4). Due to its high bicarbonate concentration, DMEM is more alkaline than some media (such as minimum essential medium, MEM); in equilibrium with 5% CO2, it is buffered to pH ~7.4. The metabolic activity of osteoblasts will cause progressive acidification, particularly in mature, bone forming cultures when cells numbers are high. To ensure that pH does not affect bone mineralisation in this assay, it may sometimes be necessary to add a small volume of OH− ions (as 6 M NaOH) to the culture medium to ensure an average running pH of 7.4.

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Fig. 4. Inhibitory effect of acidosis of bone mineralisation by cultured rat osteoblasts. In 14 day cultures, decreasing pH caused progressive reduction in the formation of mineralised nodules stained with alizarin red, with complete inhibition at pH 6.93. Values are means ± standard error of the mean (SEM) (n = 6); significantly different from control (pH 7.43) value, **p < 0.001, *p < 0.05.

5. Measuring the medium pH: Accurate measurement of the operating pH of mammalian osteoblast cultures is important for meaningful comparison of results from different laboratories. For HCO3−/CO2 buffered media, accurate pH measurement can only be achieved by the use of a standardised blood gas analyser. We use a Radiometer ABL 705 blood gas analyser (Radiometer, Crawley, UK), which features a multielectrode system to measure pH, pCO2 and pO2 in a 200 μl injected sample (cyclic time ~ 2 min). The first medium measurement taken immediately after removing the culture plates from the incubator is assumed to provide a pCO2 value that is identical for all wells and that reflects the actual pCO2 during incubation. It should be noted that opening the incubator door during experiments may cause perturbations in CO2 levels that affects the measured pH and pCO2 values. Incubators with individual, separate compartments are unlikely to prevent this problem unless the compartments in question can be sealed. 6. Statistics: Because of interassay variability, statistical comparisons should be performed only within one assay, and not between different assays.

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Acknowledgment Grant Funding: The authors gratefully acknowledge the support of Arthritis Research UK, the Biotechnology and Biological Sciences Research Council (UK) and the European Union (Framework 7 programme). References 1. Hausser, H. J., and Brenner, R. E. (2005) Phenotypic instability of SaOS-2 cells in longterm culture. Biochem. Biophys. Res. Commun. 333, 216–222. 2. Peck, W. A., Birge, S. J., Jr., and Fedak, S. A. (1964) Bone cells: biochemical and biological studies after enzymatic isolation. Science 146, 1476–1477. 3. Wong, G., and Cohn, D. V. (1974) Separation of parathyroid hormone and calcitonin-sensitive cells from non-responsive bone cells. Nature 52, 713–715. 4. Bellows, C. G., Aubin, J. E., Heersche, J. N., and Antosz, M. E. (1986) Mineralized bone nodules formed in vitro from enzymatically released rat calvaria cell populations. Calcif. Tissue Int. 38, 143–154. 5. Hoebertz, A., Mahendran, S., Burnstock, G., and Arnett, T. R. (2002) ATP and UTP at low concentrations strongly inhibit bone formation

by osteoblasts: a novel role for the P2Y2 receptor in bone remodeling. J. Cell Biochem. 86, 413–419. 6. Orriss, I. R., Utting, J. C., Brandao-Burch, A., Colston, K., Grubb, B. R., Burnstock, G., and Arnett, T. R. (2007) Extracellular nucleotides block bone mineralization in vitro: evidence for dual inhibitory mechanisms involving both P2Y2 receptors and pyrophosphate. Endocrinology 148, 4208–4216. 7. Brandao-Burch, A., Utting, J. C., Orriss, I. R., and Arnett, T. R. (2005) Acidosis inhibits bone formation by osteoblasts in vitro by preventing mineralization. Calcif. Tissue Int. 77, 167–174. 8. Utting, J. C., Robins, S. P., Brandao-Burch, A., Orriss, I. R., Behar, J., and Arnett, T. R. (2006) Hypoxia inhibits the growth, differentiation and bone-forming capacity of rat osteoblasts. Exp. Cell Res. 312, 1693–1702.

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Chapter 4 Isolation of Primary Avian Osteocytes Cor M. Semeins, Astrid D. Bakker, and Jenneke Klein-Nulend Abstract Osteocytes can be isolated from chicken calvaria using mild EDTA treatment alternating with collagenase treatment. The cell population obtained contains both osteoblasts and osteocytes. A pure population of osteocytes is obtained following immunomagnetic separation with the osteocyte-specific monoclonal antibody MAb OB7.3. Key words: Osteocyte, Osteoblast, Avian, MAb OB7.3, Phex, Immunomagnetic separation

1. Introduction Osteocytes are the most abundant cells in bone. Although individual osteocytes are buried in an isolated position within the bone matrix, they remain in contact with one another and with cells on the bone surface by long cell processes that run via small channels, the canaliculi. Gap junctions provide intracellular contact, where the cell processes of two osteocytes meet in a shared canaliculus (1). For a long time, osteocytes were outside the main stream of bone research, but this has changed as a result of increasing interest in the mechanoregulation of bone. General consensus exists that osteocytes play a pivotal role as mechanosensors and effectors in bone (2). In addition, there is growing awareness that osteocytes may also play an important role in phosphate homeostasis (3), mineral homeostasis (4), and regulation of matrix mineralization (5, 6). The anatomical location of osteocytes, deep within bone, has proved to be a major obstacle in studying the role of osteocytes in bone metabolism. Osteocytes depend entirely on the diffusion of

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_4, © Springer Science+Business Media, LLC 2012

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oxygen, hormones, nutrients, and waste via the canaliculi, for their activities and survival. Bone explants have to be very limited in size to allow sufficient transport of nutrients to and from the osteocytes and in fetal bone tissues the osteocyte contribution to the cellular component is very small. Direct isolation of osteocytes is therefore the method of choice to study osteocyte physiology (7). However, three major problems arise. First, how does one isolate live osteocytes from the bone matrix in sufficient numbers for subsequent studies? Second, how can osteocytes be separated from other cells, and how can the population be kept homogeneous? Third, how can osteocytes be recognized in culture, as the cells tend to lose some of their morphological characteristics when they are removed from their three-dimensional tissue structure? This chapter describes the isolation of osteocytes from 18-day-old fetal chicken calvaria. Calvaria of this age are used because at this developmental stage the calvaria are not yet too heavily calcified to allow collagenase to liberate matrix-entrapped cells. The isolation of these cells is facilitated by mild EDTA treatments alternating with collagenase treatments. The choice of fetal chicken calvaria is important because in chicken the periostea from both calvaria surfaces can be relatively easily dissected, which results in a collagenase-released cell population without periosteal cells. Furthermore, fetal chicken calvaria have a much broader layer of osteoid on their surface than mouse or rat calvaria. Finally, the availability of a monoclonal antibody (MAb) that specifically recognizes avian osteocytes allows the purification of osteocytes from mixed cell populations by immunomagnetic separation. The MAb used in the separation procedure (MAb OB7.3) is also used to recognize osteocytes in cell cultures.

2. Materials 2.1. Fertilized Chicken Eggs

2.2. Media and Solutions

Incubate the eggs at 38.5ºC in a humidified air atmosphere for 18 days. The incubator should have a mechanism that turns the eggs regularly through 180°, about ten times per hour. Fertilized eggs can be stored for 2–3 weeks at 14–16°C before development of the embryo is started by incubation in the incubator. 1. Hank’s balanced salt solution (HBSS). 2. Phosphate-buffered salt solution (PBS). 3. Isolation salt solution (ISS): 100 mM NaCl, 30 mM KCl, 1 mM CaCl2, 10 mM NaHCO3, 25 mM Hepes, 5 mg/ml

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glucose, 1 mg/ml bovine serum albumin (BSA), and 7 μM Nα-tosyl-L-lysyl-chloromethane hydrochloride (BDH Biochemicals). Adjust to pH 7.4 at 37°C. The Nα-tosyl-L-lysyl-chloromethane hydrochloride is added to inhibit proteases other than collagenase (8). 4. Collagenase solution: 1 mg/ml ISS. Collagenase Type I. 5. EDTA solution: 4 mM EDTA in PBS. Adjust to pH to 7.4. 6. Wash solution: 10% inactivated chicken serum in HBSS. For inactivation, heat serum for 30 min at 56°C, and centrifuge at 200 × g for 5 min. 7. Culture medium: α-MEM fortified with 2% chicken serum, 0.2 g/l glutamine, 0.05 g/l ascorbic acid, 0.05 g/l gentamycin, and 1 g/l glucose. 8. Trypsin–EDTA (TE) solution: 0.05% trypsin 1:250 and 0.27 mM EDTA in PBS. 9. Coated bead suspension: Mix DNA-conjugated beads (CELLection™ Pan Mouse IgG kit from Dynal), with MAb OB7.3 IgG (see Subheading 2.3) and PBS to a final concentration of 15 μg IgG/8 × 107 beads/ml. 250 μl of this suspension is needed for the isolation of osteocytes from 40 calvaria. Incubate overnight at 4°C gently shaking the suspension and store at 4°C until use. Wash the beads shortly before use two times with 2% inactivated chicken serum in PBS, by placing the bead suspension in a holder next to a magnet (Dynal) that attracts the magnetic beads to one side of the tube, allowing the removal of the suspension fluid. Finally, resuspend the IgGcoated beads in 250 μl 2% inactivated chicken serum in PBS. 10. Cell strainers, 40 μm. 11. Zinc fixative: Dissolve 0.5% zinc chloride and 0.5% zinc acetate in 0.1 M TRIS–acetate buffer, pH 4.5. 2.3. Monoclonal Antibody OB7.3

The antibody MAb OB7.3 was originally raised according standard procedures by injecting bone cells isolated from calvaria of 18-dayold fetal chickens into BALB/c mice (9). In bone sections, it only recognizes osteocytes embedded in osteoid or in calcified matrix (Fig. 1a, b). In cultures of cells enzymatically isolated from fetal chicken calvaria, MAb OB7.3 stains a minority of the isolated cells. The positive cells show an osteocyte-like morphology in culture (Figs. 1c, d and 2). The antibody may be requested from Prof. Dr. J. Klein-Nulend (Dept. Oral Cell Biology, ACTA, University of Amsterdam and VU University Amsterdam, Research Institute MOVE, Amsterdam, The Netherlands; http://www.j.kleinnulend@ acta.nl).

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Fig. 1. Frozen section of calvaria (a), and air-dried cell cultures (b–d) stained with MAb OB7.3. (a, b and d: Immunofluorescence, c: Phase contrast). (a): Section of 18-day-old fetal chicken calvaria. Note that only the osteocytes inside the bone matrix are stained. (b): OBmix after one day of culture. In the lower left corner, a group of osteocytes with one MAb OB7.3 negative cell is visible; in the upper right corner, an osteoblast colony with two osteocytes is present. (c, d): Purified osteocyte population. Note the contaminating elongated fibroblast-like cell in the upper left corner. Black bar: 100 μm. (Reproduced from see ref. 10 with permission of the American Society for Bone and Mineral Research).

3. Methods 3.1. Tissue Dissection

All of the following steps must be conducted under sterile conditions. 1. Remove the eggs from the incubator after 18 days. Keep one of the eggs with the blunt side upward where the air chamber is located. Crack the top of the shell and peel the shell off to the edge of the air chamber with a pair of sterile tweezers. Stab one leg of a second pair of tweezers underneath the white shell membrane and the chorioallantoic membrane. Close the tweezers and peel both membranes off in one movement.

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Fig. 2. Scanning electron microscopical pictures of isolated osteocytes. (a): Osteocyte 5 min after seeding. The osteocyte is already attached and has formed finger-like projections in all directions. (b): Osteocytes 20 min after seeding. The cell processes in the plane of the support have elongated, whereas the processes perpendicular to the support have disappeared. Note the presence of two magnetic beads. (c): Two osteocytes have made contact with each other via their cell processes after 24 h of culture. (d): Extensive network of flattened osteocytes with many branched cell processes after 48 h of culture. (Reproduced from see ref. 10 with permission of the American Society for Bone and Mineral Research).

2. Grasp the embryo under its head with a curved forceps, lift it a little above the egg, and decapitate underneath the forceps. The body of the embryo will fall back into the egg. Transfer the head to a petri dish with some HBSS placed on ice. Resterilize tweezers and forceps by dipping into ethanol (100%) and burning off the ethanol. 3. Make a cut in the back of the neck with a small pair of scissors. Hold the head by the bill with tweezers and tear the skin off in the direction of the bill. Cut the calvaria loose along the edge and cut it into two parts through the central suture. 4. Put the calvaria halves in a droplet of sterile HBSS and remove the ectocranial and endocranial periostea with small scalpels. Transfer the “bare” calvaria halves into a petri dish with HBSS and keep on ice.

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5. Repeat these actions for all eggs. Generally, we use 40 eggs in one isolation session. Too many eggs would make the dissection procedure too long, and when the number of eggs is too low, the procedure is inefficient and relatively decreases the number of isolated osteocytes. 3.2. Isolation of OBmix

Perform all incubations in the steps below in a shaking water bath set at 37°C. 1. Transfer all calvaria halves to a small flask containing 2.7 ml ISS and 300 μl collagenase solution (final collagenase concentration: 0.1 mg/ml). Incubate 10 min and discard the supernatant (the discarded supernatants predominantly contain damaged cells and erythrocytes). 2. Repeat step 1. 3. Add 3 ml collagenase solution to the calvaria. Incubate for 15 min and discard the supernatant (this contains predominantly fibroblastic cells and some osteoblasts). Wash calvaria three times with 2.5 ml PBS each time and discard the PBS washes. 4. Add 4 ml EDTA solution to the calvaria. Incubate 10 min and collect the supernatant. 5. Wash the calvaria three times, once with 2 ml PBS, and twice with 1 ml ISS. Add the washes to the supernatant from step 4. 6. Centrifuge at 4°C for 10 min at 600 × g and resuspend the cell pellet in wash solution. This cell suspension is termed Fraction 1. 7. Add 4 ml collagenase work solution to the calvaria, incubate for 45 min, and collect the supernatant. 8. Wash the calvaria three times with 1 ml PBS. Add the washes to the supernatant from step 7. 9. Centrifuge at 4°C for 10 min at 600 × g and resuspend the pellet in wash solution. This cell suspension is termed Fraction 2. 10. Combine Fractions 1 and 2, centrifuge at 4°C for 10 min at 600 × g, and resuspend the sedimented cells in culture medium. This is the OBmix population. It contains osteoblasts, 20–30% osteocytes, and a few fibroblasts (Fig. 1c, d). 11. Determine the cell concentration with a hemocytometer. 12. Seed 1−1.5 × 106 cells into a small (25 cm2) culture flask containing 3.5 ml culture medium. Generally, 40 eggs yield four culture flasks (about 4 × 106 cells in total). 13. Culture the cells for 24 h at 37°C in a humidified atmosphere of 5% CO2 in air (see Note 1).

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1. After 16–24 h, remove the culture medium from the culture flasks, and rinse three times with PBS. Add 3 ml TE solution per flask and incubate for 5 min at 37°C. Stop the trypsin activity by adding 0.3 ml chicken serum per flask and tap the flask, containing the cells, on the lab table to loosen the OBmix cells from the bottom. 2. Disperse the cells by repeated pipetting over the surface bearing the cell layer. 3. Remove cell clumps by sieving the cell suspension through a 40 μm mesh cell strainer, centrifuge the cells at 600 × g for 10 min, discard the supernatant, and resuspend the cells in 125 μl cold (4°C) 2% inactivated chicken serum in PBS. The sieving procedure is necessary to achieve a single cell suspension. OBmix cells, in particular the osteocytes, tend to form clumps of (probably) gap junction-coupled cells. These clumps however not only contain osteocytes, but also osteoblasts that will later quickly overgrow the nondividing osteocytes (see Note 2). 4. Add 125 μl coated bead suspension and incubate for 30 min in a rotator (60 rpm) at 4°C. Take good care during this step that beads and cells remain in suspension. 5. Separate the bead-bound osteocytes from the osteoblasts with the magnet. Wash the bead-bound osteocytes four times with 2% inactivated chicken serum in PBS (using the magnet to collect the osteocytes) and resuspend the osteocytes finally in 200 μl 2% inactivated chicken serum in PBS. 6. Add a fresh batch of 125 μl coated beads to the osteoblast fraction and repeat the separation procedure. 7. Combine both two bead-bound osteocyte fractions, separate the cells with the magnet and resuspend them in 100 μl culture medium. Count the number of cells per unit volume. Now, the bead-bound osteocytes can either be seeded and cultured, or can be freed from their beads and cultured (see Note 3). 8. For immediate removal of the beads, add 4 μl releasing buffer containing 50 U DNase/μl (CELLection™ Pan Mouse IgG kit) and incubate 15 min at 37°C in a shaking water bath. 9. Separate the beads from the osteocytes with the magnet. Wash the beads two times with PBS containing 2% inactivated chicken serum to remove all osteocytes from the beads and add the washes to the liberated osteocytes. Remove the last contaminating beads with the magnet. 10. Centrifuge the cell suspension, discard the supernatant, and resuspend the osteocyte pellet in culture medium. 11. If it is not necessary to use the isolated osteocytes immediately, it is much easier to seed the bead-bound osteocytes in culture

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medium in a petri dish. The next day, the beads can simply be removed by washing the cell layer. DNAse treatment is not needed. The osteocyte yield from 40 calvaria is generally about 200,000 osteocytes (Fig. 1c, d). 12. Attached osteocytes can be removed from their support by a short treatment with TE solution. After washing and reseeding, the osteocytes reacquire their typical morphology of stellate cells in secondary culture (see Note 4).

4. Notes 1. Optimizing Isolation of OB mix. The actual procedure one has to use depends heavily on the activity of the collagenase. If you start with a new batch of collagenase, you may have to adapt the procedure. For example, step 3 may have to be shortened if too many osteocytes are isolated in this step. If too many fibroblasts are still present in Fraction 1, then step 3 should be repeated. Steps 4 and 5 have to be repeated if too many cells, in particular osteocytes, are isolated in the repeated steps. The yield of OBmix cells from 40 calvaria following the protocol in Subheading 3.2 is generally 3−4 × 106 cells. About 20–30% of these cells are osteocytes (i.e., about 106 osteocytes). Most of the OBmix cells are however, present in clumps of osteoblasts, osteocytes, or mixtures of osteoblasts and osteocytes. Trying to isolate osteocytes from this population primarily results in the isolation of clumps of osteocytes often containing osteoblastic cells with high proliferative capacity. Sieving the original OBmix population (see Subheading 3.2, step 11) before immunomagnetic separation of osteocytes has been performed results in a very low osteocyte yield. Culturing the OBmix population for 24 h allows the cells of the clumps to separate themselves from each other, thereby increasing the number of single cells after the trypsin/EDTA treatment and thus increasing the osteocyte yield. 2. Reducing numbers of contaminating cells. Generally, we use 2% chicken serum or less in the culture medium. Osteocytes are postmitotic cells. Since serum strongly stimulates cells capable of mitosis to proliferate, isolated osteocytes will quickly be overgrown by contaminating cells in the presence of serum. However, a serum concentration that is too low will lead to deterioration of cell quality. We recommend that osteocyte cultures be used for experimentation as soon as possible after isolation. 3. Purity of the osteocyte population. If the isolation procedure is used routinely, it is advisable to regularly examine the purity of

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the osteocyte populations. Contamination of the osteocyte isolate may be determined by seeding a small sample of the osteocyte suspension in a culture dish. Incubate until the cells are firmly attached (4–6 h) and immunostain the cells for the presence of MAb OB7.3. Note that the removal of the beads will not remove the antibody from the cell surface!! At least 95% of the cells should be positive for the antibody (Fig. 1d). 4. Phenotypic characterization of osteocytes. In bone, osteocytes are fully defined by their location within the bone matrix. For isolated osteocytes, other markers are needed to establish their identity. (a) Stellate morphology. Avian osteocytes reacquire the stellate morphology of osteocytes in situ after isolation and attachment (10) (Fig. 2). This may also be the case for murine osteocytes (11). (b) Antibodies. Three osteocyte-specific MAbs have been described in the literature: MAb OB7.3 (9), MAb OB37.11 (12), and MAb SB5 (13). All three antibodies are specific for avian osteocytes and do not cross-react with mammalian cells. The identity of the antigen involved is only known for MAb OB7.3, the antigen is phosphate-regulating gene with homologies to endopeptidases on the X chromosome (PHEX) (3). MAb OB7.3 is the only antibody used for osteocyte isolation so far. Other antibodies are E11, an MAb that reacts specifically with highly mature osteoblasts and with osteocytes in tissue sections of rat bone (14), and antibodies against fimbrin, an actin-bundling protein that is abundantly present in osteocytes (15). The OB7.3 antigen is easily destroyed by harsh fixation procedures. For immunostaining of osteocytes in bone tissue unfixed, air-dried, frozen sections are recommended (Fig. 1a). Cell cultures can be stained by incubating live cells with MAb OB7.3 in HBSS for 30–60 min at room temperature or by washing the cells with HBSS followed by air drying and incubation with the antibody (Fig. 1b). Also a short (10 min) fixation with 2–4% paraformaldehyde at 4°C will leave enough antigen intact for a reasonable staining. We have also successfully used zinc fixative for the fixation and immunostaining of cell cultures. Zinc fixative does not appear to damage the OB7.3 antigen. A further advantage of this fixative is that cell cultures can be kept in the fixative for a long time (days) without loss of immunoreactivity of the antigen. (c) Protein products. Several proteins, such as osteocalcin and osteopontin, have been demonstrated in or around osteocytes in relatively high amounts (16). Other proteins like matrix extracellular phosphoglycoprotein (MEPE) (17),

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dentin matrix protein 1 (DMP1) (18), and Phex (19) are highly expressed in osteocytes as compared to osteoblasts. More osteocyte-specific proteins are sclerostin (20) and E11/gp38 (21), which are not expressed in osteoblasts. CD44, a membrane-bound glycoprotein that is involved in cell attachment to matrix proteins, is generally highly expressed in osteocytes (22), but is also expressed by other cells in bone. Alkaline phosphatase, a cell surface-bound enzyme, is generally low in osteocytes, in particular when compared to osteoblasts. The production and presence of some of these proteins may not be specific for osteocytes, but especially when combined with others, they can be used as additional markers for osteocyte identification. References 1. Doty, S. B. (1981) Morphological evidence of gap junctions between bone cells. Calcif. Tissue Int. 33, 509–512. 2. Klein-Nulend, J., and Bonewald, L. F. (2008) The osteocyte, in Principles of Bone Biology (Bilezikian, J. P., Raisz, L. G., and Martin, T. J., eds.), Academic Press, San Diego, California, pp. 153–174. 3. Westbroek, I., De Rooij, K. E., and Nijweide, P. J. (2002) Osteocyte-specific monoclonal antibody MAb OB7.3 is directed against Phex protein. J. Bone Miner. Res. 17, 845–853. 4. Inoue, K., Mikuni-Takagaki, Y., Oikawa, K., Itoh, T., Inada, M., Noguchi, T., Park, J. S., Onodera, T., Krane, S. M., Noda, M., and Itohara, S. (2006) A crucial role for matrix metalloproteinase 2 in osteocytic canalicular formation and bone metabolism. J. Biol. Chem. 281, 33814–33824. 5. David, V., Martin, A., Hedge, A. M., and Rowe, P. S. N. (2009) Matrix extracellular phosphoglycoprotein (MEPE) is a new bone renal hormone and vascularization modulator. Endocrinology 150, 4012–4023. 6. Holmbeck, K., Bianco, P., Pidoux, I., Inoue, S., Billinghurst, R. C., Wu, W., Chrysovergis, K., Yamada, S., Birkedal-Hansen, H., and Poole, A. R. (2005) The metalloproteinase MT1-MMP is required for normal development and maintenance of osteocyte processes in bone. J. Cell Sci. 118, 147–156. 7. Van der Plas, A., Aarden, E. M., Feijen, J. H. M., de Boer, A. H., Wiltink, A., Alblas, M. J., de Leij, L., and Nijweide, P. J. (1994) Characteristics and properties of osteocytes in culture. J. Bone Miner. Res. 9, 1697–1704. 8. Hefley, T. J. (1987) Utilization of FPLCpurified bacterial collagenase for the isolation

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of cells from bone. J. Bone Miner. Res. 2, 505–516. Nijweide, P. J., and Mulder, R. J. P. (1986) Identification of osteocytes in osteoblast-like cell cultures using a monoclonal antibody specifically directed against osteocytes. Histochemistry 84, 342–347. Van der Plas, A., and Nijweide, P. J. (1992) Isolation and purification of osteocytes. J. Bone Miner. Res. 7, 389–396. Mikuni-Takagaki, Y., Kakai, Y., Satoyoshi, M., Kawano, E., Suxzuki, 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. Nijweide, P. J., van der Plas, A., and Olthof, A. A. (1988) Osteoblastic differentiation, in Cell and Molecular Biology of Vertebrate Hard Tissues. Ciba Foundation Symposium 136 (Evered, D. and Harnett, S., eds.), John Wiley & Sons, Chichester, UK, pp. 61–77. Bruder, S. P., and Caplan, A. I. (1990) Terminal differentiation of osteogenic cells in the embryonic chick tibia is revealed by a monoclonal antibody against osteocytes. Bone 11, 189–198. Wetterwald, A., Hoffstetter, W., Cecchini, M. G., Lanske, B., Wagner, C., Fleisch, H., and Atkinson, M. (1996) Characterization and cloning of the E11 antigen, a marker expressed by rat osteoblasts and osteocytes. Bone 18, 125–132. Tanaka-Kamioka, K., Kamioka, H., Ris, H., and Lim, S. S. (1998) Osteocyte shape is dependent on actin filaments and osteocyte processes are unique actin-rich projections. J. Bone Miner. Res. 13, 1555–1568. Aarden, E. M., Wassenaar, A. M., Alblas, M. J., and Nijweide, P. J. (1996) Immunocytochemical

4 demonstration of extracellular matrix proteins in isolated osteocytes. Histochem. Cell Biol. 106, 495–501. 17. Petersen, D. N., Tkalcevic, G. T., Mansolf, A. L., Rivera-Gonzalez, R., and Brown, T. A. (2000) Identification of osteoblast/osteocyte factor 45 (OF45), a bone-specific cDNA encoding an RGD-containing protein that is highly expressed in osteoblasts and osteocytes. J. Biol. Chem. 17, 36172–36180. 18. Toyosawa, S., Shintani, S., Fujiwara, T., Ooshima, T., Sato, A., Ijuhin, N., and Komori, T. (2001) Dentin matrix protein 1 is predominantly expressed in chicken and rat osteocytes but not in osteoblasts. J. Bone Miner. Res. 16, 2017–2026. 19. Ruchon, A. F., Tenenhouse, H. S., Marcinkiewicz, M., Siegfried, G., Aubin, J. E., DesGroseillers, L., Crine, P., and Boileau, G. (2000) Developmental expression and tissue

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distribution of Phex protein: effect of the Hyp mutation and relationship to bone markers. J. Bone Miner. Res. 15, 1440–1450. 20. Poole, K. E., van Bezooijen, R. L., Loveridge, N., Hamersma, H., Papapoulos, S. E., Löwik, C. W., and Reeve, J. (2005) Sclerostin is a delayed secreted product of osteocytes that inhibits bone formation. FASEB J. 19, 1842–1844. 21. Zhang, K., Barragan-Adjemian, C., Ye, L., Kotha, S., 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 role in dendrite elongation. Mol. Cell Biol. 26, 4539–4552. 22. Nakamura, H., and Ozawa, H. (1996) Immunolocalization of CD44 and the ERM family in bone cells of mouse tibiae. J. Bone Miner. Res. 11, 1715–1722.

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Chapter 5 Isolation of Mouse Osteocytes Using Cell Fractionation for Gene Expression Analysis Christine Halleux, Ina Kramer, Cyril Allard, and Michaela Kneissel Abstract Osteocytes are the terminally differentiated cells of the osteoblastic lineage embedded within the mineralized bone matrix. They have been identified as key players in mechanotransduction and in mineral and phosphate homeostasis. In addition, they appear to have a role in mediating bone formation, since they secrete the bone formation inhibitor sclerostin. In contrast to osteoblasts and osteoclasts, which reside on the bone surface, it has been difficult to isolate and analyze cellular and molecular properties of osteocytes due to their specific location inside the “hard” mineralized bone compartment. This chapter describes a method to isolate osteocytes from newborn mouse calvaria and adult mouse long bone, followed by immediate total RNA extraction allowing to selectively study osteocytic versus osteoblastic gene expression by quantitative real-time polymerase chain reaction (qPCR). The osteocyte-enriched cell fraction isolated by this method can further be purified by FACS and selectively expresses osteocytic marker genes, such as Dmp1 and Sost. Key words: Osteocyte isolation, DMP-1, Gene expression, DMP-GFP-sorting, FACS sorting

1. Introduction Osteocytes are the terminally differentiated cells of the osteoblastic lineage embedded within the mineralized bone matrix. They are interconnected with each other and with cells on the bone surface via long cellular protrusions, the so-called dendrites that travel inside small channels, termed canaliculi, thereby forming an extensive canalicular network throughout the mineralized bone matrix. They have been identified as critical regulators of mineral and phosphate homeostasis (1) and are thought to be key players in mechanotransduction (2). Furthermore, they appear to have a role in the mediation of bone formation, since they secrete the potent bone formation inhibitor sclerostin, encoded by the Sost gene (3). 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_5, © Springer Science+Business Media, LLC 2012

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In contrast to osteoblasts and osteoclasts, which reside on the bone surface, it has been difficult to isolate and analyze cellular and molecular functions of osteocytes due to their specific location inside the “hard” mineralized bone compartment. In addition, even if available, isolated osteocytes cultured on a cell culture dish may not be representative of osteocytes being embedded in the three-dimensional structure of the mineralized bone matrix in vivo. The isolation and culture of osteocytes from chicken calvaria has been described in the past (4–6). More recently, Gu et al. isolated osteocytes from newborn rat long bone (7) and Paic et al. (8) isolated osteocytes from newborn mouse calvaria from DMP1-GFP transgenic mice (9) by fluorescent-activated cell sorting (FACS). Following surgical intervention in human cancer patients, Eisenberger et al. isolated RNA from laser-capture microdissected human osteocytes (10). Here, we describe a method to isolate osteocytes from newborn mouse calvaria and adult mouse long bone, followed by immediate total RNA extraction allowing to study osteocytic versus osteoblastic gene expression by quantitative real-time polymerase chain reaction (qPCR). The osteocyte enriched cell fraction isolated by this method can optionally be further purified by FACS and selectively expresses osteocytic marker genes, such as Dmp1 and Sost.

2. Materials 2.1. Bone Fractionation and Sorting of GFP Positive Osteocytes by FACS 2.1.1. Tissues

2.1.2. Instruments, Material

Fractionation was performed on newborn (i.e., from perinatal mice aged 5 or 6 days) wild-type (C57BL/6 J), DMP1-GFP (9) mouse calvaria expressing green fluorescent protein (GFP) under the control of the dentin matrix acidic phosphoprotein 1 (DMP1) promoter, or adult (aged 4 months) wild-type (C57BL/6J) mouse long bones. In the case of DMP1-GFP mice, FACS sorting followed the fractionation (see Subheading 3.1.4). When isolating osteocytes from newborn calvaria, we determined that calvaria from 5- or 6-day-old mice (P5 or P6) are optimal for the protocol described here (see Note 1). 1. Centrifuge Multifuge 3S-R. 2. Disposable conical polypropylene tubes (15 ml). 3. FACS Aria (Becton Dickinson), BD FACSDiva version 6.0 and higher. 4. FastPrep FP120 Homogeniser (BIO 101, ThermoSavant). 5. Inverted microscope. 6. Needle: 0.6 × 30 mm. 7. Neubauer counting chamber.

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8. Polypropylene tube (1.5 ml). 9. Petri dish (6 and 10 cm diameter). 10. Polypropylene round-bottom tube (14 ml). 11. Screw cap polypropylene tube. 12. Section material (scissors, forceps, curved forceps). 13. Shaker 56EVC (Witec AG). 14. Stainless steel beads 5 mm (Qiagen Cat# 69989). 15. Sterilization filter 0.2 μm, 500 ml. 16. Sterilization filter 0.2 μm, 50 ml. 2.1.3. Media and Solutions

1. BD Accudrop beads (BD Biosciences, Cat# 345249). 2. BD Cytometer Setup and Tracking (CS&T) beads (BD Biosciences, Cat# 641319). 3. Bovine serum albumin, BSA. 4. Dipotassium hydrogen phosphate, K2HPO4. 5. Dentin matrix acidic phosphoprotein 1 (Dmp1) probe (Applied Biosystems, Cat# Mm00803833_g1). 6. Calcium chloride, CaCl2 × 2H2O. 7. Collagenase type IV. 8. Ethylenediaminetetraacetic acid, EDTA 0.5 M solution, pH 8.0. 9. Fetal calf serum, FCS. 10. Glucose. 11. HEPES (Sigma Cat# H7523). 12. HEPES 1 M solution. 13. L-glutamine. 14. MEM alpha medium. 15. Penicillin/streptomycin. 16. Periostin (Postn) probe Mm00450111_m1).

(Applied

Biosystems,

Cat#

17. Phosphate-buffered saline, PBS pH 7.4. 18. Dulbecco’s PBS (Dulbecco’s, 1×), without Ca2+ and Mg2+. 19. Potassium chloride, KCl. 20. Propidium iodide 1 mg/ml. 21. RNeasy MinElute Cleanup Kit (Qiagen Cat# 74204). 22. Sodium chloride, NaCl. 23. Sodium hydrogen carbonate, NaHCO3. 24. Sorbitol. 25. Sclerostin (Sost) probe Mm00470479_m1).

(Applied

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26. TRIzol reagent. 27. Water ultrapure. 28. Isolation buffer: 70 mM NaCl, 10 mM NaHCO3, 60 mM sorbitol, 30 mM KCl, 3 mM K2HPO4, 1 mM CaCl2, 0.1% BSA, 0.5% glucose, 25 mM HEPES. 29. Collagenase solution 0.2% in isolation buffer: Make up just before use. Sterilize the solution by filtration through a 0.2 μm filter and store at 4°C until use. 30. EDTA solution: 5 mM EDTA, 0.1% BSA in PBS. Sterilize by filtration through a 0.2 μm filter and store at 4°C until use. 31. MEM medium: 10% FCS, 2 mM L-glutamine, 10 mM HEPES, 100 U/μg/ml penicillin/streptomycin in MEM alpha. Sterilize by filtration through a 0.2 μm filter and store for maximum 1 month at 4°C.

3. Methods 3.1. Bone Fractionation and Sorting of GFP Positive Osteocytes by FACS

1. Decapitate the pups with scissors into 70% ethanol and transfer the heads into PBS.

3.1.1. Isolation of Newborn Mouse Calvaria

4. Dissect the calvaria as indicated by the dashed lines and remove the thin layer of soft tissue (Fig. 1b).

2. Using scissors, incise on both lateral sides of the calvaria (dashed lines) from the eyes until the base of the skull (Fig. 1a). 3. Flip the skin on top of the mouse nose.

Fig. 1. Schematic representation of newborn mouse calvaria isolation. The dashed lines show where to cut the skin (a) and how to remove the calvaria (b).

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Fig. 2. FACS sorting of newborn DMP1-GFP mouse calvaria fractions. Propidium iodide dead cell exclusion (a). Propidium iodide (Pi) is a dye that quantitatively stains cellular DNA and double-stranded RNA. Pi penetrates dead cells but is efficiently excluded from intact cells (i.e., living cells). Pi positive cells are excluded for the gating procedure. DMP1-GFP mouse osteocytes express green fluorescent protein (GFP) (b). On the calvaria fractions, FACS cell sorting was performed according to the GFP expression level [no GFP expression (neg), GFP positive expression (GFP+), and intermediate GFP expression (med)].

5. Before proceeding with the fractionation, store the calvaria in PBS. It is best to start the fractionation as soon as possible (see Fig. 2 and see Note 2). 3.1.2. Fractionation of Newborn Mouse Calvaria

1. Incubate the calvaria (between 4 and 15 calvaria/tube) in 5 ml collagenase solution at 37°C for 20 min in a disposable roundbottom tube under agitation (700 rpm). 2. Collect the digest in a disposable conical tube. 3. Rinse the calvaria once with PBS and collect this rinsing solution into the conical tube containing the initial digest. Centrifuge at 800 × g for 8 min, discard the supernatant, and resuspend the cell pellet in 0.3 ml MEM medium. Determine the number of cells present in the pellet and store it on ice (Fraction 1, see also Note 3). 4. Digest the remains of the calvaria in 5 ml EDTA solution at 37°C for 15 min under agitation (700 rpm). 5. Collect the digest in a new disposable conical tube. 6. Rinse the calvaria remains once with PBS, collect this rinsing solution, and mix it with the digest of step 5. Centrifuge at 800 × g for 8 min, discard the supernatant, and resuspend the cell pellet in 0.2 ml MEM medium. Determine the number of cells present in the pellet and store the pellet on ice (Fraction 2, osteoblast-enriched fraction). 7. Incubate the remains of the calvaria in 5 ml collagenase solution at 37°C for 20 min under agitation (700 rpm). 8. Collect the digest in a new disposable conical tube. 9. Rinse the calvaria remains once with PBS, collect this rinsing solution and mix it with the digest of step 8. Centrifuge at

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800 × g for 8 min, discard the supernatant and resuspend the cell pellet in 0.1 ml MEM medium (Fraction 3). 10. Repeat step 7–9 and resuspend the cell pellet in 0.06 ml MEM medium (Fraction 4). 11. Digest the remains of the calvaria in 5 ml EDTA solution at 37°C for 15 min under agitation (700 rpm). 12. Collect the digest in a new disposable conical tube. 13. Rinse the calvaria remains once with PBS, collect this rinsing solution, and mix it with the digest of step 12. Centrifuge at 800 × g for 8 min, discard the supernatant, and resuspend the cell pellet in 0.06 ml MEM medium (Fraction 5, osteocyteenriched fraction). 14. Incubate the remains of the calvaria at 37°C for 20 min in 5 ml collagenase solution under agitation (700 rpm). 15. Collect the digest in a new disposable conical tube. 16. Rinse the calvaria remains once with PBS, collect this rinsing solution, and mix it with the digest of step 15. Centrifuge at 800 × g for 8 min, discard the supernatant, and resuspend the cell pellet in 0.06 ml culture medium (Fraction 6, osteocyteenriched fraction). 17. Repeat last steps 14–16 twice to get Fractions 7 and 8 (see Note 4). 3.1.3. Fractionation of Adult Mouse Long Bone

1. Euthanize the mice with CO2. 2. Decapitate the mice with scissors. 3. Dissect the femur and remove the skin and muscle around the femur as quickly as possible. 4. Cut away both femur ends and flush out the bone marrow with PBS using a needle. 5. Transfer the femora (we recommend 2–12 femora/tube) into 5 ml collagenase solution in a round-bottom tube. Incubate the femora at 37°C for 20 min (30 min is also appropriate) under agitation (700 rpm). 6. Collect the digest in a new disposable conical tube. 7. Rinse the rest of the femora once with PBS, collect this rinsing solution, and mix it with the digest of step 6. 8. Centrifuge at 800 × g for 8 min, discard the supernatant, and resuspend the cell pellet in 1 ml MEM medium if cell number determination is needed (Fraction 1). Store fraction on ice. (If no cell number determination is needed, skip step 8 and proceed directly to step 9, see Note 3). 9. Centrifuge Fraction 1 at 800 × g for 8 min, discard the supernatant, and resuspend the pellet in 1 ml TRIzol. Store it at −80°C until later RNA extraction.

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10. Digest the remainder of the femora in 5 ml EDTA solution in PBS at 37°C for 20 min (30 min is also appropriate) under agitation (700 rpm). 11. Collect the digest in a new disposable conical tube. 12. Rinse the rest of the femora once with PBS, collect this rinsing solution, and mix it with the digest of step 11. Centrifuge at 800 × g for 8 min, discard the supernatant, and resuspend the cell pellet in 1 ml TRIzol (unless cell determination is needed). Store at −80°C for later RNA extraction (Fraction 2). 13. Digest the remains of the femora in 5 ml collagenase solution at 37°C for 1 h under agitation (700 rpm). Repeat steps 11 and 12 to obtain the Fraction 3 (osteoblast-enriched fraction). 14. Repeat step 13 to obtain Fraction 4. 15. Digest the remainder of the femora in 5 ml EDTA solution at 37°C for 45 min under agitation (700 rpm). Repeat steps 11 and 12 to obtain the Fraction 5. 16. Transfer the remainder of each femur into 1 ml TRIzol (per femur) in a 2 ml screw cap polypropylene tube together with one stainless steel bead. Homogenize with a FastPrep Homogenizer under speed Nb. 5 for 30 s. 17. Incubate for 10 min on ice and transfer the homogenate into 1.5 ml polypropylene tube, before storing it at −80°C until RNA extraction (Fraction 6, osteocyte-enriched fraction). 3.1.4. FACS Sorting of Newborn DMP1-GFP Mouse Calvaria Fractions

1. Prepare and set up the FACS Aria cell sorter for four-way high speed sorting (nozzle 70 μm, 70 psi) using the CS&T and Accudrop beads. (BD FACSDiva software version 6.0 and higher). Use sterile filtered Dulbecco’s PBS (1×) for sorting instead of the standard BD sheath fluid. 2. Set the cooling of the injection chamber at +4°C and switch on the cooling device (+4°C) connected to the tube holder for sorted cells. 3. Place four 5 ml collection tubes containing 1 ml cold medium into the cooled holder. 4. Dilute propidium iodide (Pi) in ultrapure water to 80 μg/ml, add the Pi solution to the calvaria fractions to a final concentration of 400 ng/ml (1:200 dilution), and incubate for 20 min at +4°C in the dark to allow cell staining. 5. Dilute the Pi-stained fractions (minimum 1:4) in Dulbecco’s PBS (1×) to reduce the serum concentration when using the FACS, and to adjust cell concentration at approximately 3 × 106 cells/ml. 6. Set up the gating to exclude Pi positive events (dead cells/ dirts, PE-Texas-Red channel, filter 616/23) (Fig. 2a).

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7. Duplet exclusion can be done using FSCH/FSCW and SSCH/ SSCW plots to increase sorting purity, but reducing the final yield. 8. Define a negative (neg) intermediate (med) and positive (GFP +) GFP-expressing cell population to be sorted (FITC channel, filter 530/30). Use the four-way purity sorting mode (purity mask 32). 9. After sorting, briefly vortex collection tubes to recover the small droplets on the tube inner side, and then centrifuge each fraction at 500 × g for 5 min and immediately resuspend the cell pellet in 1 mL TRIzol reagent. Freeze the samples immediately at −80°C. 10. GFP positive sorted Fractions 1–8 are pooled as the osteocyteenriched (Ot) fraction before the RNA extraction. Similarly, sorted GFP negative Fractions 2–3 are pooled as well to constitute the osteoblast-enriched (Ob) fraction (Fig. 2b). 3.1.5. RNA Isolation

Total RNA was isolated from each fraction, or pooled fractions, using TRIzol reagent according to the manufacturer’s instructions. Contaminating genomic DNA was removed by digestion with DNaseI and the RNA was cleaned further using the MinElute Cleanup Kit, according to the manufacturer’s instructions. In addition, the eluted RNA was applied a second time to its RNeasy MinElute spin column and centrifuged for 1 min at full speed to increase the total RNA recovery. Reverse transcription and real-time PCR were then performed to quantitatively determine the mRNA levels for selected osteoblastic and osteocytic marker genes.

3.1.6. Comparison of Gene Expression in the Osteocytic and Osteoblastic Fractions

Fractions from newborn wild-type and DMP1-GFP mice as well as from adult mouse femora were analyzed (Fig. 3). Quantitative real-time PCR was essentially done as described by Keller and Kneissel (10), using 5 ng cDNA. As expected, Sost and Dmp1 expression were significantly higher in osteocytic than osteoblastic fractions of all samples tested. Periostin has been described to be differentially expressed in mouse osteoblastic and osteocytic cell lines: Kato et al. showed its expression in primary osteoblasts as well as in several osteoblastic cell lines and early osteocytic clones. In contrast, the more mature MLO-Y4 osteocytic cell line did not express periostin (11). Here, Postn was strongly expressed in both the osteoblastic and osteocytic fractions of newborn calvaria, whereas its expression tended to decrease in adult long bone and be higher in the osteoblastic than the osteocytic fraction. These data confirm the suitability and usefulness of the here described fractionation method. It is interesting to note that the Ct values of all analyzed markers were slightly lower in cell fractions from wild-type compared to those from DMP1-GFP mice, which were isolated by FACS. A possible explanation is that the RNA was

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Fig. 3. Gene expression in newborn mouse calvaria and adult mouse femur. Gene expression comparison between fractions isolated from 6-day-old wild-type (P6 wt calvaria), 6-day-old DMP1-GFP mouse calvaria (P6 DMP1-GFP calvaria), and 4-month-old wild-type mouse femora (Adult wt femur). Only the fractions isolated from DMP1-GFP mice were FACS sorted. Ob osteoblast-enriched fraction, Ot osteocyte-enriched fraction. Expression values normalized by 18 S are represented as percentage of expression in the Ob fraction and are means ± SD (n = 3 technical replicates) of a representative experiment. Ct values are indicated on the x axis.

partially degraded during the additional handling time of the FACS process. In addition, some osteocytes can be expected to be already extracted with the first EDTA treatment rounds owing to the presence of the cutting edge of the bone, thereby contaminating the osteoblastic fraction in wild-type bone, which was not further purified by an additional FACS step to remove DMP1 positive osteocytes. This interpretation is consistent with the smaller fold change in osteocyte marker gene expression between osteoblast- and osteocyteenriched fractions in wild-type relative to FACS-sorted DMP1GFP calvarial bone. Moreover, using FACS we found that GFP positive osteocytes were indeed present at lower numbers in early fractions (see Note 4). Finally, the higher fold change in osteocyte

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marker gene expression between osteoblast- and osteocyte-enriched fractions obtained from adult wild-type mice femur likely reflects spatial and temporal differences in osteocytic marker gene expression indicating higher expression of osteocyte marker genes in adult femora than in postnatal calvaria.

4. Notes 1. We have compared the fractionation of newborn calvaria from 3-, 6-, and 9-day-old mice. Based on our experience with the expression analysis of many different genes, it appears that it is very important to perform the fractionation on mice of exactly the same age. In addition, 6-day-old mice are more suitable for the protocol described here than 3- or 9-day-old mice (Fig. 4). The time of fractionation needs to be carefully adapted for mice at any given age due to changes in bone structure and thickness. 2. Until dissolution of the cell pellet in TRIzol, it is important to carry out the whole fractionation and the following FACS as rapidly as possible to minimize possible gene signature modifications due to the isolation conditions (such as cell stress, apoptosis, etc.). 3. In case no FACS sorting or cell counting is needed, each fraction is resuspended directly in 1 ml TRIzol instead of MEM medium. Direct FACS sorting into TRIzol does not provide good quality RNA.

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4. Fractionation can usually be stopped after Fraction 7 because very few cells (osteocytes) are still coming off in Fraction 8 (Fig. 5). Given the small amount of cells per fraction obtained from bone of newborn mice, it might be advantageous to pool several fractions after FACS sorting and before RNA isolation.

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For example, GFP negative cells from Fraction 2 and 3, representing the osteoblast-enriched fractions, were pooled. Likewise, GFP positive cells from Fractions 1–7, representing osteocytes, were pooled.

Acknowledgments We are most grateful to T. Grabenstaetter and G. Guiglia for excellent technical skills and to Dr. F. Raulf (Novartis Institutes for BioMedical Research, Basel, Switzerland) for giving access to his FACS device and providing valuable advice. We also would like to thank Profs. D. W. Rowe and I. Kalajzic (University of Connecticut Health Center, Farmington, CT, USA) for sharing the DMP1GFP transgenic mouse line. References 1. Feng, J. Q., Ye, L., and Schiavi, S. (2009) Do osteocytes contribute to phosphate homeostasis? Curr. Opin. Nephrol. Hypertens. 18, 285–291. 2. Bonewald, L. F., and Johnson, M. L. (2008) Osteocytes, mechanosensing and Wnt signaling. Bone 42, 606–615. 3. van Bezooijen, R. L., ten Dijke, P., Papapoulos, S. E., and Löwik, C. W. (2005) SOST/sclerostin, an osteocyte-derived negative regulator of bone formation. Cytokine Growth Factor Rev. 16, 319–327. 4. van der Plas, A., and Nijweide, P. J. (1992) Isolation and purification of osteocytes. J. Bone Miner. Res. 7, 389–396. 5. Tanaka-Kamioka, K., Kamioka, H., Ris, H., and Lim, S. S. (1998) Osteocyte shape is depen-

dent on actin filaments and osteocyte processes are unique actin-rich projections. J. Bone Miner. Res. 13, 1555–1567. 6. Kamioka, H., Ishihara, Y., Ris, H., Murshid, S. A., Sugawara, Y., Takano-Yamamoto T., and Lim, S. S. (2007) Primary cultures of chick osteocytes retain functional gap junctions between osteocytes and between osteocytes and osteoblasts. Microsc. Microanal. 13, 108–117. 7. Gu, G., Nars, M., Hentunen, T. A., Metsikkö, K., and Väänänen, H. K. (2006) Isolated primary osteocytes express functional gap junctions in vitro. Cell Tissue Res. 323, 263–271. 8. Paic, F., Igwe, J. C., Nori, R. , Kronenberg, M. S., Franceschetti, T., Harrington, P., Kuo, L., Shin, D. G., Rowe, D W., Harris, S. E., and Kalajzic, I. (2009) Identification of differen-

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tially expressed genes between osteoblasts and osteocytes. Bone 45, 682–692. 9. Kalajzic, I., Braut, A., Guo, D., Jiang, X. Kronenberg, M. S., Mina, M., Harris, M. A., Harris, S. E., and Rowe, D. W. (2004) Dentin matrix protein 1 expression during osteoblastic differentiation, generation of an osteocyte GFP-transgene. Bone 35, 74–82.

10. Keller, H., and Kneissel, M. (2005) SOST is a target gene for PTH in bone. Bone 37, 148–158. 11. 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.

Chapter 6 Studying Osteocyte Function Using the Cell Lines MLO-Y4 and MLO-A5 Jennifer Rosser and Lynda F. Bonewald Abstract We describe the culture and use of MLO-Y4 cells in studies of gene expression, response to fluid flow, and dendrite growth. We also describe how to use the MLO-A5 cells as a model of osteoblast to osteocyte differentiation and how to study their mineralization. These studies serve as a beginning point to study osteocyte functions and molecular mechanisms responsible for these functions. Key words: Osteocyte, Mineralization, Dendrites, E11, Fluid flow, MLO-Y4, MLO-A5

1. Introduction Originally, osteocytes were largely defined by their anatomical position within bone and their dendritic morphology. Later, in 1995, molecular markers were identified and they were defined as being low in alkaline phosphatase while high in osteocalcin and casein kinase II (1). Within the past two decades, selective osteocyte markers have been identified including E11/gp38/podoplanin for early, embedding osteocytes, followed by Sost/sclerostin, a marker for the mature osteocyte. In addition, several proteins that play critical roles in phosphate homeostasis, including phosphateregulating gene with homologies to endopeptidases on the X chromosome (PHEX), matrix extracellular phosphoglycoprotein (MEPE), dentin matrix protein 1 (DMP1), and fibroblast growth factor-23 (FGF23) were identified as important and selective osteocyte-expressed genes. Osteocytes are enriched in a number of proteins important in cytoskeletal function, including destrin, CapG, cdc42, E11/gp38, and are also enriched in molecules involved in muscle contractile function, including myosin heavy

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and light chains, α-actin, troponins, tropomyosins and α-actinin and the actin binding protein, Capzb (2). Osteocyte selective markers have distinctive functions. E11/ gp38 appears to play a role in dendrite formation, an essential process for osteocytes to generate their network within bone. Sclerostin is a negative regulator of bone formation. Mutation of the SOST gene causes high bone mass in humans, and deletion in mice results in high bone mass. Phex is specific for avian osteocytes but is found in late osteoblasts and osteocytes in mammals. Dmp1 and MEPE are highly expressed in osteocytes compared to osteoblasts or other cell types. Deletion or mutation of either Phex or DMP1 results in hypophosphatemic rickets, which is associated with a dramatic increase in FGF23 expression in osteocytes (2–5). Isolating primary osteocytes is difficult, and yields decrease, with increased mineralization and age of bone, thereby making it difficult to obtain sufficient numbers of cells for many experimental purposes. We, therefore, generated the MLO-Y4 and MLO-A5 cell lines to study osteocyte function (6, 7). Both cell lines were derived from a transgenic mouse in which the immortalizing T-antigen was expressed under control of the osteocalcin promoter. MLO-Y4 cells exhibit properties of osteocytes including high expression of osteocalcin, low expression of alkaline phosphatase, high expression of connexin 43 and the antigen E11, and retain a dendritic morphology, similar to that observed in primary osteocyte cultures. This cell line has proved useful to study effects of fluid flow and substrate stretching on activation of osteocyte-specific signaling pathways and on the release of signaling factors, especially prostaglandin, nitric oxide, ATP, and calcium. Other studies reported have included the role of cilia, gap junctions, and hemichannels, dendrite formation, apoptosis, autophagy, hypoxia, hormonal responses, control of osteoclast formation and activation, and effects on osteoblasts and mesenchymal stem cells. This cell line is exquisitely sensitive to fluid flow shear stress unlike other known fibroblast and osteoblast cell lines (2–5). The MLO-A5 cell line has characteristics of a postosteoblast/ preosteocyte. These cells are very large, over 100 μm, express markers of the late osteoblast such as extremely high alkaline phosphatase, bone sialoprotein, PTH type 1 receptors, and osteocalcin, and rapidly mineralize in sheets, not nodules (7). In culture, these cells start to express markers of mature osteocytes such as E11/ gp38/pdpn as they generate cell processes (8). PTH and mechanical load decreases Sclerostin expression in these cells (9), while BMP increases it (10). MLO-A5 cells have been used to study the osteoblast to osteocyte differentiation process (11), the mineralization process (8), and the effects of mechanical loading on biomineralization (12). As with any cell line, key characteristics are used to monitor the stability of the cell phenotype. For the MLO-Y4 cells, these

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characteristics are generation of dendritic processes, high E11, high osteocalcin, and low alkaline phosphatase. For the MLO-A5, the major characteristic is to form a “honeycomb”-like mineralized matrix within 7–9 days of culture. In this chapter, we describe the culture of the MLO-Y4 and A5 cell lines, their morphology, application of fluid flow shear stress, how to measure dendrite elongation, transfection for siRNA studies, and mineralization assays.

2. Materials 2.1. Tissue Culture Reagents

1. Culture medium: α-MEM with Earle’s salts, L-glutamine, ribonucleosides, and deoxyribonucleosides; supplemented with penicillin/streptomycin at 100 U/ml. 2. Fetal Bovine Serum (FBS) and Bovine Calf Serum (CS) (defined & iron-supplemented); both heat inactivated (Hyclone) (see Note 1). 3. Dulbecco’s Phosphate-Buffered Saline (DPBS) without calcium and magnesium, pH 7.4. 4. 0.05% Trypsin/0.53 mM EDTA solution for passaging cells. 5. Rat tail type I Collagen solution (Becton Dickson Bioscience, Cat.# 354236)for coating plates to maintain cells, or buy precoated plates. 6. 0.02 M Acetic acid for diluting collagen. 7. Dimethyl sulfoxide (DMSO) for freezing cells. 8. Ascorbic acid for mineralizing osteoblast cultures. 9. β-Glycerophosphate for mineralizing osteoblast cultures. 10. Tissue culture treated dishes (100 and 150 mm).

2.2. Fluid Flow Reagents

1. Flexcell® Streamer® Shear Stress Device (Flexcell) includes the following: Pump, Fluid flow chamber with hex bolts and hex wrench, glass bottle, pulse dampeners, latex tubing and quick disconnect fittings. 2. Collagen-coated microscope slides, for example Thermo Scientific Colorfrost* Plus Slides(6776214), which are 25 × 75 × 1 mm, or use Flexcell “Culture slips” (see Note 2). 3. Sterilized forceps for manipulating the slides. 4. Square Petri dishes that can hold three microscope slides. 5. Cell Culture Media for growing cells and performing fluid flow. 6. 70% ethanol and sterile PBS for sterilizing and rinsing the system, respectively. 7. Deionized water and 70% ethanol for cleaning the system.

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2.3. Dendrite Quantitation Reagents

1. 10% buffered formalin.

2.4. siRNA Reagents

1. Opti-MEM® I Reduced Serum Medium without serum (Invitrogen).

2. 0.1% Crystal Violet.

2. Oligofectamine (Invitrogen) (see Note 8). 3. Risc-Free siRNA as a negative control. 4. siRNA to 3 regions of E11 in the form of 21 base nucleotides for transient infection. siRNA “A” and “B” are based on the sequence used for human E11 siRNA (13). siRNA “C” was designed by Ambion (Austin, TX) with the help of the Cenix algorithm. A = +165 ACTGGAGGGCTTAATGAATCT +185 B = +397 AAGATGGCTTGCCAGTAGTCA +417 C = +66 AGGGACTATAGGCGTGAATGA +86 2.5. E11 Western Blotting

1. Primary antibody: 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 this monoclonal antibody and corresponding hybridoma with the Developmental Studies Hybridoma Bank (http://www.uiowa. edu/~dshbwww/) 8.1.1 (anti-murine GP38; T1alpha; podoplanin) and it is available from there on request. 2. Secondary antibody: Peroxidase-conjugated goat anti-Syrian hamster IgG- (ImmunoResearch Laboratories Inc).

3. Methods 3.1. General Maintenance of MLOY4 OsteocyteLike Cell-Line

All cell culture procedures should be performed in a Tissue Culture hood under aseptic conditions, using sterile supplies. Figure 1 shows the typical appearance of the cells in culture. 1. Maintain stock cultures in α-MEM culture medium containing 2.5% FBS and 2.5% CS at ~60–75% confluence on collagencoated plates (see Subheading 3.2) in a humidified incubator at 37°C and at 5% CO2 (see Note 3). 2. Passage at ~ 1:5 dilution (~1:7maximum), every 3–4 days (see Note 4). 3. Freeze cells in 60% α-MEM, 30% FBS, 10% DMSO, at 1 × 106 cells/ml/cryovial. 4. When defrosting cells, place the frozen vial of cells in a 37°C water bath just long enough to thaw. Transfer the cell suspension into a 15-ml conical tube with 9 ml of media + serum, invert to mix.

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Fig. 1. Appearance of the MLO-Y4 cells at 4 and 12 h after plating. The cells are attached by 4 h and already generate dendritic processes by 12 h.

5. Centrifuge at approx. 1,000 rpm, for 5–10 min. Aspirate media, gently resuspend and plate the cell pellet in media containing 5% FBS and 5% CS. This higher amount of serum, 10% total, is useful to give the cells an extra boost. The next day, check the viability of the cells. If there are a lot of floating “dead” cells, change the medium. You can revert back to medium with 2.5% FBS + 2.5% CS at this point. 3.2. Collagen Coating of Tissue Culture Treated Dishes

Perform this procedure under aseptic conditions and dry plates in a sterile hood. 1. Using a chilled pipette, dilute the concentrated, sterile collagen into previously filter- sterilized 0.02 M acetic acid to a final concentration of 0.15 mg/ml (working dilution). Use 8 and 14 ml of working dilution per 10 and 15 cm dishes, respectively.

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2. Coat the plates for 1 h at room temperature. Then, tilt the plate at an angle for several minutes and remove the solution, which can be reused approximately six times and should be kept in the refrigerator between uses. 3. For immediate use, rinse the plate with PBS to remove residual acid. Alternatively, plates can be stored. To do so, completely dry the plates with the lid off (>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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5% serum

2.5% serum

1% serum

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20 0

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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

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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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2.11. Oil Red O Staining

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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.

9 3.1.2. Immunomagnetic Purification of Osteoclasts (see Note 8)

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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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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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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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3.3. Purification of Rabbit Osteoclasts on Plastic (from Subheading 3.1)

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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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8.

9.

10.

11.

12.

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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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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 (<1 min) thaw the vial (e.g., in a 37°C water bath or by rapidly rubbing between palms), resuspend the cells in a small amount (~0.5 ml) of culture medium, and add the cell suspension to a T25 tissue culture flask. Increase the volume in the flask to 10 ml with additional culture medium and place into a tissue culture incubator (day 0). 2. On day 3, withdraw the spent medium and refeed the cells with 10 ml of fresh medium (see Note 1). 3. Culture the cells until confluent (typically 4–5 days). 4. To subculture the confluent RAW cells, withdraw the spent medium, add 10 ml of fresh medium to the flask, and scrape the cell layer into this fresh medium using a rubber scraper (see Note 2). 5. Immediately add 0.01 ml of the cell suspension to 0.09 ml of fresh medium in a microfuge tube, mix gently, and count the cells using a hemocytometer. 6. Calculate the cell concentration. Plate the RAW cells at 1.5 × 105 cells/cm2 into tissue culture dishes of the desired size. Typically, one confluent T25 flask will provide a sufficient number of RAW cells to seed two 100-mm dishes or 24-well dishes (see Notes 3 and 4). Increase these volumes

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with additional medium as needed to yield 8 ml per 100-mm dish or 0.5 ml (or 1.0) per well of a 24-well dish, and then place the cells into a tissue culture incubator. 7. To grow the RAW cells for an extended period of time, refeed the cultures every 2–3 days and subculture when they reach confluency as in steps 4–6. 3.3. RAW-OC Formation (See Note 4)

This method is based on the published procedure of Hsu et al. (10). 1. Culture RAW cells to confluency (see Subheading 3.2, steps 1–3). 2. Subculture confluent RAW cells into 24-well dishes as described in Subheading 3.1, steps 4–6 (see Notes 3–6). If the cells are to be used for cytochemical or immunological staining, replate the RAW cell suspension into 24-well dishes that contain a sterile glass coverslip in each well. If bone resorption is to be evaluated in parallel with OC development in the RAW cell cultures, replate the RAW cell suspension into 24-well dishes that contain 2–4 small disks of bone or ivory (see Subheading 3.1) per well (with or without a glass coverslip under the disks). 3. Immediately add soluble recombinant RANKL to the dishes at a final concentration of 35 ng/ml to initiate OC development (day 0) and increase the volume in the wells to 0.5 ml (or 1.0) with additional culture medium (see Note 7). 4. Culture to day 3. Briefly examine the cells under a microscope for evidence that RAW cells are beginning to fuse into multinucleated RAW-OCs. Refeed the developing RAW-OC cell cultures with 0.5 ml (or 1.0) of fresh medium containing 35 ng/ml RANKL. 5. Culture until day 5 or 6 when many multinucleated RAW-OCs have formed but have not completely covered the dish (see Note 8). The day 5 or 6 RAW-OC populations may be immediately fixed and used for cytochemical or immunological staining, harvested for biochemical or molecular studies, or analyzed for bone resorption (see other Chapters in this volume). For greater bone resorption, such cultures may be incubated until days 7–9. Alternatively, RAW-OCs can be purified further by serum gradient density fractionation (see Subheading 3.4).

3.4. Serum Gradient Purification of RAW-OC

Because not all RAW cells fuse into multinucleated RAW-OCs by day 5 or 6, those that have can be purified from the remaining mononuclear cells using serum density gradient fractionation (see Note 9). This procedure is a modification of the one we routinely use to purify in vitro formed OCs or OC-like cells from chick or human origin (15). Directions are provided for RAW-OCs formed on 100-mm tissue culture dishes. All steps are conducted at room

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temperature, unless otherwise noted, and are performed in a sterile hood using sterile solutions and supplies. 1. Remove the spent culture medium from two 100-mm dishes of day 5 or 6 RANKL-generated RAW-OCs. 2. Gently add 10 ml of MHB to each 100-mm dish to wash the cells. Remove and discard the washes. 3. Repeat step 2 to wash the RAW-OCs twice more with MHB. 4. Add 10 ml of MHB to each dish and place them into a tissue culture incubator at 37°C for 15 min. 5. Remove and discard the MHB solution from each dish. 6. Add 5 ml of freshly prepared collagenase–trypsin digestion solution to each dish and incubate at 37°C for 5 min. 7. Remove the dishes from the incubator and shake the plates gently by hand back and forth (e.g., slide the dish on a flat surface) for ~30 s to detach and loosen the interaction of cells with extracellular matrix produced by the cells. 8. Completely remove the collagenase–trypsin solution containing the released matrix material from each dish and discard (see Note 10). 9. Gently wash the adherent cells on each dish by releasing 10 ml of PBS slowly against the side wall of the dish. Completely remove and discard the washes. 10. Repeat step 9 to wash the cells on each dish with PBS twice more. 11. Add 5 ml of protease–EDTA digestion solution to each dish. Incubate at 37°C for 10–15 min (see Note 11). 12. Loosen the adherent cells on each dish by flushing the protease– EDTA incubation solution with a pipette gently over the surface of the cell layer to free the cells (see Note 12). 13. Transfer the cell suspensions from two 100-mm dishes into one 50-ml sterile centrifuge tube containing 1.0 ml FBS (to inhibit further protease action). 14. Centrifuge the cells at 100 × g for 5 min. 15. Remove and discard the supernatant. Gently resuspend the cell pellet in 15 ml of MHB by repeatedly drawing up and releasing from a pipette (not too vigorously, see Note 12). 16. Prepare 16 ml of 70% FBS in MHB (11.2 ml FBS plus 4.8 ml of MHB) in a 50-ml centrifuge tube, and 16 ml of 40% FBS in MHB (6.4 ml FBS plus 9.6 ml MHB) in another 50-ml tube. 17. Prepare an FBS gradient in a 50-ml round-bottom centrifuge tube. To do this, carefully dispense 15 ml of the 70% FBS-MHB solution (from step 16) into the bottom of the tube. Very slowly overlay this with 15 ml of the 40% FBS-MHB solution (from

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step 16), using a pipette held at a 45° angle against the side of the tube just above the 70% FBS layer and slowly releasing the 40% FBS solution in a thin stream so as not to deform the surface of the 70% FBS layer. 18. Let the tube stand undisturbed for 30 min (at room temperature) to allow the larger multinucleated RAW-OCs to settle under normal gravity and penetrate the FBS layers (see Note 13). 19. Carefully take off the top 17 ml which contains mononuclear cells, and transfer it into a 50-ml tube. 20. Then, remove the 16 ml middle layer, which contains primarily mononuclear cells and some small multinucleated RAW-OCs, and transfer this into another 50-ml tube. 21. The bottom 12 ml fraction contains predominantly large multinucleated RAW-OCs. 22. Centrifuge the purified RAW-OC bottom fraction (and the other fractions if they are also to be cultured and/or analyzed) at 100 × g for 5 min. 23. Gently resuspend the RAW-OC pellet in culture medium, count an aliquot in a hemocytometer, and plate 1,000–4,000 cells per well of a 24-well dish. Typically, the purified RAWOCs from two 100-mm dishes can be cultured in 2–10 wells of a 24-well dish (with 0.5–1.0 ml medium per well) for 6–24 h (see Note 14). The top and middle fractions from the serum gradient fractionation are typically cultured in 20–40 wells and 15–30 wells of a 24-well dish, respectively. Alternatively, RAWOCs (and the top and middle fractions, if desired) may be used immediately for analysis (see Subheading 3.2, step 5). Serum gradient fractionation routinely provides 4,000–10,000 purified RAW-OCs from one 100-mm dish (this depends on the efficiency of one’s technique and, more importantly, on the exact stage of RAW-OC used to purify the cells; see Notes 6, 8, 13, and 14). In unfractionated RANKL-generated RAW-OC cultures, multinucleated (more than three nuclei) RAW-OCs typically represent ~1% on a per cell basis and 25% on a per nuclear basis of the total cell population (Fig. 1, left panel). By contrast, serum gradient purified RAW-OCs (with more than three nuclei) typically comprise 60–90% on a per cell basis and 96% on a per nuclear basis of the total cell population in the bottom serum fraction (Fig. 1, lower right panel). On average, RAW-OCs in this bottom serum fraction contain 15–30 nuclei per cell. 3.5. Phenotypic and Functional Characterization of RAW-OCs

Standard protocols can be used to evaluate the morphological (light, scanning electron microscopy), ultrastructural (transmission electron microscopy), histochemical (general or enzymatic activity stains), or immunocytochemical staining (e.g., for OC developmental markers) characteristics of RAW cells representing pre-OCs and

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Fig. 1. RANKL-mediated RAW-OC formation and serum gradient purification. (Left ) RAW cells were cultured with 35 ng/ml murine recombinant RANKL for 6 days and then subjected to serum gradient fractionation. A well cultured in parallel was fixed and stained for TRAP activity to show the proportion of mononuclear and multinucleated TRAP+ cells that arise by day 6 in RANKL-differentiated RAW cell cultures. The cells were viewed using a light microscope and images were captured with a computer-linked digital camera. (Reduced from original magnification, ×100.) (Right ) The top, middle, and bottom fractions from the serum gradient fractionation were replated and cultured on plastic for several hours, after which the cells were fixed and stained for TRAP activity. (Upper right ) The top fraction consists entirely of mononuclear cells, some of which are TRAP+ (in contrast to untreated RAW cells which are all TRAP-, not shown). (Reduced from original magnification, ×200.) (Middle right ) The middle fraction primarily contains mononuclear cells, a portion of which are TRAP+, and some small multinucleated RAW-OC. (Reduced from original magnification, ×200.) (Lower right ) The bottom fraction consists primarily of large multinucleated RAW-OC, although a few mononuclear cells may still be present. (Reduced from original magnification, ×100).

in vitro RANKL-formed RAW-OCs (see Chapter 9, this volume). Whereas untreated RAW cells do not stain for tartrate resistant acid phosphatase (TRAP) activity, a key marker and enzyme involved in OC bone resorption, RANKL-differentiated RAW cell cultures develop both TRAP+ mononuclear and multinucleated cells (Fig. 2a, c). The RAW-OCs formed by cell fusion contain multiple nuclei clustered together, and the cells may appear either spread out or partially elongated when cultured on plastic (Fig. 2a). RAW-OCs cultured on bone or ivory (either during RANKL development or following replating of the differentiated cells) frequently display a more compact and highly motile elongated shape with numerous pseudopodial extensions (Fig. 2c). Resorption pits formed by RAWOCs are typified by clusters of multilobulated excavation cavities or long resorption tracks (which may also be multilobulated) adjacent to or underlying RAW-OCs actively engaged in bone resorption (Fig. 2c). Molecular, immunological, and/or biochemical analyses have shown that RAW-OCs express the key hallmark properties of OCs including TRAP, calcitonin receptor, cathepsin K, matrix

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Fig. 2. RANKL-mediated RAW-OC or MA-OC formation and bone pit resorption. (a, c) RAW cells were cultured with 35 ng/ ml murine recombinant RANKL for 6 days on plastic (a) or ivory (c), and then fixed and stained for TRAP activity. Note the well spread morphology of RAW-OCs on plastic (a) compared with the more compact and motile phenotype of such cells actively engaged in bone resorption on ivory (c). Abundant resorption pits and tracks were evident that were frequently composed of connecting excavation cavities. These represent periods of RAW-OC attachment and pit formation, followed by RAW-OC movement to an adjacent area of ivory for further resorption. (a) and (b) reduced from original magnification, ×200). (b, d) Murine bone marrow cells were isolated and cultured at 5.6 × 105 cells per well of a 24-well dish (1.9 cm2 per well) with 25 ng/ml of murine M-CSF and 35 ng/ml of murine RANKL for 6 days on plastic (c) or ivory (d), after which the cells (MA-OCs) were fixed and stained for TRAP activity. Like RAW-OCs, the TRAP+ MA-OCs were well spread on plastic (b) and more compact on ivory (d). Resorption pits and tracks formed by MA-OCs (d) were indistinguishable from those formed by RAW-OCs (b). (b) and (d) reduced from original magnification, ×100 and ×200, respectively).

metalloproteinase-9, integrin αvβ3, and c-src (refs. 10, 16, our unpublished data). Both the phenotypic and functional characteristics of RANKL-differentiated RAW-OCs resemble those of in vivo formed isolated murine OCs or RANKL-differentiated OCs (MAOC) formed from murine bone marrow cells in the presence of M-CSF (Fig. 2b, d). Thus, like RAW-OCs, TRAP+ MA-OCs exhibit a well-spread morphology on plastic (Fig. 2b) and a more compact, motile phenotype on bone or ivory (Fig. 2d). Multilobulated resorption pits and tracks formed by MA-OCs (Fig. 2c) also resemble those formed by RAW-OCs, with well defined margins and deep resorption lacunae (Fig. 2d). Resorption pit formation by RAW-

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OCs, in the presence or absence of modulators, can be quantified as for other OCs (see Chapters 8 – 12, this volume). In addition to these phenotypic and functional analyses, RAW-OCs provide abundant material (protein, RNA, etc.) for investigations of gene or protein expression or microarray profiling, receptors and signal transduction pathways, production of various factors (cytokines, chemokines, growth factors, and arachidonic acid metabolites), release of other substances (free radicals and enzymatic activities), cell and matrix interactions, and diverse regulatory mechanisms (particularly since RAW cells are more easily transfected than primary bone marrow cells) (see Note 15).

4. Notes 1. Some DMEM formulations may produce a visible dark precipitate that causes rapid cell death of RAW cells during culture. In such cases, we find it best to obtain new DMEM lacking Fe(NO3)3 along with a separate stock (1,000×) of Fe(NO3)3 that is stored in aliquots (1 ml) at −20 C. This iron stock is only thawed and added to DMEM at the time that complete medium is prepared (containing FBS and a/a) for current use. Typically, this complete medium is usable for 10 days to 2 weeks for cell refeeding before evidence of precipitation occurs, at which time any remaining medium should be discarded. Some investigators have reported that RAW cells also grow and form OCs well in α-MEM medium, so this may be an alternative to DMEM if problems are encountered with the latter. 2. We find that a rubber-tipped cell scraper works best because it completely contacts the surface of the tissue culture flask (or dish) and causes the least cell damage. 3. In general, RAW cells should be subcultured at a ratio of 1:3–1:6. 4. If more cells will be needed than are provided by a reasonable number of T25 flasks, the confluent RAW cells can be subcultured into T75 flasks (at a 1:3–1:6 ratio) and then grown to confluency. 5. The number of RAW cell passages affects RANKL-mediated OC formation. In our hands, RAW-OCs seem to better form after passage 4 and will no longer form in response to RANKL stimulation once they have undergone 18–20 passages from the time that they were received from the ATCC repository. The reason for this is not fully clear, although other researchers have similarly noted that not all RAW 264.7 cell lines (or passages?) will form OCs after RANKL treatment, and subclones of RAW 264.7 cells can be derived that are more or less efficient at

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RANKL-mediated OC formation (17, 18). It is also possible that particular lots of FBS may differentially influence RANKLmediated RAW-OC formation. Therefore, different lots and sources of FBS may be tested if there are difficulties encountered in trying to form RAW-OCs, although RAW-OCs have been formed even in serum-deprived conditions (19). 6. The density of RAW cells plated affects the rate and yield of RAW-OC development, as well as the subsequent analysis of RAW-OCs formed. Too low a cell density (100–500 cells/ cm2) delays RAW-OC formation and decreases the final yield. For most purposes (e.g., testing the effects of various agents on RAW-OC development), the plating density for RAW cells should be in the range of 103–3 × 104/cm2 to facilitate counting or characterization of the RAW-OCs formed and still generate a sufficient number for analysis. If RAW-OCs are to be purified by serum density gradient fractionation, the initial plating density of RAW cells should be considerably higher (1.5 × 105/cm2) so that enough RAW-OCs are obtained following their purification (see Subheadings 3.1 and 3.2). However, too high a density of RAW cells (4.5–7.5 × 105/cm2) inhibits RAW-OC formation. 7. In our experience, the potency of recombinant soluble RANKL for inducing OC formation is dependent upon its source (not only for RAW cells, but also for human monocyte, mouse bone marrow, or chicken bone marrow preparations). Thus, different commercial RANKL preparations vary significantly in the dose required, kinetics of OC formation, and final yield of bone pit resorptive OCs obtained. This is not strictly due to species compatibility issues because human or murine recombinant RANKL are similarly efficient for inducing OCs from murine RAW or bone marrow-derived cells, chicken bone marrow cells, or human peripheral blood monocytes (although all but RAW cells require M-CSF costimulation). Although we have successfully used various commercial RANKL preparations, we typically prepare soluble recombinant mouse RANKL in our own lab that exhibits high osteoclastogenic activity with murine RAW or bone marrow cells, chicken bone marrow cells, or human monocytes. At 35 ng/ml, this mouse RANKL induces multinucleated TRAP+ cell formation that is first apparent on days 3–4 of RAW cell culture. Lower RANKL concentrations delay the kinetics and final yield (and size) of RAW-OC formation. Others have used recombinant RANKL preparations in the range of 20–100 ng/ml (depending upon its source and bioactivity) to form multinucleated TRAP+ cells that usually first appear on days 3–4 of RAW cell culture (11, 18, 20, 21). However, certain recombinant mouse RANKL preparations appear to require an additional anti-RANKL antibody

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cross-linking step to induce osteoclastogenesis (16). Therefore, it is recommended that pilot studies be performed with each new source and preparation of recombinant RANKL to ascertain an appropriate dose to achieve the level of RAW-OC formation needed (see also the discussion in Chapter 7, this volume). 8. In our model of RANKL-mediated RAW-OC development, TRAP+ cells first become apparent on day 2 of culture, and multinucleated TRAP+ cells appear on days 3–4 and nearly reach a peak on days 5–6 of culture. It is important to either use the cells or subject them to serum gradient purification at this point (and not wait another day until the full peak of RAW-OC formation has occurred) because if the cells become overconfluent and overfused, they die very rapidly (<24 h) thereafter. 9. The mononuclear cells that remain in the RANKL-treated RAW cell cultures by days 5–6 can still fuse to form more RAW-OCs upon further culture, even in the absence of any additional RANKL stimulation. Therefore, such cells represent an early stage in OC differentiation and are not equivalent to the original non-RANKL treated RAW cells (in a number of characteristics). It is not advisable to simply culture the RANKL-treated RAW cell populations for any longer period of time than days 5–6 to encourage more of the mononuclear cells to fuse into multinucleated RAW-OCs because continuing cell fusion (even 1 extra day) produces cells that are too large and fragile, and which undergo rapid and massive apoptosis in the cultures once such RAW-OCs have formed. 10. It is important to completely remove this released extracellular matrix, here and in the subsequent washes, or the cells will reattach to it and become extremely difficult to detach. At this point, the cells can be seen to have begun to pull up from the dish. 11. A rounding up from the dish becomes more obvious at this point for the RAW-OCs. The cells may appear somewhat shrunken, but should still appear bright and viable. 12. Use only wide-bore pipettes or tips for any work in isolating or manipulating OCs to avoid fragmenting these large multinucleated cells. Also, care should be taken to resuspend, mix, or vortex OC preparations gently and for as little time as necessary. 13. The extent of RANKL-mediated RAW-OC formation impacts on this step as well. Therefore, if the RAW-OCs formed are relatively small (<10 nuclei per cell), they will be unable to effectively settle into the 70% portion of the serum gradient. However, if the RAW-OCs are too large (>30 nuclei per cell), they will be too fragile, prone to break, and die too quickly so

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that few viable RAW-OCs will be recovered following the serum gradient purification. 14. Even under controlled conditions of RANKL-mediated RAW-OC formation as discussed in this chapter, once such cells have formed they tend to apoptose very rapidly and the cells can be lost within 24 h if allowed to develop too long (see also discussion in Chapter 8, this volume). Short survival after formation in culture is specifically a problem with mouse osteoclasts and not seen with for example human osteoclasts. Addition of 10 ng/ml IL-1α to promote RAW-OC survival on plastic only slows the apoptotic process slightly, and a recent report indicates that it preferentially activates larger over smaller RAW-OCs (22). Therefore, we typically use RAW-OCs formed in tissue culture dishes by days 5–6 for analysis within 6–24 h (e.g., staining, RNA or protein extraction, etc.). If RAW-OCs have been formed on bone or ivory, resorption pits are usually evident by day 4 and maximal by day 6 or 7; modulators can be added at appropriate times to observe stimulatory or inhibitory effects on resorption. When RAW-OCs are purified by serum gradient fractionation and replated onto tissue culture dishes, their viability is usually extended for an additional 24 h. Alternatively, purified RAW-OCs can be replated onto bone or ivory (~800 cells per well of a 48-well dish containing one piece of ivory or bone) and cultured with 35 ng/ml RANKL and 10 ng/ml IL-1α, in the presence or absence of other modulators, for 5–6 days to ascertain effects primarily on preformed RAW-OCs (although some additional RAW-OC development also occurs during this time period since the 70% serum purified fraction still contains some mononuclear cells). Purified RAW-OCs typically do not exhibit pit formation within the first 24 h after replating onto bone or ivory. 15. Although many of the phenotypic and functional characteristics of RAW-OCs match those of RANKL-differentiated primary murine bone marrow-derived OCs or isolated in vivo formed murine OCs, this cannot automatically be assumed to be true for any particular property being evaluated. The most obvious difference is the requirement for M-CSF in RANKLstimulated OC formation from bone marrow cells (which have relatively low RANK prior to M-CSF exposure) but not for transformed RAW cells (which already make M-CSF and express high RANK levels). In addition, apoptosis/survival pathways (including ERK) may differ between primary bone marrow cells and transformed RAW cells, and various other differences have been noted. Therefore, it is important to consider that the specific attribute under study in the RAW-OC cell system may not necessarily reflect that of normal murine OC formation or function. However, because RAW cells are

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easier to obtain and culture than primary bone marrow cells, represent a pure population of pre-OCs (deficient in osteoblasts, stromal cells, lymphocytes, etc.), are more readily transfected, and provide abundant material for study, they provide a highly valuable resource for rapidly and efficiently screening and determining mechanisms underlying OC-related processes. Therefore, we recommend that RAW cell studies are subsequently followed by at least a limited number of experiments using primary murine OCs (directly isolated and/or RANKLgenerated in vitro) to confirm that these processes are likewise observed in normal murine OCs and are not unique to transformed RAW cells or RAW-OCs.

Acknowledgments We are greatly indebted to the Drs. Xuefeng Yu and Hong Zheng for their advice and many valuable contributions to an earlier version of this chapter. This work was supported by NIH Grants AR32927, AG15435, and AR32087 to P.O. References 1. Roodman, G. (1996) Advances in bone biology – the osteoclast. Endocrine Rev. 17, 308–332. 2. Mancini, L., Moradi-Bidhendi, N., Brandi, M., Perretti, M., and McIntyre, I. (2000) Modulation of the effects of osteoprotegerin (OPG) ligand in a human leukemic cell line by OPG and calcitonin. Biochem. Biophys. Res. Commun. 279, 391–397. 3. Nagai, M., Kyakumoto, S., and Sato, N. (2000) Cancer cells responsible for humoral hypercalcemia express mRNA encoding a secreted form of ODF/TRANCE that induces osteoclast formation. Biochem. Biophys. Res. Commun. 269, 532–536. 4. Hentunen, T., Reddy, S., Boyce B. et al.,(1998) Immortalization of osteoclast precursors by targeting Bcl-XL and Simian virus 40 large T antigen to the osteoclast lineage in transgenic mice. J. Clin. Invest. 102, 88–97. 5. Chen, W., and Li Y. (1998) Generation of mouse osteoclastogenic cell lines immortalized with SV40 large T antigen. J. Bone. Miner. Res. 13, 1112–1123. 6. Takeshita, S., Kaji, K., and Kudo, A. (2000) Identification and characterization of the new osteoclast progenitor with macrophage pheno-

types being able to differentiate into mature osteoclasts. J. Bone Miner. Res. 15, 1477–1488. 7. Takahashi, N., Udagawa, N., and Suda, T. (1999) A new member of tumor necrosis factor ligand family, ODF/OPGL/TRANCE/ RANKL, regulates osteoclast differentiation and function. Biochem. Biophys. Res. Commun. 256, 449–455. 8. Chambers, T. (2000) Regulation of the differentiation and function of osteoclasts. J. Pathol. 192, 4–13. 9. Schoppet, M., Preissner, K., and Hofbauer, L. (2002) RANK ligand and osteoprotegerin. Paracrine regulators of bone metabolism and vascular function. Arterioscler. Thromb. Vasc. Biol. 22, 549–553. 10. Hsu, H., Lacey, D., Dunstan, C., et al (1999) Tumor necrosis factor receptor family member RANK mediates osteoclast differentiation and activation induced by osteoprotegerin ligand. Proc. Natl. Acad. Sci. USA 96, 3540–3545. 11. Yamamoto, A., Miyazaki, T., Kadono, Y., et al (2002) Possible involvement of IkappaB kinase 2 and MKK7 in osteoclastogenesis induced by receptor activator of nuclear factor kappaB ligand. J. Bone Miner. Res. 17, 612–621.

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12. Mizukami, J., Takaesu, G., Akatsuka, H., et al (2002) Receptor activator of NF-kappaB ligand (RANKL) activates TAK1 mitogen-activated protein kinase kinase through a signaling complex containing RANK, TAB2, and TRAF6. Mol. Cell Biol. 22, 992–1000. 13. Raschke, W., Baird, S., Ralph, P., and Nakoinz, I. (1978) Functional macrophage cell lines transformed by Abelson leukemia virus. Cell 15, 261–267. 14. Shadduck, R., Waheed, A., Mangan, K., and Rosenfeld, C. (1993) Preparation of a monoclonal antibody directed against the receptor for murine colony-stimulating factor-1. Exp. Hematol. 21, 515–520. 15. Sells-Galvin, R., Cullison, J., Avioli, L., and Osdoby, P. (1994) Influence of osteoclasts and osteoclast-like cells on osteoblast alkaline phosphatase activity and collagen synthesis. J. Bone Miner. Res. 9, 1167–1178. 16. Cappellen, D., Luong-Nguyen, N., Bongiovanni, S., Grenet, O., Wanke, C., and Susa, M. (2002) Transcriptional program of mouse osteoclast differentiation governed by the macrophage colony-stimulating factor and the ligand for the receptor activator of NF-kappa B. J. Biol. Chem. 277, 21971–21982.

17. Cassady, A., Luchin, A., Ostrowski, M., et al (2003) Regulation of the murine TRAP gene promoter. J. Bone Miner. Res. 18, 1901–1904. 18. Watanabe T, Kukita T, Kukita A, et al. (2004) Direct stimulation of osteoclastogenesis by MIP-1a: evidence obtained from studies using RAW264 cell clone highly responsive to RANKL. J. Endocr. 180, 193–201. 19. Vincent, C., Kogawa, M., Findlay, D., et al (2009) The generation of osteoclasts from RAW 264.7 precursors in defined, serum-free conditions. J. Bone Miner. Metab. 27, 114–119. 20. Koseki, T., Gao, Y., Okahashi, N., et al (2002) Role of TGF-beta family in osteoclastogenesis induced by RANKL. Cell Signal 14, 31–36. 21. Shin, J., Kim, I., Lee, J., Koh, G., Lee, Z., and Kim, H. (2002) A novel zinc finger protein that inhibits osteoclastogenesis and the function of tumor necrosis factor receptor-associated factor 6. J. Biol. Chem. 277, 8346–8353. 22. Trebec-Reynolds, D., Voronov, I., Heersche, J., et al (2010) IL-1alpha and IL-1beta have different effects on formation and activity of large osteoclasts. J. Cell Biochem. 109, 975–982.

Part III Biochemical and Molecular Analysis of Bone Cells

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Chapter 14 Transfection of Osteoclasts and Osteoclast Precursors Julie C. Crockett, David J. Mellis, and Adam Taylor Abstract Osteoclasts and their precursors have traditionally been considered difficult cells to transfect using standard approaches. Here, we describe several methods for transfection of mature osteoclasts and their precursors using the Amaxa™ Nucleofector system, lentiviruses, and adenoviruses. Key words: Osteoclast, Transfection, Adenovirus, Lentivirus

1. Introduction It is often necessary to evaluate the role that specific proteins play in the formation, function, or survival of osteoclasts in vitro. This can be achieved by expressing proteins of interest in the target cells using plasmid or viral vectors or by reducing expression of an endogenously produced protein by introducing small interfering RNA molecules (siRNA) into the target cell. Retroviral vectors have been extensively used by some research groups to transfect osteoclast precursors (1–3) but mature osteoclasts cannot be transfected with this technique since mitosis is required for integration of the retrovirus into the genome of the target cell. In this chapter, we describe methods for transfection of osteoclast precursors using lentiviral vectors (4–6) and the Amaxa™ Nucleofector (7) and also describe methods for transduction of mature osteoclasts using adenoviral vectors (8), Fig. 1 illustrates the stages of osteoclast differentiation at which each technique is best employed. 1.1. Amaxa ™ Nucleofector

This instrument uses electroporation in combination with specialised buffers supplied by the manufacturer to transfect target cells. We have used the Amaxa™ system to transfect osteoclast precursors with GFP and have shown that these cells can be differentiated into multinucleated osteoclasts that expressed the transgene for

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_14, © Springer Science+Business Media, LLC 2012

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MCSF & RANKL

MCSF

Haematopoietic stem cell

Macrophage /monocyte

MCSF & RANKL

MCSF & RANKL

Committed osteoclast progenitor

• Electroporation • Lentiviral transduction • Lentiviral transduction

• Adenoviral transduction • Lentiviral transduction

• Adenoviral transduction • Lentiviral transduction

Fig. 1. A schematic representation of the stages of osteoclast differentiation, highlighting when Amaxa™ nucleofection, lentiviral transduction, or adenoviral transduction is an effective method.

Fig. 2. Gene overexpression in human osteoclasts following transfection of precursor cells using the Amaxa™ Nucleofector. Human osteoclast precursors were transfected with EGFP-Rab6 after 2 days’ culture in M-CSF and RANKL and osteoclast formation was induced by culturing the cells with M-CSF and RANKL for a further 3 days. The cells were fixed in 4% (v/v ) paraformaldehyde, stained with wheatgerm agglutinin (WGA) and analysed by confocal microscopy. E-GFP-Rab6 (righthand panel ) co-localised with WGA (lefthand panel), suggesting that EGFP-Rab6 is localised to the Golgi apparatus, in agreement with the localisation of endogenous Rab6 in osteoclasts (7) (bar = 50 μm).

more than 7 days, whilst retaining the ability to polarise and resorb dentine. We have also used the Amaxa™ Nucleofector to generate human osteoclast-like cells over-expressing EGFP-Rab6, although this approach appears to be construct specific since we have been unable to generate osteoclasts that express Rab18, RANK, or PLEKhm1 using the same technique (7). We have, however, successfully used the Amaxa™ technique to introduce siRNA into osteoclasts and knockdown various Rab proteins including Rab 18 (Figs. 2 and 3).

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mRNA level by qPCR

a 100

0

-ve Ctl

siRNA1

siRNA3 Rab18

siRNA4

GAPDH

Protein level by western blot

b

GAPDH

Rab18 -ve Ctl

siRNA1

siRNA3

siRNA4

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Rab18 Fig. 3. siRNA knock-down in human osteoclast precursors using the Amaxa™ Nucleofector. Human osteoclast precursors were transfected with 1 μg of three different siRNAs against Rab18 or siRNA against GAPDH as a positive control. The cells were incubated for 72 h in MCSF and RANKL. (a) mRNA levels were assessed by quantitative RTPCR and (b) protein abundance was determined by western blot analysis using primers and antibodies against Rab18 and GAPDH. Both qPCR and western blot confirm the knockdown of GAPDH expression and the effectiveness of siRNA #1 against Rab18.

1.2. Lentiviral Vectors

Lentiviruses are a type of retrovirus which can incorporate into the genome of the host cell but do not require mitosis and can therefore be used to transduce both dividing and non-dividing cells. Recombinant lentiviral vectors have been engineered in which the genes necessary for incorporation of the virus into the host DNA have been retained but in which the genes required for viral replication have been deleted (9). These viruses can be used to insert foreign DNA sequences into the genome of the target cell with minimal viral sequence. Although these recombinant lentiviruses are replication incompetent, it is possible to transfect these vectors into a “packaging” cell line which expresses other components that are required for DNA replication and generation of intact lentiviruses containing the foreign DNA of interest, which are released

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into the culture supernatant. The viral particles can then be harvested from the supernatant and used to transduce host cells. A number of commercially available kits are available to generate lentiviral vectors. It is important to select a three or four plasmid system whereby the structural and coat proteins required for the production of lentiviral particles are encoded on separate plasmids to reduce the amount of lentiviral sequence within the expression vector and prevent homologous recombination events between the vector and wild-type HIV. All commercial systems involve a series of cloning steps to generate the lentiviral expression vector and these steps will depend on the availability of compatible restriction sites within the expression vectors and flanking your gene of interest. The Invitrogen ViraPower™ kit utilises Gateway™ cloning to transfer genes of interest from Gateway™ cloning vectors into the lentiviral vectors. We have found the Cell Biolabs promoterless lentiviral vector useful for incorporating entire expression cassettes into lentiviral vector (e.g. expression cassettes containing a cDNA driven by a cell-specific promoter or a bidirectional expression cassette). Whichever lentiviral system is used, the process of producing, purifying, and titreing the lentiviral particles is similar. 1.3. Adenoviral Vectors

Adenoviral vectors are useful for introducing foreign DNA into mature osteoclasts since they have the capacity to transduce terminally differentiated cells and express an RGD sequence on the penton base which allows them to attach to the αvβ3 integrin which is highly expressed by osteoclasts (10). Adenoviruses reside within the cytoplasm and are not incorporated into the host genome which means that they are of less value for transducing early osteoclast precursors which need to undergo cell division and fusion to form mature osteoclasts. Recombinant adenoviral vectors containing the construct of interest can be generated using various techniques including homologous recombination and in vitro ligation (11–13). The recombinant adenoviruses used in these experiments are replication deficient since the gene of interest replaces elements E1 and E3 which are essential for viral replication. However, the HEK293 cell line endogenously expresses the genes required for adenoviral replication and can therefore be used to produce recombinant infectious adenoviral particles when transfected with a recombinant adenoviral vector. We have employed the AdenoX™ expression system 1 (Takara Clontech) to generate recombinant adenoviral vectors. This involves ligating the construct of interest into the pshuttle2 vector and then ligating the entire expression cassette into pre-linearised adenoviral DNA. Adenoviral vectors can also be generated by homologous recombination. Whichever method is employed to generate the recombinant adenoviruses, assessment of the viral titre is required before experiments can commence so that the target cells can be transduced

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Fig. 4. Adenoviral transduction of human osteoclast-like cells. Human osteoclast-like cells were generated in the presence of rh-MCSF and rmRANKL for 6 days and transduced with (a) no virus, (b) 200 pfu/cell wildtype RANK. The cells were incubated for 48 h, fixed in 4% (v/v ) paraformaldehyde and (i) immunostained with an anti-RANK antibody and (ii) counterstained with TO-PRO 3 (bar = 100 μm).

with a known multiplicity of infection (MOI). This can either be assessed by a bioassay in which the researcher looks for evidence of a cytopathic effect in cells exposed to the virus [described as plaqueforming units (PFU)/ml] or by commercially available kits in which viral proteins are detected by immunostaining of infected cells. We have successfully used recombinant adenoviruses to study the effects of mutations in TNFRSF11A on the subcellular localisation of the RANK protein within mature osteoclast-like cells in vitro (Fig. 4).

2. Materials 2.1. Primary Cells

1. Human peripheral blood mononuclear cells. 2. Mouse bone marrow macrophages.

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2.2. Cell Lines for Virus Production

1. HEK293 cells (ECACC).

2.3. Reagents

1. Recombinant mouse and human M-CSF (R&D Systems).

2. 293 LTV cells (Cell Biolabs) or 293 FT cells (Invitrogen).

2. Recombinant mouse RANKL (R&D Systems) (see Note 1). 3. Mouse Macrophage Nucleofector™ Solution for electroporation (Amaxa™). 4. AdenoX Expression System 1™ (Takara Clontech). 5. Lentiviral Expression System e.g. ViraPower™ (Invitrogen) or ViraSafe™ (Cell Biolabs). 6. Polybrene for lentiviral transduction. 7. Lentiviral purification kit, ViraBind™ (Cell Biolabs). 8. Lentiviral titration kit, QuickTitre™ Lentivirus Titre kit (Cell Biolabs). 9. Pac 1 restriction enzyme (New England Biolabs). 10. Tris–EDTA (TE) buffer (10 mM Tris–HCl (pH 8.0) and 1 mM EDTA). 11. 3 M sodium acetate, pH 5.2 (NaOAc). 12. Phenol–Chloroform–isoamyl alcohol (25:24:1; Sigma). 13. Glycogen (20 mg/ml; Roche). 14. 100% Analytical grade ethanol. 15. 4% (v/v) Paraformaldehyde in PBS. 16. Cell lysis buffer [1% (v/v) triton-X-100, 0.5% (w/v) sodium deoxycholate, 0.1% (w/v) SDS in dH2O] supplemented with 1% protease inhibitor cocktail (Sigma), 1% phosphatase inhibitor cocktail (Sigma), 1 mM EDTA, 1 mM sodium orthovanadate, 10 mM sodium fluoride. 2.4. Culture Media and Buffers

1. Complete medium #1 [α-MEM containing 10% (v/v) FCS, 100 U/ml penicillin, 0.1 mg/ml streptomycin, 2 mM L-glutamine]. 2. Complete medium #2 [α-MEM containing 20% (v/v) FCS, 100 U/ml penicillin, 0.1 mg/ml streptomycin, 2 mM L-glutamine]. 3. Complete medium #3 [D-MEM containing 10% (v/v) FCS, 2 mM L-glutamine, 0.1 mM MEM non-essential amino acids]. 4. SF medium #4 (D-MEM containing 2 mM 0.1 mM MEM non-essential amino acids).

L-glutamine,

5. Serum-free α-MEM for viral transductions. 6. Phosphate buffered saline (PBS). 7. Trypsin-EDTA [0.05% (w/v) trypsin in 0.5 mM EDTA].

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1. Amaxa™ Nucleofector II™ machine. 2. Amaxa™ cuvettes and Pasteur pipettes. 3. Centrifuges. 4. Selection of pipettes and tips. 5. Tissue culture flasks and plates (Falcon). 6. Sterile Eppendorf tubes, bijoux, 15 ml conical tubes and 50 ml conical tubes. 7. Vacuum flask for liquid nitrogen. 8. Water bath set to 37°C.

3. Methods 3.1. Transfecting Human Osteoclast Precursors Using the Amaxa™ Nucleofector

This technique can be used to transfect human osteoclast precursors and requires at least 1 × 106 cells per transfection. If cell numbers are limited, the lentiviral system can be used as an alternative (Subheading 3.3). 1. Isolate peripheral blood mononuclear cells from 30 ml of venous blood sample by ficoll-hypaque gradient density centrifugation (see Note 2). 2. Wash the cells in PBS, divide into two Falcon 75 cm2 tissue culture flasks and culture in complete medium #1 supplemented with human M-CSF (20 ng/ml) until they reach 80–90% confluence (6–7 days). 3. Add mouse RANKL (10 ng/ml) to the cultures and maintain in culture for a further 48 h. 4. Aspirate the medium and wash the cell layer in PBS which has been pre-warmed to 37°C. 5. Add about 5 ml trypsin/EDTA to the flask and incubate for 30 min at 37°C. 6. Add 10 ml complete medium #1 and gently scrape the cells and transfer into a sterile Universal Container. 7. Pellet the cells by centrifugation at 300 × g for 5 min. 8. Resuspend the cells in 5 ml complete medium #1 and incubate on ice. 9. Count the cells using a haemocytometer and prepare aliquots of 1 × 106 cells for each transfection you wish to perform. Include two extra aliquots of cells as controls (see Note 3). 10. Pellet each aliquot by centrifugation at 300 × g for 3 min and carefully remove the supernatant.

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11. Suspend the cell pellet in 100 μl Mouse Macrophage Nucleofector™ Solution (see Note 4) and transfer to an Eppendorf tube containing 2 μg of endotoxin-free plasmid DNA or 1 μg of siRNA. 12. Transfer the cell suspension–DNA mixture to Amaxa™ electrode cuvettes and electroporate in an Amaxa™ Nucleofector II™ using program Y-010. 13. Immediately after the program has finished, transfer the cells to a bijou tube using the Amaxa™ Pasteur pipette and add 2.5 ml complete medium #1 containing 20 ng/ml M-CSF and 10 ng/ ml RANKL, which has been pre-heated to room temperature. 14. Cool the cell suspension on ice for 5–10 min. 15. Seed the transfected cells into 96-well plates at 4 × 104 cells/ well and/or onto 9 mm glass coverslips in 48-well plates at 1 × 105 cells/well and/or in 6-well plates at 4 × 105 cells/well (see Notes 5 and 6). 16. Seed the sham-transfected and control cells at half of this density. 17. Replace the medium after 48 h with complete medium #1 containing 20 ng/ml human M-CSF and 10 ng/ml mouse RANKL. 18. Replace with complete medium #1 containing 20 ng/ml human M-CSF and 10 ng/ml mouse RANKL every 3 days until multinucleated osteoclast-like cells have formed (a further 4–5 days). 3.2. Generating Lentivirus

Here, the lentiviral expression vector containing your gene of interest (that will have been generated through a series of cloning steps as broadly explained in Subheading 1.2) will be transfected, together with a cocktail of plasmids that encode the replication and structural proteins for lentivirus generation, into a 293-based packaging cell line. The exact conditions for transfection and production of lentiviral particles will depend on the system; here transfection using the Invitrogen Virapower™ lentiviral packaging mix is described. 1. Seed 293 FT cells (see Note 7) in a 10-cm culture dish and culture until the cells are 70–80% confluent. 2. To 1.5 ml SF medium #4 add 36 μl Lipofectamine 2000 in a bijou tube. Incubate for 5 min at room temperature. 3. In a separate bijou tube, add 1.5 ml SF medium #4 to 9 μg Virapower™ lentiviral packaging and 3 μg lentiviral expression vector. Mix by gently flicking the tube. 4. Combine the DNA and Lipofectamine solutions, mix gently and incubate for 20 min at room temperature. 5. Add DNA/Lipofectamine to 293 FT cells in a dropwise manner and rock plate gently to distribute evenly across plate.

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6. Incubate overnight and then replace medium with complete medium #3. 7. Maintain the cells in culture at 37°C for a further 72 h. 8. Aspirate the culture supernatant containing the infectious lentiviral particles using a syringe and pass through a 0.45 μm filter. 9. Purify the viral particles using the ViraBind™ kit by adding the filtrate from step 8 onto the LTV filter and allow it to flow through the filter under gravity (see Note 8). 10. Wash the filter twice with LTV wash buffer. 11. Elute the virus into a collection tube with 2 ml of LTV elution buffer. 12. Measure the amount of lentivirus present using the Quicktitre™ lentivirus kit and follow manufacturer’s instructions (see Note 9). 13. Add glycerol to a final concentration of 10%. Aliquot the viral preparation and freeze at −80°C. 3.3. Transduction of Human Osteoclast Precursors with Lentiviral Vectors

The lentivirus based technique is useful for experiments where cell numbers are limited since the Amaxa™ Nucleofector protocol described in Subheading 3.1 requires a minimum of 1 × 106 cells per transfection. 1. Generate M-CSF-dependent human macrophages from peripheral blood as described in Subheading 3.1 steps 1–2. 2. Seed the macrophages into a 96-well plate at a density of 3 × 104 cells/well in complete medium #1 supplemented with 20 ng/ml human M-CSF and 10 ng/ml mouse RANKL and culture overnight. 3. Aspirate the medium and wash the cells in PBS. 4. Add between 1.5 × 106 and 3 × 106 TU of the lentiviral vector (see Note 10) to each well in 50 μl serum free α-MEM containing human M-CSF (20 ng/ml) and mouse RANKL (10 ng/ml) and 8 μg/ml Polybrene. 5. Culture the cells for 3 h, remove the medium and replace with complete medium #1 containing human M-CSF (20 ng/ml) and mouse RANKL (10 ng/ml). 6. Culture the cells for up to 7 days, replacing the culture medium every 2 days. 7. When multinucleated osteoclasts have formed (typically 6–7 days) terminate the experiment by aspirating the medium and adding 50 μl of 4% paraformaldehyde to each well to fix the cells.

3.4. Generating Recombinant Adenoviruses

1. Linearise the recombinant adenoviral DNA containing the gene of interest by PacI digestion for 2 h at 37°C in Eppendorf tubes. Add 60 μl 1× TE buffer and 100 μl phenol–chloroform– isoamyl alcohol and vortex gently.

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2. Centrifuge at 13,000 × g for 5 min to separate the phases. Transfer the top, aqueous layer to a clean 1.5-ml Eppendorf tube. 3. Add 100 μl 95% (v/v) ethanol, 50 μl 3 M NaOAc (pH 5.2), and 1 μl glycogen (20 mg/ml). Vortex gently. 4. Remove supernatant and wash pellets in 70% (v/v) ethanol. Centrifuge at 13,000 × g for 2 min. Remove supernatant and air-dry pellets at room temperature. Re-dissolve DNA in 10.5 μl TE buffer. Check restriction digest has been successful by analysis of 0.5 μl of the DNA on a 0.8% agarose gel. Store remaining DNA at −20°C. 5. Seed 2 × 106 HEK293 cells into each well of a 60-mm tissue culture plate and culture overnight at 37°C in complete medium #1. 6. Change the medium on the HEK293 cells at least 90 min (up to 3 h) prior to transfection. Transfect the 10 μl of linearised DNA (from step 4) into the HEK293 cells using CalPhos™ transfection kit (BD Biosciences) according to the manufacturer’s instructions (see Note 11). Include a mock transfected control. 7. After 3 h replace the medium with complete medium #1. 8. Culture the cells for between 8 and 12 days, and check the cultures each day on an inverted, phase contrast microscope to determine the proportion of cells that have detached from the plate. Do not replace the medium during this time. If necessary, add 2–3 ml more medium to the plate if the colour of the medium becomes very yellow. 9. When the majority (60–80%) of the cells have detached aspirate the contents of the dish into a 15-ml conical tube and pellet the cells by centrifugation at 300 × g for 5 min. Dislodge the cells by gentle agitation; do not use trypsin. 10. In the 15 ml conical tube, resuspend the cell pellet in 500 μl PBS and lyse the cells using a freeze–thaw technique. Using forceps, immerse the bottom of the tube (and hence the cell suspension) for approximately 2 min in liquid nitrogen held in an open vacuum flask. 11. When cell suspension is frozen, thaw by transferring to a waterbath set at 37°C for periods of 10 s at a time, interspersed with bursts of vortexing for 30 s at a time until the lysate has just melted (see Note 12). 12. Repeat the freeze–thaw cycles described in steps 10 and 11 two more times. 13. Centrifuge the lysate at 1,500 × g for 10 min, aspirate the supernatant, and store this adenovirus stock at −80°C until required.

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14. For amplification of this stock, seed a 60-mm tissue culture dish with 2 × 106 HEK293 cells. 15. The following day, defrost the adenoviral stock and add the total volume (~500 μl) to the HEK293 cells in the 60-mm plate. 16. Incubate for 3–5 days. Check the cells on an inverted microscope to check for cells lifting off. When 60–80% cells have lifted off follow steps 9–12 (see Note 13). Freeze this primary amplification stock at −80°C in 50 μl aliquots. 3.5. Determining the Titre of Adenovirus

This subheading describes the end-point dilution method for estimating the number of viral particles present in the supernatant from step 16 of Subheading 3.4. By convention adenoviral titres are expressed in plaque forming colony units (pfu)/ml (see Note 14). 1. Seed HEK293 cells into 96-well plates at a density of 1 × 104/ well in 100 μl complete medium #1 and culture overnight. 2. Thaw the adenoviral stock (Subheading 3.4, step 16) on ice and make a 1:100 (10−2) dilution by adding 10 μl virus to 990 μl complete medium #1. 3. Prepare eight serial 1:10 dilutions from step 2 by transferring 100 μl of diluted virus into 900 μl of complete medium #1 to give a “standard curve” ranging from a dilution of 10−3–10−10 as compared with the original adenoviral stock. This gives sufficient volume of each dilution for eight replicates. 4. Add 100 μl of the diluted adenovirus stock to each well of the first eight columns of the 96-well plate set up in step 1 and add 100 μl of culture medium to the remaining rows. 5. Culture the cells for a further 10 days without changing the medium. 6. Score each well for the presence of cytopathic effect (CPE) and express the results as the fraction of CPE positive wells per column. 7. Calculate the viral titre (pfu/ml) using the following equation: viral titre (pfu/ml) = 10(x+0.8) where x is the sum of the fractions of all the virus infected columns (see Note 15).

3.6. Amplification of Adenoviral Stocks (See Note 16)

1. Seed HEK293 cells into 75-cm2 tissue culture flasks and culture overnight at 5 × 106 cells. 2. Add adenovirus stock to the flask with a multiplicity of infection (MOI) of 1–5 pfu/cell. 3. Maintain the cells in culture for up to 12 days and monitor daily by microscopy until the majority (60–80%) of the cells have detached from the flask. 4. Aspirate the contents of the flask into a 50-ml conical tube and pellet the cells by centrifugation at 300 × g for 5 min.

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5. In the 50-ml conical tube, resuspend the cell pellet in 1 ml PBS and lyse the cells using a freeze–thaw technique. Using forceps, immerse the bottom of the tube (and hence the cell suspension) for approximately 1 min in liquid nitrogen held in an open vacuum flask. 6. When cell suspension is frozen, thaw by transferring to a waterbath set at 37°C for periods of 10 s at a time, interspersed with bursts of vortexing for 30 s at a time until the lysate has just melted (see Note 12). 7. Repeat the freeze–thaw cycles described in steps 5 and 6 two more times. 8. Centrifuge the lysate at 1,500 × g for 10 min, aspirate the supernatant, and store this adenovirus stock in 50 μl aliquots at −80°C until required. 9. Determine the titre (following Subheading 3.5) of these adenoviral stocks prior to moving on to Subheading 3.7. 3.7. Adenovirus Transduction of Mature Human Osteoclasts

This technique works well for transfecting mature human osteoclastlike cells that have been generated from peripheral blood using M-CSF and RANKL. 1. Generate osteoclast-like cells from human M-CSF dependent macrophages in complete medium #1 supplemented with human M-CSF (20 ng/ml) and mouse RANKL (10 ng/ml) and seed into 96-well plates at 1.5 × 104 cells/well, 48-well plates containing 9 mm glass coverslips at 3 × 104 cells/well, or 6-well plates at 2 × 106 cells/well (see Note 6) until multinucleated osteoclast-like cells have formed (approximately day 6). 2. Remove the medium and replace with between 50 and 750 μl (see Note 17) of serum free α-MEM supplemented with human M-CSF (20 ng/ml) and mouse RANKL (10 ng/ml) containing the equivalent of 200 pfu adenovirus per cell, based on the initial seeding density, and incubate at 37°C for 3 h. 3. Add an equal volume of complete medium #2 to each well supplemented with human M-CSF (20 ng/ml) and mouse RANKL (10 ng/ml) and culture the cells for a further 48 h. 4. If you want to study the effect of the gene of interest on bone resorption, follow steps 6–10. If you want to study the effect on signalling pathways in osteoclasts follow steps 11–13. 5. Wash the cells from step 5 in PBS which has been pre-warmed to 37°C. 6. Incubate the cells in trypsin/EDTA solution for 30 min. 7. Gently scrape the cells into complete medium #1 and pellet the cells by centrifugation at 300 × g for 5 min.

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8. Resuspend the cells in complete medium #1 supplemented with human M-CSF (20 ng/ml) and mouse RANKL (10 ng/ml). 9. Seed the cells into a 96-well plate containing sterile dentine slices at a density of 2 × 104 cells/well and culture the cells for up to 72 h. 10. Terminate the experiment by aspirating the medium and fixing the cells in 4% paraformaldehyde in PBS. 11. To prepare protein lysates for western blot analysis (see Note 18), wash cells in 2 ml ice cold PBS and then add 100 μl cell lysis buffer to each well, keeping the plate on ice. 12. Scrape the cells into the lysis buffer using a rubber policeman and transfer the lysate to 1.5 ml Eppendorf tube. Incubate on ice for 20 min and then centrifuge at 13,000 × g for 15 min. 13. Transfer the supernatant to a clean Eppendorf tube and snap-freeze the lysate in liquid nitrogen before transferring to a −80°C freezer for storage. 3.8. Adenovirus Transduction of Mature Mouse Osteoclasts

This technique works well for transfection of mouse osteoclasts but since these cells have a very short life span, the transfection must be performed just before the osteoclast precursors fuse to form mature osteoclasts (see Note 19). 1. Prepare M-CSF dependent mouse macrophages according to standard techniques. 2. Suspend in complete medium #1 supplemented with mouse M-CSF (100 ng/ml) and mouse RANKL (100 ng/ml) and seed into 96-well plates at 1.5 × 104 cells/well, 48-well plates containing 9 mm glass coverslips at 3 × 104 cells/well, or 6-well plates at 2 × 106 cells/well (see Note 6). 3. Continue the cells in culture for a further 3–4 days. 4. Remove the culture medium from each well and replace with 50−750 μl (see Note 17) serum free α-MEM supplemented with mouse M-CSF (100 ng/ml) and mouse RANKL (100 ng/ml) and 200 pfu/cell of adenoviral vector stock. 5. After 3 h, remove the medium and replace with complete medium #1 supplemented with M-CSF (100 ng/ml) and mouse RANKL (100 ng/ml). 6. Culture the cells for a further 48 h and fix or lyse the cells, depending on the analysis that you wish to perform (see Note 18).

3.9. Assessment of Cell Viability

Cell viability can be assessed using the Alamar blue technique and we recommend that you perform this routinely the first few times the experiments are carried out as part of optimisation. If you are using the Amaxa™ system, cell viability can be assessed in parallel with the experiment by seeding transfected cells at 4 × 104 cells/well

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(or sham-transfected and control cells at 2 × 104 cells/well) in a separate 96-well plate. For lentiviral and adenoviral transductions, it is necessary to prepare additional wells at the start of the experiment that will be specifically used to assess viability. 1. Add 10 μl Alamar blue reagent to test wells in 96-well plates either 24 h and/or 48 h after the transfection procedure. 2. Culture the cells for 3–4 h. 3. Read the fluorescence on a plate reader with the emission spectrum set to 530 nm and the excitation spectrum set to 590 nm. 4. The effect of the transfection procedure on viability can be assessed by comparing values for the transfected cells with those in the control wells (untransfected and sham transfected). 3.10. Assessment of Transfection Efficiency

Transfection efficiency can be assessed in cells that have been transfected with a GFP containing construct or where the gene of interest has been epitope tagged (see Note 20). The procedure for assessing transfection efficiency is as follows: 1. At the end of the experiment, remove the medium and fix the cells in 4% paraformaldehyde for 15 min and wash with PBS. 2. For GFP-transfected cells, obtain digital fluorescent and phase contrast images of the wells and merge these to assess the numbers of cells which are positive for GFP. Since GFP is a fluorescent protein, this can be carried out on both unfixed (i.e. throughout the culture) or fixed cells. 3. Repeat the procedure for untransfected and sham transfected cells to assess background fluorescence. 4. Work out the proportion of positive versus negative cells in the transfected versus non transfected or sham transfected controls. 5. If the gene of interest has been epitope tagged, perform immunostaining using a suitable antibody and count the number of cells that stain positive. 6. Repeat the procedure for untransfected and sham transfected cells to determine background staining. 7. Work out the proportion of positive versus negative cells in the transfected versus non transfected or sham transfected controls.

4. Notes 1. We use mouse RANKL for both human and mouse osteoclast cultures. It would also be possible to use human RANKL for human cultures, but this cannot be used to generate mouse osteoclasts.

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2. 30 ml of venous blood will generate approximately 4 × 106 M-CSF dependent macrophages over the culture expansion period. 3. Two controls should be included; untransfected control cells and sham transfected cells – which should be processed as for the cells undergoing transfection with the exception that electroporation is carried out in the absence of plasmid DNA. 4. We have also tested human macrophage Nucleofector™ solution, but found that this was associated with higher levels of cytotoxicity than the mouse solution. 5. The transfected cells are seeded at a high density (double that is normally used for these cultures) to take account of the fact that the electroporation procedure is associated with approximately 50% toxicity. 6. Mouse and human osteoclasts are generated in 96-well plates to assess osteoclast formation, 48-well plates (containing 9 mm glass coverslips) for downstream confocal microscopic analysis of fixed cells, or 6-well plates for analysis of cellular proteins by western blot analysis. 7. The 293FT cell line, stably expresses the SV40 large T antigen and must normally be maintained in medium containing Geneticin™ as per suppliers instructions. For transfection of lentiviral expression construct with the packaging mix, all antibiotics including Geneticin should be omitted from the culture medium (complete medium #3). 8. It is possible to snap-freeze the filtrate in liquid nitrogen at this point, store at −80°C and proceed with the purification steps at a later date. 9. The number of viral particles within a given volume of supernatant (termed transforming units; TU) can be estimated by transducing HeLa cells with the supernatant and determining the number of antibiotic resistant clones that develop over 2–3 weeks. For a much faster and more objective assay, this commercially available ELISA kit that detects the HIVassociated p24 protein that is associated with the virus gives absolute values for the amount of p24 present in each sample which can then be worked back to calculate TU/ml. 10. The amount of lentivirus must be optimised for each experiment, but we find that a MOI of between 50 and 100 TU/cell gives good results. 11. We have tried numerous methods to transfect the recombinant adenoviral DNA into 293 cells (including Fugene-6 and Clonfectin) and the only method that resulted in production of adenoviral particles was the CalPhos transfection kit. 12. It is important that the suspension does not warm up, hence the need to assist the thawing process by 30 s bursts of vortexing

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between the periods of submersion in the waterbath (which should be for no more than 10 s at a time). 13. The adenovirus obtained as the result of this procedure is referred to as “Primary Amplification Stock”. 14. The titre of your adenoviral stocks can be determined by various methods which should be selected based on time and budget considerations. When adenovirus is applied to HEK293 cells, the cells undergo lysis and this can be visualised as a cytopathic effect (CPE) in the cell cultures as regions of dead or absent cells. Any evidence of CPE within a culture demonstrates that there was a least one infectious unit of virus present. This can be exploited in the end-point dilution assay that is described here. In addition a plaque formation assay can be performed in which cells are grown under a layer of medium enriched agarose and areas or plaques of dead cells identified and counted, or immunohistochemistry based kits are quick but expensive. We have occasionally used the Adeno-X™ Rapid-Titre kit (Clontech), which employs an anti-hexon primary antibody and horse-radish peroxidase secondary antibody to stain infected cells. 15. The equation for calculating viral titre is a modification of the Spearman-Karber method (as described in pAdenoX User Manual, Takara Clontech). The values for 10−1 and 10−2 dilutions must be included in the calculation (which will be 1 and 1 if 10−3 dilution is 1) for the equation to be valid. This method relies on the data representing a total response curve, i.e. 100% effect at the lowest dilution and no CPE at the highest dilution. Therefore, an accurate estimation of titre will only be achieved if at the lowest virus dilution (10−3) all wells show CPE (ratio = 1) and at the highest dilution (10−10), zero wells show CPE (ratio = 0). If there is not CPE in all wells for 10−3 then the viral titre is not sufficiently high for transduction experiments. If, in this case, the end-point dilution assay was on a Primary Amplification Stock then start again with adenoviral transfection into HEK293 cells (Subheading 3.4). If the end-point dilution assay was on a later stock of virus, repeat Subheading 3.6 and use 5 pfu/cell to transduce your HEK293 cells and freeze–thaw the cells in a smaller volume of PBS (e.g. 500 μl). In all cases, ensure that CPE has occurred in 60–80% of cells before harvesting the adenovirus from the cells. If, on the otherhand, CPE is observed in the cells containing the 10−10 dilution then the adenoviral stock should be diluted (e.g. 1:10) and this diluted stock taken as 100 (i.e. undiluted) for the purposes of the end-point dilution assay – it is important to correct your calculated titre for this dilution at the end of the assay. The expected viral titres for the primary amplification stocks should be around 109 pfu/ml.

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16. The risk of generating replication competent adenoviruses increases with the number of amplification steps performed. Therefore, it is important to return to early amplification stocks when you are generating large volumes of high titre adenovirus. 17. Use the minimum volume of virus-containing medium possible to cover the bottom of your culture plate to maximise efficiency of transfection. We recommend the following: 50 μl/well for 96-well plates, 150 μl/well for 48-well plates, 750 μl/ well for 6-well plates. 18. For studying the effects on cell signalling pathways then the cells should be stimulated with an appropriate cytokine, lysed and cytoplasmic extracts prepared. In the case of experiments investigating the effects of siRNA knockdown RNA can be extracted in preparation for cDNA synthesis and quantitative RT-PCR. 19. Since mouse osteoclast-like cells have a very short life span, it is necessary to transduce the cells about 24–48 h before they fuse to form multinucleated osteoclasts. This is usually between 3 and 4 days after the cultures are initiated but the most appropriate stage of the culture should be determined prior to commencing these experiments. 20. It is possible to tag the cDNA by cloning GFP downstream to generate a fusion protein but if you are conducting functional experiments, it is important to confirm that the GFP does not interfere with the properties of your protein. Alternatively, the cDNA can be tagged with an epitope tag (e.g. FLAG tag) that can be detected with commercially available antibodies. In all cases, it is important to consider the position of the tag (N- or C-terminus) and whether it might interfere with protein processing or function. References 1. Xu, D., Wang, S., Liu W, et al. (2006) A novel receptor activator of NF-KappaB (RANK) cytoplasmic motif plays an essential role in osteoclastogenesis by committing macrophages to the osteoclast lineage. J. Biol. Chem. 281, 4678–4690. 2. Kapur, R. P., Yao Z., Iida, M. H., et al. (2004) Malignant autosomal recessive osteopetrosis caused by spontaneous mutation of murine Rank. J. Bone Miner. Res. 19, 1689–1697. 3. Maruyama, T., Fukushima, H., Nakao, K., et al. (2010) Processing of the NF-KappaB2 precursor, p100, to p52 is critical for RANKL-induced osteoclast differentiation. J. Bone Miner. Res. 25, 1058–1067.

4. Teschemacher, A. G., Wang, S., Lonergan, T., et al. (2005) Targeting specific neuronal populations using adeno- and lentiviral vectors: applications for imaging and studies of cell function. Exp. Physiol. 90, 61–69. 5. Chu, K., Cornetta, K. G., and Econs, M. J. (2008) Efficient and stable gene expression into human osteoclasts using an HIV-1-based lentiviral vector. DNA Cell Biol. 27, 315–320. 6. Zhao, H., Ross, F. P., and Teitelbaum S. L. (2005) Unoccupied alpha(v)beta3 integrin regulates osteoclast apoptosis by transmitting a positive death signal. Mol. Endocrinol. 19, 771–780.

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7. Taylor, A., Rogers, M. J., Tosh, D., et al. (2007) A novel method for efficient generation of transfected human osteoclasts. Calcif. Tissue Int. 80, 132–136. 8. Crockett, J. C., Mellis, D. J., Duthie, A., et al. (2011) Signal peptide mutations in RANK prevent downstream activation of NF-κB. J. Bone Miner. Res. 26, 1926–1938. 9. Naldini, L. (1998) Lentiviruses as gene transfer agents for delivery to non-dividing cells. Curr. Opin. Biotechnol. 9, 457–463. 10. Tanaka, S., Takahashi, T., Takayanagi, H., et al. (1998) Modulation of osteoclast function by adenovirus vector-induced epidermal growth factor receptor. J. Bone Miner. Res. 13, 1714–1720.

11. Miyake, S., Makimura, M., Kanegae, Y., et al. (1996) Efficient generation of recombinant adenoviruses using adenovirus DNA-terminal protein complex and a cosmid bearing the fulllength virus genome. Proc. Natl. Acad. Sci. USA 93, 1320–1324. 12. Mizuguchi, H., and Kay, M. A. (1998) Efficient construction of a recombinant adenovirus vector by an improved in vitro ligation method. Hum. Gene Ther. 9, 2577–2583. 13. Mizuguchi, H., and Kay, M. A. (1999) A simple method for constructing E1- and E1/ E4-deleted recombinant adenoviral vectors. Hum. Gene Ther. 10, 2013–2017.

Chapter 15 Analysis of Signalling Pathways by Western Blotting and Immunoprecipitation Aymen I. Idris Abstract This chapter describes the analysis of signalling pathways in bone cells by the use of western blotting and immunoprecipitation, including a step by step guide to cell culture techniques, protein isolation, purification, measurement, electrophoretic transfer, and detection. Key words: Signalling, Western blotting, Immunoprecipitation, Protein, Phosphorylation, Prenylation

1. Introduction Signal transduction is the process by which extracellular information is received, relayed, and translated into a cellular response (1). Systemic hormones and local cytokines regulate bone cell differentiation and activity by activating signalling transmission systems that originate at the cell surface or cytoplasm and terminate in the nucleus (Fig. 1). At the molecular level, bone-selective ligands such as receptor activator of NFκB (RANK) ligand and parathyroid hormone (PTH) bind to their respective membrane receptors and activate a complex network of receptor associated factors (RAF) to relay the initial signal across cell membrane (2, 3). The strength and duration of the signal is regulated by the action of a number of highly specialised intracellular enzymes which form signalling cascades that operate as “biological switches” (Fig. 1). Specialised enzymes such as protein kinases phosphorylate specific amino acids in the target protein, most commonly at specific tyrosine or serine residues (1, 4). This causes a change in

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_15, © Springer Science+Business Media, LLC 2012

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Fig. 1. Schematic diagrams showing a hypothetical model of post-translational modification of signalling proteins and methods that routinely used to study these changes. Signalling proteins are normally present in the cytoplasm. Upon cell stimulation, receptor associated factors (RAF) are recruited by membrane receptors. This results in structural modification of cytoplasmic signalling proteins (CSP) that in turn alter protein mobility, level, and/or activity. Some cytoplasmic signalling proteins function as transcription factors (TF) by translocating to the nucleus where they bind to DNA and induce the transcription of various genes needed for cell survival, differentiation, and activity.

configuration and a change in function. For example, studies showed that pharmacological inhibition of RANK ligand-induced phosphorylation of cytoplasmic proteins such as IκB kinase (IKK) in osteoclast is sufficient to suppress osteoclast formation in vitro and in vivo, and as a result prevent osteoclastic bone destruction in animal models of osteoporosis and breast cancer (5–7). Conversely, signalling is often de-activated by protein phosphatases which dephosphorylate the target protein rendering it inactive (4). Reflecting this fact, phosphorylation is the most common post-translational modification to occur in mammalian cells (1, 4, 8). Over the last decade, there has been a tremendous increase in understanding of the signal transduction pathways that systemic hormones, local regulatory factors, and drug treatments

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use to regulate bone cell differentiation, survival, and function. Protein prenylation is another extensively studied mode of protein post-translation modification in bone cells (9, 10). Treatment with nitrogen-containing bisphosphonates inhibits prenylation of small GTPases and leads to the accumulation of unprenylated GTPases in osteoclasts (10). Following cytoplasmic events, the propagation of the signal to the nucleus involves nuclear translocation and DNA binding of transcription factors (Fig. 1). Changes in the levels, phosphorylation status and binding partners of intracellular signalling proteins are most commonly assessed by Western blotting and immunoprecipitation and I shall describe these techniques in this chapter (Fig. 2). Electrophoretic mobility shift assay (EMSA) or mobility shift electrophoresis (Fig. 2) frequently used to study the activation and DNA binding of transcription factors in bone cells is described in Chapter 16, this volume. For the description of “kit-based” methods such as enzyme-linked immunosorbent assay (ELISA) and immunosorbent assays refer to Fig. 2 and follow instruction by suppliers of kits.

resolved protein

Immunoprecipitation

Cytoplasmic protein Transcription factor

pull down protein and run on SDS page gel add primary and purify protein secondary antibody

visualise protein

DNA Primary Antibody

resolved protein

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lyse cell

run on SDS page gel

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HRP conjugated secondary antibody Chemiluminescent substrate EMSA labelled probe*

visualise protein

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Enzyme-linked immunosorbent assay treat cells

permeabilise cells add primary antibody

add secondary add fluorescent substrate and visualise protein antibody

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probes

lyse cell and add EMSA labelled run on SDS page gel nuclear and competitive probes

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Enzyme-linked immunosorbent assay lyse cell

add cell lysate to add primary antibody add secondary antibody followed DNA-coated plate by chemiluminescent substrate

Fig. 2. Overview of the principles of protocols routinely used to study signalling transduction mechanisms in bone cells.

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2. Materials Trade names and commercially products are occasionally stated for identification purposes only and do not imply endorsement. 2.1. Immunoprecipitation

1. Standard lysis buffer: 0.1% (w/v) sodium dodecyl sulphate (SDS), 0.5% (w/v) sodium deoxycholate, 2% (v/v) protease inhibitor cocktail (see Note 1) in sterile PBS. 2. Modified lysis buffer: Standard lysis buffer containing 1 mM EDTA, 1 mM sodium orthovanadate, and 10 mM sodium fluoride in sterile PBS.

2.2. Western Blotting

1. Resolving gel: 40% bisacrylamide:acrylamide (1:29), 25% Tris (1.5 M, pH 8.8), 0.1% SDS (10%), 0.1% ammonium persulfate (10%), and 0.04 μl of TEMED, 34.8% distilled water; added in this order. 2. Stacking gel: 12.5% bisacrylamide:acrylamide (1:29), 25% Tris (0.5 M, pH 6.8), 1% SDS (10%), 0.5% ammonium persulfate (10%), and 0.1% TEMED, 60.9% distilled water; added in this order. Pre-cast SDS PAGE gels provide an alternative to making the resolving and stacking gels and can be purchased from a number of manufacturers. 3. Sample loading buffer: 12.5% Tris–hydrochloric acid (Tris– HCl) (0.5 M; pH 6.8), 10% glycerol, 20% SDS (10%), 5% β-mercaptoethanol, 2.5% bromophenol blue (0.05%), 50% distilled water. 4. Electrophoresis running buffer: dissolve 3% Tris base, 1% SDS, 1.1% glycine dissolved in distilled water. 5. Transfer buffer: 48 mM Tris base, 39 mM glycine, 0.04% (w/v) SDS, 20% methanol in distilled water. 6. 100% Methanol. 7. Pads or standard chromatography papers. 8. Nitrocellulose or polyvinylidene fluoride (PVDF) membrane. 9. Tris–HCl: dissolve 10% (w/v) Tris base in 60 ml of dH2O, adjust pH by adding HCl (1 M), make up volume to 150 ml in distilled water, and store at 4°C. 10. Tris-buffered SDS with Tween (TBST): 50 mM Tris and 150 mM sodium chloride in distilled water. 11. Blocking solution: 5% w/v dried non-fat milk in TBST. 12. Stripping buffer: 1 mM dithiothreitol (DDT), 2% SDS, and 6.35 mM Tris–HCl (pH 6.7).

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3. Methods 3.1. Immunoprecipitation

3.1.1. Preparation of Cell Lysate

Immunoprecipitation (IP) is a technique carried out to probe for the presence of proteins of interest in a cell lysate and to determine if it forms part of a protein complex (Fig. 2). The technique involves incubating an antibody directed against the protein of interest with a cell lysate and precipitating the complex of antibody bound to the target protein by agarose or magnetic beads (see Note 2). The antibody/protein complex can then be separated by SDS-PAGE and subjected to Western blotting. This allows the investigator to determine if the protein of interest is present within the cell lysate and to determine (by probing with antibodies to other proteins) if the protein of interest is part of a protein complex. 1. Culture bone cells to confluence in standard tissue culture plates in the presence or absence of test stimuli for the desired period of time. 2. Aspirate and rinse monolayer three times with ice-cold PBS. 3. Incubate adherent cells in standard lysis buffer for 10 min. Use modified lysis buffer for studies involving extraction of phosphorylated proteins (see Notes 3 and 5). 4. Vortex cell suspension for 10 s, pass supernatant through a 21-gauge needle, and incubate on ice for 10 min. 5. Centrifuge cell lysate at 14,000 × g at 4°C for 10 min and transfer the supernatant to a clean Eppendorf (at this point it is possible to proceed with the IP or store the samples frozen at −80°C for up to a year). 6. Determine protein concentration using a standard commercially available protein assay such as bicinchoninic acid (BCA) Pierce® protein assay (Pierce, UK).

3.1.2. Immunoprecipitation

1. Incubate 490 μl of cell lysate containing 250–500 μg of total protein concentration with 5 μg of polyclonal or monoclonal antibody specific for protein of interest at 4°C for 24 h (see Note 4 and Fig. 2). 2. Add 10 μl of bead of choice to the immunoprecipitation cell lysate mix and then incubate with agitation at 4°C for 6–16 h (see Note 2). 3. Briefly centrifuge sample, carefully aspirate, discard the supernatant, and recover the pelleted bead/protein(s) complex. 4. Wash pellet three to five times in ice-cold standard lysis buffer by inverting Eppendorf several times. 5. Boil sample mixture at 90°C for 5 min to dissociate protein of interest from bead/protein(s) complex.

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6. Briefly centrifuge sample, carefully aspirate, and transfer the supernatant to a fresh Eppendorf. 7. Repeat steps 1–5 to further purify or dissociate desired protein from other proteins if part of a complex. 8. Mix cell lysate with the appropriate volume of sample loading buffer and boil at 90°C for 5 min to denature protein. 9. Resolve samples on polyacrylamide-SDS gel and blot to nitrocellulose or PVDF membrane (see below). 3.2. Western Blotting

Western blotting is used to determine if a protein of interest is present within a specific cell type, to assess protein abundance and to assess phosphorylation status or other post-translational modifications such as prenylation (11). The technique uses denaturing gel electrophoresis to separate proteins present in cell extracts according to their molecular weight (Fig. 2). Once separated, proteins are transferred to nitrocellulose or PVDF membrane and incubated with a primary antibody directed against the proteins of interest. If the protein of interest is present within the lysate the primary antibody will bind to the membrane-bound protein and this can be detected by incubating the membrane with a secondary antibody linked to an amplification system such as HRP that uses HRPconjugated secondary antibody that can be visualised using enhanced chemiluminescence imager or transferred on a film (Fig. 3).

Fig. 3. Schematic of gel assembly and transfer assembly for western blotting. (a) Gel assembly; (b) transfer assembly.

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1. Culture bone cells to confluence in standard tissue culture plates in the presence or absence of test stimuli for the desired period of time. 2. Aspirate and rinse monolayer three times with ice-cold PBS. 3. Incubate adherent cells in standard lysis buffer for 10 min. Use modified lysis buffer for studies involving extraction of phosphorylated proteins (see Note 5). 4. Vortex cell suspension for 10 s, pass supernatant through a 21-gauge needle, and incubate on ice for 10 min. 5. Centrifuge cell lysate at 14,000 × g at 4°C for 10 min and transfer the supernatant to a clean Eppendorf (at this point it is possible to proceed with the WB or store the samples frozen at −20°C for up to a year). 6. Determine the protein concentration using a standard commercially available protein assay such as BCA Pierce® protein assay (Pierce, UK).

3.2.2. Preparation of SDS Polyacrylamide Gel

1. To prepare SDS polyacrylamide gels in the laboratory, assemble a glass-plate gel assembly using in-house components or those supplied by commercially available kits (see Fig. 3a). 2. Once the glass-plate assembly is in place, carefully pour resolving gel allowing about 3 cm from the top. Add a layer of distilled water on top of the resolving gel to ensure an even surface. 3. Once the resolving gel is polymerised, aspirate water and replace with a layer of stacking gel. Insert a plastic comb between the two glass-plates to create wells (see Fig. 3a). 4. Alternatively pre-cast SDS polyacrylamide gels can be used. These can be obtained in a wide range of sizes and thicknesses from a number of suppliers (see Note 6).

3.2.3. Electrophoresis

The procedure separates the denatured proteins according to their molecular weight. 1. Mix cell lysate with the appropriate volume of sample loading buffer. 2. Boil sample mixture at 95°C for 5 min to denature protein. 3. Place pre-cast gel into a vertical electrophoresis tank filled with electrophoresis running buffer. 4. Load carefully into the designated well on the pre-cast gel (see Note 7). 5. Place gel-plates assembly into electrophoresis tank filled with electrophoresis running buffer and run at constant volume of 200 V for 30–40 min (see Note 8).

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3.2.4. Electrophoretic Transfer from Gel to Membrane

The procedure transfers proteins from the SDS polyacrylamide gel to a nitrocellulose or PVDF membrane, prior to incubation with the primary and secondary antibodies and detection step. 1. Remove gel from the glass-plate assembly and immerse gel into transfer buffer for 3–5 min. 2. Meanwhile, cut nitrocellulose or PVDF membrane to size, immerse in 100% methanol, and then allow it to equilibrate in transfer buffer for 5 min. 3. Prepare a blotting sandwich by arranging successive layers of three pads pre-soaked in transfer buffer, nitrocellulose or PVDF membrane, SDS polyacrylamide gel, three pre-soaked pads (see Fig. 3b). 4. Run transfer at a constant current for 1–3 h (see Note 8).

3.2.5. Protein Detection

1. Incubate nitrocellulose or PVDF membrane at room temperature for 1 h in blocking solution containing 5% dried non-fat milk in TBST (see Note 9). 2. Rinse nitrocellulose or PVDF membrane in TBST buffer for at least 15 min while changing the buffer every 5 min. 3. Incubate membranes with the appropriate amount of primary polyclonal or monoclonal antibody at 4°C for 16–24 h with continuous gentle agitation (see Note 4). 4. Rinse nitrocellulose or PVDF membrane as described in step 2 and then incubate with the appropriate amount of the HRPconjugated secondary antibody for 1 h at room temperature or overnight at 4°C with continuous gentle agitation (see Note 4). 5. Repeat step 2 and visualise protein of interest using a standard chemiluminescence imager. The intensities of the bands can be quantified using software supplied with most commercially available imagers. 6. To probe the membrane for a different protein, remove primary and secondary antibodies by incubating the membrane in stripping buffer at 56°C for 5–10 min depending on the strength of the signal obtained with previous antibody. Revisualise blot using a chemiluminescence imager to ensure that the antibody has been completely removed. 7. Proceed with detection from step 1 above using a different antibody at step 3. Membrane can be stripped and reprobed with a different primary and secondary antibodies many times (see Note 10).

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4. Notes 1. We routinely use protease inhibitor cocktail at concentration that 1 ml of the protease inhibitor cocktail is sufficient for the inhibition of proteases extracted from 20 g of bovine liver. 2. There are numerous types of protein beads available on the market, therefore, consult the literature and manufacturer’s websites. Use the most suitable commercially available protein bead at the right pH to ensure optimum elution of desired protein from the beads. 3. Incubate cells for shorter periods (1–3 min) if working with fragile cells such as osteoclasts. 4. Prepare antibody in blocking buffer (3% bovine serum albumin or 5% low fat milk) to reduce non-specific binding during the incubation period. Avoid using large quantities of an antibody or a cocktail of two or more primary antibodies to minimise non-specific binding and antibody to antibody cross reaction. 5. Use fresh protease and phosphatase inhibitors to avoid antigen degradation during immunoprecipitation. 6. Pre-cast SDS polyacrylamide gels are convenient but expensive and most have a short shelf life. 7. Standard marker with known apparent molecular weights should be loaded alongside the sample to aid with the identification of molecular weight of protein under investigation. Load multicoloured kaleidoscope pre-stained standard marker to check the quality of transfer. 8. Consult manufacturers of gel and transfer equipment on deciding the current at which the electrophoresis and/or electrophoretic transfer can be carried out. The duration of transfer depends on the size of protein under investigation – larger proteins require more time to transfer across. 9. This step is essential to ensure blocking of non-specific binding sites. 10. When stripping primary and secondary antibodies, membrane bound proteins – in particular phosphorylated proteins – may be damaged or affected resulting in reduced sensitivity in subsequent detections. This can be avoided by using fluorescent imagers, which allow detection of multiple proteins simultaneously.

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References 1. Gomperts, B. D., Kramer, I. M., and Tatnell, M. A. (2004) Signal transduction. Elsevier. 2. Feng, X. RANKing intracellular signaling in osteoclasts. (2005) IUBMB. Life 57, 389–395. 3. Swarthout, J. T., D’Alonzo, R. C., Selvamurugan, N., and Partridge, N. C. (2002) Parathyroid hormone-dependent signaling pathways regulating genes in bone cells. Gene 282, 1–17. 4. Hubbard, M. J., and Cohen, P. (1993) On target with a new mechanism for the regulation of protein phosphorylation. Trends Biochem. Sci. 18, 172–177. 5. Idris, A.I, Mrak. E, Greig. I, Guidobono, F., Ralston, S. H., van ’t Hof, R. (2008) ABD56 causes osteoclast apoptosis by inhibiting the NFkappaB and ERK pathways. Biochem. Biophys. Res. Commun. 371, 94–98. 6. Idris, A. I., Libouban, H., Nyangoga, H., Landao-Bassonga, E., Chappard, D., and Ralston, S. H. (2009) Pharmacologic inhibitors of IkappaB kinase suppress growth and

7.

8.

9.

10.

11.

migration of mammary carcinosarcoma cells in vitro and prevent osteolytic bone metastasis in vivo. Mol. Cancer Ther. 8, 2339–2347. Idris, A. I., Krishnan, M., Simic, P., LandaoBassonga, E., Mollat, P., Vukicevic, S., and Ralston, S. H. (2010) Small molecule inhibitors of I{kappa}B kinase signaling inhibit osteoclast formation in vitro and prevent ovariectomy-induced bone loss in vivo. FASEB J. 24, 4545–4555. Cohen, P. Protein kinases--the major drug targets of the twenty-first century? (2002) Nat. Rev. Drug Discov. 1, 309–315. Zhang, F. L., and Casey, P. J. (1996) Protein prenylation: molecular mechanisms and functional consequences. Annu. Rev. Biochem. 65, 241–269. Rogers, M. J. (2003) New insights into the molecular mechanisms of action of bisphosphonates. Curr. Pharm. Des. 9, 2643–2658. Kurien, B. T., Scofield, R. H. (2006) Western blotting. Methods. 38, 283–293.

Chapter 16 Analysis of Transcriptional Regulation in Bone Cells Huilin Jin and Stuart H. Ralston Abstract Transcription is the process by which the rate of RNA synthesis is regulated. Here, we describe the techniques for carrying out promoter-reporter assays, electrophoretic mobility shift assays, and chromatin immunoprecipitation assays, three commonly used methods for studying gene transcription. Key words: Transcription, Chromatin, Immunoprecipitation, ChIP, EMSA, Promoter, Gene

1. Introduction Gene expression is regulated by the process of transcription which governs the rate at which ribonucleic acid (RNA) is produced. Transcription is carried out by RNA polymerase enzymes but they cannot initiate transcription in isolation. Instead, the activity of RNA polymerase is regulated by other proteins called transcription factors, which bind to regulatory elements in DNA, either in the gene promoter or at other regulatory sites within or around the gene. In this chapter, we describe three commonly used methods for the study of gene transcription. These are promoter-reporter assays (1), electrophoretic mobility shift assays (1), and chromatin immunoprecipitation assays (2); all of these techniques have previously been used to investigate the functional effects of polymorphisms and mutations that have been associated with osteoporosis (3–6) and other bone diseases (7). In promoter-reporter assays, DNA sequences containing putative regulatory elements are cloned upstream of a reporter gene such as Firefly luciferase and transfected into cultured cells. In order to control for transfection efficiency the cells are usually cotransfected with another vector such as β-galactosidase or Renilla luciferase. The activity of the promoter or enhancer of interest is then assessed by measuring activity

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_16, © Springer Science+Business Media, LLC 2012

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of the luciferase reporter in cell lysates with correction for activity of the control vector. Electrophoretic mobility shift assays (EMSA) are used to study the binding of transcription factors to recognition sites in DNA and/or their binding affinity for the recognition site. They involve incubating labeled double-stranded oligonucleotides containing the recognition site of interest with nuclear protein extracts. Then, the reaction mixture is subject to gel electrophoresis to assess if proteins within the extracts bind the DNA and slow migration of the oligonucleotides through the gel. Chromatin immunoprecipitation (ChIP) assays are used to study interaction of transcription factors with DNA in chromosomal fragments. These assays involve the use of specific antibodies to immunoprecipitate fragments of DNA bound to the transcription factor of interest followed by PCR amplification of the DNA sequences flanking the recognition sites in the gene of interest. The advantage of ChIP assays over EMSA assays is that they examine transcription factor binding in a chromosomal context. They can also be used to identify cis-acting sequences in the genome when combined with DNA sequencing (ChIP-Seq) and can also be used to quantify allele-specific transcription when combined with quantitative PCR.

2. Materials 2.1. Promoter-Reporter Assay

1. Endofree MaxiPrep Kit (Qiagen). 2. Spectrophotometer. 3. Dual-luciferase® Reporter Assay System, including pGL3 & pRL-TK Luciferase Reporter Vectors and associated buffers and reagents (Promega). 4. Magnetic assisted transfection kit and magnetic plate (MATra, IBA BioTAGnology). 5. Luminometer. 6. Tissue culture facilities. 7. Culture medium (aMEM medium with 10% fetal calf serum, 50 IU/ml penicillin, and 2 mM glutamine).

2.2. Nonradioactive Electrophoretic Mobility Shift Assay

1. Oligonucleotides flanking target DNA–protein binding site with or without biotin-labeling. 2. 10× Annealing buffer: 1 M Tris, pH 8.0, 5 M NaCl, and 0.5 M EDTA. 3. 5× TBE Buffer: 445 mM Tris Base, 445 mM Boric Acid, and 10 mM EDTA, pH 8.0. Dilute 1 in 10 with distilled water before use.

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4. Lightshift® Chemiluminescent EMSA Kit (Pierce, UK). 5. Positively charged nylon membrane (Pierce). 6. UV lamp or transilluminator equipped with 312 nm bulb. 7. Electrophoresis apparatus. 8. Electroblotter or capillary transfer apparatus. 9. Blotting paper and pad. 10. Polyacrylamide gel. 2.3. Chromatin Immunoprecipitation Assay

1. Nuclear extract kit (Active Motif ®). 2. 37% formaldehyde. 3. 1 M glycine in PBS. 4. PBS with protease inhibitor cocktail: One Roche protease inhibitor cocktail tablet dissolved in 10 ml PBS (keep on ice until use). 5. Lysis buffer: 50 mM Tris, pH 8.1, 10 mM EDTA and 1% (w/v) SDS. 6. Protease inhibitor cocktail in lysis buffer: One Roche protease inhibitor cocktail tablet in 10 ml lysis buffer from step 5. 7. IP Dilution buffer: 16.7 mM Tris pH 8.1, 1.2 mM EDTA, 167 mM NaCl, 1.1% Triton X 100 (v/v), and 0.01% (w/v) SDS. 8. TSE I Buffer: 20 mM Tris pH 8.1, 2 mM EDTA, 150 mM NaCl, 1% (v/v) Triton X-100, and 0.1% (w/v) SDS. 9. TSE II buffer: 20 mM Tris pH 8.1, 2 mM EDTA, 500 mM NaCl, 1% (v/v) Triton X-100, and 0.1% (w/v) SDS. 10. Elution buffer: 0.1 M NaHCO3 and 1% (w/v) SDS (make up fresh before use). 11. Buffer III: 10 mM Tris pH 8.1, 0.25 M LiCl, 1 mM EDTA, 1% (v/v) NP40, and 1% (w/v) sodium deoxycholate. 12. TE buffer: 50 mM Tris pH 8.1 and 1 mM EDTA. 13. Phenol–Chloroform–Isoamyl alcohol (25:24:1) or Quiaquick PCR purification kit. 14. Sonicator.

3. Methods 3.1. Dual Luciferase Reporter Assay

This technique involves cloning the target promoter region to upstream of the reporter gene and measuring Firefly luciferase activity in cell lysates. In the example given, a second Renilla luciferase vector is cotransfected to provide an internal control.

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The advantage of using Firefly luciferase and Renilla luciferase together is that the activities of both vectors can be measured sequentially in the same cell lysate by using different substrates for each enzyme. Here, we describe the use of the MATra magnetic transfection system to introduce a PGL-3 reporter vector and a PRL-TK control vector into human osteosarcoma TE85 cells. 1. Purify the vectors using an EndoFree Plasmid Maxi Kit according to the manufacturer’s recommendations (see Note 1). 2. Determine the DNA concentrations by UV spectrophotometry (see Note 2) and store the vectors at −20°C until required. 3. Culture the TE85 cells in medium using 75-cm2 tissue culture flasks at 37°C in a humidified atmosphere of 5% CO2. Replace the medium every 3 days until the cells are nearly confluent. 4. Twenty-four hours prior to carrying out the transfection, passage the cells and seed into 24-well plates at a density of 4 × 104 cells per well. 5. For each well to be transfected, combine 0.6 mg of PGL-3 reporter vector with 0.1 mg of PRL-TK control vector and 50 ml aMEM without serum, glutamine, or antibiotics. 6. For each well to be transfected, add 0.6 ml MATra-A reagent, to the diluted reporter vector mix in step 5 and mix thoroughly. Incubate at room temperature for 20 min. 7. Remove the culture medium from each well of the 24-well plate and replace with 500 ml fresh culture medium. 8. Add 50 ml of the vector/MATra-A reagent mix from step 6 to each well of the plate and gently mix by swirling the medium around the plate. 9. Place the 24-well plate on the Magnetic Plate and incubate for 15 min at 37°C in a tissue culture incubator set at 37°C in a humidified atmosphere of 5% CO2. 10. Remove the Magnetic Plate and continue to culture cells. Check the cells after 6 h for evidence of toxicity (e.g., detachment of cells or rounding up of cells) and if present replace the transfection mixture with normal culture medium. 11. Incubate the cells for 24 h after transfection in the presence or absence of test agents. 12. Terminate the experiment by aspirating the culture medium. 13. Wash each well of the plate three times with PBS. 14. Add 80–100 ml lysis buffer provided with the Dual-luciferase® Reporter Assay System, to completely cover each well of the tissue culture plate. 15. Wrap the plate in foil paper and store it at −80°C for at least 24 h.

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16. Thaw the plates at room temperature and transfer 40 ml of the lysate from each sample into a 96-well clear bottom plate, ensuring that there are no bubbles in the sample (see Note 3). 17. Measure Firefly luciferase activity and Renilla luciferase activity sequentially by auto-injection of 40 ml of luciferase assay reagent and 40 ml Stop & Glo® reagent (provided with the Dual Luciferase reporter assay system) into the lysate from each well using a luminometer fitted with two injectors (see Note 4). 18. Assess the activity of the promoter of interest by correcting the Firefly luciferase activity to that of Renilla activity for each sample. 19. Calculate values for each of the replicates and assess between group differences using an appropriate statistical test such as t-Test (for paired samples) or ANOVA. This technique is based on the fact that protein–DNA complexes migrate through a nondenaturing polyacrylamide or agarose gel more slowly than free DNA molecules. The EMSA technique involves mixing a crude nuclear extract or sample of purified protein with a labeled oligonucleotide probe containing the putative transcription factor binding site. The samples are then analyzed by electrophoresis through a nondenaturing polyacrylamide. Oligonucleotides that are bound to transcription factors migrate more slowly through the gel than free oligonucleotides and can be detected as a “gel shift” (see Fig. 1). Furthermore specificity of the DNA–protein binding is established by addition of

3.2. Electrophoretic Mobility Shift Assay

a

b

Probe Protein Antibody Specific competitor non-specific competitor

+ -

+ + -

+ + + -

+ + + -

+ + +

Probe (Allele 1)

+

+

+

+

+

+

+

+

Protein

-

+

+

+

+

+

+

+

Competitor(Allele 1) -

-

+

+

+

-

-

-

Competitor(Allele 2) -

-

-

-

-

+

+

+

supershift shift

shift

Free DNA probe

Free DNA probe

Fig. 1. In the EMSA, DNA–protein binding complex migrates more slowly than free DNA probe on the gel, called a shift. (a) Testing the specificity of a DNA–protein binding reaction. Addition of a protein specific antibody further shifts the band called a supershift. The DNA–protein shift is completely (or partly) eliminated by addition of an unlabeled specific competitor which contains known DNA–protein binding sequence. However, a nonspecific competitor will not alter the shift. (b). Testing the DNA–protein binding affinity between alleles. Unlabeled competitor at higher concentrations competitively binds to the protein and blocks the labeled DNA probe–protein binding, which causes a reduction in intensity of the shift. For example, unlabeled competitor with allele 2 eliminates the labeled DNA–protein shift at a lower concentration than the one with allele 1. Therefore allele 2 has higher binding affinity to this protein than allele 1.

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the specific antibody or competition experiments using oligonucleotides containing a binding site for the protein of interest or other unrelated DNA sequences. The technique described here uses a nonradioactive method of detecting the DNA–protein complexes (5–9) instead of the radioactive methods of detection that have been used previously by ourselves and other investigators (4–6). 3.2.1. Probe Design

1. Obtain a pair of complementary oligonucleotides (probes) flanking the putative DNA–protein binding site of interest with HPLC purification. 2. End-label the forward strand sequence with biotin (see Note 5). 3. Combine 2 mM of biotin-labeled forward probe with 3 mM reverse probe, 5 ml 10× annealing buffer and make up to 50 ml with distilled water. 4. Mix well and incubate at 90°C for 1 min. 5. Allow it to cool slowly to room temperature on the bench (approximately 1 h). 6. Store the resulting double-stranded probes at −20°C until required. 7. To make double-stranded competitor probe, follow the above steps 3–6 but replace the biotin-labeled forward probe with unlabeled forward probe.

3.2.2. Preparation of Nuclear Extracts

1. Culture the cells of interest in a 75-cm2 tissue culture flask until near confluent. 2. Remove the medium and wash the cells three times with icecold PBS. 3. Extract nuclear proteins from cells using the Active Motif® nuclear extract kit according to the manufacturer’s instructions. 4. Determine the protein concentration in the extract using a standard commercially available protein assay. Store the extracts frozen in 10 ml aliquots at −80°C until further use (see Note 6).

3.2.3. Setting up the EMSA Binding Reaction

1. Prepare the binding reactions according to the schedule shown in Table 1a if you wish to test the specificity of a putative DNA–protein binding, and Table 1b if you wish to test the DNA–protein binding affinity of different probes. 2. Incubate binding reactions at room temperature for 20 min.

3.2.4. Electrophoresis and Transfer from Gel to Membrane

1. Place a precast 4–6% native polyacrylamide minigel (8 × 8 × 0.1 cm), into a vertical electrophoresis tank filed with 0.5× TBE buffer (see Note 7). 2. Prerun the gel at a 100 V for 60–90 min.

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Table 1 Typical setup for EMSA binding reactions a. Binding reactions to test the specificity of putative DNA–protein binding Component

#1

#2

#3

#4

#5

Ultrapure Water (ml)

15

14

13

13

13

10× Binding Buffer (ml)

2

2

2

2

2

BSA (10 mg/ml) (ml)

2

2

2

2

2

Unlabeled specific competitor probe (4 mM) (ml)

–

–

–

Unlabeled nonspecific competitor probe (4 mM) (ml)

–

–

–

Hos TE85 protein extract (8–10 mg/ml) (ml)

–

Antibody (ml)

–

Biotin-labeled probe (0.04 mM) (ml) Total volume (ml)

1

1 –

1

–

– 1

1

1

1

1

1

1

1

1

20

20

20

20

20

b. Binding reactions to test binding affinity between different probes Component

#1

#2

#3

#4

#5

#6

#7

#8

Ultrapure Water (ml)

15

14

13

12

10

13

12

10

Binding Buffer (10×) (ml)

2

2

2

2

2

2

2

2

BSA (10 mg/ml) (ml)

2

2

2

2

2

2

2

2

1

2

4

Unlabeled specific competitor probe 1 (1 mM) (ml)

–

–

Unlabeled specific competitor probe 2 (1 mM) (ml)

–

–

Hos TE85 protein extract (8–10 mg/ml) (ml)

–

Biotin-labeled probe 1 (0.04 mM) (ml) Total volume (ml)

–

–

–

–

–

–

1

2

4

1

1

1

1

1

1

1

1

1

1

1

1

1

1

1

20

20

20

20

20

20

20

20

®

Binding buffer (10×) is supplied in the LightShift Chemiluminescent EMSA Kit (Pierce). Other components are also supplied in the kit for optimization of the binding reaction such as glycerol, Poly (dI·dC), NP-40. Always add the components in descending order as listed in the table. To reduce nonspecific binding, a short preincubation (~2 min) may be required before adding the biotin-labeled probe

3. Mix the binding reaction with 5 ml 5× loading buffer (provided with the chemiluminescent EMSA kit) by pipetting up and down several times. 4. Flush each well with 0.5× TBE buffer and load carefully 20 ml of each sample. 5. Run the gel at a 100 V until the bromophenol blue dye migrates approximately three quarters of the way down the gel.

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Well Glass plates

Stacking gel

Resolving gel

Spacer

Clip polyacrylamide stacking gel

b

SDS polyacrylamide gel

pad pad PVDF membrane

+

Fig. 2. (a) Gel assembly; (b) transfer assembly.

6. Remove gel from the cassette and immerse it in 0.5× TBE buffer for a few minutes. 7. Meanwhile, soak a PVDF membrane, filter paper, and pad in 0.5× TBE buffer for 10 min. 8. Make a blotting sandwich by arranging successive layers of one presoaked filter pad, two filter papers, PVDF membrane, gel, presoaked filter paper, and pad (see Fig. 2). 9. Remove any air bubbles by using a pencil or a glass tube to flatten the filter papers and pads during the layering process. 10. Close the cassette firmly and place it in a clean electrophoretic transfer unit. 11. Add the cooling unit and fill the tank with 0.5× TBE buffer. 12. Place the whole tank in a cold room or a container filled with plenty of ice. 13. Run the blot at 380 mA for 60 min. When the transfer is complete cross-linked the membrane for 15 min by placing face down on a transilluminator equipped with 312 nm bulb.

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The buffers referred to in this section are not listed individually in the materials section but are included in the Lightshift® Chemiluminescent EMSA kit. 1. Immerse the membrane in 10 ml blocking buffer for 15 min at room temperature with gentle shaking. 2. Prepare Conjugate/Blocking buffer by adding 66.7 ml Stabilized Streptavidin–Horseradish Peroxidase Conjugate to 20 ml Blocking Buffer. 3. Discard the Blocking buffer and add 10 ml Conjugate/ Blocking buffer. Incubate for 15 min at room temperature with gentle shaking. 4. Discard and rinse the membrane five times for 5 min each in 10 ml 1× Washing buffer with gentle shaking. 5. Transfer the membrane to a new container and incubate with 10 ml Equilibrium Buffer for 5 min with gentle shaking. 6. Remove the membrane and excess buffer. Place the membrane in a clean sheet of plastic wrap. Pour 600 ml Substrate Working buffer over the membrane and incubate for 5 min without shaking. 7. Visualize DNA–protein binding complex using a standard chemiluminescence imager. The intensity of the band can be quantified using image analysis software.

3.3. Chromatin Immunoprecipitation Assay

3.3.1. Cross-Link Chromatin

The Chromatin Immunoprecipitation Assay (ChIP) involves crosslinking cell extracts to bind chromatin-associated proteins to the DNA, shearing the DNA into fragments of between 300 and 1,000 bp by sonication, and immunoprecipitating the DNA– protein complexes using specific antibodies directed against the protein of interest. The immunoprecipitated DNA fragments are then purified and analyzed by PCR or DNA sequencing (see Fig. 3). In this example we described an adaptation of the ChIP technique developed by Naughton (9) using MG63 human osteosarcoma cells as the source of chromatin (5). 1. Culture the MG63 cells in culture medium in two 10 cm tissue culture dishes until nearly confluent. 2. Cross-link the chromatin to the protein by adding 37% formaldehyde directly to the culture medium to a final concentration of 1% (v/v) and incubate for 10 min at 37°C. 3. Add glycine to a final concentration of 0.125 M to the culture medium and incubate for 10 min at room temperature with gentle rocking. 4. Aspirate the medium and wash the cells twice with ice-cold PBS. 5. Scrape the cells from each plate into separate tubes and add 1 ml of PBS with protease inhibitor cocktail solution to each and mix by pipetting up and down.

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Fig. 3. Overview of Chromatin precipitation (ChIP).

6. Pellet the cell lysate by centrifugation at 450 × g for 4 min at 4°C (see Note 8). 7. Resuspend the cell pellets from each plate in 200 ml lysis buffer with protease inhibitor cocktail and mix by gently pipetting up and down taking care to avoid forming air bubbles. Incubate for 10 min on ice. 8. Combine the lysates (400 ml) from both culture plates into a single tube. 9. Sonicate the lysates to shear the DNA into lengths of between 300 and 1,000 bp with 12 sets of 20 s pulses at 2 Am. Keep samples on ice at all times to avoid overheating and sonicating at intervals of not less than 1 min (see Note 9). 10. Centrifuge samples at 20,000 × g for 10 min at 4°C. 11. Aliquot 100 ml of the sheared chromatin extract to a fresh 1.5 ml microcentrifuge tube and store frozen at −80°C until further use. 3.3.2. Examination of the Sheared Chromatin

This step can be used to ensure that the chromatin has been sheared into fragments of the desired size by the sonication procedure. 1. Add 5 ml 4 M NaCl to a 100 ml sample from step 11 in Subheading 3.3.1 and incubate for 4 h at 65°C. 2. Purify the DNA by adding 100 ml Phenol–Chloroform– Isoamyl alcohol (25:24:1) to each 100 ml sample. Vortex for 10 s and centrifuge at 20,000 × g for 5 min at 4°C. 3. Carefully aspirate 100 ml of the supernatant and transfer to a fresh 1.5 ml microcentrifuge tube.

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Fig. 4. MG63 cells were fixed with 1% formaldehyde and 0.125 M glycine, and then chromatin was prepared using PBS and lysis buffer as described. Chromatin was fragmented with 12 pulses at 2 Am using a sonicator in a volume of 400 mL. Each pulse consists of a 20 s sonication followed by a 60 s rest on ice. 100 mL of the sheared chromatin were extracted and precipitated as described. Sample (lane 2 ) was separated by electrophoresis through a 1% agarose gel along with a 100-bp DNA ladder (NEB, lane 1 ).

4. Add 10 ml 3 M Acetate and 220 ml ice-cold 100% ethanol to each sample and vortex. 5. Place the solution in a −80°C freezer for 15 min or a −20°C for 30 min to allow the DNA to precipitate. 6. Centrifuge the sample at 20,000 × g for 5 min at 4°C. 7. Aspirate the supernatant and wash the DNA pellet by adding 1 ml 70% ethanol and inverting the tube a few times. 8. Collect the DNA pellet at the bottom of the tube by centrifuging the sample at 20,000 × g for 5 min at 4°C. 9. Briefly air-dry the pellet for no more than 5 min at room temperature. 10. Suspend the DNA pellet in 30–50 ml TE buffer. 11. Run aliquots of sample (e.g., 5, 10, 15, and 20 ml) and a 100bp DNA ladder on a 1.5% agarose gel to visualize the size of the sheared chromatin fragments (see Fig. 4). 3.3.3. Immunoprecipitation of Chromatin Bound to Protein

1. Prepare fresh IP dilution buffer by adding one protease inhibitor tablet to 10 ml IP dilution buffer. 2. Add 800 ml IP dilution buffer to 100 ml of chromatin extract from Subheading 3.3.1, step 11 (see Note 10). 3. Remove 30 ml of the above sample and save as an input/starting control for the subsequent PCR.

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4. Add 1 ml IP dilution buffer to 50 ml Protein G-Agarose beads and mix well, preparing additional tubes for each sample of chromatin extract that you wish to analyze. Pellet the beads by centrifugation at 450 × g for 1 min at 4°C and aspirate the supernatant. 5. Wash the beads with IP dilution buffer by repeating step 4 on a further two occasions. 6. Add 1 mg mouse IgG, and 2 mg salmon sperm DNA to the washed beads and make up to a final volume of 100 ml with IP dilution buffer (see Note 11). 7. Add 900 ml of the chromatin mixture from step 2, to the washed beads containing mouse IgG and salmon sperm DNA from step 6. Incubate for at least 3 h at 4°C in a rotary mixer set at 15 rpm. 8. Centrifuge at 450 × g for 1 min at 4°C and aspirate the supernatant using a syringe and fine needle. Transfer the supernatant to a fresh Eppendorf tube for the immunoprecipitation (step 12). 9. Add 50 ml fresh Protein G-Agarose beads to a new tube and wash with IP dilution buffer as described in steps 4–5 above, preparing separate tubes for each sample that you wish to analyze and include an additional tube as a negative control. 10. Add 5 mg of the antibody specific for the protein of interest (see Note 12) and 2 mg salmon sperm DNA to the washed beads from step 9 and make up to a final volume of 100 ml in each tube with IP dilution buffer. 11. Prepare the negative control tube as described in step 10, in which mouse IgG is added in place of the specific antibody. 12. Add the samples of chromatin containing supernatant from step 8 to the mix of Protein G agarose beads, salmon sperm DNA, and antibody from step 10 and 11. 13. Incubate the samples overnight at 4°C in rotary mixer set at 15 rpm. 14. Centrifuge at 450 × g for 1 min at 4°C. Aspirate the supernatant using a syringe and fine needle and discard. 15. Wash the pellet by adding 1 ml TSE I buffer and mix by incubating in a rotary mixer at 4°C set at 20 rpm for 5 min. 16. Centrifuge at 450 × g for 1 min at 4°C. Aspirate the supernatant using a syringe and fine needle and discard. 17. Wash the pellet by adding 1 ml TSE II buffer and mix by incubating in a rotary mixer at 4°C set at 20 rpm for 5 min. 18. Centrifuge at 450 × g for 1 min at 4°C. Aspirate the supernatant using a syringe and fine needle and discard.

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19. Wash the agarose pellet by adding 1 ml buffer III and mix by incubating in a rotary mixer at 4°C set at 20 rpm for 5 min. 20. Centrifuge at 450 × g for 1 min at 4°C. Aspirate the supernatant using a syringe and fine needle and discard. 21. Wash the agarose pellet by adding 1 ml TE buffer and incubate on a rotary mixer set at 20 rpm for 5 min at 4°C. 22. Centrifuge at 450 × g for 1 min at 4°C. Aspirate the supernatant using a syringe and needle and discard. 23. Repeat steps 21 and 22 once more and proceed to recover the immunoprecipitated DNA as described in Subheading 3.3.4. 3.3.4. Recover DNA Bound to Protein of Interest

1. Add 250 ml freshly made elution buffer to each sample from Subheading 3.3.3, step 23 and incubate on a rotary mixer set at 15 rpm for 30 min at room temperature. 2. Centrifuge at 450 × g for 1 min at 4°C. Aspirate the supernatant carefully using a syringe and needle and transfer to a fresh tube. 3. Add 200 ml elution buffer to the agarose pellet remaining in the tube from step 2 and incubate on a rotary mixer set at 15 rpm for 15 min at room temperature. 4. Centrifuge at 450 × g for 1 min at 4°C. Aspirate the supernatant carefully using a syringe and needle and transfer to a fresh tube. 5. Combine both elution products together from steps 2 and 4 for each sample (450 ml). 6. Add 25 ml 4 M NaCl to each 450 ml sample from step 5 and 1.7 ml 4 M NaCl to 30 ml input control (from Subheading 3.3.3, step 3). Incubate all samples at 65°C for 6 h. (At this point samples can be stored at −20°C overnight). 7. Prepare fresh recovery buffer by mixing 10 ml 0.5 M EDTA, 20 ml 1 M Tris–HCl (pH 6.5), and 2 ml 10 mg/ml proteinase K for each sample. Add 32 ml of the recovery buffer into each test sample from step 6 and 2.12 ml into the input control. 8. Incubate all samples at 45°C for 1 h. 9. Clean the DNA using a QIAquick PCR purification kit according to the manufacturer’s instructions or perform a phenol– chloroform extraction followed by ethanol precipitation. 10. Recover the DNA from step 9 and suspend in 30 ml dH2O. Store samples at −20°C for use in subsequent PCR or DNA sequencing reactions (see Note 13).

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4. Notes 1. The following vectors are required for a transfection experiment: a Firefly luciferase reporter vector (PGL3 basic) containing the promoter sequence of interest, a negative control vector which does not contain the promoter sequence (PGL3 basic), a positive control vector which contains a promoter sequence such as SV40 that is constitutively active in Eukaryotic cells (e.g., PGL3 promoter) and a Renilla luciferase vector which constitutively expresses Renilla luciferase as an internal control (e.g., PRL-TK). 2. High quality DNA is required for the subsequent transfection. That is evaluated by the A260/280 reading from a spectrophotometry. Ensure the values are close to 1.8. 3. The exact amount of lysate used in the detection of luciferase expression depends on the activity of target promoter sequence and should be determined empirically for each experiment. 4. If using a luminometer without two injectors, predispense 40 ml of LAR II reagent into a luminometer tube. Add lysate to the luminometer tube and mix. Place the tube in the luminometer and measure the Firefly luciferase activity. Then add 40 ml Stop & Glo Reagent to the same tube either manually or using a reagent injector. Mix briefly and measure the Renilla luciferase activity from the same sample. 5. This can be carried out in house by using a commercially available Biotin 3¢end DNA labeling kit (e.g., Pierce) or ordering 5¢ end labeling when the probe is synthesized by a manufacturer (e.g., Invitrogen). 6. The yield of protein varies between cell types but for HOS TE85 cells growing in a 75 cm2 culture flask the total protein yield should be in the range of 8–15 mg/ml. Note that the frozen aliquots are best for single use and freeze/thaw cycles should be avoided. 7. Commercially available mini gels work very well for reactions with small to medium size probes (<300 bp). The percentage of the gel depends on the size of the probe and the size, number, and the charge of the proteins that bind. 8. The lysates can be stored frozen at −80°C at this point until further use. 9. You might need to vary the number of the 20-s pulse to break DNA into different sizes. The shearing fragment sizes should be 100–1,000 bp smear with an average length of 400–500 bp. Once the conditions have been optimized, keep cell number and the all above steps consistent for subsequent experiments.

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If samples from different experiments are being used ensure that the sonicator probe is cleaned with 75% ethanol between samples. 10. Depending on the abundance of the DNA–protein binding you might need to add less IP dilution buffer to the chromatin extract so that the extract is more concentrated. 11. This step is required to reduce nonspecific binding. 12. The amount of antibody used needs to be determined empirically for each experiment, depending on the abundance of the protein and the antibody’s affinity for the protein. One tube should also be prepared in which the antibody has been omitted to act as a negative control. 13. Several options are possible at this stage. For example, to verify the binding of a transcriptional factor to the gene promoter of interest in cells, a PCR reaction could be performed to amplify the target promoter region using the DNA which was enriched in above IP using antibody to the transcriptional factor. Quantitative real-time PCR assay could be used to further quantify the binding. References 1. Strachan, T., and Read, A. P. (2011) Human Molecular Genetics. Garland Science, New York. 2. Das, P. M., Ramachandran, K., vanWert, J., and Singal, R. (2004) Chromatin immunoprecipitation assay. Biotechniques 37, 961–969. 3. Morrison, N. A., Yeoman, R., Kelly, P.J., and Eisman, J.A. (1992) Contribution of trans-acting factor alleles to normal physiological variability: Vitamin D receptor gene polymorphisms and circulating osteocalcin. Proc.Natl.Acad.Sci. USA. 89, 6665–6669. 4. Mann, V., Hobson, .E. E., Li, B., Stewart, T. L., Grant, S .F., Robins, S. P., Aspden, R. M., and Ralston, S. H. (2001) A COL1A1 Sp1 binding site polymorphism predisposes to osteoporotic fracture by affecting bone density and quality. J. Clin. Invest. 107, 899–907. 5. Jin, H., van’t Hof, R. J., Albagha, O. M., and Ralston, S. H. (2009) Promoter and intron 1 polymorphisms of COL1A1 interact to regulate transcription and susceptibility to osteoporosis. Hum. Mol. Genet. 18, 2729–2738.

6. Arai, H., Miyamoto, K. I., Yoshida, M., Yamamoto, H., Taketani, Y., Morita, K., Kubota, M., Yoshida, S., Ikeda, M., Watabe, F., Kanemasa, Y., and Takeda, E. (2001) The polymorphism in the caudal-related homeodomain protein Cdx-2 binding element in the human vitamin D receptor gene. J. Bone Miner. Res. 16, 1256–1264. 7. Leupin, O., Kramer, I., Collette, N. M., Loots, G. G., Natt, F., Kneissel, M., and Keller, H. (2007) Control of the SOST bone enhancer by PTH using MEF2 transcription factors. J. Bone Miner. Res. 22,1957–1967. 8. Garcia-Giralt, N., Enjuanes, A., Bustamante, M., Mellibovsky, L., Nogues, X., Carreras, R., ez-Perez, A., Grinberg, D., and Balcells, S. (2005) In vitro functional assay of alleles and haplotypes of two COL1A1-promoter SNPs. Bone 36, 902–908. 9. Naughton, C., MacLeod, K., Kuske, B., Clarke, R., Cameron, D. A., and Langdon, S. P. (2007) Progressive loss of estrogen receptor alpha cofactor recruitment in endocrine resistance. Mol.Endocrinol. 21, 2615–2626.

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Chapter 17 Extraction of Nucleic Acids from Bone Alun Hughes, Tracy L. Stewart, and Val Mann Abstract Here, we present methods for extracting DNA and RNA from samples of whole bone tissue and culture bone cells and describe methods quantitative and qualitative measurement of the extracted nucleic acids. These protocols described provide high-quality nucleic acids suitable for downstream applications such as quantitative PCR and microarrays. Key words: RNA extraction, DNA extraction, RNA quantification, DNA quantification, RNA integrity

1. Introduction Nucleic acids can be extracted from bone for analysis of gene expression, to look for somatic mutations in the analysis of tumors or other pathological tissue, or for genotyping archive material when other sources of DNA are not available. Extraction of nucleic acids of sufficient purity, abundance, and quality from bone presents many problems, especially in maintaining the integrity of RNA that is essential for the accurate measurement of gene expression. Factors such as the speed of isolation and how samples are stored prior to isolation can greatly affect the quality of the extracts prepared. For example, if delays occur in transportation of samples from clinic to the laboratory this can cause degradation of RNA leading to inconsistent and unreliable results in gene expression analysis. Animal bone tissues must likewise be handled and stored appropriately to prevent nucleic acid degradation. High-purity nucleic acid is also essential for the success of many downstream applications. For both DNA and RNA this involves preparing samples devoid of protein or solvent contamination, as both can adversely affect the success of post-purification processing. For gene expression work, it is also important to ensure that the extracted 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_17, © Springer Science+Business Media, LLC 2012

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RNA is free from DNA contamination, to avoid false-positive results in subsequent PCR-based assays which do not differentiate between cDNA and genomic DNA. Several methods have been published for the extraction of DNA and RNA from cultured cells and nonmineralized tissues (1–4). Here, we describe methods for isolating DNA and RNA from whole bone and bone cell cultures in a form suitable for many downstream applications such as PCR, DNA sequencing, quantitative RT/PCR, and microarray analysis

2. Materials 1. Cryogenic Mill to pulverise bone samples (e.g., SPEX 6770 Freezer/Mill, Wolf Laboratories, York, UK).

2.1. Equipment

2. Sterile scalpels, scissors and bone cutters. 3. Sterile 35-mm tissue culture plates. 4. 19–20 G needles and 2-ml syringes. 5. Mortar and pestle. 6. Spectrophotometer for measuring nucleic acid yield. 7. Horizontal electrophoresis tank for measuring RNA integrity. 1. Tris-saturated phenol (pH 7.8–8.0) (Sigma).

2.2. For DNA Extraction

2. 3 M Sodium acetate, pH 5.2. 3. Tris–EDTA: 1 mM Tris/100 μM EDTA. 4. DNA extraction buffer: 40 mM sodium citrate, pH 7.0, 25 mM sodium lauryl sarcosinate, 6 M guanidinium isothiocyanate. Add 7.2 μl β-mercaptoethanol per ml of Lysis Buffer on the day of use (see Note 1). 5. Fumed Silica: Required for DNA extraction from dried or embedded bone (Subheading 3.3). ●

Suspend 50 g of unfumed silica (Sigma) in 100 ml double distilled water. Stir for 1 h and allow to settle under gravity for 1 h.

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Aspirate the supernatant and centrifuge at 6,000 × g for 10 min to pellet the glass. Resuspend the pellet in 25–30 ml doubledistilled H2O.

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Add concentrated nitric acid to 50%. Bring to boil in a fume hood and allow to cool.

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Pellet the silica by centrifugation at 6,000 × g for 10 min.

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Wash the pellet with double-distilled water until the pH of the supernatant is 7.0.

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Store the pellet as 50% slurry in double-distilled H2O (see Note 2).

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6. Silica Wash buffer: 10 M Tris–HCl, 6 M guanidinium isothiocyanate. 7. Hoechst 33258 (see Note 3). 2.3. For RNA Extraction

1. TRIzol® reagent (InVitrogen, Paisley, UK). 2. Agarose. 3. TBE buffer (50 mM Tris–HCl pH 7.8, 50 mM orthoboric acid, 2 mM Na2EDTA). 4. 6× Orange G loading buffer (0.2% (w/v) Orange G, 30% glycerol, 0.1% SDS in distilled water). 5. Ethidium bromide (5 μg/ml) (see Note 4). 6. RiboRulerTM High Range RNA Ladder (Fermentas).

2.4.General Reagents

1. DEPC-treated water (0.1% diethylpyrocarbonate in water, incubated for 12 h at 37°C then inactivated by autoclaving) (see Note 5). 2. Chloroform. 3. Isopropanol. 4. 75 and 100% Ethanol.

3. Methods 3.1. DNA Extraction from Fresh Bone

This method is suitable for extraction of genomic DNA fragments (up to 80 Kb) from fresh or freshly frozen bone. It should be noted that the resulting DNA will be derived from primary bone cells such as osteoblasts and osteocytes as well as other cells in the bone marrow space. 1. Collect the bone sample in a sterile container containing PBS and transport to the laboratory within 1–2 h (see Note 6). 2. Place the bone tissue in a clean glass petri dish. Using bone cutters dissect out a piece of bone measuring about 1 cm3 and transfer to a sterile 35-mm cell culture plate. 3. Add 1 ml DNA extraction buffer and mince the tissue with the bone cutters finely until a slurry is obtained. 4. Transfer 500 μl of the homogenised tissue into two screwtopped conical-bottomed 1.5-ml eppendorf tubes. 5. Add one volume of Tris-saturated phenol, followed by one volume of chloroform to each tube. Mix well by inverting the tube a few times or by shaking. Do not vortex (see Note 7). 6. Centrifuge the tubes at 10,000 × g for 20 min to separate the phases.

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7. Transfer the supernatant to a fresh centrifuge tube being careful not to disturb the white layer at the interface. Make a note of the volume you transfer. 8. Add 1 volume of ice-cold isopropanol and 0.1 volumes of 3 M sodium acetate to the supernatant. Mix well and place on ice for 15 min. 9. Centrifuge the tubes for at 10,000 × g for 20 min to pellet the DNA (see Note 8). 10. Aspirate and discard the supernatant, taking care not to disturb the pellet. Wash the sample with 1.75 ml ice-cold 70% ethanol and centrifuge at 10,000 × g for 5 min. 11. Repeat step 10. 12. Carefully aspirate the supernatant and allow the DNA pellet to air-dry for 5 min. Dissolve the DNA pellet in 10–50 μl water or Tris–EDTA buffer and quantitate by spectrophotometry (see Note 9). 13. Store the sample frozen at −20°C or below until further use. 3.2. DNA Extraction from Cultured Cells

1. Remove the medium from the cultured cells and discard. 2. Wash the cell layer twice with sterile PBS. 3. Aspirate the PBS and add 1 ml DNA extraction buffer per 75 cm3 flask of cells. Swirl the Lysis Buffer around the flask so that all cells are coated. Leave for 1 min, and detach the cells using a cell scraper. 4. Transfer the extracted cell layer into screw-capped conical-bottomed 1.5-ml eppendorf tubes in 500 μl aliquots. Pipette the mixture up and down to lyse the cells. 5. Follow steps 5–12 Subheading 3.1.

3.3. DNA Extraction from Dried or Embedded Bone

of

the

extraction

procedure

in

Dried bones contain little cellular material and the following method is more suitable for these specimens. The technique relies on the propensity of nucleic acids to adsorb to silica (5). Kits based upon this methodology are available (e.g., High Pure kit, Roche, Burgess Hill, UK). 1. Homogenise the bone sample using a liquid nitrogen-cooled powder mill or manually by cooling the specimen with liquid nitrogen and then grinding to a fine powder in a mortar and pestle or an electric food grinder. 2. Add 500 μl DNA extraction Buffer to a clean eppendorf and place on ice. Carefully add 100–500 mg of bone powder to the extraction buffer, ensuring that it moistens all the powder. 3. Place the sample on an end-over-end rotator in a cold room for 24 h.

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4. Centrifuge the sample at 10,000 × g for 5 min in a microcentrifuge to pellet the bone powder. 5. Aspirate the supernatant (which contains the DNA) and transfer to a fresh eppendorf tube. 6. Add 50 μl fumed silica to the supernatant. Place on an endover-end rotator in a cold room for 1 h. 7. Pellet the silica (which contains the adsorbed DNA), by centrifugation at 6,000 × g for 5 min and discard the supernatant (see Note 10). 8. Add 1 ml of silica wash buffer to the pellet and invert a couple of times to resuspend the silica. Centrifuge at 6,000 × g for 5 min and discard the supernatant. 9. Resuspend the pellet in 95% ethanol to remove salt contamination and mix by inverting 2–3 times (see Note 11). 10. Centrifuge at 6,000 × g for 5 min, aspirate the supernatant and discard. 11. Repeat steps 9 and 10 once. 12. Elute the DNA from the silica by resuspending the pellet in up to 100 μl of either 10 mM Tris–HCl (pH 8.0) or distilled deionized water (pH >7.5) (see Note 12). 13. Incubate the sample at 56°C for 10 min. 14. Centrifuge at 6,000 × g for 5 min to pellet the silica and aspirate the supernatant (which contains the DNA) into a fresh tube and store frozen at −70°C or below. 3.4. RNA Extraction from Fresh Bone

This method uses TRIzol® reagent to isolate RNA from fresh bone. The method described focuses on the isolation of RNA from bone, but TRIzol can be used to isolate DNA, RNA and protein from a single bone sample. The procedure should be performed under RNAse free conditions (see Note 13). 1. Collect the bone sample in a sterile container containing PBS and transport to the laboratory within 30 min (see Note 14). 2. Place a ~200 mg sample of bone tissue (see Note 15) into a 35-mm sterile petri dish using sterile forceps. 3. Add 1 ml TRIzol® solution and chop the tissue up into a coarse slurry using bone cutters or scissors, followed by a sterile scalpel until the tissue is homogenised (see Note 16). 4. Carefully transfer the homogenate into a 2-ml screw-cap eppendorf tube and place on ice for 5 min. 5. Centrifuge samples for 12,000 × g for 10 min at 4°C and transfer the supernatant to a clean 2-ml screw-cap eppendorf tube (see Note 17).

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6. Pass lysate slowly through a 19–20 G needle 4–6 times to shear genomic DNA and reduce viscosity of the sample. 7. Add 0.1 volume of chloroform and shake vigorously for 15 s. 8. Centrifuge at 10,000 × g for 15 min at 4°C. 9. Carefully aspirate the supernatant (taking note of the volume) into a fresh 1.5-ml conical bottom screw-top eppendorf, taking care not to disturb the interface (see Note 18). 10. Add 1 volume of isopropanol to the supernatant and mix gently by inverting the tube 5–6 times. 11. Incubate the sample at room temperature for 10 min to precipitate the RNA (see Note 19). 12. Centrifuge at 10,000 g for 10 min at 4°C to pellet the RNA (see Note 20). Aspirate the supernatant and discard. 13. Wash the pellet with 1 ml ice-cold 75% ethanol and centrifuge at 10,000 × g for 5 min at 4°C. Aspirate the supernatant and discard. Briefly centrifuge the tubes again for about 15 s and aspirate the remaining traces of ethanol from the RNA pellet. 14. Allow the pellet to air-dry for 3–5 min. Do not over-dry the pellet. 15. Dissolve the RNA pellet in 10–30 μl of water (see Note 21). 16. Quantitate and assess purity of the eluted RNA by spectrophotometry (Subheading 3.8) and integrity by gel electrophoresis (see Note 22). 17. Store the RNA frozen in aliquots of 5–10 μl at −70°C or below until further use. 3.5. Extraction of RNA from Frozen Bone

If the sample has been stored at −70°C it can be ground to a fine powder prior to the extraction protocol, either in a cryogenic mill (Glen Creston, Middlesex, UK) or by mortar and pestle. 1. Pulverise the sample to a fine powder using a cryogenic mill or mortar and pestle (see Note 23). 2. Transfer the powder into a clean 2-ml screw-cap eppendorf tube, add 1 ml of TRIzol® (see Note 24), mix well by shaking the tube vigorously, and place on ice. 3. Proceed through steps 5–17 of the RNA extraction procedure described in Subheading 3.4.

3.6. Extraction of RNA from Adherent Cultured Cells

1. Aspirate the medium from the cell culture and discard. 2. Wash the cells with sterile PBS and discard. 3. Add 1 ml TRIzol® per 75 cm3 flask of cells and allow it to spread over the cell layer. 4. Detach the cell layer using a cell scraper and transfer the resulting slurry into a clean 2-ml screw-cap eppendorf tube. 5. Proceed through steps 6–17 of the RNA extraction procedure described in Subheading 3.4.

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1. Pellet the cells by centrifugation at 400 g for 10 min. 2. Aspirate the culture medium and discard. 3. Suspend the cell pellet in 1 ml TRIzol®. 4. Proceed through steps 6–17 of the RNA extraction procedure described in Subheading 3.4.

3.8. Quantification of Nucleic Acids by Spectrophotometry

a

DNA and RNA can be quantified by spectrophotometry, which is also useful for determining if any protein or solvent contamination is present in the prepared samples. However, spectrophotometry cannot assess whether or not the nucleic acid is degraded (see Note 22). Here, we present a method that should be applicable to most spectrophotometers, but specialized spectrophotometers such as the NanoDrop (Thermo Scientific), permit rapid quantification of undiluted RNA and DNA samples using a very low sample volumes (~1 μl). The NanoDrop also provides a spectral scan that can be used to detect protein or solvent contamination. Example spectrophotometer results from samples obtained using the methods described above are presented in Fig. 1, which also highlights the importance of checking not only yield and purity, but also RNA integrity (see Note 22 and Subheading 3.9).

Agilent Bioanalyser traces Sample 1

b

Sample 2

Spectrophotometer results Sample

260/280 ratio

260/230 ratio

Total RNA conc (ng/µl)

1

1.99

2.02

255.0

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2.06

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247.8

Fig. 1. Agilent Bioanalyser and spectrophotometer readings for total RNA extracted from bone. RNA was extracted as per the protocol in Subheading 3.5 from a 10 mm (H) × 5 mm (D) core of human trabecular bone, including the optional column-based clean-up and DNase I treatment discussed in Note 22. Aliquots of the isolated RNA were assayed by (a) Agilent Bioanalyser (see Note 23 ) and (b) Nanodrop spectrophotometer. Note the similar lack of protein or solvent contamination in both samples, as indicated by the 260/280 and 260/230 ratios respectively, and the similar yield in both samples despite the degraded profile of Sample 2 on the Agilent Bioanalyser trace.

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1. Dilute RNA 1/100 or DNA 1/200 in water. 2. Using a quartz cuvette, blank the spectrophotometer using water. 3. Read the absorbance for each sample at 230, 260, and 280 nm. 4. Calculate the amount of nucleic acid present in each sample by multiplying the A260 value by 40 ng/μl for RNA or 50 ng/μl for dsDNA, then by the dilution factor used, e.g., ×100 for RNA, ×200 for dsDNA (see Note 25). 5. Calculate the A260/A280 and A260/A230 ratios to determine the levels of protein or solvent contamination respectively (see Note 26). 3.9. Evaluation of RNA Integrity by Agarose Gel Electrophoresis

It is important to evaluate the integrity of isolated RNA to validate the extraction has been successful in not only isolating an abundance of solvent- and protein-free RNA (as measured by spectrophotometry), but also in maintaining the integrity of the RNA. A quick way of achieving this is by analysing the degradation of the 18 s and 28 s ribosomal RNA on an agarose gel. Figure 1 illustrates the importance of checking RNA integrity to ensure, whilst Fig. 2 shows what undegraded high-quality RNA looks like when analysed on an agarose gel, as described below.

Fig. 2. Agarose gel electrophoresis of total RNA extracted from cultured mouse calvarial osteoblasts. RNA was extracted from one, two or three wells of a six-well tissue culture plate into 1 ml of TRIzol® reagent as per Subheading 3.6. After quantification, 1 μg of total RNA was loaded onto an agarose gel as per Subheading 3.9. The integrity of RNA can be assessed by the absence of “smearing” of the 28S and 18S ribosomal RNA bands, assuming that mRNA will be degraded if the rRNA is affected.

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1. Prepare a 1% agarose gel in TBE buffer plus 0.5 μg/ml ethidium bromide (see Note 4). 2. Mix 1–2 μg of RNA with 6× Orange G loading buffer and heat to 65°C for 5 min. 3. Mix RNA ladder (e.g., RiboRulerTM High Range RNA Ladder, Fermentas) with 6× Orange G loading buffer. 4. Load and run gel in TBE at 10 V per cm between electrodes in electrophoresis tank. 5. Image gel and visualise bands on a UV transilluminator with a digital camera.

4. Notes 1. All chemicals should be of molecular biology grade. The solutions can be stored at 4°C for up to 3 months. 2. Working solutions can be stored for up to 1 week at 4°C. Stock solutions should be stored frozen at –70°C. 3. Hoechst 33258 is a DNA specific dye that can be used to quantitate DNA. Quantification is achieved by setting up a standard curve of DNA at known concentrations and analysing the test samples in a fluorimeter at an excitation wavelength of 350– 363 nm and a detection wavelength of 410–480 nm. 4. Alternatives to the use of Ethidium Bromide in laboratories are available, such as SYBR® safe DNA gel stain (Invitrogen) and SYBR® green II which binds RNA. These dyes have reduced mutagenicity and also lower waste management costs than Ethidium Bromide. While these products can be viewed using UV illumination better results can be obtained with a Safe Imager™ Blue-Light Transilluminator system (Invitrogen). 5. There is some debate whether DEPC-treated water is required as it can inhibit subsequent enzyme-mediated reactions. Sterile distilled water filtered through 0.2 μm filter would suffice if this is a concern. 6. If the DNA extraction is not initiated immediately, freeze the sample at −20°C or below for later use. 7. Vortexing causes long strands of DNA to shear. 8. Orientate the eppendorf tube so that you can identify where the DNA pellet lies. A pellet should be visible the bottom of the tube. 9. Dilute the sample 1/200 for DNA in distilled water and read the absorbance at 260 nm and 280 nm using a quartz cuvette in a UV spectrophotometer (Subheading 3.8). Hoechst 33258 dye can also be used to quantify DNA.

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10. You may wish to keep the supernatant in a fresh tube until you are confident that the extraction has been successful. 11. Do not simply add the ethanol and immediately decant it off again; the pellet needs to be well mixed so that the ethanol can penetrate the sample and dissolve the salt. 12. The pH must be above 7.5 to elute the DNA from the silica. 13. Gloves should be worn at all times when handling samples and all solutions should be prepared to ensure that they are free of RNAses. This involves making up the solutions for RNA use with DEPC treated water using glassware, stirrers and spatulas which have been oven baked for 2 h to inactivate RNAses. Further details can be found in Sambrook’s text (6). 14. A major challenge in extracting RNA from any tissue is to isolate the nucleic acid in an intact state before it is degraded. If it is impractical to begin the extraction within 30 min, the bone should be snap-frozen in liquid nitrogen immediately and stored at −70°C or below until use. Reagents such as RNAlater® (Invitrogen) can be useful for preserving soft tissues or cell pellets for weeks, but may not permeate bone samples fully enough to inactivate all RNases. In our experience, it is best to homogenise the bone as soon as possible into the extraction buffer and then snap-freeze the homogenate for storage as above, rather than using RNA preservation buffers. 15. Too much starting material can result in a high degree of DNA and protein contamination, so you may need to scale up TRIzol® volumes if you exceed 200 mg sample weight. 16. Samples can also be homogenized using a Precellys homogeniser and MK28 tubes with large diameter metal bearings. Homogenisation at 6,500 for 2 × 45 s cycles yielded RNA of high integrity from whole bones isolated from mouse. A mortar and pestle can also be used for fresh bone, as per Subheading 3.5, by snap-freezing the fresh sample in liquid nitrogen and grinding it to a fine powder in the mortar. 17. Samples with a high fat content, such as trabecular bone samples obtained from osteoporotic femoral heads, may yield a layer of fat that floats on top of the clear phase containing the nucleic acid. This should be discarded. 18. The upper (clear) aqueous layer contains the RNA and is separated from lower (pink/purple) phase containing debris and DNA by a white layer containing protein and lipids. Do not attempt to completely remove the aqueous layer, to minimise the risk of protein or DNA contamination. 19. The samples can be stored overnight at −20°C at this point. 20. It is a good idea to mark the side of the eppendorf so you can identify where the RNA pellet lies. Small pellets can be “bulked

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up” for easier detection by adding 10–100 μg/ml glycogen to the isopropanol precipitation stage. 21. For some subsequent applications it may be necessary to perform further purification or DNA digest. If so, the RNA can be further purified on a Qiagen RNeasy Minikit with an on-column DNase digest using a Qiagen RNase Free DNase Set (Qiagen, West Sussex, UK). These additional steps are highly recommended for preparation of RNA suitable for microarray or quantitative PCR analysis. 22. Spectrophotometry cannot measure the integrity of RNA, which is an important consideration for further downstream applications; quantity and purity should be reinforced by a measure of integrity, such as gel electrophoresis (see Subheading 3.9) or by Agilent Bioanalyser. The Bioanalyser is a microfluidics, laser-detection based system that can sensitively assay fragment size and calculate the RNA Integrity Number (RIN) for quality control of RNA samples destined for quantitative techniques such as qPCR or microarrays (see Fig. 1). 23. A small amount of liquid nitrogen can be poured into the pestle to maintain low temperature. 24. Too much starting material can result in a high degree of DNA and protein contamination, so you may need to scale up TRIzol® volumes if you exceed 200 mg sample weight. 25. Nucleic acids have an absorbance maximum at 260 nM, so spectrophotometry can not determine if RNA samples are free of DNA contamination. 26. If the separation is free of protein contamination, the optical density reading at A260/A280 should be 1.8 or greater, although this is based on RNA solubilized in TE buffer (10 mM Tris–HCl pH 8.0, 1 mM EDTA). Ratios generated in water will tend to be lower on most spectrometers. The A260/A230 ratio should be >2.0 and gives an indication of solvent contamination; Solvent contamination can inhibit subsequent enzyme-based reactions. References 1. Chomczynski, P., and Sacchi, N. (1987) Single step method of RNA isolation by acid guanidinium thiocyanate-phenol-chloroform extraction. Anal. Biochem. 162, 156–159. 2. Blin, N., and Stafford, D. W. (1976) A general method for isolation of high molecular weight DNA from eukaryotes. Nucleic Acid Res. 3, 2303–2308. 3. Kupiec, J. J., Giron, M. L., Vilette, D., Jeltsch, J. M., and Emanoil-Ravier, R. (1987) Isolation of high molecular weight DNA from eukaryotic cells by formamide treatment and dialysis. Anal. Biochem. 164, 53–59.

4. Bowtell, D. D. L. (1987) Rapid isolation of eukaryotic DNA. Anal. Biochem. 162, 463–465. 5. Vogelstein, B., and Gillespie, D. (1979) Preparative and analytical purification of DNA from agarose. Proc. Natl. Acad. Sci. USA. 76, 615–619. 6. Sambrook, J., Fritsch, E.F., and Maniatis, T. (2000) Molecular Cloning. A laboratory Manual. 3rd Edition, Cold Spring Harbour Laboratory Press.

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Chapter 18 Analysis of Gene Expression in Bone by Quantitative RT/PCR Alun Hughes Abstract This chapter describes methods for quantitation of gene expression in bone cells and bone tissues using the technique of quantitative reverse transcription-polymerase chain reaction. Key words: PCR, Quantitative PCR, Gene expression, RNA, Bone disease, Osteoporosis

1. Introduction The polymerase chain reaction (PCR) is a technique that can amplify small amounts of target DNA in an exponential manner by using sequence-specific oligonucleotide primers (1). In this chapter, I describe the methodology for performing quantitative PCR, linked to a reverse-transcription step, thereby allowing one to analyse levels of gene expression in cells or tissues of interest. 1.1. Assay Choice for Quantitative PCR

Like conventional PCR, qPCR relies upon the specificity of two primers for amplification. A surrogate for actual copy number is generated by use of either a fluorophore that intercalates with the double-stranded amplicon generated during extension (e.g. SYBR Green; see Fig. 1a), or by using the principles of FRET (Förster resonance energy transfer) to generate labelled probes (sequencespecific, modified primers) that emit the expected wavelength either when they are bound in the correct position (hybridisation probes) or when they are hydrolysed by the exonuclease activity of the DNA polymerase (hydrolysis probes; see Fig. 1b). The addition of a third modified primer with hybridisation or hydrolysis probes provides extra specificity, although the sensitivity of intercalating dyes is higher due to the binding of multiple fluorophore molecules.

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_18, © Springer Science+Business Media, LLC 2012

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Sample extraction

RNA Extraction Exogenous Control

(Sections 1.3 & 3.4)

Sample preparation

cDNA Synthesis

(Sections 2.2 & 3.1)

Assay design

Analysis method

Hydrolysis Probes

(Sections 1.1 & 3.2)

SYBR Green

(Sections 1.1 & 3.3)

Absolute Quantification

Relative Quantification

(Section 1.2)

(Section 1.2)

Standard Curve (Section 1.2)

Efficiency Corrected

(Section 1.2)

Purified PCR product

Plasmid DNA

cDNA Template

(Section 3.5)

(Section 3.5)

(Section 3.6)

Delta-delta Ct (Sections 1.2 & 3.9)

Fig. 1. Quantitative assay design considerations flow chart. For discussion on the most appropriate pathway for your application, refer to Subheadings 1.1–1.3.

The choice of assay system for qPCR depends to an extent on the funding available with which to conduct your experiment. Assays with SYBR green are cheaper than those that use fluorescently labelled probes and can be useful where a single target is being assayed repeatedly in a series of experiments. Although fluorescently labelled probes are more expensive to synthesise they offer greater specificity than SYBR green and can be more cost effective for some targets. Another advantage of hydrolysis probes is that they can be multiplexed, and allow one to analyse expression of several genes in a single well using different fluorophores. Several companies provide “off-the-shelf” probe kits (e.g. TaqMan® assays from Applied Biosystems) or “do-it-yourself” hydrolysis probe style assays (e.g. the Universal ProbeLibrary system from Roche), allowing researchers to quickly and conveniently measure gene expression.

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1.2. Methods for Quantification of Transcripts

Quantitative PCR can be performed in terms of “absolute” or “relative” quantification. The majority of applications require just relative quantification, where the investigator wants to determine if there has been a change in expression of a specific target gene, relative to an internal control or “housekeeping” gene. Absolute quantification, involves measuring the exact number of transcripts of the gene of interest in relation a standard curve constructed from a reversed transcribed “decoy” mRNA, a plasmid, a purified PCR product or, less commonly, a synthetic oligonucleotide sequence (see Note 1). Relative quantification can be calculated at a basic level using the delta-delta Ct method (2), which compares two samples based on expression of a gene of interest and a reference gene, returning a fold increase or decrease value that describes their relationship. While quick and convenient, this method makes the assumption that all qPCR assays work with perfect efficiency. Modifications to the delta-delta Ct calculation have been made by to include values for the efficiencies of both the reference and gene of interest, allowing for more accurate values to be calculated (3).

1.3. Relating Expression of Target Genes to an Internal Control

Levels of expression for the gene(s) of interest can also be related to those of an endogenous reference (or “housekeeping”) RNA. The reference RNA has to be chosen carefully since expression of many housekeeping genes can vary substantially as the result of changes in cell proliferation and differentiation (4, 5). To compensate for this, multiple reference genes can be used and an average taken which is then used as a control with which to relate levels of expression of the gene(s) of interest (6). Alternatively, an exogenous target can be spiked in to samples prior to reverse transcription and used as a reference (7). The accuracy of quantification using exogenous controls is dependent upon both the accurate quantification of RNA transcript and the use of identical starting amounts or RNA for cDNA synthesis between samples. Absolute quantification is commonly normalised to the input amount of RNA prior to cDNA synthesis, or by the amount of cDNA used per reaction, resulting in data of the form copies/μg RNA or cDNA. However, this relies on accurate quantification of the RNA or cDNA and assumes identical integrity or impurities across all samples to be compared. As a result this strategy is often unsatisfactory, so it is common to use additional forms of normalisation to help improve confidence in absolute quantification results, such as the endogenous or exogenous controls used for correcting relative quantification experiments described above.

1.4. Preparation of Input RNA

The quality and purity of the starting RNA template is of critical importance to qPCR. A degraded sample can severely impair the quality of the results obtained and decrease the confidence associated in observations, while protein or solvent contamination can negatively affect the enzymatic steps involved in procuring the final

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a

SYBR Green

b

Hydrolysis probes

Q

5’

3’

5’

5’

3’

5’

3’

R

Annealing

R

3’

Q

3’

5’

3’

5’

5’

3’

5’

3’

Extension

R

Q

3’

5’

3’

5’

5’

3’

5’

3’

Fig. 2. Commonly used qPCR assays for gene quantification. Primers anneal to each strand of the template DNA, allowing binding of the DNA polymerase (indicated by the open arrow ) and subsequent elongation and synthesis of two new strands in the 5¢ to 3¢ direction. During elongation, SYBR Green binds to the minor groove of double stranded where it emits fluorescence of the expected wavelength when excited, see panel (a). Hydrolysis probes (b) require the use of a specialised primer that is labelled at the 5¢ end with a reporter (R) dye in proximity to a 3¢ quencher (Q) that inhibits the production of the expected fluorescence. The exonuclease activity of the DNA polymerases used in hydrolysis probe reaction mixes cleaves the probe during elongation, liberating the reporter from the quencher and allowing fluorescence to be detected.

data and lead to artefacts (8). The utmost care and attention must therefore be made to standardise sample preparation to generate robust and reliable data. The protocols within this chapter assumes pure, high-integrity RNA has been generated. The flow chart in Fig. 2 indicates which sections within this chapter will be appropriate for your particular application.

2. Materials 2.1. Reverse Transcription

1. High-quality RNA. 2. Diethylpyrocarbonate (DEPC)-treated water: Incubate water containing 0.1% v/v DEPC water at 37°C for 1 h and then autoclave to inactivate DEPC (see Note 2). 3. RNase-free PCR tubes and filter pipette tips.

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4. 2 μg/ml Random oligonucleotide hexamer primers. 5. 10 mM Deoxyribonucleotides in water (dNTP). 6. Reverse transcriptase (e.g. SuperScriptTM, Invitrogen, Paisley, Scotland). 7. A thermocycler compatible with use of FAM dyes, such as the Roche LightCycler®480 (see Note 3). 8. Custom oligonucleotide primers and labelled hydrolysis probes. 9. Mastermix compatible with hydrolysis probe (e.g. LightCycler®480 Probes Master Mix) or with SYBR Green (e.g. LightCycler®480 SYBR Green I Master Mix).

3. Methods 3.1. Reverse Transcription

A reverse transcription step is required to convert the RNA into complementary DNA (cDNA) prior to PCR. Some “one-step” kits exist for performing the RT reaction in the same tube as the qPCR reaction, but the method presented here is for a separate RT reaction that yields sufficient cDNA for the measurement of multiple transcripts. 1. Aliquot 2 μg of RNA into duplicate RNase-free PCR tubes and adjust total volume to 11 μl with DEPC-treated water. Store on ice. 2. Add 1 μl of 2 μg/ml random hexamer primers to each sample, flick tube to mix and pulse on a centrifuge to collect the reaction mix at the bottom of the tube (see Note 4). 3. Heat to 70°C for 10 min on a thermocycler and then immediately cool on ice for 2–3 min. 4. Add 4 μl of 5× Reaction Buffer, 2 μl of 0.1 M DTT and 1 μl of 10 mM dNTPs mix to each tube. Add 200 U of SuperScriptII M-MLV reverse transcriptase to half of the tubes or an equivalent volume of DEPC-treated water to the other to generate no-RT controls (see Note 5). 5. Mix tubes by flicking and collect the reaction mix at the bottom of the tube by pulsing briefly on a centrifuge. 6. Incubate the reaction mix at room temperature for 10 min. 7. Heat the tube to 42°C for 50 min in a thermal cycler then at 95°C for 5 min before cooling on ice for 2–3 min. 8. Adjust reaction volume to 100 μl with DEPC-treated water and store at −20°C or short-term at 4°C (see Note 6).

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3.2. Designing Primers and Probes for Hydrolysis qPCR Assays

Many companies provide off-the-shelf hydrolysis probe assay probes and primers, but it can be more economical to design your own especially if it is an assay you intend to use frequently. This method uses the online version of the open source Primer3 program by Rozen and Skaletsky (9). 1. Download mRNA sequence from Ensembl database (http:// www.ensembl.org/) or GenBank database (http://www.ncbi. nlm.nih.gov/nuccore/), making sure to note the position of the exon–exon boundaries. 2. Paste sequence in to Primer3 (http://frodo.wi.mit.edu/ primer3/). 3. Chose a “Mispriming Library (repeat library)” if a suitable species library is available. 4. Mark the two bases at the interface of adjacent exons using the “[”and “]” characters. 5. Check the tick box to design an internal oligo. 6. Set the product size range to 70–150. 7. Keep “General Primer Picking Conditions” to pick primers with an optimal Tm of 60°C. 8. Change “Hyb Oligo (Internal Oligo) General Conditions” to return a hybridisation oligo with a minimum, optimum, and maximum Tm of 65, 68, and 70°C, respectively. 9. Select a “Hyb Oligo Mishyb Library” if a suitable species library is available. 10. Click “Pick primers” and select primer and probe combinations that give short product sizes and have at least one base gap between the left primer and the hybridisation oligo. Reject assays which have a 5¢ “G” on the hybridisation oligo. 11. BLAST search primers and hybridisation oligo to insure specificity (http://blast.ncbi.nlm.nih.gov/Blast.cgi). 12. If no suitable assays can be found, repeat the process selecting a different exon–exon boundary using the “[”and “]” characters (see Note 7). 13. Order primers from a reputable supplier with HPLC purification. Order the hybridisation probe with a 5¢ FAM and 3¢ TAMRA, Dabcyl or Black Hole Quencher (e.g. BlackBerry®). 14. Proceed to perform PCR as described in Subheading 3.7.

3.3. Design of Primers for SYBR Green qPCR Assays

Quantitative PCR assays using SYBR green are more sensitive than those that employ hydrolysis probes, and can work out cheaper to perform. However, since any double-stranded product will contribute to the fluorescent signal generated, care must be taken to design very specific primers that don’t generate non-specific products or primer-dimers. The design strategy outlined below uses the

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National Center for Biotechnology Information’s Primer-BLAST tool using the Primer3 primer design programme (9): 1. Obtain the RefSeq ID for your gene via the Entrez Gene web site (http://www.ncbi.nlm.nih.gov/gene/) (see Note 8). 2. Insert the RefSeqID in to the “PCR Template” box on NCBI Primer-BLAST tool (http://www.ncbi.nlm.nih.gov/tools/ primer-blast/). 3. Set the product size range to 80–200 in the “Primer Parameters” section (see Note 9). 4. Keep “Primer melting temperatures” to pick primers with an optimal Tm of 60°C. 5. Select “Primers must span an exon–exon junction” from the “Exon junction span” drop-down menu. 6. Enter organism species in to the “Specificity check” box to allow automatic BLAST searching of your primers for specificity. 7. Click “Get Primers”. 8. Order primers from a reputable supplier with HPLC purification (see Note 10). 9. Proceed to perform PCR as described in Subheading 3.8. 3.4. Preparation of an Exogenous Control

Exogenous controls are completely foreign transcripts (e.g. from a different kingdom or synthetically generated) spiked into all samples to produce a target that can be used as an alternative to native, endogenous reference genes for normalisation. Purified sample RNA is accurately quantified and mixed with a known amount of exogenous controls (e.g. 5 × 104 copies of exogenous control per μg of sample RNA), creating a “constant” that can be used between samples for normalisation or to measure inhibition of cDNA synthesis caused by solvent or protein contamination of RNA samples. Exogenous spikes are useful when commonly used endogenous genes such as beta-actin might give erroneous results due to the effects of drug treatment, cell differentiation or tissue variation. Exogenous controls can be prepared in-house according to established techniques (10) or obtained commercially (e.g. Alien Reference RNA QRT-PCR Detection Kit by Agilent Technologies). 1. Add 105 copies/reaction of the exogenous RNA transcript to the cDNA synthesis components described in Subheading 3.1, step 1. Reduce the volume of water accordingly. 2. Design an assay that will amplify your exogenous transcript using the principles outlined in Subheadings 3.2 or 3.3. 3. Following qPCR analysis, normalise all your candidate reference genes to the exogenous control as per Subheading 3.9. Suitable endogenous controls should maintain a constant ratio across all samples.

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3.5. Preparation of an External Reference Curve for Absolute Quantification

Various methods can be used to generate a standard curve for qPCR including the generation of recombinant RNA (recRNA) molecules (see Note 11), but here I describe the method for generating a standard curve from plasmid DNA since this is considered to be a more stable and reliable method for generating standard curves (11) (see Note 12). 1. Purify the plasmid that contains the target of interest using a commercially available cDNA clean-up kits (e.g. QIAquick PCR Purification Kit, Qiagen). 2. Quantify the abundance of purified PCR product or linearised plasmid by spectrophotometry or by PicoGreen® (see Note 13). If using spectrophotometry, please note that an absorbance value of 1 at 260 nm is equal to 50 μg/ml of dsDNA. Whatever method is being used, take repeated measurements at two or three dilutions to increase accuracy. 3. Calculate the number of copies/μl of sample by the following formula: Copies/ml=

6.022 ´ 10

23

(molecules/mol) ´ DNA concentration (g/ml)

Length of plasmid or product (basepairs) ´ 660 daltons

.

4. Prepare serial dilutions of the plasmid to give a suitable range of copies for use as a standard curve in Subheadings 3.7 or 3.8 (see Note 14). 3.6. Preparation of cDNA Reference Curve for Relative Quantification

Relative quantification, unlike absolute quantification, does not require the use of standard curves generated from plasmid or recRNA sources to calculate copy number. However, to obtain the most accurate results using relative quantification it is essential to calculate the efficiency of each gene-specific assay used. The efficiency of each assay can be obtained from the slope of a series of serially diluted samples, creating a “standard curve” but using arbitrary values to define the relationship between each dilution (see Note 15). Using this method, an individual cDNA sample or pool of cDNA can be used to correct relative quantification data from errors associated with assuming a default efficiency value of 2. 1. Select a cDNA sample that you know or suspect will express your gene(s) of interest, or create a pool of cDNA to use as a reference curve. 2. Serially dilute cDNA across a range of at least four dilutions suitable to the suspected abundance of your transcript (e.g. a highly expressed gene will be detectable across a dilution range of neat, 1/10, 1/100, 1/1,000, etc. dilutions, or neat, 1/3, 1/9, 1/27, etc. can be used for genes with lower abundance).

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3.7. Performing Quantitative RT-PCR Using Hydrolysis Probes

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1. Pipette 3 μl of each cDNA sample (RT and no-RT controls) in triplicate technical reactions per gene (see Note 16) to be measured on to a white-walled plastic plate suitable for your qPCR machine (see Note 17). 2. Pipette 3 μl of each standard into the plate in triplicate, plus triplicate blank wells each containing 3 μl of water. 3. Prepare sufficient master mix for each assay plus 10% to account for losses on tips, allowing for a total reaction volume of 20 μl per well (17 μl master mix + 3 μl standard, sample or blank). An example master mix per reaction is displayed in Table 1. 4. Add 17 μl of master mix to the relevant standard, sample, and blank wells on the plate 5. Run on a qPCR machine for 40 cycles, using thermal cycling conditions of: 95°C, 10 min; 40 cycles of (95°C, 10 s; 60°C, 30 s) (see Note 18). 6. Analyse the data as outlined in Subheading 3.9.

3.8. Performing Quantitative RT-PCR Using SYBR Green Assays

These assays can be adversely affected by the formation of primerdimers or by mispriming events. Because of this SYBR Green assays should always be evaluated using a melting curve profile (i.e. a post-amplification monitoring of the fluorescence as the temperature is increased incrementally from 65 to 95°C) to ensure only a single product is generated during amplification. If multiple peaks are present in the melting curve analysis, then the specificity of the assay can be improved by increasing the annealing temperature or, especially in the case of primer-dimer, decreasing the primer concentrations.

Table 1 Example reaction components for SYBR Green and hydrolysis probe qPCR

LightCycler®480 SYBR Green l Master (2×) ®

LightCycler 480 Probes Master (2×)

SYBR Green Volume per well (ml)

Hydrolysis probes Volume per well (ml)

10

–

–

10

Forward primer (10 μM)

1

1

Reverse primer (10 μM)

1

1

Hydrolysis probe (2 μM) PCR grade water

– 5

1 4

N.B.: These are example reaction mixes assuming a 20 μl total volume and are presented as an illustration of the general primer/probe concentrations utilised in SYBR Green and hydrolysis probe assays. Other assay mixes are available and can be optimised to work with most qPCR machines, but please refer to your specific reagent supplier’s guidelines. When creating “master mixes”, remember to include 10% extra volume to account for loss of mixture in pipette tips

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Table 2 Example thermal cycling conditions for SYBR Green qPCR Cycles

Step

Temperature (°C)

Hold time

1×

Enzyme activation

95

5 min

40×

Denaturation Annealing Extension

95 Avg. primer Tm minus 2–5°C 72

10 s 15 s (amplicon length [b.p.]/25) s

1×

Denaturation Strand annealing Melting curve

95 65 65–97

10 s 1 min –

These conditions work well for SYBR Green assays using Roche’s LightCycler®480 SYBR Green I Master enzyme mix on a Roche LightCycler®480 qPCR machine and are presented for illustrative purposes. Please refer to your reagent supplier’s guidelines for appropriate conditions for your enzyme and system as recommendations can vary

1. Pipette 3 μl of each cDNA sample (RT and no-RT controls) in triplicate per gene to be measured on to a white-walled plastic plate suitable for your qPCR machine. 2. Pipette 3 μl of each standard into the plate in triplicate, plus triplicate blank wells each containing 3 μl of water. 3. Prepare sufficient master mix for each assay plus 10% to account for losses on tips, allowing for a total reaction volume of 20 μl per well (17 μl master mix + 3 μl standard, sample, or blank). An example master mix per reaction is displayed in Table 1. 4. Add 17 μl of master mix to the relevant standard, sample, and blank wells on the plate. 5. Run on a qPCR machine for 40 cycles, using the thermal cycling conditions in Table 2 (see Note 18). 6. Perform melting curve analysis to validate amplification is specific and not due to primer-dimer or mispriming events. Plot the first derivative (dF/dT) of the melting curve data generated against temperature. Specific primers will generate a single, strong peak, while non-specific primers will generate multiple peaks. Primer dimerisation will cause a “hump” evident at lower temperatures (see Note 19). 7. Analyse data as outlined in Subheading 3.9. 3.9. Analysis of q-RT-PCR Data

The analysis method below can be used with arbitrary units or with quantified standards to generate normalised data based on the use of either an endogenous or exogenous reference gene. Alternatively, the data generated from a reference curve of cDNA template can be used to calculate the efficiency of each assay as per step 1 and 2, allowing changes in gene expression to be calculated using Pfaffl’s

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modification of the delta-delta Ct method (3), or inserted in to the freely available REST© spreadsheet (available at www.gene-quantification.de/download.html) (12). In the absence of a standard curve, changes in gene expression can be roughly calculated out via the delta-delta Ct method, as per Livak and Schmittgen (2). Karlen et al. (13) provide an in-depth discussion of the number of independent biological repeats required to provide confidence in the changes observed (see Note 16), while a discussion of several appropriate ways to perform statistical analysis of qPCR data can be found in Yuan et al. (14). 1. Calculate the cycle threshold (Ct) values for each amplification curve (see Note 20). 2. Analyse the standard curve for each gene-specific assay. Calculate the Ct value of each standard curve point generated from cDNA (Subheading 3.6), plasmid or PCR product (Subheading 3.5) and plot against log copy number or arbitrary units. The efficiency of each assay can be determined by the slope of the standard curve using the formula E = 10−1/slope. 3. Calculate the mean and standard deviation (SD) for each sample based on the triplicate reactions performed. 4. Calculate the coefficient of variation (CV) for each sample: CV = SD/mean. 5. Normalise your “gene of interest” (GOI) data to your reference gene or exogenous control (Ref) by dividing the value for each sample’s GOI by the value for the corresponding Ref (see Note 21). 6. Calculate the quotient CV value: CVQ = (CVGOI2 + CVRef2)1/2. 7. Calculate the SD of the quotient: SDQ = CVQ⋅X (where X = the ratio of your GOI/Ref ). 8. Results can be displayed as GOI/Ref ± SDQ or calculated as fold change from a calibrator control as required (see Note 22).

4. Notes 1. It is technically difficult to accurately measure transcript numbers by qPCR for various reasons. First, reverse transcription is not 100% efficient and the ease with which different mRNA targets are reverse transcribed can vary greatly (15). Although PCR products, plasmid and synthesised oligonucleotides can be used to estimate target abundance, they are likely to provide an overestimate, since they do not have to undergo the RT stage.

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2. There is some debate whether DEPC-treated water is required as it can inhibit subsequent enzyme-mediated reactions. Sterile distilled water filtered through 0.2-μm filter would suffice if this is a concern. 3. Fluorescein (FAM) has a similar excitation and emission maxima as SYBR Green, which all qPCR machines currently on the market are capable of detecting. 4. Random primers provide good 3¢/5¢ coverage of most mRNA transcripts but will initiate reverse transcription of all RNA to cDNA including ribosomal RNA. Oligo-dT primers will only initiate reverse transcription of mRNA, but may not provide complete 3¢/5¢ coverage. RNA that is partially degraded is better analysed by using random primers. Gene-specific primers can also be used to ensure reverse transcription of troublesome transcripts. Nolan et al. (8) provide practical advice on how to evaluate the best priming strategy for reverse transcription. 5. The efficiency of reverse transcription varies between enzymes, so ensure that the same priming strategy and enzyme choice is used between any samples that are to be compared (15). 6. Contaminants from the reverse transcription process can negatively effect PCR reactions, so sometimes diluted samples give more accurate and reliable results. Always perform a dilution series of your cDNA in advance of quantification experiments to titrate out any PCR inhibitors. 7. The unfortunate situation may arise that no suitable intronspanning assays can be designed, in which case RNA samples will need to be treated with DNAse and no-RT controls must be included for every sample to exclude false-positive results due to amplification of contaminating genomic DNA. 8. RefSeq IDs follow the format of “NM_xxxxxx”. FASTA format sequence information can be used instead. 9. The optimal SYBR green assay length is 150 bp. Amplicon lengths of longer than 300 bp are not recommended, but can be used if essential. 10. Desalted primers also work well in most qPCR assays. 11. Plasmids or purified PCR products are not subject to the inefficiencies of reverse transcription; hence, give a value for the amount of transcript successfully reverse transcribed rather than the actual copy number in the original sample. One way of overcoming this disparity in measurement is to generate standard curves from recombinant RNA (recRNA) for each gene of interest, then subjecting each recRNA to the same reverse transcription methods as the RNA samples of interest. Methods for generating standard curves for absolute transcript quantification using recombinant RNA can be found in the papers by Fronhoffs et al. (16) or Bustin (17).

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12. Plasmids can be more reliably quantified than purified PCR products, and are more stable for long term storage. A quick and convenient method for cloning a purified PCR product into a pBAD-TOPO plasmid can be found in the paper by Whelan et al. (18), although any expression plasmid containing your transcript of interest is perfectly acceptable. There is no need to linearise plasmids prior to use as standard curve in a qPCR assay. Whelan et al. (18) compared intact via linearised plasmids and observed no significant difference in the resulting quantification. 13. Substantially more accurate quantification of plasmid or purified PCR product can be obtained by using PicoGreen®. Kits for quantification using this dye are available from many suppliers. Plasmid dilutions can be stored at −20°C. Diluted purified PCR products may be less stable and are best made fresh from concentrated stocks for each qPCR assay. 14. For example, a tenfold dilution series from 1 × 105 copies/μl to 10 copies/μl should be appropriate for must genes. 15. For relative quantification it is best to generate a reference curve using material as close to that you are assaying as possible: cDNA template generated using the same protocols for reverse transcription is ideal and will give the best estimation of reaction efficiency. 16. Technical replicates should not be confused for biological replicates. Four independent biological replicates will give sufficient confidence to detect changes or around twofold. This will vary by the relative “tightness” of your biological system and the size of changes in gene expression expected between treatments or conditions. 17. White-walled plates give substantially better fluorescent signals than clear wells, offering an improvement in data quality that far outweighs the inconvenience associated with effectively “blind” loading in white wells. 18. Conditions may vary depending on your qPCR machine and choice of enzyme, probes and primers. Use these conditions as a guideline, but always refer to manufacturer’s instructions to obtain the best results. 19. The effects of primer dimerisation on melting curves can be diminished by either increasing the annealing temperature used or by decreasing the concentration of primers used, but it is important to remember that both of these options could affect the efficiency of the PCR reaction. Primer dimerisation is not necessarily a problem if it only affects your blank wells. Melting curves displaying multiple peaks can be run on an agarose gel, imaged by ethidium bromide or similar, and evaluated for multiple bands. Some “humps” on the side of melting

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curves can look like two products, but may in fact be breathing in A/T-rich regions of the amplicon. It is often prudent to design new primers if the melting curve profiles are exceptionally noisy. 20. CP (crossing point) values obtained using Roche instruments are equivalent to Ct values. Ct values can be calculated by whatever method is available on your system: Roche systems use the Second Derivative Max which works well with sigmoidal amplification curves, while most other systems predominately use the setting of arbitrary threshold lines. No matter which method you use, the important thing is to be consistent. 21. Exogenous spikes (i.e. those described in Subheading 3.2) can be used as a “reference” gene to treat potential endogenous housekeeping genes as “genes of interest”. If the ratio between endogenous and exogenous is consistent across samples and treatments then the endogenous gene can be used as a housekeeping (or reference) gene. 22. If using this method to calculate relative expression using a reference curve of diluted sample cDNA, then units are arbitrary. Care should be taken not to infer differences between different genes if you have used this approach: The only really appropriate comparisons that can be made are of the differences of expression within individual genes. References 1. Saiki, R., Scharf, S., Faloona, F., Mullis, K., Hom, G., and Erlich, H. (1985) Enzymatic amplification of β-globin genomic sequences and restriction site analysis for diagnosis of sickle cell anemia. Science 230, 1350–1354. 2. Livak, K. J., and Schmittgen, T. D. (2001) Analysis of relative gene expression using realtime quantitative PCR and the 2-deltadeltaCT method. Methods 25, 402–408. 3. Pfaffl, M. W. (2001) A new mathematical model for relative quantification in real-time RT-PCR. Nucleic Acids Res. 29, e45. 4. Thellin, O., Zorz,i W., Lakaye, B., De Borman, B., Coumans, B, Hennen, G., Grisar, T., Igout, A., and Heinen, E. (1999) Housekeeping genes as internal standards: use and limits. J. Biotechnol. 75, 291–295. 5. Dheda, K., Huggett, J. F., Bustin, S. A., Johnson, M. A., Rook, G., and Zumla, A. (2004) Validation of housekeeping genes for nomalizing RNA expression in real-time PCR. Biotechniques 37, 112–119. 6. Vandesompele, J., De Preter, K., Pattyn, F., Poppe, B., Van Roy, N., De Paepe, A., and Speleman, F. (2002) Accurate normalization of

7.

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11.

real-time quantitative RT-PCR data by geometric averaging of multiple internal control genes. Genome Biology 3, 34. Gilsbach, R., Kouta, M., Bonisch, H., and Bruss, M. (2006) Comparison of in vitro and in vivo reference genes for internal standardization of real-time PCR data. Biotechniques 40, 173–177. Nolan, T., Hands, R. E., and Bustin, S. A. (2006) Quantification of mRNA using realtime RT-PCR. Nat. Protocols 1, 1559–1582. Rozen, S., and Skaletsky, H. J. (2000) Primer3 on the WWW for general users and for biologist programmers. In: Krawetz S, Misener S (eds) Bioinformatics Methods and Protocols: Methods in Molecular Biology. Humana Press, Totowa, NJ, pp 365–386. Bower, N. I., Moser, R. J., Hill, J. R., and Lehnert, S. A. (2007) Universal reference method for real-time PCR gene expression analysis of preimplantation embryos. Biotechniques 42, 199–206. Pfaffl, M. W., Hageleit, M. (2001) Validities of mRNA quantification using recombinant RNA and recombinant DNA external calibration

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curves in real-time RT-PCR. Biotech. Letts. 23, 275–282. Pfaffl, M. W., Horgan, G. W., and Dempfle, L. (2002) Relative expression software tool (REST©) for group-wise comparison and statistical analysis of relative gene expression results in real-time PCR. Nucleic Acids Res. 30, e36. Karlen, Y., McNair, A., Perseguers, S., Mazza, C., and Mermod, N. (2007) Statistical analysis of quantitative PCR. BMC Bioinformatics 8, 131. Yuan, Y. S., Reed, A., Chen, F., and Stewart, C. N. Jr. (2006) Statistical analysis of real-time PCR data. BMC Bioinformatics 7, 85. Stahlberg, A., Hakansson, J., Xian, X., Semb, H., and Kubista, M. (2004) Properties of the

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reverse transcription reaction in mRNA quantification. Clin. Chem. 50, 509–515. 16. Fronhoffs, S., Tozke, G., Stier, S., Wernert, N., Rothe, M., Bruning, T., Koch, B., Sachinidis, A., Vetter, H., and Ko, Y. (2002) A method for the rapid construction of cRNA standard curves in quantitative real-time reverse transcription polymerase chain reaction. Mol. Cell Probes 16, 99–110. 17. Bustin, S. A. (2000) Absolute quantification of mRNA using real-time reverse transcription polymerase chain reaction assays. J. Mol. Endocrinol. 25, 169–193. 18. Whelan, J. A., Russel, N. B., and Whelan, M. A. (2003) A method for the absolute quantification of cDNA using real-time PCR. J. Immunol. Methods 278, 261–269.

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Part IV Microscopical Techniques

Chapter 19 Histomorphometry in Rodents Reinhold G. Erben and Martin Glösmann Abstract Bone histomorphometry remains an important tool to study the pathophysiology of bone disease and the cellular mechanism by which treatments work. Here, we review the methods for embedding, sectioning, staining, and analysis of bone sections in rodents. Key words: Bone, Histomorphometry, Osteoblast, Bone remodeling

1. Introduction Bone histomorphometry is an indispensable tool for assessing the mechanisms by which bone diseases occur, the mechanisms by which therapeutic agents affect the skeleton, and the skeletal safety of therapeutic agents. While high-resolution imaging techniques such as mCT can provide excellent information about bone mass and bone structure, they cannot at present provide the researcher with insights into cellular activity in bone. When applied correctly, bone histomorphometry can yield a wealth of information about bone structure, bone formation, bone resorption, bone mineralization, as well as bone modeling and remodeling activity. This chapter focuses on histomorphometric analysis of rodent cancellous and cortical bone sites. The nomenclature for histomorphometric parameters in this chapter is based on the suggestions made by the ASBMR nomenclature committee (1). One of the “axioms” in histomorphometry is that “Your histomorphometry is only as good as your histology”. Because histological quality is a prerequisite for meaningful histomorphometric analysis, this chapter starts with a description of how mouse and rat bone specimens have to be embedded and stained to achieve high quality histological “raw” material.

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_19, © Springer Science+Business Media, LLC 2012

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We use two different methylmethacrylate (MMA) embedding protocols, a conventional method, and a modified embedding method suitable for histochemistry and immunohistochemistry. Conventional MMA embedding results in almost complete destruction of enzyme activity and antigenic determinants in the embedded tissue by covalent modification of biological molecules through radicals generated during the polymerization process. Therefore, bones embedded in conventional MMA cannot be used for histochemistry or immunohistochemistry in a reliable fashion. For partial preservation of enzyme activities and tissue reactivity to antibodies, we use a modified MMA embedding method developed in our laboratory (2). Because analysis of osteoclast numbers in mice is greatly facilitated by Tartrate Resistant Acid Phosphatase (TRAcP) histochemistry, we embed all our mouse bones using the modified method which involves performing the polymerization at low temperature. By contrast, we embed all our rat bones in conventional MMA, unless histochemical or immunohistochemical analyses are important end points. The histological quality is comparable for both methods of embedding. For sectioning we use both rotary (Microm HM 355S and HM 360) and sledge microtomes (Leica Polycut S2500). For optimal section quality, it is important to use a slide press for drying the sections. Standardization of sampling sites is also very important to achieve reproducible results and to reduce the interindividual variance between groups. We use midsagittal sectioning planes for both the rat tibia and the rat vertebra (Fig. 1). In mice, most labs prefer the distal femoral metaphysis for cancellous bone histomorphometry in the appendicular skeleton. A drawback of the distal femur is that the shape of the growth plate is more complex compared with the proximal tibia (Fig. 1). However, the strong curvature of the tibia in mice makes it more difficult to standardize the sectioning plane, outweighing the advantage of the simpler growth plate geometry. In contrast to rats, murine vertebrae need to be sectioned in a frontal plane. Otherwise, the amount of cancellous bone available for analysis is very limited (Fig. 1).

2. Materials 2.1. Sectioning, Fixation and Embedding

1. Rotary microtomes (Microm HM 355S and HM 360). 2. Sledge microtome (Leica Polycut S2500). 3. Water-cooled precision diamond band saw (Exakt, Norderstedt, Germany). 4. Slide press (Enno Vieth Mikrotome GmbH, Wiesmoor, Germany).

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Fig. 1. Sectioning planes in rat and mouse bones. (a–b) Midsagittal sectioning planes of a rat proximal tibia (a), a rat lumbar vertebral body (b), and a mouse lumbar vertebral body (c). (d–e) Exactly midsagittal (d) and slightly parasagittal (e) sectioning planes of a distal mouse femur show the complex geometry of the distal femoral growth plate in parasagittal sections. (f) Midsagittal section of a mouse tibia. (g) Frontal section of mouse lumbar vertebral body. (a) and (b) are von Kossastained 5-mm-thick sections without counterstain, (c)–(d) are 3-mm-thick von Kossa-stained sections counterstained with McNeal’s tetrachrome. Original magnifications ×12 in (a), ×20 in (b), ×25 in (c)–(g).

5. Binder environmental test chamber MK series (Binder GmbH, Tuttlingen, Germany). 6. Aminopropyltriethoxysilane. 7. Loctite 420 adhesive (Henkel, Düsseldorf, Germany). 8. Splicer (Exakt). 9. Microgrinding system (Exakt). 10. 30% H2O2. 11. Acetone–isopropanol: 50% acetone (v/v) in isopropanol. 12. Fluoromount (Serva). 13. PFA: 4% paraformaldehyde in 0.1 M phosphate buffer, pH 7.4. 14. 40% Ethanol, 70% ethanol, and 96% ethanol. 15. Propan-2-ol. 16. 0.1 M phosphate buffer, with 10% (w/v) sucrose, pH 7.4. 17. Xylene. 18. Dry benzoyl peroxide (see Note 1).

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19. MMA Solution #1: Methylmethacrylate (e.g., Merck 800590) with 20% (v/v) dibutyl phthalate. 20. MMA Solution #2: Methylmethacrylate with 20% (v/v) dibutyl phthalate, and 1% (w/v) dry benzoyl peroxide. 21. MMA Solution #3: Methylmethacrylate with 20% (v/v) dibutyl phthalate, and 3% (w/v) dry benzoyl peroxide (see Note 2). 22. MMA solution #4: Methylmethacrylate with 35% (v/v) butyl methacrylate, 5% (v/v) methyl benzoate, and 1.2% polyethylene glycol 400. 23. MMA solution #5: Methylmethacrylate with 35% (v/v) butyl methacrylate, 5% (v/v) methyl benzoate, 1.2% polyethylene glycol 400, and 0.4% (w/v) dry benzoyl peroxide. 24. MMA solution #6: Methylmethacrylate with 35% (v/v) butyl methacrylate, 5% (v/v), methyl benzoate, 1.2% (v/v) polyethylene glycol 400, and 0.8% (w/v) dry benzoyl peroxide (see Note 2). 2.2. Von Kossa and McNeal Stain

1. 5% silver nitrate in distilled water (keep in dark). 2. Sodium carbonate/formaldehyde solution: 9.5% formaldehyde (v/v) in 0.138 M sodium carbonate (see Note 3). 3. Farmer’s reducer: 5% (w/v) potassium ferrocyanide in 10% sodium thiosulfate (see Note 4). 4. Tetrachrome stock solution: 0.1% (w/v) Methylene blue chloride, 0.16% Azure A eosinate, 0.02% Methylene violet (Bernthsen) in a solution of 50% methanol, and 50% glycerol (see Note 5). 5. Tetrachrome working solution: 5% tetrachrome stock solution in distilled water.

2.3. Toluidine Blue Stain

1. Buffer I: 8.22 mM Citric acid, 2.1 mM disodium hydrogen phosphate, pH 3.7. 2. Toluidine blue stain: 2% (w/v) Toluidine blue O in Buffer I, pH 3.7 (see Note 6).

2.4. Tartrate Resistant Acid Phosphatase Stain

1. Acetate buffer: 0.2 M Sodium acetate, 50 mM L(+)-tartaric acid, pH 5.0. 2. TRAcP reagent: 0.5% (w/v) Napthol AS-MX phosphate disodium salt and 1.1% Fast Red TR salt in distilled water (see Note 7). 3. Mayer’s Hematoxylin 0.1% (w/v) Hematoxylin, 0.02% (w/v) sodium iodate, 5% (w/v) aluminum potassium sulfate (12 H2O), 5% (w/v) chloral hydrate, and 0.1% (w/v) citric acid (see Note 8). 4. 0.2 M Tris–HCl buffer, pH 9.0.

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2.5. Cement Line Stain

1. Toluidine blue with tetraborate and formic acid: 1% (w/v) toluidine blue O, 1% (w/v) sodium tetraborate, and 0.1% (v/v) formic acid in distilled water (see Note 9).

2.6. Histomorphometric Analysis

1. Microscopes: Zeiss Axioskop microscope and Zeiss SV11 stereomicroscope. 2. Digital camera: Diagnostic Instruments Spot Insight CCD camera. 3. Osteomeasure interactive 3.0 interactive image analysis software (OsteoMetrics). 4. Zeiss AxioVision 4.7 software package.

3. Methods 3.1. Fixation, Dehydration and Infiltration of Bone Samples

The fixation and infiltration steps below should be carried out at 4°C using a magnetic stirrer. 1. Carefully remove soft tissues from the bones (see Note 10). 2. Fix the tissue samples as quickly as possible by immersing in 40% ethanol (see Note 11) for 48 h or in 4% paraformaldehyde (PFA) in 0.1 M phosphate buffer, at pH 7.4, for 24 h. 3. When the fixation step is complete, wash the samples in 0.1 M phosphate buffer, pH 7.4, containing 10% (w/v) sucrose (see Note 12). 4. Dehydrate the samples by immersing in graded alcohols and xylene as detailed in steps 5–11 below, adjusting the duration of incubation depending on the bone characteristics (see Note 3). 5. Immerse in 70% ethanol for 1–4 days. 6. Immerse in 96% ethanol for 1–4 days. 7. Immerse in 100% propan-2-ol for 1–2 days and repeat twice in total. 8. Immerse in xylene for 1–4 days and repeat twice in total. 9. Immerse the samples in MMA Solution #1 (routine histology) or MMA Solution #4 (histochemistry and immunohistochemistry) for 2–4 days. 10. Immerse the samples in MMA Solution #2 (routine histology) or MMA Solution #5 (histochemistry and immunohistochemistry) for 2–4 days. 11. Immerse the samples in MMA Solution #3 (routine histology) or MMA Solution #6 (histochemistry and immunohistochemistry) for 2–4 days.

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3.2. Preparation of Glass Vials for Tissue Embedding

3.2.1. Routine Embedding

In this section we describe how to prepare glass tubes containing polymerized MMA which are used as a receptacle for embedding the bone samples. The procedure described assumes that you are using 25-ml glass vials, but smaller vials can be used depending on the sample size (see Note 14). 1. Prepare fresh MMA #3 solution at RT. 2. Pour 5 ml of the polymerization mixture into each glass vial. 3. Place an airtight cap on the top. 4. Incubate for at least 24 h at 40°C. 5. Store the vials at room temperature until further use.

3.2.2. Histochemistry and Immunohistochemistry

1. Chill the 25 ml glass vials on ice. 2. Add 600 ml of N,N-dimethyl-p-toluidine to 100 ml of freshly prepared MMA solution which has been prechilled to 4°C and stir for a few minutes. 3. Pour 5 ml of the polymerization mixture into each glass vial. 4. Thoroughly gas the vial with N2 or CO2 for 20–30 s and place an airtight cap on the top. 5. Incubate for at least 24 h at 4°C. 6. Store the vials at room temperature until further use.

3.3. Embedding for Routine Histology

This polymerization and embedding protocol described below is based on that published previously by Schenk et al. (3) with some modifications. 1. Place the infiltrated bone samples from Subheading 3.1, step 11 into the prepared glass vials, described in Subheading 3.2.1, adding one bone sample per vial along with a label to identify the sample. 2. Fill the vials with the MMA #3 solution and place a plastic cap on the vial, ensuring that it is airtight. 3. Incubate in a water bath overnight at 26°C. 4. Continue the incubation at 28°C for a further 12 h and continue the incubation for a further 3 days, gradually increasing the temperature by 0.5°C to reach a final temperature of 31°C. 5. Store the embedded tissue samples at room temperature until further use.

3.4. Embedding for Histochemistry and Immunohistochemistry

Here, we describe an embedding technique for undecalcified bone samples suited for bone histomorphometry, histochemistry, and immunohistochemistry (2). The method can also be used for soft tissues. It is important that steps 1–4 are carried out on ice (see Note 15).

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1. Chill the prepared glass vials from Subheading 3.2.2 on ice. 2. Add 400 ml of N,N-dimethyl-p-toluidine to 100 ml of MMA III solution, which has been chilled to 4°C and stir for a few minutes. 3. Fill the vials with this solution. 4. Place the infiltrated bone samples from Subheading 3.1, step 11 into the prepared glass vials, described in Subheading 3.2.2, adding one bone sample per vial along with a label to identify the sample. 5. Place a plastic cap on the vial, ensuring that it is airtight. It is not necessary to gas the tubes, since the MMA is filled right to the top. 6. Transfer the vials to a cooling unit (see Note 16) and incubate at −23°C for 16 h. 7. Gradually increase the temperature from −23 to −22°C for over 1 h. 8. Gradually increase the temperature from −22 to −20°C for over 46 h. 9. Gradually increase the temperature from −20 to −18°C for over 24 h. 10. Gradually increase the temperature from −18°C to +2°C for over 12 h. 11. Store the tissue blocks at −20°C until further use (see Note 17). 3.5. Preparation of Micromilled Cross-Sections for Routine Histological Analysis

The usual sites for cortical bone histomorphometry in rats and mice are the femoral and tibial shaft. Because histochemical or immunohistochemical staining is rarely required for cross-sections, we embed all specimens routinely in conventional MMA as described in Subheading 3.2. In rats, we usually use the tibial shaft, sampled at 2 mm proximal to the tibiofibular junction. In mice, we use the femoral midshaft defined by equal distances from the proximal and distal end of the bone. The slides are then stained with toluidine blue after etching the surface with H2O2 (4) as described below or analyzed for fluorochrome labels without staining as described in Subheading 3.6: 1. Immerse acetone-cleaned glass slides in aminopropyltriethoxysilane for 5 min, rinse in distilled water, and allow them to dry. 2. Prepare 3–5 150 mm thick cross-sections of bone at the desired site using a water-cooled precision diamond band saw. 3. Glue the sections onto the 3-aminopropyltriethoxysilanecoated glass slides with Loctite 420 adhesive using a splicer (Exakt). 4. Grind the sections to a final thickness of 15–20 mm with the microgrinder.

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5. Wipe the micromilled section with acetone–isopropanol. 6. Immerse the sections in 30% H2O2 and agitate for 5 min. 7. Thoroughly rinse in tap water. 8. Immerse the sections in toluidine blue solution for 60 min. 9. Rinse the sections in distilled water. 10. Allow the sections to air-dry for 120 min. 11. Wipe the sections with acetone–isopropanol. 12. Cover-slip sections with DePeX and proceed to histological analysis. 3.6. Preparation of Micromilled Cross-Sections for Analysis of Fluorochrome Labels 3.7. Von Kossa Stain

1. Follow steps 1–4 of Subheading 2.3. 2. Cover-slip sections with Fluoromount and proceed to histological analysis.

We use this stain for image analysis of bone structure, because it gives a very high contrast between bone which is stained black and bone marrow which is unstained. For optimal histological quality, it is advantageous to avoid dehydration at the end of the staining protocol and we therefore mount the sections using an aqueous mounting medium. 1. Immerse the sections in 2-Methoxyethyl acetate for 20 min and repeat three times in total. 2. Immerse the sections in 70% ethanol for 5 min. 3. Immerse the sections in 40% ethanol for 5 min. 4. Immerse the sections in distilled water for 5 min and repeat three times in total. 5. Immerse the sections in the dark with 5% silver nitrate solution for 5–10 min (see Note 18). 6. Wash the sections in distilled water and repeat three times in total. 7. Immerse the sections in sodium carbonate/formaldehyde solution for 2 min. 8. Rinse the sections in tap water. 9. Immerse the sections in Farmer’s reducer for 30 s. 10. Rinse the sections in running tap water for 20 min. 11. Cover-slip in Kaiser’s glycerol gelatine.

3.8. Von Kossa/McNeal Stain

This protocol describes a combination of von Kossa’s stain with McNeal’s tetrachrome stain as counterstain (3). It combines good cellular detail with a very clear distinction between mineralized and

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unmineralized bone. We use this stain for the assessment of cellular and osteoid parameters especially in mice. 1. Follow the protocol for von Kossa staining as described in Subheading 3.7, steps 1–10. 2. Immerse the slides in fresh tetrachrome working solution for 20–60 min. 3. Rinse the slides in distilled water. 4. Rinse the slides in propan-2-ol twice. 5. Rinse the slides in xylene for 5 min. 6. Cover-slip slides with DePeX mounting medium and proceed to histological analysis. 3.9. Toluidine Blue Stain

The toluidine blue stain at acid pH described previously by Baron et al. (5) gives very good cellular detail. If applied properly, it also gives a good distinction between mineralized and unmineralized bone. It is our favorite stain for the histomorphometric measurement of cellular parameters in rats. 1. Remove the MMA from the sections by follow steps 1–4 of protocol in Subheading 3.7. 2. Immerse the slides in toluidine blue stain for 10 min. 3. Immerse the slides in Buffer I for 1 min and drain the slide rack on blotting paper. 4. Repeat step 3. 5. Immerse the slides in n-butanol for 1 min and repeat three times in total. 6. Immerse the slides in n-butanol/xylene for 1.5 min. 7. Immerse the slides in xylene for 1 min and repeat twice in total. 8. Mount in DePeX, add a cover slip, and proceed to histological analysis.

3.10. Tartrate Resistant Acid Phosphatase Staining

This protocol is used for histochemical detection of TRAcP in undecalcified sections of bone which have been embedded using the modified MMA protocol described in Subheading 3.4. The stain identifies osteoclasts since they express very high levels of TRAcP (see Note 19). 1. Immerse the sections in 2-Methoxyethyl acetate for 20 min and repeat three times in total. 2. Immerse the slides in acetone for 5 min and repeat twice in total (see Note 20). 3. Immerse the slides in distilled water for 5 min and repeat twice in total.

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4. Immerse the slides in 0.2 M acetate buffer, pH 5.0, for 20 min at room temperature. 5. Add approximately 100 ml TRAcP-reagent to each sections and incubate for 2–4 h at 37°C in a humidified chamber. 6. Rinse in distilled water. 7. Counterstain with Mayer’s hematoxylin for 2 min. 8. Rinse in distilled water. 9. Wash in running tap water for 5–10 min. 10. Mount in Kaiser’s glycerol gelatine. 3.11. Cement Line Stain

This protocol describes a modification of the surface-staining technique described by Schenk et al. (3) for demonstration of cement lines in undecalcified sections of rat and mouse bones. It is not necessary to remove the embedding material prior to this procedure. 1. Immerse the slides in 0.1% formic acid for 45 s. 2. Rinse the slides in distilled water. 3. Incubate the slides in 20% methanol for 60 min. 4. Rinse the slides in distilled water. 5. Immerse the slides in toluidine blue solution for 3 min. 6. Rinse the slides in distilled water. 7. Air-dry the slides for 120 min. 8. Cover-slip with DePeX and proceed to histological analysis.

3.12. Cancellous Bone Histomorphometry

Standard sites for cancellous bone histomorphometry in rats are the proximal tibial or femoral metaphyses and the lumbar vertebral bodies, but the choice of site depends on the age of the animal and whether or not the animal is osteopenic (see Note 21). Normally, cancellous bone histomorphometry is performed within the secondary spongiosa. To exclude the primary spongiosa, bone within a certain distance from the growth plate is excluded from the analyses (Fig. 2). There is no definitive rule for this, but suggested criteria are discussed below. In the rat proximal tibial metaphysis, 1 mm distance from the growth plate is appropriate in 2- to 3-month-old rats. Beyond about 5 months of age, 0.5 mm can be used. If necessary, this value can be reduced to 0.25 mm in rats beyond 9–12 months of age. In the murine distal femoral metaphysis and proximal tibial metaphysis, 0.25 mm can be used for all ages beyond 4 weeks of age. In lumbar vertebrae, 0.25 mm distance from the cranial and caudal growth plate each can be used for all ages in mice, and 0.5 mm for all ages in rats. Under some circumstances such as in experiments that involve the use of antiresorptive drugs, the area beneath the growth plate excluded from the analysis can be larger (see Note 22). To exclude endocortical bone remodeling activity, cancellous bone within 0.25 mm from the endocortical bone surface should be excluded from the analyses, regardless of the

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Fig. 2. Measuring areas for cancellous bone histomorphometry. (a–b) Large measuring areas for automatic image analysis encompass most of the secondary spongiosa in a sagittal section of a distal mouse femur (a), and a frontal section of a mouse lumbar vertebral body (b). The distance of the measuring field from the growth plates and the endocortical bone surfaces is approximately 0.25 mm in both bones. (c) Typical measuring fields for interactive measurements at ×200 magnification in a rat proximal tibia. The size of the individual measuring fields is 0.5 × 0.5 mm. A distance of 0.5 mm is kept from the growth plate. Von Kossa-stained sections of mouse and rat bones. Original magnifications ×25 in (a) and (b), ×12 in (c).

sampling site (Fig. 2). Unless there is an impairment in bone mineralization (see Note 23), we follow a two-step analysis strategy in our lab for almost all samples. In a first step, we measure bone structural parameters such as bone area or trabecular width by automatic image analysis in a large area encompassing more or less the whole secondary spongiosa in a specific site (Fig. 2). Because this is a very rapid analysis requiring only about 3 min per section, two sections can be analyzed to reduce the variance. We use a Diagnostic Instruments Spot Insight CCD camera in combination with a Zeiss Axioskop microscope with a × 2.5 objective or a Zeiss SV11 stereomicroscope for image acquisition. For image analysis, we use the Zeiss AxioVision 4.7 software package.

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In a second step, we assess bone turnover by measurements done with an interactive image analysis system. We prefer to employ a microscope with a drawing attachment and a digitizing tablet equipped with a cursor with a LED light source for the interactive measurements. The measurement area is defined by a square grid in one of the oculars. Thus, with the help of the LED-illuminated cursor and the drawing attachment, features within the section can be traced by looking into the microscope. The advantage of this setup is that it is faster, more accurate, and also less strenuous for the eyes compared with measurements on the screen. We employ a Zeiss Axioskop microscope with a drawing attachment, and the OsteoMeasure interactive image analysis software for the latter setup. Typical measuring areas are shown in Fig. 2. 3.13. Assessment of Bone Structural Parameters

Bone structural parameters should be assessed using von Kossastained sections. The method described here is only applicable in samples with a low amount of osteoid (see Note 23). Depending on the sample size, we use a Zeiss Axioskop microscope with a ×2.5 objective or a Zeiss SV11 stereomicroscope for image acquisition. The images are processed and analyzed using Zeiss AxioVision 4.7 software. To reduce the variation, we measure two sections spaced at least 100 mm from each other. The mean value of both measurements is used for further calculations. Analysis of bone structural parameters involves making only four primary measurements (Table 1), but from these various other parameters in both two dimensions (2D) and three dimensions (3D) can be derived.

Table 1 Primary and derived structural parameters of bone Primary parameters Total area (T.Ar; the measuring area) Bone area (B.Ar) Bone perimeter (B.Pm), a The number of individual bone trabeculae (N.Tb) 2D derived parameters Bone area (B.Ar/T.Ar) = B.Ar/T.Ar × 100 (%) Number of trabeculae per tissue area (N.Tb/T.Ar) = Tb.N/T.Ar (no./mm²) Trabecular area (Tb.Ar) = B.Ar/N.Tb (mm²) Number of trabeculae per bone area (N.Tb/B.Ar) = N.Tb/B.Ar (no./mm²) Bone perimeter (B.Pm/T.Ar) = B.Pm/T.Ar (mm/mm²) Trabecular width (Tb.Wi) = B.Ar/B.Pm × 2,000 (mm) 3D derived parameters Bone volume (BV/TV) = B.Ar/T.Ar × 100 (%) Bone surface (BS/TV) = B.Pm/T.Ar × 4/p (mm²/mm³) Trabecular thickness (Tb.Th) = B.Ar/B.Pm × 2000 × p/4 (mm) Trabecular number (Tb.N) = 4/p × 0.5 × B.Pm/T.Ar (no./mm) Trabecular separation (Tb.Sp) = 1/Tb.N × 1000 − Tb.Th (mm)

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Two-dimensional parameters such as number of trabeculae per tissue area, trabecular area, and number of trabeculae per bone area indirectly reflect the connectivity of the trabecular network. The calculation of the 3D parameters bone surface, trabecular thickness, trabecular thickness, and trabecular separation from 2D data is based on the factor 4/p. This factor is based on the assumption that the structure is isotropic (1). This model assumption is obviously wrong for the standard sampling sites in mice and rats (Fig. 1). Therefore, one has to be aware that calculation of histomorphometric 3D parameters in rat and mouse cancellous bone is always associated with some error. Histomorphometry has largely been superseded by mCT analysis for analysis of 3D bone structure. Therefore, we usually express structural histomorphometric data in 2D only. In addition, we routinely calculate Tb.Th, Tb.N, and Tb.Sp for comparison with structural data obtained by mCT analysis. 3.14. Analysis of Dynamic Histomorphometry in Mice

Histomorphometry is an indispensable tool for the assessment of local turnover mechanisms within bone. For mouse studies, it is usually sufficient to analyze fluorochrome labeling as described in this section and osteoclast numbers as described in Subheading 3.15 because together these reflect bone remodeling activity. Osteoid and mineralization parameters are analyzed only when needed. Although older mice do have a bone remodeling activity, remodeling-based parameters are normally not assessed in mice. Dynamic bone histomorphometry is based on the analysis of fluorochromelabeled bone specimens and is used to assess the rate of osteoblastic bone formation and bone mineralization. We routinely use double calcein labeling for dynamic histomorphometry (Fig. 3). Other labs

Fig. 3. Calcein double labeling for measurement of bone formation. Unstained 3-mm-thick section viewed under blue excitation shows calcein double labels (arrows) in murine cancellous bone. Mineralized bone is marked by asterisks. Original magnification ×400.

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use two different labels for double labeling. However, unless the information contained within the individual labels is used (6, 7), two different labels do not offer any advantage in our hands. It is very important to use appropriate marker intervals. In order to avoid large label escape and skewed sampling errors, the marker interval has to be as short as possible. For a detailed explanation of these errors, the reader is referred to other textbooks (6, 8). In 3to 4-week-old mice we use a marker interval of 1 day, in 6- to 12-week-old mice 2 days, and in older mice 2 or 3 days (6). In rats older than 3 months, we use a 5-day marker interval (6). The measurements are made on unstained sections, using a ×20 objective in most cases, together with an interactive image analysis system. Primary measurements are the bone perimeter, the mineralizing bone perimeter (M.Pm), and the mineral apposition rate (MAR). There are two possible definitions for M.Pm. M.Pm is either defined as the double-labeled perimeter (M.Pm = Db.Lb. Pm), or as Db.Lb.Pm + one half of the single-labeled perimeter (M.Pm = Db.Lb.Pm + 0.5 × S.Lb.Pm). The latter expression is the mathematically more correct term, and is the standard in humans (1). However, because nonspecific fluorochrome labeling is a problem in rats and mice, we only use the definition M.Pm = Db.Lb. Pm. This definition underestimates the true M.Pm, but reduces the likelihood of errors caused by nonspecific labeling. The MAR (mm/day) is defined by the mean distance between the labels, divided by the marker interval. The MAR can be measured indirectly by tracing along the individual labels, or directly by a twopoint distance measurement. When broader, “fuzzy” labels are present, the interlabel distance is best measured from mid-point to mid-point of the labels (8, 9). We mostly use the indirect method, because Db.Lb.Pm and MAR are measured at once, which saves time. The surface-based bone formation rate (BFR) is defined by multiplying M.Pm with MAR, BFR/B.Pm = BFR/BS = M. Pm/100 × MAR (mm²/mm/day or mm³/mm²/day). There are other ways to express BFR, using different referents such as the tissue volume or the time (1). However, we routinely use only BFR/B.Pm or BFR/BS, because it is the best reflection of the intensity of bone formation per unit of bone perimeter/surface. Dynamic histomorphometric parameters are a functional readout of matrix synthesis by the osteoblast and the subsequent mineralization of that matrix. Therefore, these parameters are clearly superior to any morphological assessments of osteoblastic activity such as osteoblast perimeter or osteoblast number. Special problems may arise in assessing dynamic histomorphometry when antiresorptive drugs are being used due to the inhibition of bone turnover which may result in the absence of double fluorochrome labels. How should BFR be reported in the case of a lack of double labeling? To exclude a technical problem due to missing labels, other sections and sites should be analyzed for the presence of

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double labels. The presence of a double label anywhere in the skeleton indicates that the animal was properly labeled. When the animal was properly labeled and double labels are absent within the sampling site, we report the M.Pm and the BFR as zero, whereas MAR is treated as missing value. 3.15. Assessment of Osteoclast Numbers and Bone Resorption in Mice

Osteoclasts are generally more difficult to recognize in mice compared with other species, because a large proportion of murine osteoclasts are mononuclear and because Howship’s lacunae are shallow. For reliable measurements of osteoclast numbers in mice, it is important to use TRAcP stained sections. We measure osteoclast parameters in mice on TRAcP-stained sections with a ×20 objective. Only TRAcP-positive nucleated cells in contact with bone are counted as osteoclasts, not osteoclast profiles lacking a nucleus or TRAcP-positive cells within bone marrow (Fig. 4). The most commonly used primary and derived measurements of osteoclast number and bone resorption are shown in Table 2. Osteoclasts can only occur on bone surfaces. Therefore, osteoclast numbers are best expressed per millimeter of bone perimeter (N.Oc/B.Pm). In fact, it can be very misleading to express osteoclast number per tissue area (N.Oc./T.Ar) in osteopenic animals. When there is a lot of osteoid, it can be useful to express osteoclast numbers per millimeter of mineralized bone perimeter, with Md. Pm = B.Pm − osteoid perimeter (O.Pm) since osteoclasts can only

Fig. 4. TRAcP staining of osteoclasts in murine bone sections. TRAcP-stained 3-mm-thick section of a mouse lumbar vertebra shows red-stained osteoclasts. Nuclei appear as lighter stained areas. Osteoclast profiles in contact with bone but without nucleus (arrow) or TRAcP-positive cells not in contact with bone (arrowheads) are not counted as osteoclasts. The section is counterstained with hematoxylin. Original magnification ×400.

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Table 2 Measured and derived indices of bone resorption Primary parameters Bone perimeter (B.Pm) Number of osteoclasts (N.Oc) Osteoclast perimeter (Oc.Pm)a Derived parameters Osteoclast number (N.Oc/B.Pm) = N.Oc/B.Pm (no./mm) (surface referent) Osteoclast number (N.Oc/Md.Pm) = N.Oc/(B.Pm − O.Pm) (no./mm) Osteoclast number (N.Oc/T.Ar) = N.Oc/T.Ar (no./mm²) (tissue referent) Osteoclast perimeter (Oc.Pm/B.Pm) = osteoclast surface (Oc.S/BS) = Oc. Pm/B.Pm (%) a

The contact perimeter between the osteoclast and the bone surface

resorb mineralized bone. Therefore, N.Oc/Md.Pm (no./mm) is the best estimate of osteoclastic bone resorptive activity in situations where osteoid perimeter differs between different groups of animals. Because the TRACP stain does not permit reliable quantification of osteoid, Md.Pm has to be calculated as follows: Md. Pm = B.Pm − B.Pm × O.Pm/B.Pm. The proportion of osteoid-covered bone perimeter (O.Pm/B.Pm) has to be quantified separately. It is worthwhile expressing osteoclast number per mineralized perimeter only in the presence of pronounced differences in osteoid perimeter between the groups of animals. Additional parameters can be the number of nuclei per osteoclast, and mean osteoclast size. However, the latter parameters make sense only in the case that osteoclast morphology is altered. 3.16. Assessment of Bone Mineralization in Mice

We do not routinely assess osteoid or osteoblast parameters in murine bone samples. These measurements add useful information only when bone mineralization is impaired. We perform these measurements on sections stained with von Kossa and counterstained with McNeal’s tetrachrome, using a ×20 objective. This stain gives a very reliable and clear distinction between mineralized bone and unmineralized osteoid tissue (Fig. 5). Primary and derived measurements are summarized in Table 3. Increases in osteoid perimeter and osteoid area are not necessarily associated with impaired bone mineralization, but can also be caused by increases in bone formation in the absence of disturbed bone mineralization. Rather, increased osteoid width and, more specifically, increased OMT are indicative of impaired bone mineralization. However, many mouse models of impaired bone mineralization are characterized by severe impairments of bone mineralization (Fig. 5), resulting in the complete absence

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Fig. 5. Osteoid in murine bone sections. Von Kossa/McNeal staining provides the most reliable discrimination between black mineralized bone and pale-stained osteoid. (a) Thin osteoid seams (arrows) covered by osteoblasts in distal femoral cancellous bone of a wild-type mouse. An osteoclast showing one nucleus is marked by an arrowhead. (b) Severe osteoidosis (asterisks) in distal femoral cancellous bone of a Hyp mouse. Hyp mice are hypophosphatemic and are characterized by a loss-of-function mutation in the Phex gene (phosphate-regulating gene with homologies to endopeptidases on the X chromosome), which indirectly controls secretion of the phosphaturic fibroblast growth factor-23 in a negative fashion; 3-mm-thick sections. Original magnification ×400.

Table 3 Measured and derived indices of bone mineralisation Primary parameters Bone perimeter (B.Pm) Bone area (B.Ar) Osteoid perimeter (O.Pm) Osteoid area (O.Ar) Osteoblast perimeter (Ob.Pm) Osteoid width (O.Wi)a Derived parameters Osteoid perimeter (O.Pm/B.Pm) = Osteoid surface (OS/BS) = O.Pm/B.Pm × 100 (%) Osteoblast perimeter (Ob.Pm/B.Pm) = Osteoblast surface (Ob.S/BS) = Ob.Pm/B.Pm × 100 (%) Osteoid area (O.Ar/B.Ar) = Osteoid volume (OV/BV) = O.Ar/B.Ar × 100 (%) Osteoid width (O.Wi) = O.Ar/O.Pm × 1,000 (mm) Osteoid maturation time (OMT) = O.Wi/MAR (days) Osteoid thickness (O.Th) = O.Wi × p/4 (mm) a

Usually O.Wi is calculated from the osteoid area and perimeter, but it can be measured directly by a two-point distance measurement by sampling osteoid seams every 50 mm

of fluorochrome double labels and immeasurable values for MAR. Therefore, OMT cannot be determined in these mouse models. However, OMT is generally a very sensitive parameter to pick up subtle impairments in bone mineralization.

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3.17. Assessment of Dynamic Histomorphometry in Rats

The procedures here are identical to mouse samples as described in Subheading 3.16.

3.18. Assessment of Static Histomorphometry in Rats

Static histomorphometric assessment of bone turnover in rats is different from mice because rat osteoclasts are easily recognized in bone sections due to their typical morphological and staining characteristics (Fig. 6) and because remodeling-based parameters can provide useful information about changes in activation frequency and remodeling periods in rats. In view of this, it is not necessary to analyze TRAcP-stained sections in rats but we routinely measure eroded perimeter in rats since this is necessary for the calculation of the resorption period. We use toluidine blue-stained sections and a ×20 objective for this type of measurement in rats. The advantage of toluidine blue over von Kossa/McNeal is that remodeling units within mineralized bone are still visible (Fig. 6). Although the von Kossa/McNeal stain is more reliable for distinguishing between osteoid and mineralized bone, information about the structural organization of mineralized bone is lost due to the deep black staining of mineralized bone. Therefore, we prefer the toluidine blue stain for analysis of structural parameters in rat bone. Primary parameters are bone perimeter, bone area, osteoid area, osteoid perimeter, osteoblast perimeter, osteoclast number, osteoclast perimeter, and eroded perimeter. Rat osteoblasts are generally more flattened compared with murine osteoblasts (Fig. 6). Therefore, a clear-cut morphological definition of rat osteoblasts is difficult. We exclude only very flat cells from the measurement of osteoblast perimeter (Fig. 6). Moreover, we count only nucleated cells as osteoclasts, not osteoclast profiles (Fig. 6). For the measurement of eroded perimeter, we use a minimum depth of 3 mm for resorption cavities as cutoff value to exclude very shallow

Fig. 6. Toluidine blue staining of rat cancellous bone sections. (a) Toluidine blue at acid pH stains mineralized bone light blue, so that structural details within bone are still visible (remodeling unit is marked by arrows ). (b) Osteoblasts sitting on a thin osteoid seam (arrows) in rat cancellous bone. Osteoclasts are labeled by arrowheads. Five-micron-thick sections of rat lumbar vertebrae stained with toluidine blue at acid pH. Original magnification ×400.

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resorption sites. Most derived parameters are identical to the ones listed in Table 1 for mice, with the addition or eroded perimeter (E.Pm/B.Pm) = Eroded surface (ES/BS) = E.Pm/B.Pm × 100 (%). 3.19. Assessment of Remodeling-Based Parameters in Rats

For the calculation of remodeling-based parameters it is necessary to determine wall width, which is the mean width of completed remodeling packages (Fig. 7). We use cement line-stained sections viewed under polarized light, and a ×20 objective for this measurement. Wall width can either be measured by tracing the bone surface and the reversal line of the remodeling package, or by two-point distance measurements. In the latter case, the standard is to make four evenly distributed distance measurements between the reversal line and the bone surface, perpendicular to the bone surface, for each remodeling unit (Fig. 7). To arrive at reliable values, we always measure the width of at least 15 remodeling units in one sample. Wall width can be converted to 3D wall thickness (W.Th) by multiplication with p/4. After determination of wall width, several derived parameters can be calculated (Table 4). The principle behind these calculations is that the surface extent of a certain activity is proportional to the time period occupied by this activity. In rodents, only the active formation period is used, not the formation period based on the adjusted apposition rate. The latter includes so-called “Off” periods (see Note 24). The relative proportion of cancellous bone remodeling compared

Fig. 7. Measurement of wall width for remodeling-based histomorphometric parameters. Wall width is defined as the mean distance between the bone surface and the scalloped reversal line in individual completed remodeling units. A smooth arrest line, indicating a temporary stop in osteoblastic bone formation within a remodeling unit, is marked by an arrowhead. Five-micron-thick section of a rat lumbar vertebra surface-stained with cement line stain. Original magnification × 400.

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Table 4 Derived remodeling based parameters in rats Active formation period (FP) = W.Wi/MAR (days) Resorption period (Rs.P) = FP × E.Pm/O.Pm (days) Active resorption period (Rs.P act.) = FP × Oc.Pm/O.Pm (days) Reversal period (Rv.P) = FP × (E.Pm − Oc.Pm)/O.Pm (days) Remodeling period = FP + Rs.P (days) Total period = FP × B.Pm/O.Pm (days) Activation frequency (Ac.F) = 1/Tt.P (no./year)

Fig. 8. Morphology of cross-sections of murine cortical bone and direct measurement of cortical thickness. (a) Toluidine blue-stained microground cross-section of a mouse femoral midshaft. (b) Binary image of the same section showing the intercepts of 90 radii originating from the center of gravity of the bone section with the cortical bone ring. Original magnification × 25.

with modeling increases with age especially in the appendicular skeleton (10). Therefore, remodeling-based parameters are usually measured in rats >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)

8.

9.

10.

11.

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

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_20, © Springer Science+Business Media, LLC 2012

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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

8–12 h (overnight)

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

8–12 h (overnight)

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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Cryo-sectioning -24°C

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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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

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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 <10 days) can be processed without the decalcification step. 3.5. Decalcification of Mineralized Tissues

1. Following fixation, immerse the bone samples in decalcification solution. 2. Place the samples at 4°C for 2–3 weeks replacing the decalcification solution weekly. 3. Check whether decalcification has been completed by X-ray photography. 4. Once decalcification is complete, proceed to embedding.

3.6. General Approach to Embedding and Analysis of Mineralized Tissues by TEM

Special care has to be taken in the embedment of calcified tissues since epoxy resins do not easily penetrate such tissues. Bones and/ or teeth obtained from very young animals can easily be embedded in resin without giving problems with cutting and/or staining. Special attention has to be paid to embedding of bones from older and larger animals or man.

3.7. Embedding of Small Tissue Samples

Tissue samples obtained from young animals (up to 1 week old) of approximately 3–5 mm in thickness are put into glass (plastic dissolves in propylene oxide!) vials which can be closed and they are embedded as follows: 1. Immerse the sample in 70% ethanol for 3 × 5 min. 2. Replace with 80% ethanol for 3 × 5 min. 3. Replace with 90% ethanol for 3 × 5 min. 4. Replace with 96% ethanol for 3 × 5 min. 5. Immerse the tissue in 100% ethanol and close the bottle. Incubate for 10 min. Repeat the 10 min incubation in 100% ethanol 3 times in total. 6. Replace the ethanol by propylene oxide. Incubate in a closed vial for 30 min. Change the propylene oxide and repeat this once. 7. Dissolve the epoxy resin in propylene oxide at a 1:1 concentration. Immerse the tissue samples in the epoxy resin–propylene mixture, close the vials and leave overnight while gentle shaking. 8. Replace with pure epoxy resin and with the vials left open, place on a shaker, and leave gently shaking for 5 h. 9. Immerse the tissue samples in fresh epoxy resin in plastic molds, label, and leave overnight in an oven at 40°C. 10. Transfer to an oven set at 60°C to allow polymerization of the resin.

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It is possible to embed tissue samples of the size of a lower jaw of a full-grown mouse with good results. Larger samples have to be reduced in size or have to be cut in smaller pieces in order to obtain sufficient penetration of the epoxy resin. 1. Follow the procedure as indicated under Subheading 3.7 but increase the duration of each ethanol dehydration step to 10 min and each propylene oxide impregnation steps to 20 min. 2. Immerse the samples in propylene oxide:epoxy resin at a ratio of 3:1 for 3 h. 3. Immerse the samples in a 1:1 mixture of propylene oxide:epoxy resin for 3 h. 4. Immerse the samples in a 1:3 mixture of propylene oxide:epoxy resin overnight. 5. Immerse the samples in pure epoxy resin, with the vials open and shaking for 6 h. 6. Embed the samples in fresh epoxy resin and leave overnight in an oven at 40°C. 7. Proceed with polymerization as described in Subheading 3.7, step 10.

3.9. Sectioning Mineralized Tissues

Sectioning of the tissue is best performed using diamond knives, which are available both for semi-thin and ultrathin sectioning (See Notes 5 and 6). 1. Use a glass knive to trim the block. 2. Proceed with semi-thin and ultrathin sectioning using a diamond Histo-knife with the cutting angle set at 6°. 3. Make sure that the microtome manually cuts at a very low speed (1 mm/s). 4. In order to avoid wetting of the surface of the tissue block, keep the water level of the trough as low as possible.

3.10. Methylene Blue Staining of Semi-Thin Sections

The methylene blue staining solution stains most tissue components. It is an excellent stain for general purposes. Due to its metachromatic properties, some components (e.g., cartilage and granules of mast cells) stain purple. 1. Cut semi-thin sections of approximately 1–2 μm thickness. 2. Collect the sections on a drop of water on a glass slide and dry the sections on a heating plate (60–70°C), leave the sections on the plate for an hour or longer. In order to avoid wrinkles of sections of larger tissue samples sections are best dried on a plate at 50°C. Dry these sections overnight before staining. 3. When the sections are dry, add a drop of the filtered methylene blue staining solution to the section.

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4. Leave the staining solution for approximately 15 s (depending on the type of tissue and thickness of the section). 5. Wash the section extensively with a jet of distilled water. 6. Dry the washed section and cover it with a drop of epoxy resin. 7. Cover the section with a cover glass and leave the section for several hours on a hot plate (or in a stove at 60°C) to polymerize the resin. 3.11. Modified Goldner’s Masson Stain

The Goldner Stain (1) stains nonmineralized bone (e.g., osteoid) red, mineralized bone green, and calcified cartilage very light green. It also provides a very good staining for cells associated with bone and cartilage (Fig. 1). All staining procedures are carried out on a hot plate at 68–70°C. 1. Cut sections of approximately 2.5 μm thickness. 2. Collect the sections on a coated (e.g., Vectabond, Vector Laboratories, #SP-1800) slide and dry them on a hot plate. 3. Wet the sections with distilled water and shake off excess of water.

Fig. 1. Light micrograph of a section of mouse calvaria bone stained with the Goldner Masson’s Trichrome staining. The mineralized bone (B) is green and the osteoid (OS) is red. Section was obtained from epoxy resin-embedded undecalcified bone. Bone was fixed in 4% formaldehyde and 1% glutaraldehyde in 0.1 M sodium cacodylate buffer. OB osteoblast. ×3,000.

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4. Stain 3 min with ferric chloride on a hot plate. 5. Rinse quickly with warm tap water and dry on a hot plate. 6. Stain 25 s with hematoxylin on a hot plate. 7. Rinse quickly with warm tap water and dry on a hot plate. 8. Stain 8 min with ponceau-acid fuchsin staining solution on a hot plate. 9. Rinse quickly with 0.5% acetic acid. 10. Stain 3 min with 0.5% phosphomolybdic acid on a hot plate. 11. Rinse quickly with 0.5% acetic acid. 12. Stain 3–5 min with light green SF yellowish on a hot plate. 13. Rinse quickly with 0.5% acetic acid. 14. Dry the sections on a hot plate and cover them with epon. 3.12. Von Kossa

The Von Kossa staining procedure (2) results in a black staining of mineralized tissue parts. If methylene blue is used as a counterstain, the other tissue components stain in different intensities of blue. 1. Cut sections of 2 μm thickness. 2. Collect the sections on coated slides (see Subheading 3.11, step 2) and dry them on a hot plate. 3. Incubate the sections with 0.5% silver lactate for 20 min at ambient temperature. 4. Rinse the sections with double distilled water. 5. Incubate the sections with 0.5% hydrochinon for 2 min at ambient temperature. 6. Rinse the sections in double distilled water. 7. Incubate the sections with 5% sodium thiosulfate pentahydrate for 2 min at ambient temperature. 8. Rinse the sections in double distilled water. 9. Sections can be counterstained with methylene blue (see Subheading 3.10). 10. Dry the sections and cover them.

3.13. Staining of Ultrathin Sections with Uranyl Acetate

1. Centrifuge the uranyl solution (10 min, 2,000 × g). 2. Put drops of uranyl acetate on a strip of parafilm. 3. Float the grids on top of the drops (sections facing the solution). 4. Stain the sections for 4–8 min in the dark. 5. Rinse the sections extensively with double distilled water. 6. Air-dry the sections and stain them with lead nitrate.

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3.14. Staining of Ultrathin Sections with Lead Nitrate (3)

1. Centrifuge the lead solution (10 min, 3,000 × g). 2. Put drops of lead nitrate on a strip of parafilm. 3. Place a few sodium hydroxide pellets around the drops. 4. Float the grids on top of the drops of the lead solution. 5. Cover the drops and pellets with a cover of a petri dish. 6. Stain the sections for 2–4 min. 7. Rinse the sections extensively with double distilled water. 8. Air-dry the sections.

4. Notes 1. The fixatives mentioned in this chapter give reproducible and reliable results (Figs. 2 and 3). The recipes given are for large volumes. If your laboratory does not have a high-throughput of samples, the amounts can be reduced appropriate to your needs. One of the advantages is that tissue samples can be stored for a very long time in the aldehyde mixture without noticeable loss of ultrastructural details. Essential is that the formaldehyde solution is not older than 1 week before its use for the initial fixation of tissue samples.

Fig. 2. Low power electron micrograph of a mouse metatarsal fixed in 4% formaldehyde and 1% glutaraldehyde in 0.1 M sodium cacodylate buffer. OC osteoclast, M mineralized cartilage, asterisk indicates an area where the bone is absent due to ultrathin cutting of the mineralized matrix. ×5,000.

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Fig. 3. Electron micrograph of osteoblasts (OB) separated from mineralized bone (B) by a layer of osteoid (OS). Calvaria bone of a 10 d old mouse was fixed in 4% formaldehyde and 1% glutaraldehyde in 0.1 M sodium cacodylate buffer. Undecalcified bone ×16,000.

2. Apart from the fixative mentioned in this chapter numerous others mentioned in the literature could be used which may give very good results. Whatever fixative used, it is crucial to avoid the use of phosphate buffer. Fixation carried out in phosphate-buffered fixatives, in particular if the fixative is somewhat older, results in the formation and precipitation of mineral crystallites (Fig. 4). These crystallites are formed at the conjunction of mineral-containing tissue (e.g., bone) and the surrounding soft tissue but also within the cells, in particular in vacuoles and mitochondria. Due to a relatively high calcium concentration at these sites, calcium-phosphate crystals are rapidly formed. 3. Postfixation with osmium/ferrocyanide results in strongly contrasted plasmamembranes. If this is not desired, postfixation should be carried out in osmium without ferrocyanide (Subheadings 2.2 and 2.3). 4. Considerations mentioned under Note 2 are also relevant for the choice of medium used to culture bone explants. Different commercially available culture media contain different concentrations of phosphate; the higher the concentration level the higher the chance for precipitation of crystals as soon as calcium is liberated. During our investigations on bone resorption, we have experienced this. We found that blocking the

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Fig. 4. Electron micrograph of a section of undecalcified bone obtained from a patient suffering from osteopetrosis. Note the presence of large crystallites (arrows) due to a phosphate-buffered fixative. Bone tissue was fixed in 4% formaldehyde and 1% glutaraldehyde in 0.1 M phosphate buffer. B bone, CYT cytoplasm, N nucleus. ×26,500.

activity of certain proteolytic enzymes resulted in the occurrence of large areas of nondigested demineralized bone matrix adjacent to osteoclasts. This was clearly shown for bone explants cultured with M199 (4). By using other media (e.g., BGJb medium), precipitation of crystals may occur at these sites. 5. A problem recognized for a long time is the loss of mineral due to handling and cutting of mineralized tissue. One has to be aware that during cutting of sections and their collection on water, mineral may dissolve. This is particularly the case if sections are collected on water with a low pH. Landis and coworkers (5) suggested using other techniques to overcome this problem. 6. Mineralized tissues are characterized by a high electron density in mineral-containing parts of the sections. Yet, quite often it appears as if mineral is absent due to the absence of a strong electron density in such sections (Fig. 5). This problem is often due to the thickness of the section; the thinner the section, the lower the electron density will be. If one wishes to compare the same area in a section with and without mineral, it is possible to decalcify ultrathin sections. To accomplish this, sections collected on grids are floated on a drop of 0.1 M EDTA in buffer for 10 min. The sections are washed, counterstained, and examined. In sections thus treated, most mineral is dissolved but decalcification of mineral enclosed by epoxy resin (e.g., free in vacuoles) proves to be very difficult.

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Fig. 5. Electron micrograph of a section of undecalcified bone obtained from a control patient. Note the electron translucent area (asterisks) in the bone (B) part of the section. A high electron density (see area indicated by B in Figs. 2–4) characterizes a thicker section of the same area. Bone sample was fixed in 4% formaldehyde and 1% glutaraldehyde in 0.1 M sodium cacodylate buffer. N nucleus, CYT cytoplasm. ×15,500.

References 1. Gruber, H. E. (1992) Adaptations of Goldner’s Masson trichrome stain for the study of undecalcified plastic embedded bone. Biotech. Histochem. 67, 30–34. 2. Rungby, J., Kassem, M., Eriksen, E. F. and Danscher, G. (1993) The von Kossa reaction for calcium deposits: silver lactate staining increases sensitivity and reduces background. Histochem. J. 25, 446–451. 3. Reynolds, E. S. (1963) The use of lead nitrate at high pH as an electron opaque stain in electron microscopy. J. Cell Biol. 17, 208.

4. Everts, V., Korper, W., Jansen, D. C., Steinfort, J., Lammerse, I., Heera, S., Docherty, A. J. P. and Beertsen, W. (1999) Functional heterogeneity of osteoclasts: matrix metalloproteinases participate in osteoclastic resorption of calvarial bone but not in resorption of long bone. FASEB J. 13, 1219–1230. 5. Landis, W. J., Paine, M. C. and Glimcher, M. J., (1978) Use of acrolein vapors for anhydrous preparation of bone tissue for electron microscopy. J. Ultrastr. Res. 70, 171–180.

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Chapter 24 Scanning Electron Microscopy of Bone Alan Boyde Abstract This chapter described methods for Scanning Electron Microscopical imaging of bone and bone cells. Backscattered electron (BSE) imaging is by far the most useful in the bone field, followed by secondary electrons (SE) and the energy dispersive X-ray (EDX) analytical modes. This chapter considers preparing and imaging samples of unembedded bone having 3D detail in a 3D surface, topography-free, polished or micromilled, resin-embedded block surfaces, and resin casts of space in bone matrix. The chapter considers methods for fixation, drying, looking at undersides of bone cells, and coating. Maceration with alkaline bacterial pronase, hypochlorite, hydrogen peroxide, and sodium or potassium hydroxide to remove cells and unmineralised matrix is described in detail. Attention is given especially to methods for 3D BSE SEM imaging of bone samples and recommendations for the types of resin embedding of bone for BSE imaging are given. Correlated confocal and SEM imaging of PMMA-embedded bone requires the use of glycerol to coverslip. Cathodoluminescence (CL) mode SEM imaging is an alternative for visualising fluorescent mineralising front labels such as calcein and tetracyclines. Making spatial casts from PMMA or other resin embedded samples is an important use of this material. Correlation with other imaging means, including microradiography and microtomography is important. Shipping wet bone samples between labs is best done in glycerol. Environmental SEM (ESEM, controlled vacuum mode) is valuable in eliminating “charging” problems which are common with complex, cancellous bone samples. Key words: Mineralisation, Morphology, Osteoporosis, Mouse genetics, Micropetrosis

1. Introduction Scanning Electron Microscopy (SEM) uses a finely focussed beam of electrons scanned across a sample surface. There are many choices of imaging modes and levels of automation. I will first describe features of the microscopes and methods important for SEM imaging of bone before going into more detailed protocols in the method sections.

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_24, © Springer Science+Business Media, LLC 2012

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1.1. Instrumentation and Imaging Modes

SEM instruments currently in production use digital scan patterns, i.e. the beam is moved to a point, stopped, a measurement of one of the available signals is made at that point, and then the beam is moved on to the next point, eventually building up an image from all the scanned points. Digital scanning is important if valid stereological measurements of properties are to be made. This is a major application of SEM imaging in bone research by probing variations in mineralised tissue matrix properties in terms of mineral content. Scanning can also have another meaning namely: moving the sample. Stage automation allows imaging of very large samples at high resolution by scanning an XY matrix of fields. These can be used to build up a large montage image, which are particularly useful in correlating SEM with other, lower resolution, imaging methods including “clinical” and laboratory optical and X-ray methods. Imaging methods for bone are backscattered electron (BSE) imaging, secondary electron (SE) imaging, and imaging using X-rays emitted from the sample. The instrument specifications required depend largely on the imaging method used and careful consideration should be given to these when purchasing or using an instrument for making quantitative measurements. For the most useful application of SEM in bone and teeth backscattered electrons are imaged and it is critical to use an EM with genuine digital scanning (not a digitised analogue signal as is the case with many instruments currently in the field) with fully automated operation, in a very stable system where the signal does not fluctuate with time on the short- or the long-term time scale. To date, SEMs with tungsten filament sources have been the most stable. In addition, the SEM must have a BSE detector which can view the sample surface at normal incidence, which means using a detector that surrounds the electron beam. It will usually be a solid-state annular detector in the “roof” of the SEM specimen chamber. Please note that “in-column” detector systems will give a lower solid angle for collection of the backscattered electrons. BSE imaging allows the study of relative mineral content at less than 1-μm3 volume resolution by using high energy electrons reflected from the sample (1). The strength of this signal is proportional to the mean atomic number “Z” of a mixture of elements. Fortunately, bone and other calcified tissues have a composition with more high Z elements than other tissue matrices, allowing Z-dependent compositional imaging and topography-dependent 3D imaging. Although BSE is the single most useful imaging mode in bone SEM, by far the commonest imaging method in biomedical SEM is the use of the low energy SE signal. In this imaging mode most electrons emanate from the sample coating (usually gold or another high Z material) on a dried sample surface. This means one is looking at the gold coating, not directly at the specimen. In addition, the specimen itself is most likely to be deformed due to shrinkage during sample drying. Despite all this, useful

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information can be obtained and methods to do so are given below under Subheading 3. A third commonly used signal in SEM is that of the X-rays emitted under electron bombardment, particularly the characteristic X-rays which permit elemental identification. Historically, this is where SEM of hard tissues began. The electron image was used only to locate where the electron beam was hitting the sample. Unfortunately, the X-ray emission volume is far larger (deeper and wider) than that of either the BSE or SE image used for location purposes (1). It is possible to measure the X-ray count and the electron beam current on one spot on the specimen, but it cannot be determined whether the X-rays came from that spot or from locations deeper in or aside from the target area (see Note 1). To avoid this problem you can use either thin sections so that the electron beam has little chance for sideways scattering, or you can isolate a small part from the tissue by a combination of micro-dissection with EDX in the SEM (2). Neither of these two approaches is implemented today in systems that can be described as SEMs, so we will leave them aside. A number of other methods are available in the SEM, but have yet to find more general application. They are briefly mentioned here, to help you understand what they can do, but we do not discuss these in the method sections. Specimen current imaging measures the difference between the current flowing to earth through the sample at different locations due to differences in backscattering. It might well have extremely useful applications in bone implant research but is not instituted as a major feature in modern biological SEMs. Light emitted from a sample under electron bombardment is called cathodoluminescence (CL) (see Note 2). This has found little application in the general realm of biomedical SEM but may yet make a comeback due to the wide range of applications of fluorescent tags in immunohistochemistry. It does have specific applications in the bone field in the study of mineralisation-front-seeking labels such as tetracycline (3). However, combining confocal LM fluorescence imaging with SEM is a strong competitive method and as this is more widely available this is described in the methods section. No special sample preparation is required for CL, but there is for sample imaging (4). Environmental SEM (controlled vacuum SEM) allows examination of wet or damp samples without coating, but wet SEM has no particular advantages for the study of bone and bone cells. Low kV and video rate scanning also permit the study of uncoated samples: combined with real-time stereoscopic visualisation by deflecting the electron beam, this has been used to study ultramicrotomy of bone and dental tissues in the SEM (5). Control of the sample chamber vacuum also permits the study of imperfectly coated, dry samples.

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1.2. Types of Samples and Interpretation of SEM Data

A large amount of information can be obtained by examining bone in the SEM. Correct interpretation of this data is important and below I give a general account of the type of information that can be extracted and the types of samples that are best suited for a particular purpose. This section will set the scene for the methods that are described in Subheading 3.

1.2.1. Unembedded Bone, with 3D Surface Detail

Success will be greatest with tabular bones, for example, the endocranial surface of the rodent calvarium, where osteoblasts, osteoclasts and ex-osteocytes liberated by osteoclastic action will be easily identified (Fig. 1a, b). Beware of identifying thinner exosteoblasts as putative resting or bone lining cells since cells may be flattened by preparative protocols. They may be so thin that the lamellar bone matrix pattern beneath them is imaged instead of the cells, particularly when using BSE. Adipocytes are common bone lining cells in established bone. Often only the sides of the fat cells contacting the bone matrix remain in the SEM sample, when again the bone matrix pattern beneath them is seen. When imaging natural surfaces with osteoid but without cells on the surface it is possible to visualise the newest osteoid packets (that do not contain any mineral as yet) and to judge the relative maturation ages of the more recently mineralised packets (due to their lower mineral content) by virtue of their reduced electron backscattering properties (Fig. 2). Surfaces where all cells and matrix are removed are referred to variously as anorganic, deproteinised, or as mineralising front preparations (6, 7). Such samples are the easiest to prepare and are extremely rich in information relevant to bone biology, as most

Fig. 1. (a) Neonatal rat parietal endocranial osteoblasts, CPD, gold sputter coated, 10 kV SE, Field width 87 μm. (b) Osteoclasts liberate live osteocytes. Nenatal rat parietal endocranial surface showing several osteoclasts and formerly encapsulated osteocytes liberated by osteoclastic resorption. CPD, gold sputter coated, 10 kV SE SEM. Field width 167 μm.

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Fig. 2. Bone packet distribution by BSE-SEM: more recent packets have darker grey levels because they are less well mineralised. Note that they do not fit the prior resorption fields. Sample of vertebra, L4 of 61-year male. Scale bar in image.

activity occurs at surfaces. Due to the relatively large depth of field of SEM images, it is possible to study cancellous surfaces with great efficiency, and if a sample is cut at the right thickness and studied from both sides, probably more than 80% of the real bone surface (and all the cut surface) can be characterised (Fig. 3). We can distinguish various bone surface activity states in anorganic or superficially anorganic preparations (7, 8) as below: Mineralising surfaces are those where the collagen replicas in the mineralising front appear to be discontinuous because mineralisation was still proceeding. There are sub-classes in this category, including 1. Actively mineralising lamellar bone, where the branching bundle pattern characterising all lamellar bone matrix is recognisable, but the mineralised segments are separated. 2. Nearly completely mineralised mineralising lamellar bone, where mineralised segments are only slightly discontinuous. 3. Mineralising woven bone, recognised by the lack of association of the mineral with collagen bundles, and by the presence of innumerable small micro-calcospherites. 4. Mineralising Sharpey fibre bone where the insertion sites of the extrinsic, penetrating fibres are below the level of the intervening intrinsic matrix. 5. Calcifying growth and articular cartilage mineralising fronts and fibro-cartilaginous mineralization (9). None of these are true “bone,” yet they are found within bone organs.

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Fig. 3. Very low magnification (wide field) 1 kV SE image. Samples can be imaged without coating or with imperfect coating at this very low voltage. 4-mm thick slice of L4 vertebra of 30-year female. 2.7-mm grid background.

Mineralised or resting surfaces are those where the collagen fibre bundles and fibrils of the bone matrix appear to be intact in spite of the fact that the collagen has been completely removed: this proves that mineralisation stopped at the current limit provided by the extent of the matrix. Prolonged resting surfaces are those where the level of mineralisation has extended beyond the collagen per se and the image of the fine detail of the collagen is partially obscured because the overlying “ground substance” is mineralised. Resorbing or resorbed surfaces are characterised by the presence of excavations made by osteoclasts. Many small excavations or pits clustered together constitute a “Howship’s lacuna.” Resorbing and resorbed surfaces can be sub-categorised between extremes where the osteoclast must have been alternately resorbing and translating (with obvious linear tracks or trails of adjacent shallow scoops) and deep pits excavated directly into the surface. The latter evidence of active resorption in depth will be found in association with rapid removal of bone during growth and drift. Good examples are found in jaw bones in association with growing teeth, or with tooth sockets moving through the alveolar bone. Others are seen in the aggressive resorption associated with cancers in bone. Very recent or still active resorption can be diagnosed if there is demineralised matrix in the pit. In anorganic samples, a recently excavated, or still active, pit will have a rougher texture, compared

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with pits where the demineralised collagen fringe made by the osteoclast has been cleaned up. This cleaning up process, which is usually by cells other than osteoclasts, leaves the pit with a smooth bottom. Surfaces generated by fracturing bone may be studied in their own right to learn something about mechanisms of bone failure. However, breaking bone by fracturing a whole bone organ is not recommended as a preparative protocol because the wet tissue tears rather than breaks. This results in the ripping of collagen fibrils and bundles out of the surface. These artefacts can be partially prevented by freezing the bone to liquid nitrogen temperature before cracking it by driving wedges into appropriate grooves cut before freezing. Even smoother breaks may be generated if bone is made anorganic before being frozen, because then the collagen has no influence on the fracture process. The smoothest broken surfaces will be generated by substituting anorganic bone with ethanol, which freezes as a glass under liquid nitrogen. Dried whole bone that contained osteocytes can also be fractured, but remarkably, osteocytes are rarely found on surfaces of bone which was broken dry. To view natural surfaces inside bone organs, it will be necessary to cut away some bone. It is best to polish the cut surface to remove the tags of torn bone created by the cutting process itself, and this is best done after dehydrating the tissue in alcohol to harden it (see Subheading 3). A special class of unembedded calcified tissue is the slice of bone, or of dentine, used as a resorption assay sample (10). 1.2.2. Resin-Embedded Block Surfaces, Topography-Free, Polished or Micromilled: qBSE

To use the BSE SEM signal to study tissue composition (density: mean Z: BSE coefficient) as against surface topography it is necessary to get rid of the latter component as far as possible. This can only be done in embedded samples where prior water-filled space is filled with an embedding resin. We use PMMA, but the inclusion of styrene results in a resin that is better suited to withstand electron bombardment (Causton, personal communication, 1980). An excellent optical flat finish can be obtained by diamond micro-milling, but this is a tedious process and requires expensive equipment and skilled labour, so we will not consider it further here, albeit that it is by far the best procedure.

1.2.3. Resin Replicas or Internal Casts of Space in Bone Matrix

Bone is a typical connective tissue in that most of bone tissue is matrix and cells are the minority component. The osteocytic lacunar and canalicular space compartment in mature bone can be as little as 1% by volume. The 99% of solid mineralised bone matrix is not transparent to electrons and will prevent us seeing the cells which were the living part of the tissue. It will therefore be convenient to etch away the bone to leave the osteocytes and their canalicular processes, or to see the blood vessel canal space. This can be

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achieved by destroying the calcified bone matrix in resin-embedded bone. The remaining cast demonstrates the 3D volume within which the osteocytes resided (and indeed they still reside in the cast). Remember that the cast does not show the shape of the osteocyte body or its canalicular processes: it instead show the limits of the fluid filled space in which they sat, perhaps making quite a loose fit.

2. Materials 2.1. Chemicals

1. SEM fixative: 2.5% Glutaraldehyde and 1% formaldehyde in 0.15 M sodium cacodylate buffer at pH 7.3. To make a 10% formaldehyde solution, dissolve 2.5 g paraformaldehyde in 25 ml of water, heating to 60°C whilst stirring. This MUST be done in a fume hood. Do NOT allow the solution to boil. The solution will be milky. Clear it by adding one or two drops of 1 M NaOH. To prepare the final fixative add 2 ml of 25% glutaraldehyde solution (Agar Scientific, http://www.agarscientific. com) to 2 ml of 10% formaldehyde, 15 ml of 0.2 M sodium cacodylate and 1 ml water, to make 20 ml of fixative. 2. Phosphate-buffered saline. 3. Industrial methylated spirit (IMS). 4. Chloroform water. Prepare chloroform water by adding a few millilitres of chloroform to distilled water in a conical flask, which then place in an ultrasonic cleaning bath for 30 min. Pipette or pour off the supernatant at the time of use. You can store it forever. 5. Tergazyme, an alkaline solution of bacterial pronase (Alconox Inc., New York, NY, USA). 6. NaOCl bleach solutions. These are best obtained fresh from the domestic grocery store, where there is a much greater stock turnover than in the chemical supply houses. 7. C-dag (Agar Scientific). 8. Carbon putty (Agar Scientific). 9. Plasticine. 10. Freon 12 (dichloro-difluoromethane). 11. HMDS (hexamethyldisolazane).

2.2. Instrumentation

Instrumentation for SEM viewing is not included here, general considerations on equipment have been discussed in Subheading 1.1.

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1. Low speed saw (Buehler Isomet http://www.buehler.co.uk, or Bennett LabCut, Agar Scientific). 2. Straight tungsten carbide milling tool. 3. Tungsten carbide plain cylinder milling tool (e.g. ISO 500 104 107175 023 (H259) from dental supply houses). 4. Rotary polisher (e.g. TexMet 2500 from Buehler http://www. buehler.co.uk or Plano-Cloth M from MetPrep Ltd, sales@ metprep.co.uk) with aqueous 1- and 6-μm diamond suspension. 5. Specimen levelling press (e.g. from Agar Scientific). 6. Carborundum paper (various grades down to at least 1,200 grit). 7. Critical Point Dryer (e.g. from Agar Scientific). 8. Sputter coater for gold and/or carbon (e.g. from Agar Scientific). 9. Freeze drying apparatus, vacuum equipment with a temperature-controlled stage at approximately −65°C, such as for example the Edwards Speedivac Pearse tissue dryer. 2.3. Software

1. PSP5 from Jasc Software Products (free software). 2. MeX software (Alicona, http://www.alicona.com/home/ products/Mex/MeX.en.php). 3. Tview.exe (http://www.skyscan.be). 4. AnaglyphView (http://www.alicona.com/home/support_ sales/Downloads.en.php), freely available. 5. Auto-Montage from http://www.syncroscopy.com.

3. Methods We will consider a variety of methods that will allow examination of bone cells, bone tissue with or without cells, and resin casts. A number of methods are general to many of the procedures and these are described first. 3.1. Cutting Bone and Bones 3.1.1. Large Bones

3.1.2. Small Bones

Use a band saw to reduce bulk frozen body parts including bones in size whilst still frozen. Do not use a band saw if you need to reduce the size of the bone before starting on an embedding procedure (see Note 3). Use a milling tool to pare away the tissue from one side to reveal the internal microarchitecture of trabecular (cancellous, spongy) bone regions. Milling is the preferred way to do this as the cut

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surface can follow the anatomical curvatures of long bones. Practice this technique with bones which are not a part of your major experiment since you cannot put back what you have cut away. 3.1.3. Large Slabs of Large Bones

1. Use a low-speed diamond saw under water or IMS cooling and lubrication. Sparse trabecular bone such as in human lumbar vertebral bodies from aged individuals should be cut as 3-mm thick slabs. Denser cancellous bone such as the distal epiphysis in equine cannon bone should be cut at 1.5–2 mm. 2. Wash slabs with water. 3. Substitute water with IMS to harden the bone slabs. 4. Hand polish the slabs on wet-and-dry silicon carbide papers placed on a backing glass sheet, using IMS as lubricant: usually 1,200 grade paper will provide a good enough polish. Wear tight rubber gloves or finger stalls.

3.2. Fixation of Cells or Tissue for SEM

The first step preceding fixation will be to wash away any proteinaceous fluid such as enriched culture medium or serum or blood. Then cells must be preserved against any morphological change due to fixation and drying. Mixed aldehyde fixation adds stability to cell membranes to withstand the rigours of solvent substitution and drying. 1. Wash samples with phosphate-buffered saline at 37°C. 2. Submerge samples in fixative (2.5% glutaraldehyde and 1% formaldehyde in 0.15 M sodium cacodylate buffer at pH 7.3), initially at 37°C, for 2–48 h. You may want to consider addition of tannic acid (see Note 4), but be aware that this may cause some slow demineralisation. 3. If necessary to hold the tissue before drying, it is best to transfer it to a solution of the same buffer used to support the fixative. 70% ethanol is a good alternative to using a buffer as it prevents both tissue swelling and shrinkage.

3.3. Drying Samples 3.3.1. Freeze Drying

Freeze drying (FD) is the sublimation of frozen water ice (see Note 5). This method requires availability of a suitable tissue freeze drying apparatus, such as for example the Edwards Speedivac Pearse tissue dryers. 1. After fixation, wash samples briefly with distilled water. 2. Place in chloroform water for 30 min. 3. Prepare a freezing mixture difluoromethane). See Note 6.

of

Freon

12

(dichloro-

4. Freon 12 gas is condensed by running it through a section of copper tube cooled by boiling liquid nitrogen in a large thermos vacuum vessel, allowing it to collect in a suitable container

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such as a Pyrex glass crystallising dish. The Freon 12 will solidify at −155°C. 5. Remove the dish of solid Freon 12 from the LN2 and melt a little of it, for example, using a stainless steel instrument. 6. Place the samples in the liquid Freon 12 and hold them ducked under for a moment. 7. Then transfer the frozen samples with the attached layer of Freon 12 liquid to the liquid nitrogen where they will be held until the next step. 8. Finally, transfer all the samples to a holder which will fit your tissue dryer and pump overnight at −65°C. 9. Raise the temperature in steps until ambient. 10. Remove and mount samples and coat for SEM examination. 3.3.2. Critical Point Drying

Critical point drying (CPD) involves the conversion of a liquid to a gaseous phase by “going round the critical point in the phase diagram” where and when there is no surface tension barrier to cause shrinkage (see Note 7). This requires availability of a CPD machine. 1. Most commonly, tissue water is substituted with ethanol. 2. Ethanol is substituted by admitting liquid CO2, at room temperature, to a committed pressure vessel, the CPD chamber. Liquid CO2 is allowed to escape intermittently to flush dissolved ethanol from the chamber. 3. The chamber is sealed and its temperature is raised to 35–40°C, above the critical temperature and automatically also above the critical pressure for CO2. 4. The high pressure CO2 gas is allowed to escape, leaving the samples dry and ready for mounting and coating.

3.3.3. Air Drying from Volatile Solvents

Air drying from volatile solvents (AD) produces results similar to those obtained with CPD. This is a much simpler technique and involves the evaporation of a very low surface tension solvent used to substitute tissue water after substitution with an intermediate solvent such as ethanol. Dry cleaning solvents, such as Freon 113 (11) are excellent for this purpose but not currently favoured because they are halogenated hydrocarbons (see Note 8). In common use today is hexamethyldisilazane (HMDS). 1. Transfer the fixed and ethanol-substituted samples to HMDS. Allow time for this to exchange. 2. Lift out the samples, place on black filter paper to blot dry and allow the liquid to evaporate (which will take 5–10 min depending on sample size), preferably in a fume cupboard. Alternatively, pipette off surplus HMDS from the vessel holding the sample(s)

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and allow to dry. HMDS is very volatile and you might expect to allow 5–30 min depending on the sample thickness. 3. Mount and coat for SEM (see Note 9). 3.4. Coating

When we want to understand the shape of cells and are not interested in imaging the substrate we prefer a heavy metal (high Z) coating such as sputtered or evaporated gold. Imaging can be done by either SE or BSE.

3.4.1. Sputtering of Gold

Sputter coaters will be found in any biological SEM lab. Follow the instructions provided with the individual coating unit or from the technical staff in the lab. Importantly, we always start with a short burst of coating lasting only one or a few seconds at a low coating current, and allow the sample surface to cool again before continuing with the main coating operation. This avoids overheating and potentially melting the coatable surface.

3.4.2. Evaporation of Gold

This is achieved by wrapping gold wire around a 0.5-mm diameter at a hairpin bend in a tungsten wire filament, through which a large current is passed. Pre-prepare the filament by melting the gold on to it on a separate run before placing the samples in the vacuum chamber. Then when you reheat it the filament will only warm to red heat while the gold boils off, thus saving thermal damage to the samples. Tilt and rotate the samples during coating to get better coverage of more surface. When imaging both the cell surface and the substrate underneath, carbon coating is preferred as it allows imaging of the cell surface under low kV SE SEM conditions (e.g. 1 kV), but makes the cells transparent when imaging the underlying matrix using BSE at higher kV (e.g. 20 kV).

3.4.3. Carbon Coating

To coat samples with carbon by evaporation, use an arc struck by passing a heavy current between two carbon rods, one of which is a 1/8th-in. rod thinned to 1/16th-in. over ¼-in. Whilst evaporating, tilt and rotate or otherwise move the samples to aim to get some carbon on all surfaces. The required equipment would be usually found in labs specialising in TEM work and is used for evaporating carbon on to glass sheets (see Note 10).

3.5. Morphological Imaging of Cells on Bone in 3D

We will now consider procedures for specific types of samples, starting with cells on a surface. The types of sample we shall consider here are those with cells on a hard substrate. The latter might be flat bone, for example, the endocranial surface of rat or mouse calvarium, a metal or ceramic implant material or a glass cover slip with cells grown in tissue culture conditions. If cells in situ on a substrate are your interest, then you have the most serious problem in sample preparation from the whole of the biomedical field, since the cells will shrink and the substrate

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will not and the relative deformation will lead to artefacts. Mostly this will consist of separation between cells in the (in real life continuous) sheet on the hard substrate, as well as flattening of the cells onto the substrate. Compare this with SEM of all other biological soft tissue systems where cells, or sheets of cells, shrink together and with their matrix and where the preparative artefacts are often missed or overlooked. The inevitable volume shrinkage is also generally ignored (12, 13). 1. Preserve the samples as described in Subheading 3.2. 2. Dry the sample using one of the methods described in Subheading 3.3. 3. Coat the sample as described in Subheading 3.4, use gold for SE imaging and carbon for BSE imaging. 3.6. Examining Undersides of Bone Cells in Contact with Substrate

To view the contact sites of osteoblasts and osteoclasts with bone: 1. Take a well-prepared FD, CPD or AD sample of a flat bone (e.g. rodent calvarium prepared as in Subheading 3.3). 2. Apply a tiny strip of adhesive tape such as double-sided adhesive tape (Sellotape) or conductive carbon tape to a hemispherical headed 1/8th-in. aluminium rivet, touching the surface with a rolling motion to strip off the surface cells. 3. Coat the resulting cell-free bone surface with gold as described in Subheading 3.4 for SE imaging in SEM.

3.7. Morphological Imaging of Bone Alone in 3D

Probably the most common and the most attractive use of SEM imaging in the bone field is to look at the calcified bone proper as a 3D object. Here I describe the preparation of bone and other calcified tissue matrices for 3D morphological imaging. This involves cutting bones, removing cells and soft tissue and uncalcified bone matrix and finally making samples anorganic. Speed of fixation is important in all branches of microscopy where we want to preserve cells. However, in many applications of SEM to study the bone itself this is less important and we often use samples that have been deep frozen as a whole organ successfully (see Note 11). First we have to reduce the samples to a suitable size as described in Subheading 3.1 and then we remove cells from the bone to reveal the bone matrix itself. There are several methods to do this and they are all described below. A range of imaging methods will be explained and illustrated further below, one trick being to lodge square-cut pieces of prepared bone in cubic indents in a lead block to achieve an “invisible” background in BSE imaging mode, making the samples apparently float in space (Fig. 4).

3.7.1. Removing Cells and Soft Tissue and/or Uncalcified Bone Matrix with Tergazyme

Maceration with Tergazyme, an alkaline solution of bacterial pronase, removes most soft tissue but leaves osteoid. Such samples are very robust. It is important that the samples are not previously

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Fig. 4. These transverse section blocks (~2.5 mm × 3.7 mm) of a longitudinal beam cut from an 18-year-old horse distal third metacarpal bone are coated with carbon and simply lodged in square indents punched in a lead block, which is not seen because the high signal level appears as a white background. 20 kV BSE.

fixed in formaldehyde or glutaraldehyde but the procedure usually works with tissue fixed (and stored) in 70% ethanol. 1. Clean samples of bulk muscle by careful dissection. 2. Open internal structure as required by cutting and polishing large slabs or using a straight tungsten carbide milling tool for small bones. 3. Immerse samples in a 4% Tergazyme solution in water at 50°C for periods of 1–2 weeks for mouse long bones and up to several months for large slabs of human or horse bone. A reliable sign of success with young long bones will be the separation of the epiphyses caused by the removal of the uncalcified growth plate cartilage. Tergazyme treatment will usually remove hyaline articular cartilage to the level of the mineralising front of the calcified cartilage. 4. Rinse in distilled water. 5. Dehydrate in IMS and allowed to air dry. 6. Coat with carbon for BSE SEM, obtaining 3D images by tilting (see Subheading 3.9.2). Make sure that slices of trabecular

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bone are carbon coated from both sides. Mount them on tiny pyramids of conductive carbon putty so that they can be turned over and remounted to look at the other side. 3.7.2. Boiling Bones to Remove Cell, Fats, and Non-mineralised Matrix

Boiling bones by simmering in a large pot of water is a commonly used preparation technique for large anatomical specimens. Although this is a process difficult to control it can work well. Adding detergent can help to disperse the fat content of the bone organ (see Note 12). Normally one will expect non-mineralised matrix to be removed first, leaving a mineralising front preparation of the bone.

3.7.3. Burying Bones to Remove Cells and Soft Tissues and Using Buried Bone

Delightful instructions in a nineteenth-century textbook of equine anatomy tell you to place the bones in a pile of rotting horse manure, which will provide the requisite microbiota, proteolytic enzymes and temperature to remove cells and soft tissues. No doubt such samples will be superficially anorganic while remaining very robust. Pieces of bone buried in well-drained soil can be very clean, and “ready to go” for SEM and have been used extensively by forensic scientists and archaeologists. Often the internal trabecular architecture will appear to be cleaned well enough for SEM study and can show fine detail to distinguish forming, resting, or resorbing surfaces. However, it is an amazing fact that such bones can be largely substituted from within and contain innumerable minute cavities and tunnels caused by soil fungi and bacteria.

3.7.4. Making Samples Anorganic Using Hypochlorite

Small animal bones can be cleaned of soft tissue and made anorganic by treatment with sodium hypochlorite (NaOCl) bleach solutions (see Note 13). 1. Dilute the concentrated NaOCl solution which is usually labelled as 14% available chlorine with water (1:1). This solution is very corrosive: wear rubber gloves and be extremely careful not to splash. 2. Place the defleshed samples in this solution at room temperature using loosely covered containers from which gas can escape. Using a rocking or tilting table will be an advantage. Label the containers so that others know the hazard. Small rodent bones will be clean within 24 h, and although intact, will be exceptionally delicate to handle when wet. 3. Wash the samples several times in distilled water to reduce the residual NaCl salt concentration. This is best done by pipetting off the solution and pipetting in distilled water. 4. Lift the samples with a small spatula from the container and place them on black filter paper to blot dry. If they are too fragile to handle, pipette off as much fluid as possible and let them dry in the container. Very small bones can be sucked into specially fashioned glass pipette tips (see step 6 below) to help transfer to blotting paper.

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5. Dry bones in air. Using an oven (37–50°C) will speed up the process. Remaining salt is drawn back and crystallises inside the sample during air drying and does not show at the sample surface. If you have left the bones in solution at step 2 for long enough there will be no organic matrix left and they cannot shrink on drying. 6. Mount the samples on carbon conductive tape. As samples may be extremely delicate and difficult to handle use of a custommade tool will help. Melt the end of a bacteriological glass pipette in a bunsen flame to make a small solid sphere. Rub this against your clothing and hold it over the samples, which will jump on to the glass because of the electrostatic charge. Now place them close to the carbon conductive tape, where they will transfer. Samples will sink into carbon conductive tape and become ever more firmly attached. This useful process can be speeded using an oven as in step 5. 7. Carbon coat for 3D BSE SEM imaging as described in Subheading 3.4.3. 3.7.5. Making Samples Anorganic Using Hydrogen Peroxide

Bones refractory to cleaning with Tergazyme can be treated using a hydrogen peroxide solution to finish off the cleaning process. H2O2 will remove non-mineralised collagen in strong preference to mineralised matrix, leaving samples with mineralising front morphology which are nevertheless quite robust. 1. Dilute the 30% H2O2 stock solution with up to six times as much water, depending on how often you are going to inspect progress (overnight in the former and up to some days for the latter). 2. Store bones and solution in pots with loose or perforated lids to allow the escape of gas. As H2O2 solutions are extremely corrosive, take great care and wear protective gloves. Label containers clearly to warn others of the danger in a multi-user environment. 3. Wash bones with distilled water and air dry on black filter (blotting) paper, in an oven at 37–50°C, or dehydrate to IMS and air dry from there. 4. Mount the dry samples on carbon conductive tape and carbon coat for 3D BSE SEM imaging as described in Subheading 3.4.3.

3.7.6. Making Samples Anorganic Using Sodium or Potassium Hydroxide

Treating bones with strong solutions of KOH or NaOH at 50°C will remove all soft tissue and matrix overnight, so if you are in a great hurry, you can try it. The solutions are dangerous to handle. Use gloves and care. You will have to use several water washes by pipetting to remove the alkali. The samples will be anorganic and delicate, as with hypochlorite treated tissue. Allow the bones to

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air dry from water or IMS and proceed as described under Subheading 3.7.4, step 6 onwards. 3.7.7. Making Samples Anorganic Using Sodium Peroxide

Solutions of Na2O2 at 50°C are very aggressive (and must be handled with extreme care), but are the fastest means of cleaning bones to leave anorganic samples. As with NaOCl or alkalis the samples are very delicate. Allow the bones to air dry from water or IMS and proceed as described under Subheading 3.7.4, step 6 onwards.

3.8. Bone and Dentine Slices Etched by Osteoclasts In Vitro

Bone slices may be used for in vitro studies of osteoclastic activity, including the harnessing of osteoclasts to etch the bone structure. Osteoclasts (OCs) seeded on bone are masters in revealing the underlying lamellar structure. No method invented by man has done better to date. To study whether there are functional osteoclasts in a mix of cells or to measure their activity, dentine slices are chosen in preference to bone because of the greater continuity of this tissue (10) (see Note 14). The presence of OC pits in the surface can be seen by reflected light microscopy, by staining for transmitted light microscopy (14), and by confocal reflection microscopy, when the depth and volume of the pits can also be determined (see other Chapters on osteoclast formation and resorption in this volume on alternative quantification methods). There are several SEM imaging options for pits (Fig. 5a, b).

Fig. 5. (a) Resorption pits made by mouse osteoclasts in sperm whale dentine, cells removed, coated carbon, 20 kV BSE image. Field width = 900 μm. Resorbed – or just demineralised! – areas are darker. In this example, there are extensive areas where the surface has been attacked by osteoclastic acid but the depth of damage is very slight. (b) Resorption pits made by chick osteoclasts in sperm whale dentine. After removing cells, sample was stained briefly with silver nitrate solution. Silver is deposited in the demineralised collagen fringe of the resorption pits, and where this is thickest, in the most recently formed pits, the signal (due to both SE and BSE) is greatest. Gold sputter coated, 10 kV image with standard Everhart–Thornley biased scintillator “SE” detector. Field width = 194 μm. In the author’s opinion, assays should use a 3D approach to measure pit depth and volume (see refs. 14–17).

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3.8.1. Morphological SE SEM Imaging of GoldCoated Samples

As with all SE imaging, edges parallel with the electron beam direction (normal to the sample surface) appear to be bright: this is due to the increased depth of surface from which SE can escape, and most SE are captured by the strong positive field applied to the grid at the front of the SE detector. Take stereo pairs.

3.8.2. Topographical BSE SEM Imaging of CarbonCoated Samples

This can be A–B mode where signals from opposing detector segments of a 2-sector BSE annular detector (or pairs of opposing detector segments of a 4-sector annular detector) are differenced. These images convey a strong sense of depth, but real pits will appear to be bumps if the image is rotated, so be careful to keep the rotation sense. Using map notation, it is generally preferred to have the apparent direction of illumination (the direction of a positive signal detector) North or North West: the same applies for SE imaging. An attractive alternative with a four sector annular BSE detector is to set one sector off or negative. This should be that corresponding to South East (bottom left).

3.8.3. Compositional A + B BSE SEM Imaging of Carbon-Coated Samples

This is the preferred and a very sensitive method for simply seeing pits and measuring their area. The method partially relies upon the fact that in vivo the demineralised collagen fringe made during the acid secretion phase of the osteoclastic activity is removed by “other” cells which are not present in the in vitro assay. Thus all pits and would-be pits show a layer of decalcified matrix, which appears dark in compositional BSE imaging (Fig. 5b). A practical point here is that this “darkness” must be distinguishable from dark regions of topographic BSE contrast originating from the roughness of the cut surface. Therefore use smooth slices (see Note 14). 1. To prepare a bone or dentine slice for SEM, remove all the cells, but preferably not the demineralised collagen fringe made by OC acid. Do not use a fixative, but immerse the slices in distilled water, which will disrupt the cells by osmotic shock. 2. Gently rub the slice between wet thumb and forefinger. 3. Wash and air dry the samples. 4. Mount samples the right way up on carbon conductive tape. 5. Coat with carbon by evaporation. 6. Image using 20 kV BSE SEM (see Note 15 and Fig. 5b). 7. Use compositional BSE SEM imaging to identify pits and sites where OCs are about to make pits. The method delineates the areas hit by OC acid, but remember it does not measure the work function of OCs as this depends upon progress perpendicular to the surface under attack, i.e. the depth of the pit (15). Measuring the pit volume is necessary to measure the rate of work of OCs. To do this use stereoscopic imaging.

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8. Decide whether you want the volume with or without the “fringe.” The latter is simpler, since now both cells and demineralised matrix can be removed by brief treatment with H2O2. 9. Take two images of each pit, or pit complex with a tilt angle difference of 10° and proceed to stereophotogrammetry (10, 16). 10. Commercial software, such as Mex, is available for processing digital SEM images to extract this information. 3.9. BSE SEM Imaging Methods

Bone (and tooth) samples for SEM truly resemble the original object far more closely than is the case for soft tissue material. Thus it is worth making an effort to show the 3D structure in such a way as to enhance information and make an impact on the viewer. With a digital SEM and BSE imaging of 3D macerated samples, one can usefully make image sets of the same field of view, varying either (a) the focus level, or (b) the angle of view seen by a single detector, or (c) the sector of view seen by different detectors.

3.9.1. Mechanical Scanning Through Focus

Although the SEM is renowned for its great depth of field, this is in fact severely limited. Image sets of trabecular bone in wide field or of forming or resorbing surfaces at higher magnification can be processed to extract the best focus found in any one part of any image of the stack, and retained in a composite all-in-focus image (17, 18). To make an all-in-focus image: 1. Take several images of the same field, changing the focus level by vertical Z movement of the sample, NOT by altering the lens currents. For example, should you have a 3-mm deep sample, take 11 images with a focus shift of 250 μm between each. If your field is 500-μm deep, take 11 images at 50-μm Z separation. This is trivial with stage automation. 2. Process the image stack using Auto-Montage software to derive the all-in-focus image. Please see ref. 17 for illustrations of all the steps and the resulting image. 3. Use the same processing to derive a 3D contour map showing which areas had the same distance/height.

3.9.2. Mechanical Tilting

By mechanically tilting the sample a number of times whilst recording one field of view, one can display images in sequence or build a movie file to show 3D relationships by exploiting motion parallax (18). Simple examples of these movies can be retrieved from, for example the Bone Research Society Web site (http://www.brsoc. org.uk/gallery/). 1. Bring the field of view to the eucentric (or a “compucentric”) position for the SEM stage. This means that you have found

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one working distance and a matching scan rotation value such that the image appears not to move when the sample is tilted. For the finest work, use 1° tilt angle differences and almost never less than 2.5°. 2. Display the image sequence back and forth using suitable software. The tview.exe programme from http://www.skyscan.be intended for use with microtomography image stacks is very good. Otherwise build an AVI or a GIF file using, for example Animation Shop in Paint Shop Pro 5 (free software). GIF files can be implanted directly into PowerPoint presentations and “go with the slide” (that means you do not have to remember or to track the file name). For simple stereo-pair imaging we would use 6° tilt angle difference. In this case, stereo pairs can be displayed as GIF files alternating the view every quarter second. Alternatively, they can be processed to make anaglyph (red– green or red–cyan) versions which are very nicely displayed on computer monitor screens and data projectors. The procedure for making anaglyphs using PSP5 is as follows: 1.

Assume that the sample was tilted “sideways” so that the two images constituting the stereo pair will require no rotation. The SEM scan rotation will have been set such that features move only in X or Y when the X or Y stage controls are used. You may need to rotate both images 90° if parallaxes lie in the wrong, i.e. the top to bottom, direction.

2.

If the images are recorded nicely in the first instance, features near the centre of the field of view will overlap in the two images, i.e. there is no parallax in X (stereo-parallax, the interocular axis direction, horizontal) or Y (vertical in the image plane). Jump to step 14.

3.

To overlay two images to remove Y parallax and to minimise X and choose the height which will appear to be that of the display screen, open both images. Let us suppose that they are TIF format – it does not matter as long as the format is known to PSP5.

4.

Copy one of them [for argument’s sake, the right, Rt, more tilted view] and close it again.

5.

Edit: Paste as new Layer: [Ctrl L].

6.

View: Toolbars: Layer Palette.

7.

Adjust Layer1 proportion using the slider to approximately 50%. You now see both images overlaid, and the target tool. Move this, which moves the Rt., Layer1 image, until features near the centre of the field of view overlap.

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8.

Use the crop tool to remove areas which do not overlap or which you do not want.

9.

Adjust Layer1 proportion using the slider to 100%.

10. File: Save as [F12]: use as file name the original file name+_b.TIF. 11. Adjust Layer1 proportion using the slider to 0%. 12. File: Save as [F12]: use as file name the original file name+_a.TIF. 13. Close the image. 14. Open filename _a.TIF [This is the right eye image]. 15. Open filename _b.TIF [This is the left eye image]. 16. Colours: Channel combining: Combine from RGB. 17. Choose Red Channel Source filename_b: Green channel source filename_a: Blue Channel source filename_a: OK. You now have your anaglyph image. 18. Save image as Anag_filename.JPG using Options: LZW compression: 10. If you are going to use it as a PowerPoint slide, you are best advised to reduce its size to 940 pixels wide by 710 high, copy it [CtrlC] and paste it [Ctrl V] directly into a blank slide. Do not use borders in Powerpoint since you thereby lose the image resolution you have so arduously striven to attain. Data projectors are as good as they are right out to the edge of the field of view. As an alternative, you can use AnaglyphView software following instructions as given in the programme. 3.9.3. Changing the BSE Detector Location (Discotheque Illumination)

1. Using a four sector overhead BSE detector, record four images of each field of view using only one of the detector sectors during image capture. Save them as filename1.tif, etc. where 1 indicates the detector segment used. An image is reconstructed using adjacent sector images as the red, green and blue components of an RGB image. 2. Using PSP5, open three images filename1.tif, filename2.tif, filename3.tif. 3. Colours: Channel combining: Combine from RGB. 4. Choose Red Channel Source filename1.tif: Green channel source filename2.tif: Blue Channel source filename3.tif: OK. You now have your colour coded for direction of apparent illumination image, in which colour now genuinely codes for sample surface slope and orientation (Fig. 6). 5. Save as filename123.jpg. The same remarks apply if you want to use your image in PowerPoint (see Subheading 3.9.2, step 18).

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Fig. 6. 20-kV BSE image of macerated, carbon coated, 3-mm thick section of L4 vertebral body from 89-year female. Field width = 4.7 mm. The same field was recorded with separate quadrants of an “overhead” solid-state BSE detector. Three images are used as red, green, and blue components of an RGB image. The result is that different slopes are seen as different colours and intensities. This method is described in ref. 18.

3.9.4. Changing the BSE Detector Location (In, Off, and Far Illumination)

An alternative procedure is to use three detectors (see Note 16), but here we will take it as the same detector positioned successively, to imitate a light source moving in a straight line across the sample (for an example see http://www.brsoc.org.uk/gallery/. picture osteoporotic bone). 1. Record a symmetrical overhead view with all detector segments on and positive: this will used as the Red component of the final RGB image. 2. Save it as filename_in.tif. 3. Move the detector off axis so that the electron beam can pass to one side of it, rather than through its centre. 4. Record a second image after increasing the detector gain – it is no longer collecting such a large solid angle of BSE. 5. Save it as filename_off.tif. 6. Move the detector further off axis and further increase the detector gain. 7. Record a third image. Save it as filename_far.tif. 8. Reconstruct an RGB image using PSP5.

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9. Open three images filename_in.tif, filename_off.tif, filename_ far.tif. 10. Colours: Channel combining: Combine from RGB. 11. Choose Red Channel Source filename_in.tif: Green channel source filename_off.tif: Blue Channel source filename_far.tif: OK. Again, colour genuinely codes for sample surface slope and orientation, red for overhead, green oblique from one side, and blue at grazing incidence from the same side. The image is easily interpreted (18). 12. Save as filename_IOF.jpg. See Subheading 3.9.2, step 18 about use of the image as a powerpoint slide. 3.10. BSE Imaging of Embedded Bone

3.10.1. Creating a Polished Surface (see Fig. 7)

Making flat block surfaces suitable for confocal microscopic imaging and production of casts: You may be fortunate enough to borrow PMMA bone blocks from another investigator, who has done all the painstaking graft of embedding and orienting and sectioning the bone for conventional bone “histomorphometry” (see Chapter 19 for details about PMMA embedding and histomorphometry). There are large numbers of human iliac crest bone biopsies which have been prepared with great care. Although a softening agent may have been added to make sectioning easier, these blocks are still fine for SEM study. We usually initially image such re-used blocks in the as-received condition since we can then match the BSE SEM image exactly to the last microtomed and stained LM section. To remove the rough, “ploughed field” effect generated by the microtome knife, we refinish the surface by polishing and then carbon coat the blocks. 1. If necessary, cut off surplus unwanted PMMA using a band saw whilst thinking about orienting the block to generate the cut and polished surface that you want to examine in the SEM (as well as keeping your fingers away from the blade). It is best NOT to clamp the blocks in a vice and to use a hand held hacksaw because you can fracture hard tissue in the block by squeezing it. Afterwards, write the specimen ID on the block. 2. Grind back the sides of the block at right angles to the expected block surface plane using successively finer grades of carborundum paper wheels under continuous cold water lubrication. Looking through the sides of the block will help you to visualise the block surface plane that you require. Begin to grind in to the plane that you want in the block surface. A finish of 1,200grit paper is acceptable and provides a good surface upon which you must again write the sample ID using a pencil or a diamond scribe. Do not use marker pens or you may lose the label. 3. Grind the base of the block to be parallel to the required top surface. Again 1,200 grit is the best finish, and do not forget to write the ID in pencil on the base of the block too.

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4. Polish the 1,200 grit finished top surface with an aqueous suspension of 6-μm diamond on a polishing cloth on a rotary polisher. 5. Rinse block in distilled water in an ultrasonic bath to release any attached 6-μm diamond. 6. Check finish under dissecting microscope. Go back to 1,200grit silicon carbide paper if needed. 7. Polish the 6-μm diamond-finished surface using 1-μm diamond on a separate polishing cloth. Be careful to keep the cloths used for different grade diamonds in clearly labelled containers. 8. Rinse in distilled water in ultrasonic bath to release any attached diamond, and rinse again. 9. Blot dry with clean tissue. 10. Check finish under dissecting microscope. Remember never to touch the top surface of the polished block. 11. Mount the samples on a suitable plate for the SEM you will be using as described in steps 12–18. 12. For our Zeiss DSM 962, we cut 80 cm × 80 mm plates from 3-mm thick aluminium alloy sheet. Current SEMs provide for 100 mm × 100 mm and 125 mm × 125 mm sample arrays. 13. Before or after mounting, use fine artist’s water colour brush to paint tracks to provide a conductive path to earth around the tops of the sides (see Note 17). Use suitably diluted colloidal graphite (“C-dag”) so as to reach both top and bottom edges of the block surface at minimally three places. 14. Prepare a large plastic Petri dish which will hold the ~4-in. diameter CC tape reel by covering the lid with strips of ordinary office grade 25-mm wide double-sided adhesive tape, leaving the non-stick cover on the tape. 15. Lay out lengths of 8-mm wide carbon conductive (CC) tape (also from Agar Scientific) on this non-stick surface. Cut this across in 1.5-mm wide strips (tags) using a single edged razor blade. 16. Attached samples to the plate using CC tags which will run over the side from the C-dag earthing regions to the bottom of each block. Three pieces per block will provide stable mounting. 17. For experiments with large numbers of nearly identical blocks, CC tape “tram lines” can be laid out on the plate (raft) to facilitate mounting. Cut 8 mm CC tape on the non-stick Petri surface in 2-mm wide, long strips. Apply these strips on the raft at a separation equal to the width of the blocks. Apply C-dag during or after mounting the array, making sure to connect the

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block top surface via the mounting tape to the metal plate (see Note 17). 18. Allow the C-dag to dry: a few minutes in our lab. 19. Apply a patch of “Post It” note paper to a bare patch on the raft where you will attach the BSE standard, so as to prevent carbon deposition in this area. (CC tape adhesive will strip evaporated carbon coatings from a substrate). 20. Coat the samples with carbon by evaporation. 21. Remove the “Post It” paper and attach the already carboncoated and CC-taped standards to be used in qBSE imaging to the raft before placing it in the SEM (19–21). 3.10.2. Making Casts from PMMA or Other ResinEmbedded Samples

After imaging PMMA blocks, the mineralised tissues can be dissolved to produce internal space casts, either globally of the marrow and canal space compartment (Figs. 3 and 7), or at a finer scale of the osteocyte lacunar-canalicular system. All calcified tissue matrices and mineral are destroyed in this process (see Notes 18 and 19). 1. Place the polished blocks in deep dishes. It does not matter too much if they have been coated with carbon for earlier qBSE imaging. In our hands, this will always have been the case and we can afterwards marry the plane qBSE image of the block face to the top surface of the prepared cast. 2. Add strong (e.g. 5 N) HCl. Add a warning sign for other lab users. 3. Place on a rocking or rotating shaking table for from one to several days. 4. Wash with water and add full strength NaOCl bleach. 5. Keep repeating, alternating, steps 2–4, until there is no bone mineral or matrix left. To begin with, it will appear that nothing is happening to the samples and the bone will seem to be a continuous tissue. This will happen if all the osteocyte lacunae and canaliculi are well filled with PMMA, and all hang together. To disperse these finer elements and leave only the major blood vessel canals and marrow spaces as casts, the slice must be washed with a gentle jet of tap water from a bacteriological pipette to hose away the smaller elements. They can also be dispersed by ultrasonication. 6. Finally, the cast must be rinsed with distilled water and dried. It is best to use freeze drying to get the ice to hold the delicate blood vessel canal casts in situ and prevent surface tension relocation which may happen with simple air drying.

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Fig. 7. Illustration of the process from resin embedding to polished surface for BSE imaging. All steps are described in Subheading 3.10.1.

To freeze dry the casts:

3.11. Correlation of SEM with Other Imaging Means

1.

Hold the samples above boiling liquid nitrogen until frozen, and then allow them to enter the liquid nitrogen. Pyrex glass crystallising dishes are ideal containers. Alternatively, place them in a dish and freeze them in contact with carbon dioxide ice (Cardice).

2.

Transfer all the samples in one batch in their container (covered with some liquid nitrogen if this is what you are using) to a chamber which can be pumped with a rotary vacuum pump, close the chamber and pump. To begin with, the nitrogen will boil off, and diminish ice condensation during the initial pump down phase. Next the ice which was the water supporting the cast will sublime off.

3.

Allow the samples to warm to room temperature.

4.

It may be difficult to apply an evaporated or a sputtered gold coating to a complex 3D cast that is good enough for SE imaging, yet gold will be necessary to get some signal to differentiate the surface of the cast. Therefore coat the samples first by carbon evaporation.

5.

Then apply a gold coating by evaporation or sputtering.

6.

Image using BSE using tilted views for 3D SEM (see Note 20 and Fig. 8a, b).

Study of the bone sample by SEM can be usefully correlated with other 3D digital imaging methods, including CSLM, X-ray microtomography, point projection microradiography (Faxitron), and

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Fig. 8. (a) PMMA cast left after acid and bleach dissolution of bone, showing compact bone at left and trabecular medullary bone at right. Note the gradual transition between the two. Osteocytes were dissociated from the cast by ultrasonication. Samples from 12-year-old horse, third metacarpal diaphysis, 1.25 kV SE image of gold sputter coated sample. Scale bar in image. (b) Transverse section of 2-year equine third metacarpal shaft, embedded in PMMA and converted to a cast by acid and bleach dissolution of bone. This is primary osteonal (plexiform) bone formed at the external surface of the cortex. At the centre is seen a cutting cone coming towards you and you can see the imprints of the original osteoclasts excavating the older bone. The osteoclasts are still inside the plastic but we cannot see them in this mode. Osteocytes were dissociated from the cast by ultrasonication, but those nearest to a free bone surface, i.e. those having canaliculi contact with a cast of marrow space, are retained and are the smallest features decorating the cast. This is available as a movie sequence to show 3D in the on-line edition. Field width = 1.937 mm. 20 kV BSE SEM of gold-coated sample.

flat bed optical scanning, and we routinely use all of these. For illustrations see the references mentioned below. 3.11.1. Correlated Confocal (or Other LM) with SEM Imaging of PMMA (or Other Resin)-Embedded Bone

Block surfaces prepared for BSE SEM, but before mounting and carbon coating, can be imaged using confocal autofluorescence microscopy (CSLM) to provide excellent histology of bone-organ cells and the two types of image married together to make one (22–24). CSLM is far less tolerant of block surface levelling errors than even BSE SEM.

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1. At the CSLM, the block is temporarily mounted on a glass slide using two strips of plasticine and a levelling press such as is used in metallurgical LM. 2. For high resolution work (and confocal is not confocal at low aperture), place glycerol on the PMMA block surface and apply a 170-μm thick glass cover slip. Glycerol is an excellent refractive index match to PMMA and does not attack PMMA. 3. Apply immersion oil to the top surface of the cover slip (assuming an upright microscope configuration). Immersion oil does attack PMMA, so keep it clear. Be careful not to mix oil and glycerol at the edges of the cover slip, so deliberately use oversized cover slips. 4. Focussing is best done using the reflected light confocal image to determine the level of the top surface of the block since this peaks very sharply at focus. Try to obtain images in the top 1–2 μm, as well as slightly deeper. 5. Name the images with a scheme which includes sample series ID and block number, objective lens magnification, alphabetical field identifier and depth of focus below the block top surface at the centre of the field of view in microns (e.g. ID123_63xa_02.TIF). You will use the topmost focus image in matching to the SEM scene, but may have a more interesting and continuous image at 3-μm deep. We try to reserve the blue channel of an RGB image for the reflected light channel, which usually gives the best match to SEM imaging. The green and red channels can then be used for different bands of autofluorescence, or for fluorescence from, for example, calcein and alizarin intra-vital mineralising front labels. All likely scenes of interest are recorded. 6. After CSLM, remove the cover slip, rinse the block surface with distilled water to remove glycerol and blot dry. 7. Rejoin the prior protocol (Subheading 3.10) at the mounting and coating stage step 11 (see Note 20). 8. Display the CSLM images on a monitor next to the SEM, and find and record the same fields by compositional BSE imaging. Using topographic BSE imaging (the difference image between opposing detector halves) can help to match to the reflection confocal image in particular. 9. Process the paired and matched CSLM and BSE SEM images (see Notes 21 and 22). Free software which we developed for this purpose is available from the author (“Honza’s programme”, ref. 22). See Fig. 9. 3.11.2. X-Ray Microtomography (XMT or mCT)

X-ray microtomography (XMT or μCT) equipment is commercially available from several suppliers, all surprisingly much more expensive than an excellent automated digital SEM, and all providing

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Fig. 9. Overlay of 20 kV BSE SEM imaging and CLSM confocal imaging. (a) BSE; (b) Same image processed according to our usual method using halogenated dimethacrylate standards (see refs. 19–24) to demonstrate variations in the degree of mineralisation of bone. The look up table explains the scale, where value 0 is signal level from monobrominated and 255 from monoiodinated standard. (c) Confocal image showing tetracycline labels which appear orange against a yellow– green background of autofluorescence and (d, e) overlays with different weights of SEM vs. CSLM contributions. Width of field of SEM image is 713 μm. Iliac crest biopsy from juvenile male with nutritional osteomalacia. Case 19 from the study by Schnitzler et al. 1994 (Schnitzler CM, Pettifor JM, Patel D, Mesquita JM, Moodley GP, Zachen D 1994. Metabolic bone disease in black teenagers with genu valgum or varum without radiologic rickets: a bone histomorphometric study. J Bone Miner Res 9:479–486).

noisy data at thousands of times worse volumetric resolution. However, the great advantage of XMT is that the sample remains intact and a complete 3D data set is obtained which can be visualised by “re-sectioning” in any plane. The best XMT laboratory apparatus correctly reconstructs linear absorption coefficients for every voxel and is pioneered in our department (http//www. microtomography.com). PMMA block surfaces can be found in the XMT stacks and exactly matched to qBSE images. 3.11.3. Faxitron Point Projection Microradiography

Faxitron point projection microradiography uses a finely focussed electron beam to provide a tiny X-ray source so that projective enlargement to fivefold can be usefully employed (see Chapter 29). Faxitron images of bone sections within PMMA in the 0.5- to 1.5-mm thickness range (thinner is better) give first class matching to BSE SEM montage images.

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3.11.4. Flat Bed Scanner Optical Imaging

Flat bed scanner optical imaging is the cheapest digital imaging method available. Macerated bone slices or embedded blocks or casts or whole arrays of samples on SEM rafts can be used to obtain 3D reference images, in the latter case supplying some support posts on the raft to hold the samples above the scanner table. With most flat bed scanners, it is only necessary to move the sample 25 mm sideways and to record a second image to have an excellent stereo pair. Although all these methods have inherently lower resolution than that possible with SEM, the wide field images are very useful for understanding where one is within a large sample.

3.11.5. Nano-indentation

Nano-indentation. Excellent correlation is also obtainable with nano-mechanical testing for Young’s modulus (stiffness) and microhardness. We use the same PMMA blocks prepared for qBSE imaging and the existing BSE image to locate particular areas of interest. Then a dense array, typically at 20-μm centres, of test points is selected and scanned using UMIS equipment. Using the tiny residual indents for location, we then re-image the same fields using higher resolution qBSE SEM to obtain measurements of electron backscattering which can be correlated with stiffness on a point by point basis (19, 24, 25, 26).

3.12. Shipping Bone Samples

We have had many experiences of international air freight shipments of bone samples ending in disasters during transport, for example, due to thawing of frozen material, brittle fracture of lids or seals of pots at low temperatures, leakage of fixative solutions, etc. Now we recommend that all tissue be fixed in suitably buffered formaldehyde for 24 h, and then substituted (via 70 and 100% ethanol, though it can be direct) to glycerol for transport. Glycerol has a very low vapour pressure and samples simply cannot dry. Samples can be sent in minimal fluid volume which greatly reduces transport costs and risks. The temperature during transport is also no problem, since the freezing temperature of glycerol is never likely to be reached, and it would not matter if it were. 1. After glycerol substitution, wipe off surplus glycerol from bone slices between finger and thumb whilst wearing rubber gloves, and place in Zip lock polythene bags with a label written in pencil on card inside the bag. 2. Batch smaller samples into larger ZipLock bags with some paper tissue to mop up any possible leakage. 3. Pack and ship to destination. 4. On arrival, and when ready to proceed, remove the glycerol with ethanol or IMS, and proceed to maceration or embedding protocols as required.

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4. Notes 1. It is important to warn against use of EDX with bulk bone samples for to determine Ca:P ratios. If you do use EDX, use the smallest over-voltage ratio that you can to keep the penetration volume small, although this cannot prevent the excitation volume for P X-rays being much larger than that for Ca (1). Never trust EDX data from near interfaces, and interfaces of bone with space are never far away in real tissue from the real bone world. In addition, you almost certainly will have altered the elemental composition during sample preparation since freezing relocates solutes to ice crystal boundaries which do not respect cell boundaries, and all chemical fixation protocols are likely to change pH, and redistribute Ca++, Mg++, phosphate, etc. 2. For CL, the microscope must able to capture light rather than electrons (beware that many systems do not). Second, you should be careful to not extinguish the CL light before it has illuminated your problem. The volume from which light is excited in the sample is greater than for any other signal you will collect. Therefore, as with X-rays, the area and volume from which you collect CL will be greater than the electron beam resolution possible with SE and BSE imaging. You might want to reduce this problem by reducing the beam voltage and hence electron beam penetration, but do not. Radiation damage is increased as a cube law function with decreasing voltage. Stay with 20 kV or higher and you will do less radiation damage. Finally, the CL light has to escape the sample to be of use. Metallic and carbon surface coatings will interfere, therefore use a transparent conductive coating. Very little work has been done in this area, but two approaches have been described: an evaporated coating of indium oxide, or drying down a solution of an organic anti-electrostatic spray such as Duron (available from Agar Scientific). A special area of interest concerning the potential of CL in bone research is to include bright CL emitters (phosphors, scintillators in the sample embedding protocol). Organic scintillators can be included in the embedding medium to highlight space within bone, or, potentially, to study the continuity of space. 3. Whether cut frozen, or fresh, or fixed, or thawed, a coarse method such as using a band saw should be avoided where possible in the preparative step preceding embedding of bone. This is because the tiny bone shards (swarf) become impacted in the marrow and canal space in the bone and survive there during the embedding process. The same is seen with human iliac crest trephine core bone biopsy samples where the results

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from the peripheral regions have to be discarded. It is therefore best to cut, or to re-cut, the bone using a water-lubricated slow speed diamond saw. You can also use IMS as cutting coolant and lubricant if the bones are already “in spirit.” If, however, the cut surface is to be prepared by maceration to look at the bone proper, then the fragmented bone can be washed away during preparation and polishing of the cut surface. Obviously, out of place bone fragments can be edited out of a BSE SEM image if you are not also interested in the marrow histology. 4. Postfixation with tannic acid has been suggested to provide greater stability of the tissue during drying (27) and help prevent shrinkage. A simple procedure is to fix samples in glutaraldehyde first as per protocols given, then postfix for 1 h in 1% tannic acid in cacodylate buffer before proceeding to drying as per protocols given. 5. A main problem with FD is the generation of holes through cell membranes by the growth of ice crystals during the freezing process. Glutaraldehyde and formaldehyde fixation will diminish this problem. Even more important is to keep the ice crystals invisibly small, and this can be done by rapidly freezing the samples in chloroform water. The logic is that clathrates are formed, and much larger number of smaller ice crystals nucleate, starting at a lower freezing temperature. 6. You may alternatively use a liquid nitrogen slush prepared by a dedicated slusher if you have access to such a machine (28). 7. Unfortunately, the soft tissue (cellular) component of the bone samples may shrink during fixation, will inevitably shrink during ethanolic (or other) substitution, will shrink again in liquid CO2, and again as this is warmed in its conversion to the gaseous phase, and will shrink again after the sample is at atmospheric pressure, and a little bit more when being coated and in the SEM chamber (12). So you may well ask, “Why bother?” That these well-recognized problems are almost universally ignored depends on the success of the method for SEM soft tissue samples. Here, the whole sample shrinks together, and this shrinkage can be accounted for by changing the stated magnification. However, when considering a bone (or similar hard tissue) sample, remember that the substrate will not shrink at all if it consists of metal or glass, and only a little if it consists of calcified bone, whereas the cells do shrink and will be stretched over the substrate, thinned down as the consequence, and usually pulled apart at mutual junctions. 8. That the entire release from all the activities of the entire bone research community in a year might be the same as in cleaning one suit of clothes has been ignored in taking this leap!

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9. Cells will be well flattened on the substrate after this procedure, which must be remembered when interpreting the images. 10. You are advised against using one of the “carbon string” coaters often found in SEM labs since these usually only accommodate small sample arrays, and the evaporation procedure only lasts for a fraction of a second. Thus it is not possible to rotate and tilt the samples to help ensure that carbon lands up on the surface from all directions. 11. So long as we resign ourselves to not studying cells, there is no reason not to deep freeze whole body parts to hold tissue prior to dissection and sectioning. All cells are badly damaged by ice crystal growth during conventional freezing, but the design of bone matrix is such as to prevent such damage. Cartilage matrix also survives. 12. Fat can be also be removed prior to simmering by immersion in trichloroethylene. Over-treatment will gelatinise even the calcified bone matrix and leave the bone partly anorganic. 13. Fatty bones may remain fatty or waxy after hypochlorite cleaning. This can be avoided by first defatting them, for which soaking in IMS in a 50°C oven is a simple solution. 14. The dentine slice will ideally be cut from a shaped “beam” of dentine prepared from a large mammal tooth. Initial reduction of the tooth (tusk) can be done dry with a band saw if you do not mind the smell of burning dentine. Your local Geology Department will have very large, high speed, water-cooled saws intended for rock samples, so get them to help. Most large mammals are protected by CITES regulations, but their teeth can be obtained gratis from customs authorities who have confiscated them in enforcing these laws. We have preferred sperm whale and walrus tusk dentine, but elephant tusk dentine (ivory: avoid the cementum!) is used by many other labs. In the latter case, it is common practice to cut an extremely large area slice, and to punch out discs to suit the size of multiwell culture dishes. The two sides of the as-cut slice are unequal. You must know which is which and choose the smoother side. The reason for this difference is that the piece of tooth from which the slice is cut is not perfectly hydrated. During the cutting process, which is done in and under water, the slice which is being cut off is rehydrating during the cutting and will usually warp away from the diamond cutting wheel: this is the rough side. The side that remains on the solid beam is polished to an extent, and will be the “out” side of the next slice to be taken: this is the smooth side. Mark each sample with a graphite pencil on the rough side and place the slice rough side down in the tissue culture well with OCs on top.

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15. The prepared “SEM” sample is also highly suited to reflection mapping mode confocal microscopy, though not all confocal microscopes work well in reflection mode. 16. Some SEM manufacturers are now offering what they call 3 or 5 segment BSE detectors. These actually comprise a 2 or 4 sector annular detector surrounding the electron beam with another rectangular chip to the side, off axis. Now you can choose two opposing sectors or sector pairs and the off axis detector to provide your three input images. 17. It cannot be over-emphasised how important it is to ensure earthing of the block top surfaces. Blocks in any one experimental series will usually be of nearly the same thickness. If some require thickening, one can add a slice of PMMA to the base of the block by wetting it with chloroform or acetone and then pressing the block into it. This must be done before final finishing of the block. The block sides will need to be refinished too. Samples embedded in resins other than MMA, for example, in glycol methacrylate or in epoxy resin can equally well be used. Although they cut well, epoxies do not polish well. 18. Etching strictly involves removal of all components in a system but at different rates. Here we attempt to remove only matrix and mineral and to leave the resin intact. The PMMA in fully mineralised bone matrix is so sparse that it disperses when collagen and mineral which support it are removed. In newly formed bone packets the PMMA remains. This can be exploited to demonstrate the 3D distribution of such newer, less mature bone in the SEM. 19. If you want to study the casts of the osteocyte lacunae and canaliculi, those that were closest to the nearest bone surface will remain attached to the cast. If you want a sample of osteocyte lacunae and canaliculi deep to natural surfaces, cut the PMMA block into thin slices and glue them to other pieces of PMMA using acetone or chloroform, or, alternatively, cut thin slices of bone before embedding so that you use the remaining bulk PMMA outside the real sample to support the final cast. 20. You can also perform the correlative CSLM or other LM imaging after the BSE SEM, but then have to contend with the light losses due to reflection and absorption by the carbon conductive coating. 21. As a first alternative to CSLM, try block surface staining for conventional LM. Toluidine blue in 70% ethanol will stain the top microns of PMMA blocks in less than a minute. Wash off with distilled water. Good illumination for thick bone block samples will be obtained by “epi-fluorescence,” i.e. making the bulk sample fluoresce with violet, blue or green illumination, thus backlighting the stained layer on the top of the block.

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22. Also try wide field conventional fluorescence LM. Here you can visualise mineralising front labels like calcein, etc., because you will get the sharper focus information from the topmost layer, deeper layers being progressively more blurred due both to the lack of focal plane discrimination in wide field fluorescence and scattering in depth.

Acknowledgments I thank Maureen Arora for her patient help in carrying out many of the procedures described in this chapter on many thousands of samples. Her employment has been funded by the Horserace Betting Levy Board. References 1. Howell, P. G. T and Boyde, A. (2003) Volumes from which calcium and phosphorus X-rays arise in electron probe emission microanalysis of bone: Monte Carlo simulation. Calcif. Tissue Int. 72, 745–749. 2. Boyde, A. and Shapiro, I.M. (1980) Energy dispersive X-ray elemental analysis of isolated epiphyseal growth plate chondrocyte fragments. Histochem. 69, 85–94. 3. Boyde, A., Reid, S. A. (1983) Tetracycline cathodoluminescence in bone, dentine and enamel. Histochem. 77, 525–533. 4. Boyde, A., Reid, S. A. (1983) Simple collectors for cathodoluminescence in the SEM made from aluminium foil. J. Microscopy 132, 239–242. 5. Boyde, A. (2008) Low kV and video-rate, beam-tilt stereo for viewing live-time experiments in the SEM Chap. 7 pp. 197–214 and colour plates 4–11. In: Schatten H, Pawley JB (eds) Biological Low Voltage Scanning Electron Microscopy Springer New York. ISBN 978-0387-72970-1. 6. Boyde, A. (1984) Methodology of calcified tissue specimen preparation for scanning electron microscopy. In: Methods of Calcified Tissue Preparation, pp251-307: Dickson GR (Ed), Elsevier, Amsterdam. 7. Boyde, A. (1972) Scanning electron microscopic studies of bone. In: The Biochemistry and Physiology of Bone, 2nd edn, Vol.1, pp259–310 Bourne GH (Ed) Academic Press, New York. 8. Boyde, A., Jones, S. J. (1996) Scanning electron microscopy of bone: instrument, specimen

9.

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and issues. Microscopy Research and Technique 33, 92–120. Boyde, A., Jones, S. J. (1983) Scanning electron microscopy of cartilage. In: Cartilage I: 105–148, Hall BK (Ed), Academic Press, New York. Boyde, A., Ali, N. N., and Jones, S. J. (1984) Resorption of dentine by isolated osteoclasts in vitro. Brit. Dent. J. 156, 216–220. Boyde, A., and Maconnachie, E. (1983) Not quite critical point drying. In: Science of Biological Specimen Preparation, pp71–75 : Revel JP, Barnard T, Haggis GH (Eds) SEM Inc, AMF O’Hare, IL. Boyde, A., and Maconnachie, E. (1979) Volume changes during preparation of mouse embryonic tissue for scanning electron microscopy. Scanning 2:149–163. Boyde, A., Bailey, E., Jones, S.J., and Tamarin, A. (1977) Dimensional changes during specimen preparation for scanning electron microscopy. Scanning Electron Microscopy 1, 507–518. Boyde, A., Ali, N. N., Jones, S. J. (1985) Optical and scanning electron microscopy in the single osteoclast resorption assay. Scanning Electron Microscopy 3, 1259–1271. Boyde, A., and Jones, S. J. (1991) Pitfalls in pit measurement. Calcif. Tissue Int. 49, 65–70. Boyde, A. (1973) Quantitative photogrammetric analysis and qualitative stereoscopic analysis of scanning electron microscope images. J. Microscopy 98, 452–471. Boyde, A. (2004) Improved depth of field in the scanning electron microscope derived from

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A. Boyde through focus image stacks. Scanning 26, 265–269 Boyde, A. (2003) Improved digital SEM of cancellous bone: scanning direction of detection, through focus for in-focus and sample orientation. J. Anat. 202:183–194. Ferguson, V. L., Bushby, A. J., and Boyde, A. (2003) Nanomechanical properties and mineral concentration in articular calcified cartilage and subchondral bone. J. Anat. 203, 191–202. Howell, P. G. T., Davy, K. M. W., and Boyde, A. (1998) Mean atomic number and backscattered electron coefficient calculations for some materials with low mean atomic number. Scanning 20, 35–40. Boyde, A., Travers, R., Glorieux, F. H., and Jones, S. J. (1999) The mineralization density of iliac crest bone from children with osteogenesis imperfecta. Calcif. Tissue Int. 64, 185–190 Boyde, A., Lovicar, L., and Zamecnik, J. (2005) Combining confocal and BSE SEM imaging for bone block surfaces. European Cells & Materials 26, 33–38. Doube, M., Firth, E. C., and Boyde, A. (2005) Registration of confocal scanning laser micros-

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copy and quantitative backscattered electron images for the temporospatial quantification of mineralization density in 18-month old thoroughbred racehorse articular calcified cartilage. Scanning. 27, 219–226. Doube, M., Firth, E. C., and Boyde, A. (2007) Variations in articular calcified cartilage by site and exercise in the 18-month-old equine distal metacarpal condyle. OsteoArthritis &Cartilage 15, 1283–1292. Bembey, A. K., Oyen, M. L., Bushby, A. J., and Boyde, A. (2006) Viscoelastic properties of bone as a function of hydration state determined by nanoindentation. Philosophical Magazine 86 (33–35 SPEC. ISSUE), 5691–5703 Oyen, M. L., Ferguson, V. L., Bembey, A. K., Bushby, A. J., and Boyde, A. (2008) Composite Bounds on the Elastic Modulus of Bone. J. Biomechanics 41:2585–2588. Levanon, D., and Stein, H. (1999) Tannic acid and thiocarbohydrazide as structural reinforcement agents in the preparation of rabbit knee articular cartilage for the scanning electron microscope. Histochem. J. 31, 71–73. Severs, N. J. (2007) Freeze-fracture electron microscopy. Nature Protocols 2, 547–576.

Chapter 25 Fluorescence Imaging of Osteoclasts Using Confocal Microscopy Fraser P. Coxon Abstract In order to understand osteoclast cell biology, it is necessary to culture these cells on a physiological substrate that they can resorb in vitro, such as bone or dentine. However, this creates problems for analysis by fluorescence microscopy, due to the depth of the sample under investigation. By virtue of its optical sectioning capabilities, confocal microscopy is ideal for analysis of such samples, enabling precise intracellular localisation of proteins in resorbing osteoclasts to be determined. Moreover, by taking a series of images in the axial dimension, it is possible to create axial section views and to reconstruct 3D images of the osteoclasts, enabling the spatial organisation of the structures of interest to be more easily discerned. Key words: Confocal microscopy, Osteoclast, Resorption, Dentine, Fluorescence

1. Introduction Imaging of osteoclasts, both in vitro and in vivo, is crucial to furthering our understanding of the activity of this complex cell type. Simple transmitted light microscopic techniques enable basic parameters such as osteoclast size, morphology, and multinuclearity to be determined, while fluorescence microscopy of osteoclasts cultured on plastic or glass enables subcellular localisation of proteins to be determined. However, when cultured on such substrates, osteoclasts are unable to polarise and therefore behave very differently from osteoclasts in vivo. To simply but very effectively model the in vivo environment, osteoclasts can be cultured on a mineralised substrate in vitro, such as cortical bone or dentine, on which the osteoclasts will polarise and resorb the substrate. The morphology of these osteoclasts is similar to osteoclasts in vivo, with a rounded shape (cell depth can be 30 μm or more), whereas osteoclasts cultured on glass or plastic spread so extensively that the only significant depth is the region containing the nuclei (Fig. 1). 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_25, © Springer Science+Business Media, LLC 2012

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Fig. 1. Comparison of osteoclast morphology on glass and on dentine. (a) zx section of a resorbing rabbit osteoclast cultured on dentine; membrane stained with an antibody to the vitronectin receptor; dentine surface stained using AF-ALN. Dentine surface marked with arrow; resorption pit with asterisk. (b) zx section of a rabbit osteoclast cultured on glass; stained using wheat germ agglutinin-Alexa Fluor488, visualising the plasma membrane and perinuclear Golgi (central area of cell shown only). Note the dramatic difference in depth of the cells on these different substrates.

Imaging osteoclasts on mineralised surfaces using conventional fluorescence microscopy it is possible to assess, for example, cytoskeletal polarisation by staining for polymerised actin (see Chapter 10), but for more discriminate imaging the depth of the cell creates significant problems. The limited depth-of-focus of high numerical aperture microscope objectives restricts clear imagery to a shallow depth-zone, with out-of-focus light scattered or reflected from above and below the focal plane significantly reducing contrast. These problems are compounded by the autofluorescent nature of mineralised substrates. Confocal microscopy minimises these problems since by design it restricts both the field of illumination and the light collected to a single point in the same focal plane (hence the name confocal). This effectively results in optical sectioning of the specimen of interest, removing out-offocus light from the image and enabling subcellular structures to be resolved in the axial (z) dimension, even within such thick specimens. In addition, the autofluorescence emitted from mineralised substrates is considerably reduced by the monochromatic laser excitation source on confocal systems, compared to excitation through band pass filters on wide-field systems (Fig. 2). Furthermore, confocal microscopy enables three-dimensional (3D) structures to be reconstructed by taking serial optical sections along the axial dimension (a “z-stack”), without the necessity for physical sectioning. Therefore, the difference between confocal microscopy and conventional fluorescence microscopy can be considered the optical equivalent of the difference between computed tomography and X-rays. Imaging of live cells is also possible, potentially enabling four-dimensional data sets to be captured, although speed constraints restrict the effectiveness of this to sectional, rather

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mirror PMT detector 1 PMT detector Alexa Fluor 647 PMT detector 2 PMT detector Alexa Fluor 488 BP 505-530

PMT detector 3

NFT 650

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Fig. 2. Typical configuration setup of a confocal microscope for 3-channel imaging of typical green, red, and near infra-red fluorophores (e.g. AF 488, AF 555, and AF 647, respectively). HFT = primary dichroic beam splitter, defects light only of the specified wavelengths onto the specimen; NFT = secondary dichroic, defects light shorter than the specified wavelength; BP = band pass filter, transmits light of the specified wavelength range. Dashed arrows = excitation light; solid arrows = emitted light.

than 3D imaging (see Subheading 3.10). Finally, the laser excitation that is fundamental to confocal microscopy also opens up the possibility of a number of more advance imaging applications, including fluorescence resonance energy transfer (FRET) and fluorescence recovery after photobleaching (FRAP), outlined briefly in Subheadings 3.9 and 3.10. It is important to remember, however, that confocal microscopy is a form of light microscopy, and the resolution is therefore limited by the wavelength of light. To discern intracellular structures at the highest resolution, electron microscopy is required. 1.1. Principles of Confocal Microscopy

The principle of confocal microscopy was developed and patented in 1957 by Minsky (1). However, it was not until 1987 that laser scanning confocal microscopes became commercially available and were first used for fluorescence imaging in biological specimens (2). The optical sectioning that is the cornerstone of confocal microscopy is achieved by the use of pinholes. However, there are other important differences compared to conventional fluorescence microscopy that are essential to this type of microscopy. In a conventional wide-field microscope, the entire specimen is bathed in light from a mercury or xenon source, with the required

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wavelengths of light isolated by the use of band-pass filters that transmit a fairly narrow range of wavelengths. In contrast, the illumination in a confocal microscope is achieved by scanning focused beams of light, usually from a high-powered laser source that delivers monochromatic wavelengths, on to the specimen to image a single pixel at a time (a requirement defined by the use of confocal pinholes). To build up a 2D image, the laser is scanned across the region of interest in a raster pattern by means of two highspeed vibrating mirrors driven by galvanometer motors. One important result of this is that it tends to slow down image acquisition rate compared to conventional wide-field microscopy. The emitted fluorescence is collected by the objective, passed back through the same scanning mirrors as the excitation light (a process known as descanning), and focused at the detector pinhole aperture. It is the diameter of the pinhole, which is user-controlled, that determines the effective thickness of the resulting optical section (Fig. 3). However, the use of a pinhole means that the emitted fluorescent signal is usually extremely weak and necessitates the use of highly sensitive photon detectors (photomultiplier tubes or PMTs). These do not require spatial discrimination, since the image is scanned pixel-by-pixel, and can respond quickly to a continuous flux of varying light intensity. The sensitivity of these detectors is regulated by setting the “gain,” which refers to the electron amplification within the detector.

Fig. 3. Effect of pinhole size on axial resolution; Images show the F-actin ring of a resorbing rabbit osteoclast cultured on dentine, stained using TRITC-phalloidin. The left hand image was taken using the optimal pinhole setting of 1 Airy unit, resulting in an optical section of 1 μm. The right hand panel shows the same field of view with the image collected using the pinhole at its widest setting, resulting in an optical section of 13 μm. Note the sharpness of the left panel compared to the out of focus light that has blurred the right panel. Stress fibres in contaminating stromal cells can also be seen.

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The use of confocal microscopy to visualise osteoclasts cultured on a physiological substrate (dentine) was first described more than 20 years ago, in which sites of attachment to the substrate were analysed by immunostaining for vinculin (3). Another study shortly afterwards further analysed the distribution of the vitronectin receptor and cytoskeleton-associated proteins in osteoclasts cultured on bone (4). Subsequently, confocal microscopy has proved particularly instrumental in identifying the membrane domains and intracellular vesicular trafficking routes in resorbing osteoclasts, information that could not have been determined from studies using conventional fluorescence microscopy. The localisation of LAMPs (lysosomal-associated membrane proteins) at the ruffled border by immunostaining offered the first evidence of the lysosomal nature of this osteoclast-specific structure (5). Furthermore, studies of the trafficking of viral proteins in osteoclasts confirmed that the ruffled border lacked properties of normal plasma membranes, and also identified the polarisation of the basolateral surface into two distinct domains (6). Seminal studies then utilised confocal microscopy (in particular, axial views of resorbing osteoclasts in vitro), together with electron microscopy to elegantly demonstrate the transcytotic trafficking of vesicles containing degraded bone matrix from the ruffled border to one of these membrane domains on the opposing side of the osteoclast, which accordingly became known as the “functional secretory domain” (7, 8). These studies have been bolstered by studies of uptake of dextran from the ruffled border into the transcytotic pathway (9) and other studies demonstrating uptake of resorbed matrix by osteoclasts (10). Other studies have utilised similar in vitro cultures of osteoclasts on physiological substrates such as ivory or bovine bone to demonstrate the intracellular localisation of TRAcP and cathepsin K (11), the localisation of cathepsin K in the resorption pit (12), the localisation of cytoskeleton-associated proteins such as Pyk2 and p130Cas (13, 14), the ruffled border targeting of the V-ATPase (15) and disruption of this by knockdown of Rab7 (16). More recently, with the development of fluorescent proteins such as GFP to tag proteins of interest, and the development of methods to transfect osteoclasts (see Chapter 14, this volume), confocal microscopy has been utilised to study the localisation of intracellular proteins for which antibodies suitable for immunostaining are not available (17, 18). Such tagged constructs enable protein localization to be monitored in real time using live cell imaging, (see Chapter 26, this volume); a good example of the utility of this approach is the study of dynamics of the actin ring in live GFPactin-expressing osteoclasts cultured on hydroxyapatite discs (19). Finally, an application for which we have used confocal microscopy is to analyse the cellular uptake of bisphosphonates from the surface of dentine, using fluorescent alendronate analogues (10)

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Fig. 4. Intracellular uptake of AF-ALN by resorbing osteoclasts. Rabbit osteoclasts were cultured on AF-ALN-coated dentine (green) for 24 h then fixed and counterstained for the vitronectin receptor (red ). (a) Images shown are 1 μm optical sections; the arrow denotes the position of the xy scan. Note the abundance of the AF-ALN in the resorption pit and the accumulation of AF-ALN into punctate intracellular structures throughout the depth of the cell. (b) Fluorescence intensity (AF-ALN only shown) displayed by false-colouring to a look-up table (8-bit image; blue = 0, red = 256). (c) Depth coding to denote the axial position of a fluorophore in a single 2D image. In this case, the axial position of the AF-ALN staining in the same cell is denoted by false-colouring to a look-up table (blue = bottom of resorption pit; red = top of osteoclast).

(Fig. 4) and purified, fluorescent risedronate analogues (20). Such compounds are also useful reagents for visualising the surface of dentine in osteoclast cultures post-fixation and crude preparations can be synthesised relatively easily (see Subheading 3.1). Rather than describing specific protocols that may have limited appeal, this chapter provides an overview of universal considerations when analysing osteoclasts by confocal microscopy, and includes details of how to generate and analyse 3D data sets, spectral unmixing of fluorophores, and some of the advanced fluorescent probes that are opening up even more applications for confocal microscopy.

2. Materials 2.1. Equipment

There are several types of confocal microscopes, although the discussion here is restricted to the most widely used, laser scanning type (LSCM), the main manufacturers of which are Zeiss, Leica, and Olympus. Other types include the spinning disc (Nipkow disk) systems, which employ a “multiple beam” scanning approach to enable much faster scanning than LSCM, and are therefore preferred for imaging faster events at full resolution (such as live cell imaging applications).

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Historically, the most common LSCM set-up has been a 3-laser system with excitation wavelengths of around 488, 543, and 633 nm, which are ideal for imaging specimens stained with fluorophores emitting in the green, red, and near infra-red (NIR) regions of the electromagnetic spectrum, respectively (see Note 1). Less common, partly due to their high cost, are blue-diode lasers that can provide an ultra-violet (UV) excitation wavelength (usually 405 nm) suitable for fluorophores that emit in the blue region (such as DAPI). In any case, such lasers are not ideal for imaging cells on dentine, due to the extremely high autofluorescence of dentine at these wavelengths (see Note 2). We routinely use a Zeiss LSM510 META system, equipped with an Argon laser able to emit at 458, 488, and 514 nm, and two Helium–Neon lasers providing 543 and 633 nm excitation (Fig. 2). These gas-based lasers, which were universally used on confocal systems, have begun to be supplanted by more efficient solid-state lasers, while systems equipped with four lasers (including UV and infra-red wavelengths) are becoming increasingly common. Systems are available with two or three PMTs; the latter enable simultaneous 3-channel imaging, whereas the former require sequential scanning, utilising a single PMT to detect more than one fluorophore. The objective lens is a crucial consideration, since it determines the magnification, field of view and resolution, and its quality determines the efficiency of light transmission and the contrast and aberrations of the resulting image. To achieve the highest resolution, oil immersion lenses (usually 40–63×) with high numerical aperture are required. These are optimised for use with 0.17-mm thick coverslips, and require oil having an identical refractive index to the glass (1.518). One caveat of such lenses is that their working distances are extremely short. 2.2. General Reagents

1. α-Minimum essential medium (α-MEM) supplemented with 10% foetal calf serum, 100 U/ml penicillin, 100 μg/ml streptomycin, 1 mM L-glutamine. 2. Foetal calf serum (FCS). 3. Phosphate-buffered saline (PBS), pH 7.2. 4. 4% paraformaldehyde in PBS (optionally containing 2% sucrose). 5. Triton permeabilisation buffer: 0.5% Triton X-100 in PBS, pH 7.2. 6. Saponin permeabilisation buffer: 0.1% saponin (Calbiochem, UK) in PIPES buffer, pH 6.8. 7. Sterile discs of dentine or bovine cortical bone (for details see, for example Chapter 10). 8. Hydroxyapatite-coated glass, such as BioCoat Osteologic™ (BD Biosciences), can be used as an alternative to bone or dentine.

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9. Secondary antibodies; we use affinity purified antibodies raised in goat, conjugated to Alex Fluor 488, 555, or 647 (Invitrogen). 10. Glass slides. 11. Anti-fade mounting reagent such as VectaShield (Vector Labs) or SlowFade® Gold (Invitrogen). 12. Nail varnish. 13. 0.17-mm Thickness glass coverslips (widely available). 14. Fluorescence-free immersion oil (we use Zeiss Immersol 518F). 2.3. Reagents for Synthesising Fluorescent BPs

1. Alexa Fluor-488 carboxylic acid 2,3,5,6-tetrafluorophenyl ester (AF-488; Invitrogen). 2. Bisphosphonate with free amino group, such as alendronate (available from several suppliers, including Discovery Fine Chemicals, Wimborne, UK). 3. 0.1 M Sodium bicarbonate, pH 9. 4. 1 mM Calcium chloride solution. 5. 1 mM EGTA. 6. Dimethyl sulfoxide.

2.4. Useful Probes for Staining Osteoclast Structures

1. Nuclear dyes. The most common of these is DAPI for historical reasons, but this has the drawback that it excites in the UV spectrum, and therefore requires a UV laser, which can be prohibitively expensive and is not universally used on confocal systems. Moreover, the short wavelength emission of DAPI is prohibitive for imaging on dentine, due to extremely high autofluorescence. However, there is now a huge range of alternative nuclear dyes available with excitation and emission spectra across the visible and near infra-red wavelengths; we most frequently use TO-PRO-3 (Invitrogen), which emits in the NIR wavelength and is compatible with a 633-nm laser or similar. 2. Lysotracker-Red or Lysotracker Green (Invitrogen); Lyso-ID® Green (Enzo Life Sciences). These are acidotropic dyes that selectively accumulate in acidic vesicles, i.e. endosomes and lysosomes, and are therefore useful markers of these organelles (Fig. 6). These probes are designed for use in living cells, but are also compatible with aldehyde fixation. 3. Wheat germ agglutinin coupled to the fluorophore of choice (Invitrogen); this plant lectin binds to glycosylated proteins, and therefore stains the plasma membrane, Golgi apparatus and secretory pathway. If only plasma membrane staining is desired, then cells should not be permeabilised prior to staining. 4. MitoTracker probes (Invitrogen; available in green, orange, red, and NIR emission wavelengths). This reagent can be used to label mitochondria in live cells, and in the case of MitoTracker

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Green without the need for a wash step due to its lack of fluorescence in aqueous solution. It is also compatible with aldehyde fixation, enabling mitochondria to be visualised in fixed cells along with immunostaining for other antigens of choice. 5. Magic red cathepsin K substrate (ImmunoChemistry Technologies). This enables cathepsin K activity to be visualised in living cells. This is a cell-permeable substrate sequence, LR, specific for cathepsin K, linked to a red fluorophore (Magic Red™), which fluoresces once cleaved by active cathepsin K (21). Such substrates specific for other cathepsins have been used in studies of phagosomes in macrophages, for example (22). A similar probe (Cat K 680 FAST) is now available from VisEn Medical. 6. Fluorescent phalloidin conjugates. The fungal toxin phalloidin binds to polymerized F-actin and can therefore be used to easily visualise the actin cytoskeleton in fixed cells (Figs. 2 and 6). This is available conjugated to a wide range of fluorophores from numerous suppliers; we mostly use conjugates with fluorescein or rhodamine produced by Sigma. 7. Anti-alpha tubulin antibodies (such as T5168, Sigma). Antibodies to tubulin, the basic protein component of microtubules, are an excellent tool to visualise the microtubule network in fixed cells. There are also direct probes for the microtubule network available, such as Tubulin Tracker™ Green (Oregon Green® 488 Taxol, bis-acetate) from Invitrogen, which is a fluorescent conjugate of the microtubule-binding protein taxol, and is designed to work in live cells. 2.5. Software for Image Analysis

While post-acquisition image analysis can be performed in the image capture software, this software will not usually offer a great deal of functionality. Dedicated image analysis software for this purpose is worth considering; good examples include Imaris (Bitplane), Volocity (PerkinElmer), Image-Pro (Media Cybernetics) or the widely used open source software ImageJ (http://rsb.info. nih.gov/ij/index.html).

3. Methods 3.1. Synthesis of Fluorescently Labelled Alendronate

Fluorescently-labelled bisphosphonates are excellent tools for visualising the surface of bone or dentine in confocal microscopic images (10). This relatively simple synthesis of labelled alendronate is based on the reaction between the primary amine group of alendronate with the succinimidyl ester of the fluorophore, to create a stable amide linkage between the alendronate and the fluorescent tag (23). Alternatively, there are also commercially available

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fluorescent bisphosphonate analogues, such as OsteoSense 680 (VisEn Medical), which fluoresce in the NIR spectrum and are compatible with widely used NIR lasers (such as Helium–Neon 633 nm laser) on confocal systems. For economic reasons the protocol described uses a ratio of 1:10 AF-488:ALN, therefore in the final preparation only a small proportion of the alendronate will be labelled with the fluorophore, but sufficient for effective labelling of mineralised surfaces (see Note 3). 1. Mix the 1 mg vial (1.1 μmol) of the amine-reactive probe Alexa Fluor-488 carboxylic acid 2,3,5,6-tetrafluorophenyl ester (AF488; Invitrogen) dissolved in DMSO with 11 μmol of alendronate [dissolved in 0.1 M sodium bicarbonate, pH 9.0 (i.e., a 1:10 molar ratio)]. 2. Adjust the volume to 1 ml with 0.1 M sodium bicarbonate, pH 9.0 and incubate the solution for 2 h at room temperature with mixing. 3. To precipitate alendronate, add 20 μmol of CaCl2 and centrifuge the mixture (14,000 × g, 10 min). 4. Wash the precipitate five times in 1 ml of distilled water. To bind the Ca2+ (and hence resolubilise the alendronate), add 20 μmol of EGTA to the precipitate. 5. Add 100 μl PBS until all alendronate-AlexaFluor-488 (AF-ALN) has dissolved then mix the solution for 30 min. The final solution (referred to as AF-ALN) will contain both labelled alendronate and free alendronate. The labelled alendronate is extremely stable, and will last for years aliquoted and stored at −20°C in the dark. 6. This compound will bind to bone and dentine extremely avidly, thereby efficiently labelling the mineral surface at concentrations of 10 μM or even lower. Dilute in dH2O and incubate fixed, permeabilised cells with AF-ALN for 10 min (Figs. 4 and 6; see Subheading 3.3). 3.2. Culture of Osteoclasts for Confocal Microscopy

Osteoclasts from a variety of sources can be used. Mature osteoclasts isolated from neonatal rats (4) or rabbits (9, 10) have the advantage that they are authentic osteoclasts, and their morphology and behaviour most closely resembles osteoclasts in vivo. Alternatively, osteoclasts can be generated in vitro from cultures of mouse marrow cells or from human peripheral blood mononuclear cells as described in other chapters in this volume. In these cases, the osteoclasts can either be generated directly on the substrate of choice (e.g. glass coverslips or dentine), or can be generated on plastic or collagen-coated dishes and then transferred to dentine discs for resorptive studies. We prefer to generate human osteoclasts (at least partially) on plastic before transferring to dentine, since such cultures tend to be more robust and reproducible

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(see Chapter 11, this volume). It is also possible to culture authentic human osteoclasts by isolating them from giant cell tumour (osteoclastoma) samples (7). These osteoclasts have a normal phenotype and function, since they are not the neoplastic cell type in these tumours, but sourcing the material is a limiting factor. 3.2.1. Culture of Osteoclasts on Glass

Although glass is the most convenient substrate for imaging osteoclasts by confocal microscopy, the morphology of osteoclasts on this substrate is completely different to when cultured on a physiological substrate such as dentine. On glass the cells are much flatter and fail to form the membrane domains that are characteristic of resorbing osteoclasts, including the sealing zone/actin ring and the ruffled border (Fig. 1). If culturing on glass, we most commonly use 9-mm diameter coverslips, sterilised in 70% ethanol and inserted into 48-well plates. A more physiologic substrate is hydroxyapatite-coated glass, on which osteoclast resorptive activity (or more accurately in this case, the ability to acidify the extracellular environment) can be assessed (19).

3.2.2. Culture of Osteoclasts on Dentine

Dentine is our preferred physiological substrate for osteoclast cultures. While bovine cortical bone is also highly suitable, the presence of Haversian systems can sometimes complicate interpretation of the images. 1. Before sterilisation in 70% ethanol, label dentine discs with a non-symmetrical identifier on one side with pencil. When inserting into 96-well pates, ensure the marked side faces down. This marking will prevent any confusion over which side the cells are on when handling the discs after culture. 2. Prior to seeding cells, pre-soak dentine in culture medium for 30 min. Details of seeding protocols of osteoclasts on to dentine are available in various chapters in this volume dealing with osteoclast generation or isolation. For imaging purposes, however, it is sometimes better to culture osteoclasts at slightly lower density than for quantification of resorption, to achieve optimum image clarity. 3. If resorbing cells are to be studied, 24–48 h is the ideal incubation time for mature osteoclasts. The osteoclasts will start resorbing the dentine within a few hours of seeding, and within 24–48 h should have resorbed sizable pits and trails on the dentine. It is useful to prepare extra “sentinel” dentine discs, which can be removed to check for resorbing activity before fixing the remainder of the experiment. 4. Before fixation, label the cells with fluorescent probes (Subheading 2.4) as appropriate. If analysing cells overexpressing fluorescent protein conjugates, for example, it is possible to reduce the cytoplasmic background in highly expressing cells

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by incubating in a mild permeabilisation buffer (0.1% saponin in PIPES buffer, pH 6.8) for 15 min prior to fixation or live cell imaging. 5. At the end of the desired culture period, rinse cultures in PBS and fix for 10 min in 4% paraformaldehyde in PBS, or the fixation method most compatible with the antibody to be used for immunostaining. Sucrose (2%) can be included in the fixative to prevent osmotic fluxes that may cause non-specific endocytosis which could alter cell morphology. 6. Store in PBS at 4 C until ready for staining and mounting. 3.3. Staining of Osteoclasts

1. If immunostaining for intracellular antigens, it is essential to permeabilise the cells prior to staining; we routinely incubate for 15 min with 0.5% (v/v) Triton X-114 in PBS. If staining with wheat germ agglutinin for the cell membrane only, then this step should be omitted. If staining the dentine with fluorescent BPs, permeabilisation is essential to ensure that the compound penetrates beneath the osteoclasts. 2. Staining can be carried out with the osteoclasts left in the 96-well plate, using ~50 μl per well. Alternatively, to save on reagents, the discs can be removed and placed on a piece of Parafilm in a humidified chamber during the staining, in which case <50 μl per disc can be used. Staining routines should be optimised for each antibody; in most cases we block for 1 h with 10% goat serum in PBS (the species of the secondary antibodies that we use), then incubate with primary and secondary antibodies for 1 h at room temperature, or overnight at 4 C, in PBS containing 5% goat serum. 3. There is a huge choice of fluorophore-conjugated secondary antibodies available. We tend to use affinity purified antibodies raised in goat, conjugated to Alex Fluor fluorophores (Invitrogen), which have a high quantum yield, good photostability and pH-independent fluorescence (unlike some fluorophores such as fluorescein). For our LSM510 confocal, which has laser lines at 488, 546, and 633 nm, Alexa Fluor 488, Alexa Fluor 555, and Alexa Fluor 647-conjugate antibodies are the best options. 4. Organelle probes such as phalloidin or fluorescent BPs (see Subheading 3.1) can be included with the secondary antibody incubation, or stained separately afterwards. 5. After staining, the discs should be washed at least four times in PBS, then transferred to a glass slide. The easiest way to do this is to push a 25-gauge needle into the disc and then gently remove on to the slide using forceps. 6. Mount the discs under a cover slip. It is possible to put several discs on each slide, then add a drop of VectaShield as an anti-fade

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mountant (see Note 4). After placing a suitably sized coverslip over the top, seal the edges with nail varnish (see Note 5). Slides may be stored in the dark until analysis by confocal microscopy. 3.4. Confocal Microscopy of Osteoclasts

1. Find the sample to be scanned through the eyepiece using the 10× objective, before further analysis using higher magnification objectives. For osteoclast imaging we most commonly use a 40× oil immersion objective (Zeiss Plan-NEOFLUAR) with a high numerical aperture (1.3); higher magnification objectives are rarely required due to the size of the osteoclasts. 2. Select the desired scanning configuration, then set the pinhole to 1 Airy unit for the optimal image resolution; this setting will usually be stored in the configuration setup (Fig. 3; see Note 6). The optical section depth will depend on the lens being used and the fluorophore being imaged, for example with a 40× lens (see step 3 below), this will be 0.9 μm for a green emitting fluorophore such as fluorescein. If scanning multiple fluorophores, 1 Airy unit should be set for the capture of the fluorophore with the longest wavelength, then the pinholes for the other fluorophores adjusted to ensure an identical optical section in each case. 3. Select the desired data depth; either 8-bit (256 levels of intensity) or 12-bit (4,096 levels of intensity; see Note 7) and the image resolution (512 × 512 pixels is usually the default setting; see Note 8). 4. Image using sequential line scanning. In this setup, the laser will scan each line with each laser in turn (i.e. only one laser on at a time), before moving to the next line. This sequential rather than simultaneous scanning reduces the chances of bleed-through between channels during acquisition. 5. Select the ideal focal plane in the specimen to be imaged while continually scanning. One issue if analysing osteoclasts on dentine discs is that it is difficult to ensure that they are completely flat when mounted on to slides. Therefore when imaging by confocal, it may not be possible to see all the dentine surface in the field of interest within the optical section (see Note 9). 6. After the initial scan, select the specific region required for the final scan (often available as a crop tool). This area will then be scanned at the same user-defined resolution (e.g. 512 × 512 pixels). Care should be taken to avoid unnecessary oversampling (see Note 10). 7. Optimise the detector settings (using the detector gain and offset controls) for each fluorophore to be captured in turn (see Note 11). It is usually recommended to this by single scanning, altering the gain and offset to improve the image

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between scans, rather than continuous scanning, to avoid excessive exposure of the sample to the excitation wavelengths prior the final scan. Capture of a brightfield image (e.g. phase contrast) can also be set as one of the channels, and can be useful for demonstrating cell morphology of cells cultured on coverslips. However, cells cultured on bone or dentine cannot be visualised using transmitted light. 8. Capture the final scan. At this point, it may be helpful to use averaging, particularly if one or more of the fluorophores is weak, and consequently there is a lot of noise in the image. This works by taking a user-defined number of repeat scans, then averaging the fluorescence intensity from these scans for each pixel in the image. Since the positive signal should appear in every scan, while the noise will appear randomly, averaging should significantly reduce noise without affecting the genuine signal, thereby improving signal to noise ratio. On our system, as a general rule, if the detector gain needs to be above 600 then averaging is helpful, and the higher it is above this value, the greater the number of scans that should be used to improve the image. 3.5. Visualisation of the Data Post-acquisition

1. It is important to remember that the PMTs detect only photons, and that any colour represented in images from a confocal microscope are pseudocolours that can be user selected for each channel scanned (see Note 12). In 3-channel imaging, green, red and blue are almost universally used, with blue usually reserved for the near infra-red wavelength if using such a fluorophore (see Note 13). 2. When analysing a single fluorophore, fluorescence intensity can be displayed as a “heat map” rather than greyscale, which can help to bring out dim features more clearly (Fig. 4b). 3. All the parameters used to scan an image are embedded in the image file, which means that it is straightforward to insert scale bars, measure point to point distances and to determine the area of structures of interest using freeform selection tools. 4. Fluorescence intensity can be measured at selected points, or across a line of interest (“profile” views; Fig. 5b). This can be particularly useful for revealing the presence of low levels of a particular fluorophore that is not evident to the naked eye due to the additional presence of a different fluorophore. 5. Colocalisation analysis. Confocal software enables colocalisation between two or more fluorophores to be determined on a pixel-by-pixel basis (Fig. 5a). Thresholding of each channel enables “gating” of the pixels exhibiting colocalisation, similar to the way in which cell populations are sorted using FACS. It is important to remember the limitations of resolution when

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Fig. 5. Colocalisation of fluorophores in confocal images. (a) Rabbit osteoclast cultured on glass with AF-ALN for 4 h (left panel) and with acidic vesicles stained using lysotracker red (right panel ). Threshold intensities correlating to positive staining were selected for each fluorophore, as indicated in the scatter plot. Pixels in region 3 (i.e. those exhibiting colocalisation of both fluorophores) are shown in the lower left panel. (b) Assessment of colocalisation by measuring intensities across a region of interest. Human osteoclast on glass stained with wheat germ agglutinin-Alexa Fluor 488 (green) and TRITC-phalloidin (red ). Intensities are shown across the region denoted by the arrow in the image. Note that not only is there a high degree of spatial colocalisation, but also that the intensity of each fluorophore across the region of interest is extremely similar.

interpreting such data, however, particularly in the axial dimension. For example, although two proteins localising to small intracellular vesicles may appear to be colocalised, they could actually reside on separate structures, since structures such as lysosomes may be only 100 nm in diameter, while axial

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resolution is typically only 500 nm, i.e. insufficient to resolve individual lysosomes of this size. 6. For more powerful analysis than may be available in the manufacturer’s software, there are numerous specialised post-acquisition image analysis software packages (see Subheading 2.5). 3.6. Generating Three-Dimensional Data Sets

Collecting a “z-stack” of images in order to reconstruct 3D images or sectional views (in any plane) of the sample requires a microscope stage that is motorised in the axial dimension to enable the extremely small increments of position (often sub-micron) to be achieved. Therefore, this is a universal requirement of confocal microscopes (see Note 14). Generating z-stacks is extremely useful when analysing resorbing osteoclasts on a physiological substrate such as dentine. However, it is generally not necessary for osteoclasts cultured on glass coverslips, due to the lack of depth of the cells. 1. First, the top and bottom positions of the sample to be scanned need to be defined (for resorbing osteoclasts, this would usually be from the top of the cell of interest to the bottom of the resorption pit) by focusing through the sample while continuously scanning. This should be carried out as swiftly as possible to reduce photobleaching. Importantly, when z scanning, the setting for the PMTs has to be optimised considering the entire depth of the specimen to be scanned. For example, if imaging F-actin, by far the highest signal in resorbing osteoclasts will be from the F-actin ring, with much weaker signals from the cortical F-actin close to the plasma membrane (see Note 15). 2. Once the top and bottom of the sample have been specified, the step interval needs to be set. This will depend on the axial resolution of each image (i.e. the depth of the optical section), which itself is dependent on the objective being used and the pinhole diameter (see Subheading 2.1). According to Nyquist sampling theory, the optimal interval will be half that of the optical slice. For resorbing osteoclasts analysed with a 40× objective, this could easily require 50 or more scans (depth of the specimen may be 25 μm or more); a 40× objective would give an optical slice of around 1 μm for fluorescein using 1 Airy unit. 3. Apply the desired frame averaging to reduce noise in the sample. Scan time and potential photobleaching is now even more of a major consideration, bearing in mind the number of scans required in the stack. 4. Start scanning. The sample will be scanned from the top to the bottom, generating a gallery of images; note that this may take several minutes to complete.

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1. Once the data from the z-stack has been generated, the software will enable immediate sectional views (xz and yz) in the axial dimension to be displayed (Fig. 4; see Note 16). Such views are useful, for example, in determining the potential localisation of intracellular proteins at the ruffed border. 2. By altering the pitch and yaw of the data set being analysed, it is possible to “cut” such sections through any part of the sample, as desired. 3. Since a volumetric data set is collected during a z-stack scan, it is possible to generate 3D reconstructions. A useful way of viewing the data from the whole image series in a single xy 2D image is to create a maximum intensity projection (MIP) view (Fig. 6). This method applies the maximum intensity of the fluorescence throughout the z-stack at each pixel (rather than the sum of the fluorescence, which would result in many saturated areas and loss of image clarity). This can be useful for determining ruffled border localisation of proteins, in which case staining will be delineated by the actin ring. 4. Some confocal image capture programs enable instant MIP views of the data from any angle. We use Zeiss Zen acquisition software, which enables this simply by rotating to the desired angle by dragging the image (Fig. 6a). Such images can be processed further by isosurface rendering, in which the surfaces of stained structures within the image are detected and assigned an opaque colour (see Note 17). This can be particularly useful to identify the surface of stained dentine in 3D views (Fig. 6c). 5. Depth coding can be used to denote the axial position of a fluorophore in a single 2D image. In this case, only a single fluorophore can be displayed, since the position in the axial dimension is denoted by false-colouring to a look-up table (Fig. 4c). Note that fluorescence intensity information will be lost if using this option. 6. Three-dimensional data can also be displayed as red–green anaglyph images for viewing with red–green glasses. This option is often available within the image acquisition software, and provides a more vivid illustration of the axial dimension within the image (Fig. 5b). Since this relies on the use of red and green images to encode the depth information, it can only be applied to a single fluorophore in an image. 7. The reconstructions outlined above show the data from a single point of view. Creating a 3D animation using multiple points of view can give the viewer a far better visual impression of the spatial organisation of the stained structures in a sample. This feature is most powerful in specialised post-acquisition imaging software, such as Imaris or Volocity. Movies can be

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Fig. 6. Representation of 3D confocal data sets. Images show osteoclasts cultured on dentine and stained with AF-ALN (green), TRITC-phalloidin (red ) and the vitronectin receptor (blue). (a) Left panel shows single xy image at the level of the dentine surface; middle panel shows a MIP image off the same cell, combining images from the top of the cells to the bottom of the resorption pit; right panel shows a MIP image from an angle closer to the plane of the dentine. (b) Anaglyph stereo images (for viewing with red–green glasses) of the osteoclast shown in (a); left panel shows the F-actin, middle panel shows the vitronectin receptor; right panel shows the AF-ALN. (c) Different osteoclast showing F-actin and AF-ALN only; left panel shows MIP image; middle panel shows MIP image from an angle closer to the plane of the dentine; right panel shows isosurface rendering from the same angle as the image in the middle panel; note the increase clarity with the rendered image.

constructed in which the sample can be “flown through” and different channels switched on or off at desired times or cut away at selected positions. 3.8. Spectral Imaging

On many confocal systems it is possible to distinguish between fluorophores that have closely overlapping emission spectra, by using a technique known as spectral imaging, for which a number of different strategies have been developed. On the Zeiss LSM510

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META system that we use, this is achieved by using an array of detectors, each of which detects only a small specific range of wavelengths of light (10 nm per detector). In this way, it is possible to build up a fluorescence emission profile for each pixel that has been scanned, then identify the fluorophore by comparing this to prescanned reference spectra for the fluorophores present in the sample (see Note 18). Since autofluorescence also has a characteristic emission spectrum, this can be set up as a reference and then removed from the final image after unmixing the emission spectra. This approach also opens up the possibility of scanning and distinguishing more than 3 fluorophores in a sample. Remarkably, it has proved possible to detect an unprecedented 90 different fluorescence spectra in a single sample of labelled neurons using an Olympus FV1000 confocal microscope (24). One slight drawback of this approach using our LSM510 META is that the light is split into several detectors at any one time, therefore reducing the sensitivity of the scan. However, with recently developed confocal systems, such as the Zeiss LSM710, spectral imaging is now possible without any loss of sensitivity. 3.9. Live Cell Imaging Using Confocal Microscopy

Live imaging of osteoclasts is possible using confocal microscopy, although there are a number of factors that need to be considered when using this technique rather than wide-field fluorescence microscopy. The major disadvantages of confocal microscopy are the relatively slow scan times and phototoxicity, which is caused by the generation of hydrogen peroxide by the scanning laser. However, improvements in optical performance and detector sensitivity have helped to reduce these problems by enabling much quicker scan times. Particular applications of confocal microscopy in live cells include dynamic studies of fluorescence recovery after photobleaching (FRAP) and the use of photoactivatable fluorophores (e.g. paGFP; see Subheading 3.10 below) (25), respectively, which rely on a high-power scanning laser to bleach and activate fluorescent probes, respectively, in user-defined regions of interest. FRAP has been used in osteoclast studies to elucidate the role of c-src and Pyk2 in podosomes dynamics within the F-actin ring, by expressing GFP-actin in osteoclasts from mice lacking these proteins compared to those from wild-type mice (26, 27). Please refer to Chapter 26, this volume, for more information on live imaging of bone cells.

3.10. Advanced Uses and Probes for Confocal Microscopy

Huge advances have been made over the last few years in the development of fluorescent probes for confocal microscopic approaches. While these have mostly yet to be utilised in the study of osteoclasts, in the future they should prove enormously useful to furthering our understanding of the cell biology of the osteoclast. The development of variants of GFP has led to novel techniques such as fluorescence resonance energy transfer (FRET). This technique has become widely used to determine if two proteins are interacting in intact cells, and works on the principle that a

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donor fluorophore in an excited electronic state may transfer its excitation energy to an acceptor fluorophore through non-radiative means, but only when the fluorophores are within of ~10 nm of each other (reviewed in ref. 28). The most common FRET pair used are the GFP variants CFP and YFP (cyan and yellow emission, respectively). Photoactivatable fluorophores enable an overexpressed protein to be visualised only in a user-defined region of interest. In the case of paGFP, illumination by a 405-nm laser causes a structural alteration, activating the paGFP, which can then be visualised using the normal excitation/emission set up for GFP (29). This is a particularly powerful probe in live cell imaging studies, enabling chase experiments or tracking of individual subcellular vesicles, for example, to be carried out. Probes have been developed, such as Kaede, that alter their fluorescence emission on exposure to UV light. Kaede fluoresces green, but after UV illumination, it is cleaved to a stable form that fluoresces bright red (30). Since photoconversion is efficient using standard mercury lamp illumination and a typical UV (DAPI) filter set, a UV laser is not required. 3.11. Non-confocal Methods for Achieving Optical Sectioning of a Specimen

There are several alternatives to confocal microscopy that are also able to achieve optical sectioning of fluorescently labelled biological samples, which will be mentioned briefly here. First, it is possible to use deconvolution, which utilises mathematical algorithms to eliminate out of focus light in z-stacks of images obtained using conventional fluorescence microscopy. Generally, this method is preferred for thin and relatively dim samples. Another technique that uses conventional wide-field illumination is structured illumination (such as the Zeiss ApoTome), which employs a grid system to achieve optical sectioning approaching confocal quality. This system has the advantage that it is relatively cheap, but it lacks the versatility of the confocal microscope. Finally, multiphoton microscopy, which relies on excitation of the fluorophore only at a specific, selected depth within the sample, usually achieved by the convergence of two laser lines, each with wavelengths twice that required for excitation of the fluorophore. Only where these two beams meet will the energy equal that required for excitation of the fluorophore of interest. However, the equipment is extremely expensive and is really specialised for deep tissue imaging, particularly in living tissues, as the long wavelengths used cause much less photodamage than does LSCM.

4. Notes 1. Far-red fluorophores are invisible to the naked eye, so cannot be seen when examining the stained specimen through the eyepiece. It is therefore advised to reserve this channel for a

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stain which does not need to be visualised to identify the preferred region for scanning. 2. Bone and dentine autofluorescence mainly at short wavelengths (i.e. in the blue–green of the visible spectrum). Therefore, if possible it is better to stain osteoclasts cultures on such substrates with longer wavelength fluorophores (such as red and near infra-red wavelengths), particularly if the staining is expected to be relatively weak. 3. We have also labelled ALN using a near infra-red fluorophore, Alex Fluor 647, enabling us to choose which is the most appropriate on the basis of the other fluorophores to be used in the sample. 4. VectaShield is available in two forms, one which remains liquid and one which sets upon incubation at room temperature for 20 min (VectaShield HardSet). While both can be used, the latter makes slides with dentine discs mounted on them easier to deal with. Invitrogen also produce a mountant that hardens after 24 h (ProLong® Gold). 5. The thickness of the dentine discs means that there will be a similar-sized gap between the slide and coverslip, so quite a lot of nail varnish can be required. This can be more exaggerated if the discs are not completely flat, which can also result in the discs being able to move slightly under the coverslip during analysis. One way round this is to attach other coverslips to the slide using nail varnish, around the region that the dentine will be mounted, to reduce this gap. 6. The Airy unit refers to the diameter of the Airy disk of a fluorophore, which is the inner, intense light circle of the diffraction pattern from this source of light. This setting will allow ~95% of the light from the Airy disk through, while preserving the best confocal image. 7. When selecting data depth, consideration should be given to how the images will be analysed. For instance, if they are for visual purposes only, then 8-bit (256 shades of grey) is sufficient, as this is beyond the resolving power of the human eye. If fluorescence intensities are to be measured, then it is worth considering capturing 12-bit data (4,096 shades of grey). 8. When selecting image resolution, the impact on scan time and file size are also considerations. Most important, however, is to ensure that the image will be at a resolution suitable for publication (usually printed at 300 dpi). 9. If the dentine surface is not flat, it is recommended to take a series of images in z that will scan the entire dentine surface of interest. This data can then be merged in a maximum intensity projection (MIP) image.

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10. The ability to zoom in and scan selected areas with confocal microscopes introduces the likelihood of oversampling (i.e. capturing an image at a resolution that is beyond the optical capabilities of the microscope). Zeiss confocal microscopes obviate this problem by having an “optimal” function that sets the image resolution to the maximum optically possible with the zoom setting selected. If this function is not available, then for visible light and high numerical aperture objectives (>0.8) a pixel size of ~0.1–0.2 μm is ideal. 11. Care should be taken to ensure that the detector gain and offset is set to utilise the full dynamic range, but without losing intensity data at the top or bottom of the scale. Most systems have a range indicator which shows background (0) levels as blue pixels and maximum intensity (256 for 8-bit images) as red pixels, enabling these settings to be easily optimised. 12. Blue and red offer poor contrast against a black background (the human eye is most sensitive to green light), therefore if imaging a single fluorophore, grey-scale should be used. An extremely effective alternative that can bring out dim features within images is to pseudocolour the image to a rainbow or spectrum look-up table, in which the colour denotes the fluorescence intensity (Fig. 4b). 13. For colocalisation studies of two fluorophores, green and red have historically been the pseudocolours of choice, with colocalisation indicated by yellow pixels. However, this can create problems for people with colour blindness. A useful alternative is to use green and magenta, which shows areas of colocalisation as white. 14. Some confocal systems, such as the Leica SP5, enable axial views to be scanned “live,” without the need for generating a whole 3D dataset before reconstructing the desired view. This achieved by using a galvanometer-driven stage allowing extremely rapid stage control in the axial dimension, and is particularly useful for live cell imaging applications. 15. When imaging deep into tissue, quenching of the fluorescent signal by the tissue above can occur. It is possible to compensate for this by setting a different (higher) detector gain at the bottom of the sample from that at the top (known as auto z correct function on Zeiss systems). The software will then increase the gain through linear increments as it moves from the top to the bottom of the sample. However, this quenching is rarely a problem when imaging osteoclasts cultured on dentine. 16. When scanning in this way, it is important to note that the axial resolution is unlikely to be as high as the xy resolution, leading to “rugby ball” artefacts in axial views, where structures

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appear more elongated than they actually are. Increasing the NA and decreasing the diameter of the pinhole will increase the z-resolution. In practice, the maximum resolution in z (axial) that can be realised in a confocal microscope system is about 0.5 μm; 2–3× worse than in the xy-dimension. 17. Isosurface rendering results in loss of fluorescence intensity information, since its purpose is simply to recreate a surface that exists in the structure under examination. If intracellular structures are to be visualised it is possible to create cut-away images. 18. It is also possible to unmix fluorophores without reference spectra, if it is known that there are individual pixels in the image that contain only one of the fluorophores. The spectra can then be obtained from these pixels, either manually or automatically: in the case of the Zeiss LSM510 META this is done by use of the automatic component extraction (ACE) facility. References 1. Minsky, M. (1988) Memoir on Inventing the Confocal Scanning Microscope. Scanning 10, 128–138. 2. White, J. G., and Amos, W. B. (1987) Confocal Microscopy Comes of Age. Nature 328, 183–184. 3. Taylor, M. L., Boyde, A., and Jones, S. J. (1989) The Effect of Fluoride on the Patterns of Adherence of Osteoclasts Cultured on and Resorbing Dentin - A 3-D Assessment of Vinculin-Labeled Cells Using Confocal Optical Microscopy. Anatomy and Embryology 180, 427–435. 4. Lakkakorpi, P. T., Helfrich, M. H., Horton, M. A., and Väänänen, H. K. (1993) SpatialOrganization of Microfilaments and Vitronectin Receptor, Alpha-V-Beta-3, in Osteoclasts A Study Using Confocal Laser Scanning Microscopy. J. Cell Sci. 104, 663–670. 5. Baron, R., Neff, L., Brown, W., Courtoy, P. J., Louvard, D., and Farquhar, M. G. (1988) Polarized Secretion of Lysosomal-Enzymes – Co-Distribution of Cation-Independent Mannose-6-Phosphate Receptors and LysosomalEnzymes Along the Osteoclast Exocytic Pathway. J. Cell Biol. 106, 1863–1872. 6. Salo, J., Metsikko, K., Palokangas, H., Lehenkari, P., and Väänänen, H. K. (1996) Bone-resorbing osteoclasts reveal a dynamic division of basal plasma membrane into two different domains. J. Cell Sci. 109 , 301–307. 7. Nesbitt, S. A., and Horton, M. A. (1997) Trafficking of matrix collagens through bone resorbing osteoclasts., Science 276, 266–273.

8. Palokangas, H., Mulari, M., and Väänänen, H. K. (1997) Endocytic pathway from the basal plasma membrane to the ruffled border membrane in bone-resorbing osteoclasts. J. Cell Sci. 110, 1767–1780. 9. Stenbeck, G., and Horton, M. A. (2004) Endocytic trafficking in actively resorbing osteoclasts. J. Cell Sci. 117, 827–836. 10. 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. 11. Vääräniemi, J., Halleen, J. M., Kaarlonen, K., Ylipahkala, H., Alatalo, S. L., Andersson, G., Kaija, H., Vihko, P., and Väänänen, H. K. (2004) Intracellular machinery for matrix degradation in bone-resorbing osteoclasts. J. Bone Miner. Res. 19, 1432–1440. 12. Xia, L. H., Kilb, J., Wex, H., Li, Z. Q., Lipyansky, A., Breuil, V., Stein, L., Palmer, J. T., Dempster, D. W., and Brömme, D. (1999) Localization of rat cathepsin K in osteoclasts and resorption pits: Inhibition of bone resorption and cathepsin K-activity by peptidyl vinyl sulfones. Biol. Chem. 380, 679–687. 13. Bruzzaniti, A., Neff, L., Sandoval, A., Du, L., Horne, W. C., and Baron, R. (2009) Dynamin reduces Pyk2 Y402 phosphorylation and SRC binding in osteoclasts. Mol. Cell Biol. 29, 3644–3656. 14. Lakkakorpi, P. T., Nakamura, I., Nagy, R. M., Parsons, J. T., Rodan, G. A., and Duong, L. T. (1999) Stable association of PYK2 and

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F.P. Coxon p130(Cas) in osteoclasts and their co-localization in the sealing zone. J. Biol. Chem. 274, 4900–4907. Toyomura, T., Murata, Y., Yamamoto, A., Oka, T., Sun-Wada, G. H., Wada, Y., and Futai, M. (2003) From lysosomes to the plasma membrane: localization of vacuolar-type H+-ATPase with the a3 isoform during osteoclast differentiation. J. Biol. Chem. 278, 22023–22030. Zhao, H., Laitala-Leinonen, T., Parikka, V., and Väänänen, H. K. (2001) Downregulation of small gtpase rab7 impairs osteoclast polarization and bone resorption. J. Biol. Chem. 276, 39295–39302. 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. Pavlos, N. J., Xu, J., Riedel, D., Yeoh, J. S., Teitelbaum, S. L., Papadimitriou, J. M., Jahn, R., Ross, F. P., and Zheng, M. H. (2005) Rab3D regulates a novel vesicular trafficking pathway that is required for osteoclastic bone resorption, Mol. Cell Biol. 25, 5253–5269. Saltel, F., Destaing, O., Bard, F., Eichert, D., and Jurdic, P. (2004) Apatite-mediated actin dynamics in resorbing osteoclasts. Mol. Biol. Cell 15, 5231–5241. 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 Coxon, F. P., Taylor, A., Van Wesenbeeck, L., and Van Hul,W. (2009) Plekhm1 is involved in trafficking of cathepsin K-containing endosomal vesicles in osteoclasts. Bone 44, S248.

22. Erwig, L. P., McPhilips, K. A., Wynes, M. W., Ivetic, A., Ridley, A. J., and Henson, P. M. (2006) Differential regulation of phagosome maturation in macrophages and dendritic cells mediated by Rho GTPases and ezrin-radixinmoesin (ERM) proteins. Proc. Natl. Acad. Sci. USA. 103, 12825–12830. 23. Thompson, K., Rogers, M. J., Coxon, F. P., and Crockett, J. C. (2006) Cytosolic entry of bisphosphonate drugs requires acidification of vesicles following fluid-phase endocytosis, Mol. Pharmacol. 69, 1624–1632 24. Livet, J., Weissman, T. A., Kang, H. N., Draft, R. W., Lu, J., Bennis, R. A., Sanes, J. R., and Lichtman, J. W. (2007) Transgenic strategies for combinatorial expression of fluorescent proteins in the nervous system. Nature 450, 56–62. 25. Lippincott-Schwartz, J., Altan-Bonnet, N., and Patterson, G. H. (2003) Photobleaching and photoactivation: following protein dynamics in living cells, Nature Cell Biol. 5, S7–S14. 26. Destaing, O., Sanjay, A., Itzstein, C., Horne, W. C., Toomre, D., De Camilli, P., and Baron, R. (2008) The tyrosine kinase activity of c-Src regulates actin dynamics and organization of podosomes in Osteoclasts. Mol. Biol. Cell 19, 394–404. 27. Gil-Henn, H., Destaing, O., Sims, N. A., Aoki, K., Alles, N., Neff, L., Saniay, A., Bruzzanitti, A., De Camilli, P., Baron, R., and Schlessinger, J. (2007) Defective microtubule-dependent podosome organization in osteoclasts leads to increased bone density in Pyk2(−/−) mice. J. Cell Biol.178, 1053–1064. 28. Kenworthy, A. K. (2001) Imaging protein-protein interactions using fluorescence resonance energy transfer microscopy. Methods 24, 289–296. 29. Patterson, G. H. and Lippincott-Schwartz, J. (2002) A photoactivatable GFP for selective photolabeling of proteins and cells. Science 297, 1873–1877. 30. Ando, R., Hama, H., Yamamoto-Hino, M., Mizuno, H., and Miyawaki, A. (2002) An optical marker based on the UV-induced greento-red photoconversion of a fluorescent protein. Proc. Natl. Acad. Sci USA. 99, 12651–12656.

Chapter 26 Live Imaging of Bone Cell and Organ Cultures Sarah L. Dallas and Patricia A. Veno Abstract Over the past two decades there have been unprecedented advances in the capabilities for live cell imaging using light and confocal microscopy. Together with the discovery of green fluorescent protein and its derivatives and the development of a vast array of fluorescent imaging probes and conjugates, it is now possible to image virtually any intracellular or extracellular protein or structure. Traditional static imaging of fixed bone cells and tissues takes a snapshot view of events at a specific time point, but can often miss the dynamic aspects of the events being investigated. This chapter provides an overview of the application of live cell imaging approaches for the study of bone cells and bone organ cultures. Rather than emphasizing technical aspects of the imaging equipment, we have focused on what we consider to be the important principles that are of most practical use for an investigator setting up these techniques in their own laboratory, together with detailed protocols that our laboratory has used for live imaging of bone cell and organ cultures. Key words: Live cell imaging, Extracellular matrix, Osteocytes, Bone cells, Dynamic imaging

1. Introduction Much of our understanding of mineralized tissue biology has come from the use of static imaging approaches, such as light and electron microscopy, combined with chemical and biochemical analysis and/ or molecular genetic approaches (1–4). However, the biological processes occurring in mineralized tissues, such as bone formation, remodelling and fracture healing, are dynamic events that have the added dimension of time. In contrast to static imaging, live cell imaging enables visualization of temporal changes in living specimens, such as cells, tissues or whole embryos, and allows quantitation of cellular, subcellular and tissue behaviour as a function of time. Live cell imaging techniques have been applied to the study of embryonic development/morphogenesis, stem cell biology, and to obtain quantitative insights into various cellular processes, as well as into assembly and reorganization of the extracellular matrix (5–14). Current technologies, using fluorescent molecular and pH-sensitive 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_26, © Springer Science+Business Media, LLC 2012

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dyes or recombinant fluorescent proteins, can label virtually any intracellular or extracellular structure. Together with the application of imaging techniques such as FRET (fluorescence resonance energy transfer), FRAP (fluorescence recovery after photobleaching) and laser confocal microscopy, it is possible to obtain biophysical, biochemical, spatial/temporal and kinetic information on cells, subcellular components and proteins (reviewed in refs. 9, 11, 14–19). These approaches have enhanced our understanding of many processes fundamental to morphogenesis, development and in vivo cell function. Technology for imaging of living cells is now advancing at a staggering rate and it is likely that these approaches will soon be standard in most mineralized tissue research laboratories. It is difficult in this chapter to present a “one size fits all” protocol for live imaging of bone cell and organ cultures, since the specifics of the experiments will be dependent on the configuration of the imaging equipment available to the investigator and the specifics of the biological events being investigated. However, in this introduction, we will first provide an overview of general considerations for live cell imaging that are relevant for most standard live imaging systems. This is followed by detailed protocols in the materials and methods sections that our laboratory has used for live imaging of bone cells and bone organ cultures. Rather than emphasizing technical aspects of the imaging equipment, etc., we have highlighted what we consider to be the important principles that are of most practical use for an investigator setting up these techniques in their own laboratory. 1.1. General Considerations for Live Imaging of Cultured Bone Cells and Bone Organ Cultures

The main considerations for live cell imaging experiments include ensuring that the microscopy equipment is suitable for the biological events being imaged, ensuring that the cells or tissues can be maintained at physiological temperature, pH, osmolarity, etc., minimizing potential phototoxic effects of the imaging process, selecting appropriate imaging probes (and being aware of their strengths and limitations), minimizing focal drift during the time lapse acquisition, and handling the large amounts of data that are generated by time lapse imaging experiments.

1.2. General Considerations About Equipment

There are many different microscope configurations suitable for live cell imaging in bone cell and organ cultures from manufacturers such as Zeiss, Nikon, Olympus, and Leica. These include widefield epifluorescence, confocal, and multi-photon microscope systems among others. It is beyond the scope of the current chapter to review the advantages and disadvantages of each of these systems, but regardless of the system, there are certain requirements of the microscope configuration and computer software that are optimal for live cell imaging, as summarized below: Incubation cabinet: We recommend a fully enclosed 37°C incubation cabinet that surrounds the microscope, including the

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specimen stage and lenses. This minimizes focal drift due to alterations in the ambient room temperature that could change the air temperature between the lenses and the sample. For long-term mammalian cell culture it is preferable to use bicarbonate buffered medium, therefore a 5% CO2 environment is required. This is generally achieved via a humidified CO2 hood that covers the sample, since filling the entire incubation cabinet with humidified CO2 could potentially corrode microscope components. Humidification is important to prevent changes in osmolarity due to evaporation of the culture medium. Camera: For live cell imaging with fluorescent probes, a high resolution, high-sensitivity CCD camera is essential to collect the maximum signal while minimizing exposure times. Highresolution cameras can be “binned” by adding together pixel arrays to increase the signal-to-noise ratio, which reduces the exposure times needed. Our laboratory routinely uses 2 × 2 binning for time lapse imaging of bone cells. Monochrome cameras are more sensitive than colour cameras, which should be avoided. Imaging of very rapid biological events (e.g. less than a second) requires a high temporal resolution camera capable of capture rates of 100 frames/s or more and high sensitivity to capture sufficient signal within a very short time. High quality lenses: High numerical aperture (NA) lenses are required for maximum collection efficiency of the emitted light. However, this generally means a shorter working distance and many of these lenses are configured only for imaging through a 0.17-mm glass coverslipped surface. Therefore, coverslip bottomed culture chambers are required. Water immersion lenses can also be used, which may increase the working distance while still retaining a reasonably high numerical aperture. Optics should be configured for brightfield and differential interference contrast (DIC) as well as fluorescence illumination. Highly stable light source: A highly stable light source is essential for live cell imaging. This is particularly important for long-term (more than a few hours) fluorescence imaging and in experiments where quantitative analyses are to be performed. A standard mercury arc lamp or metal halide lamp source may not have sufficient long-term stability, which can lead to “flashing” of one or more image frames within an image sequence (due to an increase in intensity of all the pixel values). Because this is caused by fluctuations in the lamp intensity, the problem is exacerbated with longer exposure times (e.g. when imaging a fluorescent probe that is not very bright). Such flashing will make it very difficult to obtain quantitative data; therefore a highly stable light source is recommended, such as the Exfo Exacte metal halide lamp (Exfo Life Sciences, Inc., Mississauga, ON, Canada). Recently, LED-based illumination systems have come on the market, which may also have good stability for live imaging.

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High accuracy motorized x, y, and z stages: Accurate motorized x, y, and z stages are essential to enable imaging of multiple fields (e.g. control and treated cells) and to allow acquisition of information from multiple z-planes. Linear encoded stages should be used. The microscope should be mounted on an anti-vibration table to isolate the system from external vibrations. Software requirements: Software should be configured to drive all motorized components of the microscope system (e.g. x, y, stage, z motor, light source, filter wheel, camera shutter, condenser turret, etc.). The software should support “multidimensional imaging” (i.e. allowing you to program time lapse settings, multiple stage positions, multiple wavelengths, multiple z-planes, etc., within a single user interface). There are many microscope configurations from different vendors that satisfy the above conditions. The widefield epifluorescence system routinely used by our laboratory is illustrated and described below in Fig. 1.

Fig. 1. Widefield epifluorescence microscopy system used for live cell imaging of bone cells in our laboratory. The system consists of a Nikon TE2000E inverted microscope (Nikon, Inc., Melville, NY) with a 50-nm accuracy linear encoded motorized z stage, a Prior x, y stage and stage controller (Prior Scientific Inc., Rockland, MA) and CFI Plan Apochromat lenses (×4, 0.2 NA, 15.7 mm WD; ×10 DIC, 0.45 NA, 4 mm WD; ×20 DIC, 0.75 NA, 1 mm WD; ×40 DIC, 0.95 NA, 0.14 mm WD). The microscope system is interfaced with a Photometrics Coolsnap HQ cooled CCD camera with 12-bit grey-scale resolution (Roper Scientific, Ottobrunn, Germany) and an Exfo Exacte highly stable metal halide lamp source (Exfo Life Sciences, Inc., Mississauga, Ontario, Canada). The system is enclosed in a customized “Cube in a Box” incubation system and 5% CO2 is delivered via a CO2 hood connected to a humidifier and “The Brick” gas mixer (Life Imaging Services, Reinach, Switzerland). Image acquisition and all hardware components are controlled by the Metamorph software (Molecular Devices, Sunnyvale, CA). The microscope system is mounted on an anti-vibration table to isolate the system from external vibrations.

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When performing live cell imaging, a compromise exists between obtaining a high enough signal-to-noise ratio for quantitative measurements and obtaining sufficient image resolution, while at the same time avoiding phototoxic effects to the cells (for review, see Frigault et al. (11)). To preserve cell viability, the investigator may have to accept a lower image quality and resolution than would be appropriate for equivalent images of fixed specimens. Photodamage is mainly caused by the oxygen-dependent reaction of free-radical species with cellular components. These free radicals are generated when fluorescent proteins or dyes are excited. For live cell imaging, it is important to minimize the amount of excitation light by optimizing the efficiency of the light path through the microscope and maximizing collection of the emitted signal (e.g. by using high NA lenses). The potential for phototoxicity to the cells is dependent on several factors including (1) the number of fluorophores being imaged and their excitation wavelength(s) (wavelengths towards the blue and ultraviolet end of the spectrum are more phototoxic than those towards the red end); (2) the intensity of the light and the exposure time (it is preferable to use a lower intensity of light with a longer exposure rather than a high intensity for a shorter exposure); (3) the number and location of fields to be imaged and the number of z-planes to be imaged (we have found it preferable to ensure that imaging fields are well separated, i.e. not overlapping, and spaced at least three field diameters apart), (4) the duration of the experiment and the time interval between image acquisitions (larger time intervals may be needed if the experiments are of long duration so that cells do not accumulate phototoxic damage); (5) the intensity and cellular localization of the fluorophore(s) being imaged (fluorophores with a nuclear localization are more likely to cause photodamage to DNA and the higher the concentration of fluorophore probe, the higher the potential damage). Oxygen free radical scavengers can also be used in the culture medium to protect the cells from free radicals. With these considerations in mind, our routine approach for long-term (24–72 h) time lapse imaging has been to image a maximum of 2–3 image fields per cm2 imaging area, ensuring that these are well separated (i.e. non-overlapping and at least three field diameters apart). We routinely image in DIC plus 1–2 fluorescent channels (avoiding blue fluorophores if possible) and we use between 5 and 11 z-planes, spaced 1–1.5 μm apart, depending on the thickness of the specimen. To reduce phototoxicity we generally reduce the light intensity to 12 or 25% of the maximal Exfo Exacte metal halide lamp output. In addition, if possible, neutral density filter(s) may be placed in the light path to cut down the light intensity. The camera is operated in 2 × 2 binning mode and exposure times are generally between 30 and 200 ms. These conditions should be optimized for each probe. Table 1 summarizes

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Table 1 Suggestions for optimal imaging of live cell specimens using widefield epifluorescence To improve efficiency

• • • •

Use high NA apertures Use filter sets that are optimal for each fluorophore Send 100% of the light to the camera (detection) port Remove DIC prisms and analyzer when imaging in fluorescence (if possible)

To improve sensitivity and signal to noise

• Avoid colour cameras • Use a high resolution, high sensitivity CCD camera • Use binning on high resolution cameras (e.g. 2 × 2)

To minimize light exposure/ phototoxicity

• Keep excitation light levels as low as possible • Avoid fluorophores towards the blue end of the spectrum • Minimize the number of probes being imaged simultaneously • Use oxygen radical scavengers (e.g. ascorbic acid) • Locate the cells of interest using DIC, not fluorescence (if possible) • Minimize exposure of cells to excitation light during selection of imaging fields • Use ND filters to reduce the intensity of the excitation light • Make sure that imaging fields are well separated (if possible)

Modified from Frigault et al. (11)

some suggestions for optimizing imaging using a widefield epifluorescence microscope, which can be used as a starting point/guide when setting up an imaging protocol. 1.4. Selection of Imaging Fields

When selecting imaging fields for time lapse experiments it is important to minimize the amount of time spent viewing the sample under fluorescent illumination, since this could bleach the probe and/or cause phototoxicity. If multiple fields are being selected, it is preferable (within the constraints of the biology being imaged), to make sure that they are spaced well apart to avoid cells being exposed to excitation light from adjacent or overlapping fields. Selection of imaging fields can be more challenging if the events to be imaged have not yet started. For example, we routinely image assembly of extracellular matrix proteins, such as fibronectin, where there is little or no fibronectin probe incorporation at the start of the movie, but by 24–48 h an extensive fibronectin matrix has been deposited (see Subheading 3.3.2). Therefore, at the start of the movie acquisition you do not really know where the “action” will take place. It is also difficult to estimate exposure times for fluorescently labelled structures that have not yet formed. In this situation, we recommend estimating exposure times from a

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b

a

X2

X2

X1

X1

Fig. 2. Schematic diagram illustrating the importance of selecting appropriate time intervals for the dynamic events imaged. In this example, a cell moves from position X1 to position X2 within a 15-min time period. If the motion trajectory approximates that shown in (a), a 15-min time interval between image acquisitions will give a reliable estimate of the cell motion and further refinement of the time interval will not significantly enhance the accuracy of the measurements. In contrast, if the motion trajectory approximates that shown in (b), a 15-min time interval will not give a reliable estimate of the cell motion, since a considerable amount of motion is being missed in between the image acquisitions. In this case, shorter time intervals would be required to generate accurate measurements of the motile properties of the cell.

prior experiment or static culture that represents how the cultures will appear at the end of the imaging period. 1.5. Selection of Time Intervals

Selection of time intervals between image acquisitions is dictated by the biological process being observed, the number of individual movie fields being imaged and the number and wavelength of the fluorochromes. Our laboratory has focused on imaging of bone cell dynamics and assembly dynamics of bone extracellular matrix proteins. These events take hours or days for completion, therefore acquiring images every 15–30 min is appropriate. However, if the event takes only minutes or seconds to complete, then shorter time intervals will obviously be needed. When you are setting up timelapse experiments you should initially select an interval that seems appropriate for the timescale of the event. You should then take images with a smaller time interval to make sure that the interval you have selected is not underestimating or missing important dynamic events (see Fig. 2 for a schematic representation of this concept).

1.6. Types of Probes and Their Limitations

There are many possible fluorescent probes and dyes that can be used in almost limitless combinations for imaging of cells, intracellular and extracellular proteins and molecules. These include; fluorescent antibodies, fluorescently labelled proteins, fluorescent dyes and probes for labelling cell nuclei, organelles and membranes, probes for assessing pH and ion flux, probes for monitoring enzyme activity, etc. (see Note 1). A major advance in live cell imaging was

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the discovery and development of green fluorescent proteins (GFPs) and their derivatives by Roger Tsien, Martin Chalfie and Osamy Shimomura as molecular tools for imaging cells and proteins, for which they won the Nobel prize in chemistry in 2008 (reviewed in Wiedenmann et al. (19)). It is important to remember that each type of probe has its own strengths and limitations and each potentially provides different information. For example, fluorescent antibodies can be used to localize where a particular extracellular matrix (ECM) protein has been incorporated into the matrix and then to monitor where the labelled population of fibrils subsequently goes (20). However, these probes may be less useful for looking at incorporation of new protein into the ECM. Fluorescently labelled purified ECM proteins may be better for monitoring incorporation of new protein into the ECM (see Subheading 3.3.2), but may not recapitulate all the intracellular steps involved in ECM assembly. GFP fusion constructs can also be used to image assembly of extracellular matrix proteins and may do a better job of revealing intracellular steps in the assembly process (21), but the incorporation of approximately 27 kDa of extra sequence into the protein of interest may alter its intracellular trafficking and/or function. Since each probe has advantages and disadvantages, the most informative approach is to use multiple types of probe to address key biological questions. Regardless of the characteristics of the probe, one must always bear in mind that any type of imaging probe has the potential to perturb the cell and alter its normal function and this must always be taken into consideration when interpreting live imaging data. 1.7. Focal Drift

Changes to the microscope focal plane (i.e. the sample drifting out of focus) are a frequently encountered problem in time lapse microscopy, particularly for long-term imaging applications. Focal drift can occur for numerous reasons including (1) changes in the air temperature between the lens and specimen and/or changes in the temperature of the room in which the instrument is housed, (2) changes in the volume of medium in the culture chamber (e.g. evaporation/leakage), (3) changes in humidity, (4) mechanical instability of the z motor due to gear slippage, etc., (5) culture chambers that are not securely mounted on the microscope, (6) “sample drift” (e.g. an organ culture breaking free of its mount and/or contraction or rolling up of cell layers or organ cultures). Without a feedback device to continuously monitor and correct the focus, a fully enclosed incubation chamber that surrounds the microscope (see Fig. 1) is recommended to minimize temperature fluctuations. The entire system should be brought to operating temperature for at least 24–48 h prior to initiating time-lapse imaging experiments, to ensure temperature stabilization. Make sure that the culture chamber is securely mounted on the microscope stage in a manner that does not allow lateral or axial movement.

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Many of the newer microscope systems have features such as Perfect focus (Nikon Instruments, Inc., Melville, NY) or Definite Focus (Carl Zeiss Microimaging, Inc., Thornwood, NY) which correct for focal drift during the experiment and are advantageous, especially for long-term live imaging. 1.8. Data Storage and Data Handling

Data storage is an important consideration when performing live cell imaging, as these experiments generate very large data files. A typical experiment in our laboratory, in which we might image 12–20 separate fields in DIC, two fluorescent channels with 5–9 z-planes, and 192 time points, would generate around 30–50 Gb of image data. Therefore, considerable planning needs to be done regarding handling the storage and backup of data (see Note 2). In Subheading 3.3, we give three examples of protocols that our laboratory has used for live imaging of bone cells and bone organ cultures. These are (1) live imaging of primary osteoblasts using a GFP transgene to monitor differentiation together with alizarin red dye to monitor mineral deposition, (2) live imaging of an osteoblast-like cell line to monitor the dynamic process by which fibronectin is assembled into the bone ECM, (3) live imaging of neonatal mouse calvaria using a GFP transgene to image the motile properties of osteocytes. The specific materials and tissues required are listed below.

2. Materials 2.1. Animals, Cells, and Tissues

1. Neonatal (1–10 days old) transgenic mice expressing the topaz variant of green fluorescent protein (GFPtpz) under control of an 8 kb fragment of the dentin matrix protein-1 (Dmp1) promoter (22). These mice were kindly provided by Drs. David Rowe and Ivo Kalajzic, University of Connecticut Health Center. This transgene has been extensively characterized and shown to give selective expression of GFP in osteocytes, with very low or undetectable expression in osteoblasts (22, 23). 2. 2T3 osteoblast cell line: This clonal osteoblastic cell line was derived from primary osteoblasts isolated from transgenic mice expressing SV40 T-antigen under control of the bone morphogenetic protein-2 (BMP2) promoter. The cell line mineralizes in culture and behaves similarly to primary osteoblasts in a number of assays (24). 3. Primary calvarial osteoblasts isolated from 5 to 7 days old wildtype or Dmp1-GFP transgenic neonatal mouse calvaria as described elsewhere (25). See also Chapter 2 for procedures for isolation of primary osteoblasts.

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2.2. Cell and Tissue Culture Reagents

Unless stated otherwise, all tissue culture media and reagents are purchased from CellGro (Mediatech, Inc., Manassas, VA) or Gibco (Invitrogen Corporation, Carlsbad, CA). Fetal bovine serum is purchased from Hyclone (Logan, Utah) and is heat inactivated. 1. Growth medium for 2T3 cells: Minimum Essential Medium Alpha (α-MEM), supplemented with 10% fetal bovine serum (FBS), 2 mM L-glutamine (LG), 100 U/ml penicillin/streptomycin (P/S). 2. Growth medium for primary osteoblasts: α-MEM, supplemented with 10% FBS, 2 mM LG, 100 U/ml P/S and 30 μg/ ml gentamicin. 3. Culture medium for mineralizing primary osteoblasts: α-MEM supplemented with 10% FBS, 2 mM LG, 100 U/ml P/S, 30 μg/ml gentamicin, 50 μg/ml L-ascorbic acid (add fresh from stock solution on day of use, see step 6) and 0.5–4 mM β-glycerophosphate (β-GP). 4. Medium for culture of neonatal mouse calvaria: BGJb medium supplemented with 5–10% FBS, 2 mM LG, 100 U/ ml P/S, 50 μg/ml L-ascorbic acid (add fresh from stock solution on day of use, see step 6) with or without 5 mM β-GP (see step 7). 5. Dulbecco’s phosphate-buffered saline (PBS), pH 7.2. 6. Ascorbic acid stock solution: 10 mg/ml in α-MEM or BGJb medium (no additives). Sterilize through 0.2-μm filter and store in single use aliquots at −20°C, protected from light. 7. β-glycerophosphate (β-GP) stock solution: 500 mM in α-MEM or BGJb medium (no additives). Sterilize through 0.2-μm filter and store in aliquots at −20°C. 8. Alizarin red vital dye stock solution: 10 mg/ml in 0.9% NaCl2 + 2% NaHCO3, pH 7.4. Sterilize through 0.2-μm filter and store at 4°C protected from light. Make a working stock of 0.1 mg/ml in media for dilution into to the cell medium. 9. 1× Trypsin/EDTA solution (Cellgro, Cat# 25 052C1: 0.05% Trypsin, 54 mM EDTA in Hank’s balanced salt solution without calcium or magnesium). 10. Tissue culture grade human plasma fibronectin (Invitrogen, Cat# 33016-015). This will be used for generating a fluorescent fibronectin probe using a fluorescence labelling kit (see Subheading 2.3, step 1 and Subheading 3.3.2). Since unlabelled fibronectin is also needed, store some single use aliquots of fibronectin at −80°C at 1 mg/ml in PBS.

2.3. Other Buffers, Solutions, and Reagents

1. Alexa 555 protein labeling kit (Molecular Probes, Invitrogen Corporation, Carlsbad, CA). 2. Autoclaved silicone grease.

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3. Autoclaved distilled water. 4. 4% Paraformaldehyde in PBS, pH 7.2: Make as a 2× stock by adding 40 g paraformaldehyde to 250 ml of dH2O. Heat to 60°C in a fume hood and add up to ten drops of 1 N NaOH to help dissolve. When dissolved, cool to room temperature. Add 100 ml of 10× PBS and adjust volume to 500 ml with dH2O. Adjust pH to 7.4 using 1 N HCl and filter. Aliquot and store at −20°C for up to 2 months. Dilute with dH2O to 1× before use and discard any left over. 5. 10% Neutral-buffered formalin (available ready made from most vendors of histology supplies). 2.4. Equipment

1. Nikon TE2000E inverted widefield epifluorescence microscope system configured for live cell imaging (see Fig. 1 for specifications of the equipment). 2. Computer workstation configured with Metamorph offline software (Molecular Devices, Downingtown, PA), ImageJ (free download from NIH) and AutoQuantX AutoDeblur + AutoVisualize deconvolution software (Media Cybernetics, Inc., Bethesda, MD). 3. 2- and 4-Well Lab-Tek chambered coverglass slides (Nalgene Nunc International, Rochester, NY, Cat# 155380 and 155383). 4. T75 and T150 tissue culture flasks. 5. Disposable sterile plastic pipettes for tissue culture (2-, 5-, 10-, and 25-ml sizes). 6. 90-mm Sterile Petri dishes (non tissue culture treated). 7. Sterile microtips (10-, 200- and 1,000-μl size). 8. 4 mm × 35 mm Sterile Petri dishes with sterile tissues or Whatman paper placed in them. 9. 50-ml Falcon tubes. 10. 0.2-μm Sterile filters. 11. Sterilized dissection instruments (straight and curved forceps, straight and curved spring-loaded scissors, straight and curved 3.5″ scissors, large forceps, 4.5″ scissors). 12. Plastic 22 mm × 22 mm coverslips. 13. Single-hole punch. 14. Standard hemocytometer or Coulter counter for counting cell number. 15. Dissection microscope with fibre optic illuminator.

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3. Methods 3.1. Preparation and Culture of Primary Osteoblasts

Use aseptic cell culture technique for all steps and perform all cell isolations/manipulations in a sterile laminar flow hood. All media and solutions to be used for cell and tissue culture should be sterile and should be warmed to 37°C prior to using. 1. Primary osteoblasts are isolated from 5 to 7 day old neonatal calvaria of Dmp1-GFP transgenic mice. The cells are isolated using four sequential 20-min digestions with 0.2% collagenase and 0.05% trypsin in HBSS or α-MEM medium without additives, according to methods described by Kalajzic et al. (25). The first digest is discarded and digests 2–4 are pooled as the osteoblastic population. An extensive description of techniques for isolation of primary osteoblasts is also provided in Chapter 2 and therefore this procedure will not be described here. 2. Plate the primary osteoblasts into T75 culture flasks at 2 × 106 cells per flask in 20-ml growth medium. 3. Culture at 37°C in a 5% CO2 humidified incubator until the cells reach confluency (2–3 days), then wash 1× with PBS and trypsinize the cells with 3 ml trypsin/EDTA solution with incubation for 5 min at 37°C. 4. Transfer the cell suspension to a 50-ml Falcon tube containing 20-ml growth medium for primary osteoblasts and pipette up and down several times to resuspend the cells. 5. Count the cell number using a hemocytometer or coulter counter and dilute appropriately in growth medium to plate for experiments at 2 × 104 per cm2 growth area (see Note 3).

3.2. Preparation of Neonatal Mouse Calvaria

Use sterile dissection instruments for all steps and re-sterilize instruments in 70% ethanol between steps (make sure any ethanol on the instruments is completely dry before using on the bone tissues). Work quickly to avoid any drying out of the tissues during the dissection procedures. 1. Euthanize 2–4 neonatal Dmp1-GFP mice of age 4–10 days by rapid decapitation. 2. Without removing the skin, dip each head very rapidly (<1 s) in 70% ethanol, holding with forceps vertically by the nose. 3. Immediately wash off the ethanol vigorously in a Falcon tube containing 45 ml sterile PBS. 4. Wash again in 45 ml sterile PBS. 5. Store the heads in a 90-mm Petri dish containing 30-ml media for neonatal mouse calvaria. 6. Dissect one head at a time by placing it in a 90-mm Petri dish and reflecting and removing the skin over the calvarium using

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curved scissors and forceps. As you do this, avoid touching the bone surface with the side of the skin that has the hair (the inside of the skin is sterile, but the outside surface is a potential source of contamination). 7. Place the dissection instruments back into 70% ethanol and, using a fresh set of instruments, dissect out the calvarium. This is done by holding the nose of the animal and pushing the point of a pair of curved spring scissors through the skull at the back of the occipital bone. Three cuts are then made as illustrated in Fig. 3. 8. Holding by forceps at the edges, lift out the calvarium and wash sequentially in three 90-mm Petri dishes containing 30 ml sterile PBS. 9. Store individual calvaria in wells of a 12-well plate containing 2-ml media for neonatal mouse calvaria. 10. Repeat steps 6–9 for each mouse head. 11. When you have dissected all of the calvaria, transfer one calvarium at a time to a 90-mm Petri dish containing 30 ml PBS. 12. Under a dissection microscope with a fibre optic illuminator, use fine forceps to remove loose adherent connective tissues, muscle, and large blood vessels/sinuses.

Fig. 3. Diagram illustrating the dissection procedure for isolation of mouse calvarial bones. The parietal, frontal, interparietal, and occipital bones and the sagittal suture are labelled. The dashed arrows labelled 1, 2, and 3 indicate the sequence of cuts that should be made and their direction (microCT image kindly provided by Dr. Yasuyoshi Ueki, University of Missouri, Kansas City).

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13. Carefully remove the periosteum on both halves of the calvarium on the top and bottom surfaces. This is done by gently scraping adjacent to the sutures to expose the edge of the periosteum and then gripping the edge with forceps and gently peeling back. The periosteum comes off like a membrane (see Note 4). Avoid contacting the bone surface in the middle of the parietal bones with the forceps, as this is where you will be imaging. 14. Cut each calvarium into two halves down the central (sagittal) suture and place back into medium for storage prior to mounting for imaging. 3.3. Time-Lapse Imaging Techniques 3.3.1. Time Lapse Imaging of Mineralization Dynamics and Osteocyte Transition in Primary Osteoblasts from Dmp1-GFP Transgenic Mice

In this model system, primary osteoblasts from Dmp1-GFP transgenic mice are cultured under mineralizing conditions. Since the Dmp1-GFP transgene is selective for early osteocytes (22, 23), GFP expression is imaged as an indicator of late osteoblasts beginning the transition to the osteocyte phenotype. By using low concentrations of alizarin red as a vital dye for calcium, the dynamics of mineralization can be imaged simultaneously with osteoblast to osteocyte differentiation (26) (see Fig. 4). 1. Plate primary osteoblasts isolated from Dmp1-GFP transgenic mice (see Subheading 3.1) into 4-well Lab-Tek chambered coverglass slides at 2 × 104 cells per cm2 growth area (see Note 5). Keep in mind that the GFP transgene is not appreciably expressed in osteoblasts and therefore GFP-positive cells will not be observed until the cells begin to differentiate after several days of culture. 2. Culture in a 37°C humidified 5% CO2 incubator for 2–3 days in 0.8-ml growth medium per well until the cells reach confluence. 3. Change the medium to 0.8-ml medium containing 50 μg/ml ascorbic acid, and 0.5 mM β-GP (see Note 6). Culture for a further 5–7 days (changing medium every 3 days) until you see small clusters of GFP-positive cells appearing. These demarcate the location where mineralization will occur. 4. Change the medium to 0.8-ml mineralizing medium containing 50 μg/ml ascorbic acid, and 4 mM β-GP, with 0.5 μg/ml alizarin red dye. 5. In the sterile hood replace the plastic lid of the Lab-Tek slide with a sterile 24 mm × 50 mm glass coverslip and use a small dab of sterile silicone grease on each corner to keep the coverslip in place (this is necessary because the DIC optics of the microscope are not optimal for imaging through plastic). 6. Place the slide onto the microscope stage (inside the microscope incubation cabinet). Place the humidified CO2 hood over the specimen. Place four 35 mm Petri dishes containing sterile tissues or Whatman paper soaked in sterile distilled water

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Fig. 4. (a) Still frames from a time-lapse imaging series of mineralizing primary osteoblast cultures from Dmp1-GFP transgenic mice in which alizarin red was used as a vital dye for mineral deposition. The movie was started when clusters of GFP-positive cells had formed in the cultures and images were acquired every 20 min for 48 h in DIC and two fluorescent channels. Triple merged images of DIC, GFP, and alizarin red are shown. Bar = 100 μm. (b) Quantitation of mineralization dynamics from the time-lapse sequence shown in (a). Mineralization was quantified using ImageJ software by thresholding of alizarin red image stacks followed by measurement of the mineralized area (red line).The number of GFP-positive cells were counted (green line). Note the deposition of mineral beginning 10 h after addition of 4 mM β-glycerophosphate and increasing until 40 h. Mineral deposition occurs specifically where there are clusters of GFP-positive cells and mineralization is accompanied by an increase in the number of GFP-positive cells. These data suggest that the cells responsible for mineral deposition are already transitioning towards the osteocyte phenotype.

inside the CO2 hood, positioned around the specimen to help maintain humidity. Allow the specimen to equilibrate for a minimum of 1 h before initiating the time lapse acquisition (see Note 7). 7. During the equilibration period, the parameters of the time-lapse image acquisition can be specified within the Metamorph software and imaging fields can be selected, but the final focusing of each field will be done after the sample is fully equilibrated. 8. Within the Metamorph software, select the 20× DIC magnification setting. 9. Use the “multi-dimensional acquisition” (MDA) interface of the Metamorph software to set up parameters for the imaging experiment. This allows the user to designate the number of wavelengths, light illumination settings, exposure times, camera binning, duration of imaging, time interval between acquisitions, x, y, and z co-ordinates of the fields to be imaged, etc.

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10. Within the MDA interface. Set the duration of the experiment as 72 h and the time interval between acquisitions to 20 min. 11. Select the appropriate illumination settings for the wavelengths you are imaging. In this example, there will be three imaging channels, including DIC, GFP, and Alizarin Red. The GFP is best imaged with an optimized GFP filter set, but can also be imaged using an FITC filter set. Alizarin red can be imaged using a TRITC filter set. We recommend using 12–25% of the light output from the Exfo Exacte metal halide lamp and using an ND4 or ND8 filter in the light path if possible, but this will be dependent on the brightness of the probe (see steps 13–15 below for setting up the exposure times). 12. Next, the co-ordinates of the fields to be imaged need to be entered. First, locate the plane of focus of the cells under DIC illumination to minimize photodamage. In this model system, clusters of GFP-positive cells indicate the locations where mineral will be deposited, therefore change to the GFP filter set to locate GFP-positive cell clusters. In live camera mode (with 2 × 2 binning), move to the position of the imaging field and add the co-ordinates to the list of stage positions. Keep the light intensity as low as possible while selecting fields containing clusters of GFP-positive cells and minimize the amount of time spent illuminating the cells. 3–5 imaging fields per well should be selected to ensure adequate spacing of fields and minimal phototoxicity. We normally select one from the top of the well, one from the middle and one near to the bottom (see Note 8). 13. Select an exposure time for the GFP between 10 and 200 ms, using 2 × 2 camera binning. Make sure that there is a sufficient difference between the signal intensity in a background area of the image and the GFP-positive cells. This would ideally be a fivefold difference, but in the case of live cell imaging, a twofold difference may be a reasonable compromise to minimize phototoxicity. If the signal is too low, you should consider increasing the intensity of the excitation light. Make sure that there are no overexposed areas of the image (i.e. places where the pixel intensity is already at its maximal value) (see Note 9). 14. Change to DIC illumination and adjust the polarizer to optimize the DIC image. Repeat the procedure in step 13 for setting the exposure time for the DIC image (exposure times for DIC are usually between 1 and 10 ms). If the DIC image is overexposed, it may be necessary to use a neutral density filter(s) in the light path. 15. Setting exposure times for alizarin red follows the same principle as for GFP, but is more challenging, as there is no mineral

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at the start of the movie, therefore there are no image features to use for setting the exposure times. This is an example where it is helpful to perform a prior experiment with a static culture containing similar amounts of mineral as would be expected to form in the time-lapse experiment, in order to estimate the approximate exposure times. On our Nikon system, we have had good success with an exposure time of 100 ms using the TRITC filter set, with an ND8 filter in line and 25% light output from the Exfo Exacte metal halide lamp. 16. Specify imaging parameters in the z-axis using the z series menu tab. Set the mode to “range around current.” In this mode, the software will use the z co-ordinate from the stage position you have entered as the centre z position and will collect images above and below that plane of focus. Set the acquisition to collect five z-planes, spaced 1–1.5 μm apart (see Note 10). Set the parameters to acquire the z series for one wavelength at a time (this will be quicker and will reduce wear on the filter wheel changer). 17. Once all the parameters for the movie acquisition are set and the specimen is fully equilibrated, each stage position should be revisited once more for final focusing (under GFP illumination) prior to starting the movie acquisition. Overwrite the co-ordinates of any positions for which the focus needs to be adjusted. 18. Start the movie acquisition and monitor at least the first 3–4 image acquisitions to make sure that the acquisition is running properly and there is no appreciable focal drift. 19. Monitor the images periodically during the time-lapse acquisition to check for focal drift and refocus if necessary. 20. At the end of the imaging period, collect detailed z-stacks for each of the positions, if desired (see Note 10). This can be done by moving to each stage position, focusing through the specimen and entering co-ordinates for the top and bottom. One can then either specify the number of z-planes or the spacing of the z-planes (e.g. 0.3–0.5 μm is appropriate for 20× magnification). 21. When you have completed all imaging, fix the specimen in 4% paraformaldehyde or 10% neutral buffered formalin for 5–10 min, so that additional stains, such as immunostaining for osteocyte markers, etc., can be performed (see Note 11). 22. During the movie acquisition and when reviewing the data, it is also important to monitor for signs of phototoxicity to the cells (see Note 12). Figure 4 shows selected still frames from an example of timelapse imaging of mineralizing primary osteoblast cultures from Dmp1-GFP transgenic mice.

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3.3.2. Time-Lapse Imaging of Fibronectin Assembly in 2T3 Osteoblast-Like Cells

In this model system, 2T3 osteoblast-like cells are cultured in the presence of fluorescence-labelled purified fibronectin as a probe for monitoring assembly of fibronectin. The cells will assemble fibronectin from an exogenous source on their cell surface and generate extensive fibronectin fibrillar networks over a 24–48 h period (see Fig. 5). 1. Prepare labelled fibronectin probes ahead of time using an Alexa 555 protein labeling kit from Molecular Probes (Invitrogen Corporation, Carlsbad, CA) (see Note 13). Use 1 mg purified fibronectin for labelling and perform the labelling according to the manufacturer’s instructions, with the exception that the elution buffer from the kit should not be used because it contains sodium azide. Instead, use tissue culture grade PBS for all the column equilibration and elution steps. Filter sterilize the probe through a 0.2-μm filter and store labelled probe in single-use aliquots at −80°C protected from light.

Fig. 5. (a) Representative still frames from a time-lapse imaging series of fibronectin fibril assembly in 2T3 osteoblast-like cells. Images were acquired every 15 min for 48 h. Fibronectin was imaged in the red fluorescence channel using an Alexafluor555-labelled probe (shown) and cells were imaged in DIC (not shown). Note that the assembly of fibronectin begins with the assembly of short fibrils on the cell surface. These then coalesce to form the more extensive fibrillar networks seen at 24 h and 48 h. Bar = 100 μm. (b) Quantitation of fibronectin fibril assembly dynamics from the time-lapse sequence shown in (a). Quantitation was performed using ImageJ software by thresholding of FN image stacks followed by measurement of the fibril area.

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2. Take a T75 flask of 2T3 cells at 90–100% confluence (if using 100% confluent cells, use when they have only just reached confluence). Wash 1× in PBS and trypsinize with 2 ml trypsin. 3. Transfer the cell suspension to a 50-ml Falcon tube containing 20-ml growth medium for 2T3 cells. 4. Count the cell number in a hemocytometer or coulter counter and plate the cells into a coverslip bottomed 4-well Lab-Tek chamber slide at 4 × 104 cells per cm2 growth area in 0.8-ml growth medium. 5. If imaging assembly in subconfluent cells, incubate overnight at 37°C in a humidified 5% CO2 incubator to allow cells to adhere. If imaging assembly in confluent cells, culture for 2–3 days to allow cells to reach confluence. Change medium every 2–3 days. 6. Change to fresh medium containing 50 μg/ml ascorbic acid and in the sterile hood replace the plastic lid of the Lab-Tek slide with a sterile 24 mm × 50 mm glass coverslip (see Subheading 3.3.1, step 5). 7. Place the slide onto the microscope stage for equilibration for at least 1 h prior to imaging as described in Subheading 3.3.1, step 6. 8. During the equilibration period, the parameters of the time lapse image acquisition can be specified within the Metamorph software and imaging fields can be provisionally selected, but the final selection/focusing of each field will be done after the sample is fully equilibrated. 9. See Subheading 3.3.1 for details of how to set up the imaging parameters within the Metamorph software using the MDA interface. Use 20× magnification and set the duration of the experiment to 48 h, with a time interval between acquisitions of 20 min. Designate two wavelengths, DIC and TRITC (for imaging of the Alexa 555 probe). Use the TRITC illumination setting at 25% of the light output from the Exfo Exacte metal halide lamp, with an ND4 filter in the light path. 10. Locate the plane of focus of the cells using DIC illumination to minimize photodamage. Select the provisional positions of the imaging fields, selecting 3–4 image fields per well, spaced apart (i.e. at the top, middle, and bottom of the well). These will be refined later after you have added the fluorescent fibronectin probe. 11. Make up the probe at 10× concentration in growth media containing 50 μg/ml ascorbic acid. The final concentration (i.e. at 1×) will be 5 μg/ml Alexa555 labelled fibronectin and 5 μg/ ml unlabelled fibronectin. Make sure that you thaw the frozen stock vials of fibronectin rapidly in a 37°C waterbath to avoid formation of aggregates.

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12. Add the 10× probe directly to the slide on the stage by removing 80-μl culture media and adding back 80-μl media containing the 10× probe (the final concentration will be 1×) (see Note 14). 13. Allow 15–20 min for the probe to begin incorporating on the cell surface and then revisit each of the provisional positions you selected earlier. Change to TRITC illumination and confirm that you can see small fibrils beginning to assemble on the cell surface. Use these for focusing and enter the new coordinates of the field. If you cannot see small fibrils, search for a better field in the vicinity of the position and update the new co-ordinates. Work as quickly as possible to minimize exposure of the cells and also because the fibronectin assembly happens relatively quickly and you will miss important early dynamic events if field selection takes too long. 14. Select an exposure time for TRITC between 10–200 ms, using 2 × 2 camera binning as described in Subheading 3.3.1, steps 13–15. When setting exposure times, take into account the fact that there will be a significant increase in brightness of the fibronectin fibrils as the movie progresses from incorporation of a small amount of fibronectin on the cell surface at the beginning of the movie to formation of an extensive fibrillar network at the end of the movie (see Fig. 5). 15. Change to DIC illumination and adjust the polarizer to optimize the DIC image. Repeat the procedure in Subheading 3.3.1, step 14 for setting the exposure time for the DIC image. 16. Set the parameters for the z-series as described in Subheading 3.3.1, step 16. 17. Once all the parameters for the movie acquisition are set, revisit each of the stage positions once more for final focusing (under TRITC illumination) prior to starting the movie acquisition. Overwrite the co-ordinates of any positions for which the focus needs to be adjusted. 18. Start the movie acquisition and monitor at least the first 3–4 image acquisitions to make sure that the acquisition is running properly and there is no appreciable sample drift. 19. Monitor the images periodically during the time-lapse acquisition to check for focal drift and refocus if necessary. Also monitor for signs of phototoxicity (see Note 12). 20. At the end of the imaging period, collect detailed z-stacks for each of the positions, if desired, as described in Subheading 3.3.1, step 20. 21. When you have completed all imaging, fix the specimen in 4% paraformaldehyde or 10% neutral-buffered formalin for 5–10 min, so that additional stains, such as immunostaining

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for other ECM proteins, osteoblast markers, etc., can be performed (see Note 11). Figure 5a shows selected still frames from time lapse imaging of fibronectin assembly in 2T3 osteoblast cultures. 3.3.3. Time-Lapse Imaging of Osteocytes in Calvaria from Dmp1-GFP Transgenic Mice

In this model system, neonatal calvaria from transgenic mice expressing the Dmp1-GFP transgene are imaged. Neonatal mouse calvaria are very thin (30–50 μm), which makes them amenable to imaging and the Dmp1-GFP transgene is selective for osteocytes (22, 23) which enables imaging of partially and fully embedded osteocytes within the isolated bone explant. In addition, our laboratory has identified a surface motile Dmp1-GFP-positive cell type that appears to represent a subpopulation of osteoblasts that has already begun transitioning towards the osteocyte phenotype (27) For imaging up to 24 h, mount half calvaria from Dmp1-GFP mice (see Subheading 3.2) in a 2- or 4-well Lab-Tek chambered coverglass slide under a sterile plastic coverglass with a viewing window. These are prepared by punching holes into 22 mm × 22 mm plastic coverglasses and trimming them to fit into the wells. The coverglass is adhered to the bottom of the glass slide using sterile silicone grease (see Fig. 6). Make sure that the silicone grease is not touching the bone specimen. Use 600- or 1,200-μl culture medium per well for the 4- or 2-well chamber slides, respectively. For longer-term imaging, the calvaria can be maintained in a perfusion culture chamber with a coverglass bottom (see Note 15). 1. Replace the plastic lid of the Lab-Tek slide with a sterile 24 mm × 50 mm glass coverslip (see Subheading 3.3.1, step 5). Place the slide onto the microscope stage for equilibration for at least 1 h prior to imaging as described in Subheading 3.3.1, step 6. 2. During the equilibration period, set the parameters of the time-lapse image acquisition in the Metamorph software (see Subheading 3.3.1 for details of how to do this using the MDA interface). Set the magnification to 20× and the duration of the experiment to 24 h, with a time interval between acquisitions of 15 min. Designate two wavelengths, DIC and GFP. Set the GFP illumination setting to 12 or 25% of the light output from the Exfo Exacte metal halide lamp, with an ND4 or ND8 filter in the light path. 3. Locate the plane of focus of the osteocyte lacunae using DIC illumination to minimize photodamage and move to the centre of the parietal bone. Switch to GFP illumination and select the provisional positions of the imaging fields containing “in focus” osteocytes. Select three image fields per half calvarium, spaced at least 2–3 field diameters apart. Work quickly so as to minimize exposure of the cells to the excitation wavelength.

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culture medium

half calvarium plastic coverglass with viewing window

alternative “T-bar”mount

perfusion culture chamber

Fig. 6. (a) Schematic diagram illustrating mounting of a half calvarium in a well of a Lab-Tek chambered coverglass for time-lapse imaging. The half calvarium is mounted under a plastic coverslip with a viewing window. The parietal bone is positioned under the viewing window, as this is the area that will be imaged. Sterile silicone grease is used to adhere the coverslip to the bottom of the well. (b) An alternative mount for the system in (a) in which the half calvarium is held under a “T-bar” mount. (c) Perfusion chamber system from Warner Instruments (Hamden, CT) that can be used for perfusion culture of cells and tissues.

4. Select an exposure time for GFP between 10–200 ms, using 2 × 2 camera binning as described in Subheading 3.3.1, step 13. 5. Change to DIC illumination and adjust the polarizer to optimize the DIC image. You should be able to clearly visualize the osteocyte lacunae. Repeat the procedure in Subheading 3.3.1, step 14 for setting the exposure time for the DIC image. 6. Set the parameters for the z-series as described in Subheading 3.3.1, step 16. For imaging of calvaria, we generally collect 7–9 z-planes with a spacing of 1.5–2 μm as the sample is thicker than a cell monolayer. 7. Once all the parameters for the movie acquisition are set, revisit each stage position once more for final focusing (under GFP illumination) prior to starting the movie acquisition. 8. Start the movie acquisition and monitor at least the first 3–4 image acquisitions to make sure that the acquisition is running properly and there is no appreciable sample drift.

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Fig. 7. Still frames from a time-lapse imaging series of Dmp1-GFP-positive osteocytes in a 7-day-old neonatal mouse calvarial explant. Images were acquired every 20 min for 12 h using the green fluorescence channel for GFP (shown) and DIC for imaging of the bone explant (not shown). The still images illustrate the motile properties of osteocytes within their lacunae. In particular, note that the two osteocytes marked with an asterisk show motions of their dendrites, which adopt various configurations at different times during the time-lapse imaging period. Bar = 30 μm.

9. Monitor the images periodically during the time-lapse acquisition to check for focal drift and refocus if necessary. Monitor also for signs of phototoxicity. 10. At the end of the imaging period, collect detailed z-stacks for each of the positions, if desired, as described in Subheading 3.3.1, step 20. 11. When you have completed all imaging, fix the specimen in 4% paraformaldehyde or 10% neutral buffered formalin, leaving it mounted in the slide if possible, so that additional stains, such as immunostaining for osteocyte markers, actin cytoskeleton, nuclei, etc., can be performed (see Note 16). Figure 7 shows selected still frames from time-lapse imaging of Dmp1-GFP-positive osteocytes in neonatal mouse calvarial explants. 3.4. Post-acquisition Processing of Image Stacks, Generation of Movies and Quantitation of Dynamic Events

After acquisition of a time-lapse image series, a significant amount of image processing is usually required to generate movies that best depict the biology being investigated and to facilitate quantitation of dynamic events. Below we summarize the main principles of some of the processing steps that may be required.

3.4.1. Generation of an In-Focus Image Stack from Multi-dimensional Imaging Datasets

Using the Metamorph software, the image data set can be viewed in the “review multi-dimensional data” interface. This provides an overall view of all the images for a given wavelength and imaging field in the form of a matrix with the time points along the top and the z-planes down the side. The user can click on any image to review it and can scroll through by time point or by z-plane. To generate an in-focus image stack from the image matrix, there are several options, including. 1. A maximum or minimum z-projection can be generated from all the z-planes for each time point or from a subset of z-planes (see Note 17). The z-projected images are then loaded using the “load images” function to generate a new image stack.

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2. The “best focus” algorithm can be used for automated selection of a single “best focus” image plane from the z-planes for each time point. The selected images are then loaded to generate a new image stack. 3. The user can manually select a best focus image from the z-planes for each time point. This is often a useful approach if there is a small amount of focal drift, such that the plane of best focus shifts up or down to a different z-plane during the acquisition. 4. A combination of automated and manual selection of z-planes can be done, whereby the software is used for initial selection of the best focus image, but this is manually corrected by the user if there is a small amount of focal drift during the acquisition. Using one of these approaches, an in-focus image stack is generated, which can then be converted to a movie file (see Subheading 3.4.5) or can be processed further. The ImageJ software, available as a free download from NIH also has plugins such as “Smart Projector” that can optimize for each time point the projection of z-stacks acquired in a time-lapse series. 3.4.2. Scaling, Contrasting, and Background Subtraction

Scaling and brightness/contrast adjustments are performed to enhance the visual appearance of the image stack. A 12-bit image consists of 4,096 possible grey levels and a 16-bit image has 65,536 possible grey levels. However, much of the important image information may be contained within a narrow range of pixel intensities towards the lower end of the greyscale (especially for probes that are not very bright). Additionally, most computer monitors can only display 256 grey levels. The Scale Image command in Metamorph allows you to select a smaller range of grey values within a 12- or 16-bit image that contain the important image information and rescale their intensities to better display them within the range of grey levels of the monitor. This allows you to see differences in greyscale values that might otherwise be impossible to discriminate visually. This function only affects the display of the data on the monitor and does not alter the actual pixel intensity values of the original image data. However, if scaled image stacks are converted to 8-bit images or exported in certain file formats that involve image compression, the original pixel intensity information may be lost. Brightness and contrast adjustments can also be made to image stacks to enhance their visual display and background subtraction operations may be required to correct for an uneven background in the image. It is important when making comparisons, such as comparing treated versus control movies, that all scaling, brightness/contrast and background subtraction adjustments are applied identically to the image stacks that are going to be compared. It is

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also important to save and archive all the original data files, so that it is always possible to go back to the original unprocessed data if necessary for analysis. Our laboratory has established a file naming system for our processed image stacks that provides information within the filename to specify what processing steps have been performed. For example, the filename “Exp15_p10_TRITC_ Zmax3-5_BGsubtr_contr_8bit” would indicate that the stack comes from experiment 15, stage position 10 viewed under TRITC illumination and that z-planes 3–5 were used for a maximal z-projection. The stack has been processed by background subtraction and has been contrast adjusted and converted to an 8-bit stack. 3.4.3. Stack Registration

Often when scrolling through an image stack there appears to be a “wobble” in the stack, whereby the entire image shifts slightly in the x and y planes. This “rigid body motion” does not represent true motion of cells, but rather the entire imaging field has moved slightly relative to the previous image, due to stage shift, vibration, etc. If quantitative measurements of linear distances are required (e.g. cell motion trajectories, fibril displacements, etc.) it is important to first align the stack to correct for rigid body motion so that accurate measurements of the true motion can be obtained. The autoalign function in Metamorph performs stack alignment. Alternatively, the ImageJ software has plugins available online, such as “turboreg” and “stackreg” that are useful for aligning image stacks. These algorithms can also compensate for small movements in the whole sample, such as rotation or small motions of a calvarium within its mount, etc.

3.4.4. Pseudocolouring and Merging RGB and DIC-Fluorescence Images

The time-lapse fluorescence images are acquired with a monochrome camera and are therefore greyscale images. If colour images are desired, the greyscale images must be pseudocolored. Most imaging softwares, such as Metamorph, have functions for pseudocoloring and converting image stacks to RGB format. Pseudocolouring can also be performed in ImageJ using look-up tables to specify the colour to be assigned to the grey/white pixels. Merging of fluorescent images acquired with different coloured probes may be needed to look for co-localization, etc. Again, this is a standard function in most imaging softwares. It can also readily be performed in ImageJ using the “RGB merge” function in which the user specifies a source stack for the red, green, or blue channels of the RGB image. Alignment of the image pairs may also be necessary if there is a slight shift in the x, y position of image features between different filter sets. Merging fluorescence images with DIC images is done by “overlaying” the two image stacks, since the DIC image cannot be assigned to the red, green, or blue RGB channel. The two images are overlayed, with the user specifying the pseudocolouring for the fluorescence image and the degree of transparency of the DIC and fluorescence images.

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3.4.5. Generating Movie Files and File Compression

Once the image stacks have been processed, they should be exported in a movie file format, such as .avi or .mpeg files or quicktime movie (.mov) files. These export formats allow the user to specify the frame rate, file compression settings, etc. The files can be very large (>100 Mb) without file compression and many journals impose limits on the sizes of movie files that they will publish as online supplementary data. Therefore file compression is often needed. However, the drawback of file compression is that it often results in loss of data and/or reduced image resolution. There are many options available for file compression in programs such as Quicktime and Windows Moviemaker available as free downloads over the internet. The key is to experiment with different methods to obtain a reasonable balance between reducing file size without significant loss of image quality. Note that it is not recommended to use compressed movie files for quantitative measurements. These should be performed on image stacks that retain the original data, such as the original pixel intensities, etc.

3.4.6. Quantitation of Dynamic Events

Time-lapse image stacks are very data rich. To maximize the impact of the work it is important to find ways to quantify the biology depicted in the movie stacks (15). The majority of softwares that drive time-lapse imaging systems include quantitative analysis tools in the basic package or as add-on modules. ImageJ also has many tools for analysis of image stacks. Parameters that can be quantified include tracking of cell motion trajectories and measurement of velocities, thresholding and cell/particle counting, linear and areal measurements, co-localization of fluorochromes, measurement of displacements and strain values, fluorescence intensities, etc. It is preferable to use tools that can automate these analyses as much as possible, but there are some situations where the analysis is best done manually. For example, cell tracking is straightforward if a nuclear tracking dye is used during imaging, as the nuclei have high contrast and can be readily tracked using automated tracking functions (the trade off is an increased risk of phototoxicity – see Subheading 1.3). DIC images, especially in postconfluent cultures, may be more difficult to track automatically, as each cell or nucleus may not be easily distinguished. In this case, manual tracking may be necessary. Below, we have described some simple quantitative analyses performed on the time-lapse movie experiments described in Figs. 4, 5, and 7. 1. For time-lapse experiments like that shown in Fig. 4, mineral deposition can be quantified by thresholding and particle counting of the alizarin red image stacks using ImageJ, to give an output of mineralized area in pixels, which can be converted to μm2 using the appropriate conversion factor for the camera and objective. The number of GFP-positive cells can be manually counted as a function of time using the “cell counter”

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plugin in ImageJ (Author: Kurt De Vos). The GFP expression can also be measured by thresholding and particle counting to give a measurement of the area of GFP-positive cells. When performing quantitative analyses it is preferable to work on the raw image data if possible or otherwise to make sure that all images are processed identically prior to quantitation. This type of quantitative analysis reveals the dynamics of mineralization as a function of time and shows that induction of GFP expression precedes and/or accompanies deposition of mineral. These data provide experimental evidence that mineral deposition is integrated with the osteoblast to osteocyte transition and suggest that the cell responsible for mineral deposition is already beginning to express markers of the osteocyte (we have shown that the Dmp1-GFP-positive cells also express the osteocyte marker, E11/gp38). 2. For the time-lapse experiment shown in Fig. 5, deposition of a fibronectin fibrillar matrix was quantified by thresholding and particle counting of the Alexa555 image stacks using ImageJ, to give an output of fibronectin fibril area as a function of time. The data show rapid assembly of fibronectin during the first 24 h, followed by a more gradual increase up to 48 h. We have previously reported a correlation between cell and ECM fibril motions in primary osteoblast cultures and shown that ECM fibril dynamics change as a function of ECM maturation (8). 3. For time-lapse experiments such as that shown in Fig. 7, We have measured parameters such as the percentage of osteocytes showing dendrite motions, and showing motions of their cell bodies within their lacunae (Dallas et al., unpublished observation) (28) When performing these measurements it is usually not necessary to count every cell within an imaging field, but rather a representative sample of cells (e.g. 20–40 cells per field). In this case, it is important to have an unbiased method of selecting cells for counting, such as using a grid and selecting cells that intersect the grid points, etc. In similar movies, we have also measured motion trajectories of surface motile Dmp1GFP-positive cell populations using the “MTrackJ” plugin for ImageJ (© Erik Meijering, Biomedical Imaging Group, Erasmus MC – University Medical Center Rotterdam). This analysis revealed that these cells travel on the bone surface at velocities of 4–5 μm/h, equating to 5-cell diameters within a 24-h period. More sophisticated computational analyses of cell and fibril motions can be applied with softwares, such as MatLab, developed for the engineering field, using techniques such as particle image velocimetry to model cell and fibril motions as flow patterns (29). These analyses can generate information on motion trajectories, positional fate of ECM proteins, vector field maps depicting directions of

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motion, strain fields, etc. These approaches have shown that in the avian embryo, in addition to ECM fibrils being affected by local cell motions, fibril dynamics appear to be affected by the global tissue motions that occur during morphogenesis of the embryo (29). It is important to archive time-lapse imaging data, as the image stacks are very data rich. As new biological questions are developed, the data can be re-interrogated in the future to extract additional quantitative end points and to test new hypotheses. The impact of the data can be further increased by making these datasets available to other investigators to analyze from a different biological perspective. In summary, as more and more laboratories begin to incorporate live imaging approaches into their studies, these techniques will become standard research tools and the truly dynamic nature of bone cell function will be fully appreciated.

4. Notes 1. A wide variety of fluorescent probes and reagents are available for investigating biological events in living cells. In particular, Molecular Probes (Invitrogen Corporation, Carlsbad, CA) carries fluorescent antibodies, kits for fluorescently labelling proteins of interest, fluorescence reagents for assessing cell viability, dyes for cell and nuclear tracking, probes for labelling of organelles, probes for assessing pH and ion flux, probes for monitoring enzyme activity, etc. In addition, the Living Colors selection of cDNA constructs (Clontech, Mountain View, CA) includes a comprehensive set of GFP-derived fluorescent protein vectors that can be used as reporter constructs or to generate fusion constructs with a protein of interest for monitoring gene expression and protein localization in vivo, in situ, and in real time. 2. Potential solutions for data storage may include the use of external plug in 1Tb hard drives, or preferably, the use of a Hard Drive RAID enclosure (for upwards of 2Tb of data storage) that would provide adequate storage capacity and fault tolerance. The RAID enclosure can be directly attached to the image acquisition computer by USB or preferably via the ISCSI connection, attached by a separate local network to handle only the data traffic to the raid enclosure. The ISCSI network can also be attached to the building network for Data backup and permanent Archive. In our experience, it is better to avoid acquiring images directly onto a networked server, since a network outage could interrupt data collection and/or crash the acquisition. 3. Alternatively, the cells can be frozen at 2 × 106 cells per cryovial in 1 ml of complete α-MEM + 40% FBS + 10% dimethyl

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sulfoxide (DMSO). A single vial should be defrosted and plated into a 90-mm Petri dish. Upon confluency, the cells should be trypsinized and used for experiments without further passaging. The plating density for experiments will be 4 × 104 cells per cm2 growth area, which is higher than that for the freshly isolated cells. 4. Whether or not the periostea are removed depends on the imaging application. For imaging of osteocytes, we generally strip off the periostea. If desired, the stripped calvarial bones can also be digested for two 20-min digestions with 0.05% trypsin and 0.2% collagenase in α-MEM (no additives) to remove osteoblasts that remain on the bone surface. This provides a clearer image of osteocytes, but you may miss interactions with surface motile cells. It is also possible to image without removing the periostea, which we have done when imaging osteoblasts using a DsRed transgene expressed in osteoblasts under control of the 3.6 kb type I collagen promoter. 5. It is important to use coverslip bottomed culture vessels for imaging with most high NA lenses. Nalgene Nunc (Rochester, NY) manufactures a range of coverslip bottomed chamber slides. Other alternatives include coverslip-bottomed Petri dishes. Coverslip-bottomed multi-well plate formats are also available from some manufacturers. 6. We have found that using a lower concentration of 0.5 mM β-GP prior to imaging is useful because it helps the cells to differentiate and express the Dmp1-GFP transgene, but mineralization will not take place until the β-GP concentration is increased to above 2.5 mM. 7. We recommend equilibration of the specimen for 1–2 h prior to imaging if this is possible within the time constraints of the biological events being imaged. This is because the most common reason for focal drift during time-lapse imaging is that the specimen is not fully equilibrated to the temperature of the incubation cabinet, etc. If you are not able to equilibrate for this amount of time, then you should monitor the focus periodically during the image acquisition and refocus the microscope if necessary. 8. Avoid the edges of the wells when selecting imaging fields as it is difficult to obtain good DIC images near the edges due to the meniscus of the culture medium affecting the optics. If your application requires high-resolution DIC imaging, it may be preferable to use a closed perfusion culture system with coverslip glass viewing windows so that the meniscus effect is avoided. 9. Our camera is a 12-bit monochrome camera, which gives 4,096 grey levels. When setting exposure times for movie

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acquisitions, we recommend that the maximal pixel intensity in the initial images does not exceed 2,000, to allow sufficient room to accommodate an increase in brightness of the signal during the movie acquisition. 10. We have found that in cell monolayer cultures, five z-planes spaced 1–1.5 μm apart is a reasonable compromise for live cell imaging that allows you to collect information from above and below the focal point you have set, in case of sample drift, etc. A more detailed z series would obviously give better resolution, but this would be at the expense of increased phototoxicity. Therefore, our approach has often been to collect a more detailed z-stack of the specimen at the end of the time-lapse imaging to enable more detailed 3D reconstructions of the sample. In that way the motile history/dynamic properties of the cells can be correlated with the 3D structure of the sample at the end of the experiment. 11. We recommend fixation of the specimens after performing live cell imaging to allow maximum data collection from the experiments. For example, immunostaining for markers of osteocytes, mineralization related proteins, etc., can be performed. In this way the motile history of the cells can be correlated with the cells’ expression of specific markers at the end of the movie. Other analyses can also be performed, such as playing the movies in reverse to “back track” the origin of a cell that expresses a particular marker. 12. Phototoxicity is a potential problem when performing timelapse imaging, especially with fluorescent probes (see Subheading 1.3). Cells can also be damaged if the 5% CO2 atmosphere is not maintained, if there is evaporation or leakage of medium from the culture vessel, overheating, etc. A healthy culture should show many cell divisions and the cells should not become rounded and detached. There are many commercially available kits that can be used to monitor cell viability/apoptosis, etc., in live cell cultures. These can be used to optimize imaging conditions when establishing protocols for live imaging experiments. 13. Molecular probes has a range of protein labelling kits, with colours ranging from the blue to the far red end of the spectrum. We have performed experiments in which we have labelled existing fibronectin fibril networks with a green Alexa488 fibronectin probe, followed by labelling newly assembled fibronectin with a red Alexa555 fibronectin probe. Depending on the biological question, the probes can either be left in the culture medium to monitor continued assembly (we have found that the background fluorescence is generally not too high) or they can be used in a “pulsed labelling” protocol to label populations of fibrils (with washing of the unbound probe) and track the fibril dynamics.

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14. The fibronectin probe is added directly to the equilibrated slide on the microscope stage because assembly of the probe on the cell surface will begin within 15–20 min of addition. Therefore, if the probe were added in the tissue culture hood, there would be insufficient time to equilibrate the sample and select imaging fields, etc., before the assembly process starts. Although the incubation cabinet on the microscope is not a fully sterile environment, we have found that by using clean technique (sterile tubes, tips, etc.), and minimizing the time that the culture slide wells are exposed we have not experienced problems with contamination within the 48 h time frame of these experiments. 15. With an inverted microscope system and high NA lenses, it is generally not possible to image calvaria cultured at the air– medium interface, as is the usual culture method for organ cultures, since the working distances of the lenses are too short. We have had good success imaging up to 24 h by mounting the calvaria at the bottom of the well under a plastic coverslip with viewing window. For longer-term imaging of calvaria, imaging at the air–medium interface may be possible with specialized longer working distance lenses. Alternatively, a closed perfusion culture system can be used. These are available commercially and in particular, Warner Instruments (Hamden, CT) offers a large variety of imaging and recording chambers for live cell and organ culture applications with various microscope configurations. 16. Intact calvaria can readily be stained as “whole mount” specimens, which can provide very nice images of intact osteocytes and/or can be used for 3D imaging of osteocytes. The undecalcified calvaria can be stained with alizarin red to label the mineral. Decalcified calvaria can be immunostained for various osteocyte markers and can also be stained with phalloidin to label the actin cytoskeleton (which provides very nice images of the osteocytes and their dendrites) and/or DAPI which stains the actin cytoskeleton and nuclei. By following dynamic imaging with whole mount immunostaining, the same fields can be located and the motile history of the cells can be correlated with their expression of specific markers. 17. The maximum intensity or minimum intensity z-projections create a 2D output image for which each pixel contains the maximum or minimum value, respectively, over all images in the stack for that pixel location. Some softwares also support average intensity or sum slice z-projections, in which each pixel displays the average intensity or sum intensity over all images in the stack for that pixel location.

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References 1. Faibish, D., Gomes, A., Boivin, G., Binderman, I., and Boskey, A. (2005) Infrared imaging of calcified tissue in bone biopsies from adults with osteomalacia. Bone 36, 6–12. 2. Huitema, L. F., and Vaandrager, A. B. (2007) What triggers cell-mediated mineralization? Front. Biosci. 12, 2631–2645. 3. McKee, M. D., Addison, W. N., Kaartinen, M. T. (2005) Hierarchies of extracellular matrix and mineral organization in bone of the craniofacial complex and skeleton. Cells, tissues, organs 181, 176–188. 4. Murshed, M., Harmey, D., Millan, J. L., McKee, M. D., and Karsenty, G. (2005) Unique coexpression in osteoblasts of broadly expressed genes accounts for the spatial restriction of ECM mineralization to bone. Genes Dev. 19, 1093–1104. 5. Eils, R., and Athale, C. (2003) Computational imaging in cell biology. J. Cell Biol. 161, 477–481. 6. Kulesa, P. M. (2004) Developmental imaging: Insights into the avian embryo. Birth Defects Res. C. Embryo Today 72, 260–266. 7. Friedl, P. (2004) Dynamic imaging of cellular interactions with extracellular matrix. Histochem. Cell Biol. 122, 183–190. 8. Sivakumar, P., Czirok, A., Rongish, B. J., Divakara, V. P., Wang, Y. P., and Dallas, S. L. (2006) New insights into extracellular matrix assembly and reorganization from dynamic imaging of extracellular matrix proteins in living osteoblasts. J. Cell Sci. 119, 1350–1360. 9. Dallas, S. L., Chen, Q., and Sivakumar, P. (2006) Dynamics of assembly and reorganization of extracellular matrix proteins. Curr. Top. Dev. Biol. 75, 1–24. 10. Zamir, E. A., Rongish, B. J., and Little, C. D. (2008) The ECM moves during primitive streak formation--computation of ECM versus cellular motion. PLoS biology 6, e247. 11. Frigault, M. M., Lacoste, J., Swift, J. L., Brown, C. M. (2009) Live-cell microscopy - tips and tools. J. Cell Sci.122, 753–767. 12. Mavrakis, M., Pourquie. O., and Lecuit, T. (2010) Lighting up developmental mechanisms: how fluorescence imaging heralded a new era. Development 137, 373–387. 13. Xie, Y., Yin, T., Wiegraebe, W., He, X. C., Miller, D., Stark, D., Perko, K., Alexander, R., Schwartz, J., Grindley, J. C., Park. J,, Haug. J, S., Wunderlich, J. P., Li, H., Zhang, S., Johnson, T., Feldman, R. A., and Li, L. (2009) Detection of functional haematopoietic stem

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cell niche using real-time imaging. Nature 457, 97–101. Lo Celso, C., Wu, J. W., Lin, C. P. (2009) In vivo imaging of hematopoietic stem cells and their microenvironment. J. Biophotonics 2, 619–631. Hamilton, N. (2009) Quantification and its applications in fluorescent microscopy imaging. Traffic 10, 951–961. Sekar, R. B., Periasamy, A. (2003) Fluorescence resonance energy transfer (FRET) microscopy imaging of live cell protein localizations. J. Cell Biol. 160, 629–633. Day, R. N., Schaufele, F. (2005) Imaging molecular interactions in living cells. Mol. Endocrinol. 19, 1675–1686. Parsons, M., Vojnovic, B.,and Ameer-Beg, S. (2004) Imaging protein-protein interactions in cell motility using fluorescence resonance energy transfer (FRET). Biochem. Soc. Trans. 32, 431–433. Wiedenmann, J., Oswald, F., Nienhaus, G. U. (2009) Fluorescent proteins for live cell imaging: opportunities, limitations, and challenges. IUBMB life 61, 1029–1042. Czirok, A., Zamir, E. A., Filla, M. B., Little, C. D., Rongish, B. J. (2006) Extracellular matrix macroassembly dynamics in early vertebrate embryos. Curr. Top. Dev. Biol. 73, 237–258. Ohashi, T., Kiehart, D. P., Erickson, H. P. (1999) Dynamics and elasticity of the fibronectin matrix in living cell culture visualized by fibronectin-green fluorescent protein. Proc. Natl. Acad. Sci. US 96, 2153–2158. Kalajzic, I., Braut, A., Guo, D., Jiang, X., Kronenberg. M, S., Mina, M., Harris, M. A., Harris, S. E., and Rowe, D. W. (2004) Dentin matrix protein 1 expression during osteoblastic differentiation, generation of an osteocyte GFP-transgene. Bone 35, 74–82. Yang, W., Lu, Y., Kalajzic, I., Guo, D., Harris, M. A., Gluhak-Heinrich, J., Kotha, S., Bonewald, L. F., Feng, J. Q., Rowe, D. W., Turner, C. H., Robling, A. G., and Harris, S. E. (2005) Dentin matrix protein 1 gene cis-regulation: use in osteocytes to characterize local responses to mechanical loading in vitro and in vivo. J. Biol. Chem. 280, 20680–20690. Ghosh-Choudhury, N., Windle, J. J., Koop, B. A., Harris, M. A., Guerrero, D. L., Wozney, J. M., Mundy, G. R., and Harris, S. E. (1996) Immortalized murine osteoblasts derived from BMP 2-T-antigen expressing transgenic mice. Endocrinology 137, 331–339.

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25. Kalajzic, I., Kalajzic, Z., Kaliterna, M., Gronowicz, G., Clark, S. H., Lichtler, A. C., and Rowe, D. (2002) Use of type I collagen green fluorescent protein transgenes to identify subpopulations of cells at different stages of the osteoblast lineage. J. Bone Miner. Res. 17, 15–25. 26. 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. 27. Dallas, S. L., Veno, P. A., Bonewald, L. F., Rowe, D. W., and Kalajzic, I. (2007) Dynamic Imaging of Fluorescently Tagged Osteoblast

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Part V Imaging Techniques

Chapter 27 Analysis of Bone Architecture in Rodents Using Microcomputed Tomography Robert J. van ‘t Hof Abstract This chapter describes the use of microcomputed tomography scanning for analysing bone structure, focussing on rodent bone. It also discusses sample preparation, the correct set-up of the scanner, and the impact of some of the important scanner settings. Key words: X-ray, 3D, Rodent, Computerised tomography, Imaging CT, Micro-CT

1. Introduction Radiological techniques, such as plain X-ray and dual energy X-ray absorptiometry (DXA), are widely used for the investigation of patients with bone disease in routine clinical practice and similar techniques can been used to examine the skeleton in animal models of bone disease. For example, the Piximus DXA scanner (GE Healthcare) has been used to measure bone density in mice and rats and follow the changes in bone density that result from ovariectomy. Similarly, radiological analysis of the skeleton in mice and rats can be undertaken using the Faxitron instrument. Although plain X-rays are adequate to detect gross morphological changes in the skeleton of rodents, they do not have sufficient resolution to detect subtle changes in bone structure or bone density. Similarly, DXA analysis has low sensitivity for detecting the changes in bone density that occur after ovariectomy, especially in mice. This is because most of the changes occur in the trabecular bone and DXA scanners cannot separate trabecular from cortical bone. For example, we have found that the Piximus scanner shows bone loss at the proximal tibia of about 5–10% 3 weeks after ovariectomy in mice (which is barely statistically significant using ten animals per group) while

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_27, © Springer Science+Business Media, LLC 2012

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analysis of a similar experiment using mCT analysis showed a highly significant 30–40% decrease of trabecular bone at the same site. The current generation of mCT scanners are the current method of choice for skeletal phenotyping of rodent models of bone disease. In contrast to histomorphometry, mCT is a nondestructive technique, and specialised instruments for mCT imaging of live animals are now available, which allow the researcher to conduct serial studies during skeletal growth and ageing in mice and rats. 1.1. Micro-CT Analysis

Micro-CT involves taking a series of X-ray images of the sample at different rotations, and then using computer algorithms to reconstruct a 3D image stack (1, 2). The process of mCT analysis can, therefore, be divided into three different stages: 1. Acquiring the X-ray projection images 2. Computerised reconstruction of the 3D stack of images from the projection images 3. Analysis of the 3D image stack

1.2. Acquiring the X-Ray Projection Images

Two different strategies are used to obtain images at different rotations. In most standard desktop mCT systems, the sample stage containing the specimen to be analysed is rotated, whereas the in vivo systems use a gantry to rotate the X-ray source and camera around the sample stage containing the specimen which is fixed in stationary position. One consequence of this difference between in vivo and standard systems is that the in vivo systems tend to be less flexible in the range of resolutions, as the source and camera are in a fixed position relative to one another. The in vivo systems also tend to have a lower maximum resolution. The best in vivo mCT systems have a best resolution of 9–10 mm, whereas the desktop systems normally have a best resolution in the 2–5-mm range and systems with <1-mm resolution are available.

1.3. Image Reconstruction

Most reconstruction software use the same Feldkamp cone-beam reconstruction algorithm (3), although faster algorithms have recently been developed. This algorithm is based on filtered-back projection, but it is beyond the scope of this chapter for its details to be discussed here. An important requirement for valid reconstruction is that the X-ray beam is not fully absorbed at any point in the sample; there should always be some transmission at each pixel of the X-ray camera.

1.4. Analysis of the 3D Image Stack

Many different software packages are available for quantitative analysis of the resulting 3D image stack. Many standard image analysis packages are available that could theoretically handle 3D image stacks, but most users employ specialist software packages that are provided by the manufacturers of mCT systems. The most

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widely used mCT machines are those manufactured by Scanco and Skyscan and these instruments have similar capabilities. In this chapter, however, the examples given are those based on the Skyscan CTAn software, as our lab uses Skyscan mCT systems.

2. Materials 2.1. mCT Scanner

The most commonly used mCT systems for the study of bone disease are manufactured by Scanco and Skyscan. Other systems are also available, however, manufactured by GE Healthcare and XRadia. The Skyscan 1172 or Scanco mCT35 are generally used for scanning specimens ex vivo, whereas live animals can be scanned in vivo using the Skyscan 1076 or the Scanco vivaCT 40 systems.

2.2. Computing Equipment

Reliable and powerful computer systems are required for mCT analysis to control the scanner, reconstruct the images, and analyse the data. The Skyscan systems are designed to work with standard Microsoft Windows®-based computers, whereas the Scanco systems are designed to work with 64-bit OpenVMS Unix workstations. Although image reconstruction using the Feldkamp conebeam algorithm is very computing intensive, it is ideally suited to parallel processing using a cluster of computers. We use a cluster of four dual-processor workstations and a special cluster version of the Skyscan reconstruction software (NRecon). This allows us to reconstruct a dataset in less time than the scan time of a specimen in most cases. Similar cluster software is available for Scanco systems. The mCT scanners can produce large amounts of data; at roughly 1 GB of data for a 5-mm scan of a mouse proximal tibia, our Skyscan 1172 can easily produce 40–50 GB of data per day. Sufficient data storage and back-up capacity is, therefore, required. We use a dedicated file server with 12 TB of storage capacity and a second system with 6 TB capacity. For data archiving, we use a high-speed, high-capacity tape system (LTO4 Ultrium) that can save approximately 1 TB of data per tape. Although high-capacity external hard drives are very attractively priced these days, they are in general not very reliable for long-term storage. Analysis of the data is again very computing intensive, and requires the handling of large datasets. Because of the size of the datasets, it is beneficial to use analysis workstations that run a 64-bit operating system, such as OpenVMS or 64-bit Windows, as these allow for a larger amount of RAM to be installed. We currently use 64-bit workstations running Windows 7 that are fitted with 12 GB of RAM and Core i7 processors running at 3.2 GHz. The whole process of acquiring the X-ray projection images, reconstruction, and analysis requires the data to be moved between

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the file servers and the different types of workstations used. Because of the size of the datasets, this can place severe demands on standard computing networks which are designed to handle relatively small datasets, such as text documents and e-mails. The network can then become a significant bottleneck in the analysis project, and saturation of the network with mCT data can lead to irritation in colleagues unable to reliably use their computer for their normal daily work. We, therefore, use a dedicated sub-network connecting the systems used for mCT using a dedicated network switch. It is important to select a professional switch with sufficient data throughput capacity, as many cheap home or office switches struggle to handle the data streams generated by mCT scanners. 2.3. Other Materials

1. Sample holders. 2. 4% formalin in phosphate-buffered saline (PBS). 3. PBS. 4. 70% ethanol. 5. Parafilm. 6. Dremel hobby tool. 7. Diamond wafering blade.

3. Methods 3.1. Ex Vivo Analysis of Bone Architecture

There are several scanning parameters to be set up, but the optimal settings depend on the type and size of the sample, and upon what needs to be analysed.

3.1.1. Voltage

The first parameter to decide on is the X-ray voltage since this determines the spectrum of X-ray energies. Low voltage shifts the distribution to low-energy X-rays while higher voltages shift the spectrum to high-energy X-rays. For soft tissue imaging, low voltages around 30–40 kV are generally used, whereas for small animal bone samples voltages should typically be set in the 50–60-kV range. Since individual X-ray generators may produce slightly different spectra even at the same voltage, it is best to determine the optimal voltage empirically by trying a small range of voltages with a test sample, and choose the setting that gives the best contrast. We generally use a 60-kV setting for our X-ray source for scanning bone samples. Even at this setting, the X-ray spectrum contains some low-energy rays that negatively affect sample contrast for bone samples, but this part of the spectrum can be removed by the insertion of a 0.5-mm aluminium filter which is one of the standard filters supplied with the Skyscan 1172 system.

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The next parameter to decide upon is the scan resolution. This depends very much on the nature of the sample and what the researcher is trying to detect. Mouse bone architecture can be imaged very well at a resolution of 4–5 mm (see Note 1). This is sufficient resolution to measure the fine trabecular structure, as most mouse trabecular structures have a width in the range of 30–60 mm. As Fig. 1 shows, reducing the resolution below 10 mm makes it almost impossible to reliably visualise mouse trabeculae.

Fig. 1. Effect of resolution on image quality. A mouse proximal tibia was scanned at different resolutions. (a) Shows an overview of a slice and the square indicates the area represented in (b–f). (b) 2.5 mm, (c) 5 mm, (d)10 mm, (e) 20 mm, (f) 30 mm. The image quality of the 5-mm scan in (c) is good enough for analysis, and even the 10-mm scan (d) provides a reasonable amount of detail. The 20- and 30-mm scans, however, are to be blurred and lack sufficient detail and contrast for meaningful analysis of trabecular architecture. (g) Scan of a human bone biopsy at 30 mm showing sufficient detail for meaningful analysis.

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Table 1 Influence of scan resolution settings on commonly used mCT measurements Resolution (mm)

BV/TV (%)

Tb.Th (mm)

Tb.Sp (mm)

Tb.N (mm−1)

SMI

Conn.Dn (mm−3)

Mouse tibia

2.5 5 10 20 30

9.05 9.17 9.75 10.64 7.55

45.40 47.94 62.27 90.66 111.41

279.89 288.79 362.93 501.52 708.19

2.15 1.91 1.57 1.17 0.68

1.91 2.10 2.27 2.51 2.67

459.69 269.88 93.26 61.31 37.41

Human biopsy

10 20 30

15.89 15.43 18.36

159.80 174.44 210.45

736.41 774.31 767.90

0.99 0.88 0.87

1.03 1.14 1.24

5.88 4.66 3.92

Specimen

The proximal tibia from a 3-month-old female mouse and a human bone biopsy were scanned at a range of different resolutions and the main trabecular bone measurements performed. BV/TV: percentage trabecular bone volume; Tb.Th: trabecular thickness; Tb.Sp: trabecular spacing; Tb.N: trabecular number; SMI: structure model index, an indicator whether the trabecular structures are rod-like or plate-like. A lower value indicates a more plate-like structure; Conn.Dn: connectivity density

However, rat and human trabeculae are considerably thicker, and 10–20 mm is usually sufficient for rat samples and 20–30 mm for human bone biopsies (Fig. 1g). One could of course always scan at the maximum resolution of the scanner; however, this usually leads to much longer scan times and larger datasets that take much longer to reconstruct and analyse. Halving of the voxel size (a voxel is the 3D equivalent of a pixel), for example from 10 to 5 mm, leads to an eightfold increase in file size to image the same volume. Datasets can get so large that standard computers would no longer be able to handle them or take days or weeks to perform the conebeam reconstruction. It is, therefore, best to use the minimum resolution acceptable to analyse the details of interest. Table 1 shows the effect of scanning resolution on some of the standard measurements for bone. There is relatively little difference between the bone volume of a mouse proximal tibia assessed at 2.5 and 5 mm. The 10- and 20-mm scans, however, tend to overestimate the amount of trabecular bone. There is also a substantial overestimate of trabecular thickness and trabecular spacing with increasing voxel size, and an underestimate of trabecular number and connectivity. Most of these effects can be explained by the fact that at lower resolutions thinner trabeculae are no longer detected. Also the partial volume effect (when a voxel is part bone and part soft tissue) leads to smearing of the structures, resulting in overestimation of thickness. The results in Table 1 show that in most cases a resolution around 5 mm is sufficient for mouse bone, with only a minor benefit of higher resolutions. Although similar trends can be seen when scanning human bone biopsies (Table 1), the resolution is much less critical, as the trabeculae in human bone are substantially thicker.

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The final important parameter at this stage is the rotation step between the individual projection images. A larger step size reduces the number of projection images acquired, and thereby scan time, dataset size, and reconstruction time. However, it also negatively affects image quality as Fig. 2 shows. For a 5-mm mouse scan, a step size in the range of 0.4–0.6° usually gives sufficient image quality for bone analysis. An additional step to improve image quality is averaging of several images at each rotation step. This reduces image noise, but can substantially increase scan time. For standard analysis, image averaging is not necessary. On our Skyscan 1172 system, a scan of a mouse proximal tibia using our standard settings of 60 kV, 0.5-mm Al filter, 0.6° rotation step, and 5-mm resolution take on average 11–12 min.

Fig. 2. Effect of rotation step on image quality. A mouse proximal tibia was scanned at 5-mm resolution using different rotation steps. (a) 0.2°, (b) 0.6°, (c) 2°, (d) 4°. The image quality increases with decreasing step sizes. Although (a) shows the best signal-to-noise ratio, the image acquisition is three times longer and data size three times larger than (b), which is still good enough for analysis. The noisy image quality in (c) and (d) makes reliable thresholding of the image virtually impossible.

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3.1.4. Sample Preparation

Although samples from many species can be analysed using mCT, the examples here are based on the analysis of mouse bones. The two most commonly used sites for mCT analysis of mouse bone are the proximal tibia and the distal femur since they are easy to dissect out and mounted in the scanners. In many cases, the investigator wants to perform histological analysis of samples after scanning. If this is required, the bones should be dissected, fixed overnight in 4% buffered formalin, washed in PBS, and stored in 70% ethanol prior to scanning. We then cut of the distal femur and proximal tibia from the bones using a Dremel hobby tool with a diamond wafering blade. Although mCT scanning is non-destructive, there is a risk of the samples drying out in the scanner due to the heat that is generated. We, therefore, wrap the samples in Parafilm® prior to scanning to minimise the risk of this occurring since it is virtually transparent to the X-ray beam. While other plastic films can also be used, PVC-containing films should be avoided since the chlorine atoms in these can cause attenuation of the X-ray beam. The reconstruction algorithm requires that the sample does not move during the scan, and the samples therefore need to be kept in place using a sample holder. Again, care should be taken that the sample holder is relatively transparent to X-rays. We routinely use holders made from 1-ml syringes and 5-ml pipettes (Fig. 3a). These can hold several samples above one another. In combination with the batch scanning options in the Skyscan 1172, this means that we can normally load 5–6 samples in a holder and

Fig. 3. Sample holders and batch scan of multiple samples. (a) Shows a collection of sample holders made from a centrifuge tube, 1-ml syringe, and a 5-ml pipette. (b) Shows the sample holder fitted to the sample stage. (c) Shows a scout view of the sample holder with five samples. The bottom two samples have already been set up for scanning.

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scan these without further user intervention being needed. This normally takes about 1 h of scan time, leaving the operator free to do something else. Once the sample has been loaded in the scanner (Fig. 3b) and scanner parameters have been set, a scout scan is performed showing the position of the individual samples (Fig. 3c). Once the samples’ positions have been set and the samples labelled, the scan is started. 3.1.5. Reconstruction

After the scan has finished, the projection images are loaded into the reconstruction program NRecon. This allows selection of the part of the scan to be reconstructed, and set the reconstruction parameters (see Note 2). One of the options in the reconstruction software is compensation for beam hardening. Beam hardening is a mCT artefact resulting from the fact the X-ray tubes used in these systems do not produce X-rays of a single energy, but a spectrum of energies. When the X-ray bundle interacts with the sample, the lowest X-ray energies are absorbed first. The remaining X-ray energies in the beam, therefore, have a higher average energy while passing through the rest of the sample. The result of this is that the outside layer of a sample appears to have higher X-ray attenuation. Beam hardening correction software tries to correct for this. The optimal setting needs to be assessed empirically for each scanner and type of sample, and for mouse bone samples the values tend to be in the 10–20% range. Another reconstruction option is smoothing. Although this results in images with less noise, it can impair the detection of fine detail in the image. Because of this, we usually do not activate smoothing. Ring artefact reduction is another correction option which tries to reduce artefacts resulting from the fact that the sample is rotated. A higher precision setting increases the setting increases the reconstruction time; but on modern computer systems, this is not a big problem, and we usually select a setting around 10. Once the parameters are set, a preview of a single slice is performed. This allows the researcher to determine whether the settings are correct and to select a region of interest. It is not useful to keep large image areas outside the sample, as these would only increase the dataset size. If the preview looks correct, a destination folder is chosen for the reconstructed images and the reconstruction started. Once reconstruction is complete, the dataset can be viewed and analysed.

3.1.6. Analysis

A very important facet of the analysis is the selection of a volume, the volume of interest or VOI that can be reproducibly identified in all samples, and contains a reasonable amount of trabecular or cortical bone (see Note 3). Mouse bones contain relatively little trabecular bone, and most of this is located close to the growth plates in the proximal tibia and distal femur metaphysis. A good landmark in mouse long bones is the growth plate. As Fig. 4 shows,

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Fig. 4. Selecting volumes for analysis. The boxes in the longitudinal sections of mCT scans of a mouse femur (a) and tibia (b) indicate the best areas for measuring trabecular bone volume. Landmarks, such as the growth plate cartilage and mineralised cartilage, are also indicated. (c, d) Examples of reference levels, where the last bridge of mineralised cartilage has been broken. (e, f) Slices 600 levels below the reference in the volume used for measuring cortical bone parameters. The shape of the tibia makes measurements from this bone harder to interpret than from the femur.

in the mouse tibia and femur, the mineralised cartilage can be identified relatively easily, and it forms a bridge across the bone. We select as a reference point the level at which this bridge breaks (Fig. 4c, d). For the tibia, the volume analysed is then 200 slices starting 20 levels distal to the reference point. A similar method can be used for the distal femur (Fig. 4a), but here the volume to be analysed should start 20 slices proximal from the reference level. Another landmark that can be used is the cartilage of the growth plate. However, the distance of the starting level from the reference

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level needs to be larger to avoid the primary spongiosa, in the region of 50 slices proximal to the reference level. The next step is separation of trabecular bone from cortical bone. This is done manually by drawing in a number of layers (usually, around 6) distributed through the selected volume (Fig. 5a). The software then interpolates to create a separation for the layers in between. Next, the bone tissue is separated from the soft tissue using thresholding. The actual threshold value depends on settings during the scan and the reconstruction stages (Fig. 5b, c). However, finding a good setting is usually fairly straightforward. Once the threshold has been decided on, the same value should be used for all samples

Fig. 5. Identifying trabecular bone. The trabecular bone is manually separated from the cortical bone as indicated by the blue area in (a). (c) Shows the presence of noise after thresholding the unfiltered image in (b). (e) Shows the much cleaner effect of the same operation on a median-filtered image in (d). Noise is indicated in (c) by yellow arrows.

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within an experiment. Thresholding can often be improved by applying noise reduction filters to the images before the threshold operation. We routinely use a median filter (radius size 1–2) as this very effectively suppresses noise without affecting the edges of structures (Fig. 5d, e). Filters based on averaging tend to soften the edges, and this can negatively affect the threshold result. After thresholding has been performed, the image can be further cleaned by removing small fragments. As the trabecular bone is a highly connected structure, any unconnected small objects are unlikely to be bone and can therefore be removed using a despeckling operator. The final operator used is the 3D analysis operator; this performs all the measurement and calculations for the final output data. Table 1 lists the most commonly used parameters. The Skyscan CTAn software allows the user to create macros which run all the operators in sequence. Although the parameters mentioned above are suitable for analysis of trabecular bone, the cortex is not very well-defined at this level. For the analysis of cortical bone, we routinely use a volume of 100 slices, starting at a distance of about 600 slices from the reference level (Fig. 4e, f). The thresholding and analysis are then performed as described for trabecular bone. However, one additional operator is usually necessary to deal with pores in the cortex which can negatively influence the average cortical thickness (see Note 1). The hole fill operator is run to close these pores before running the analysis operator. Although the main parameter to be studied is cortical thickness, other parameters, such as periosteal and endosteal circumference, porosity and shape parameters, such as eccentricity and minimal and maximal bone diameter, may also be analysed. Most of these parameters can be measured on a single 2D slice. It is possible to measure the mineral density of the bone tissue by mCT. If this is required, a calibration phantom should be scanned and the image reconstructed using the same scanning parameters as the bone samples. Skyscan provides standard mouse (2-mm diameter) and rat (4-mm diameter) hydroxyapatite phantoms, each containing a resin loaded with hydroxyl apatite at 250 and 750 mg/cm3, and this provides a linear calibration, which can then be used to calculate mineral content from the X-ray attenuation data. This density is different from that measured in a DEXA scanner. DEXA scanners measure the average density of a bone, so this is lowered by the contribution of soft tissue. In contrast, the mineral density measured using mCT represents the mineral content of the mineralised tissue only. It is very important to have optimised the beam hardening correction when measuring density. Averaging of images during image acquisition and small rotation step sizes can also be useful for density measurements as this suppresses noise in the attenuation values, leading to a more accurate measurement.

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3.2. Analysis of Focal Bone Lesions Using mCT

Analysis of bone architecture in mice requires relatively highresolution images as described in section Subheading 3.1, but there are other applications that do not need such high resolution. One of these is the detection of periarticular erosions, metastases, and focal lesions, such as those occurring in models of inflammatory arthritis, cancer, and Paget’s disease of bone. The lesions are usually large enough to be detected at a resolution of around 15–20 mm. Detection of bone erosions in the mouse collageninduced arthritis model is fairly straightforward. The arthritis affects hind and rear paws, and the individual paws can easily be imaged in the Skyscan 1172. Scanning should be performed at standard settings of 50 kV, 0.5-mm Al filter, and 0.6° rotation step but with a 17-mm resolution. Reconstruction should be performed as described previously, but analysis should be performed slightly differently as the purpose of the scans is visualisation of lesions rather than quantification of the trabecular bone structure. Image filtering and thresholding are performed as described for trabecular bone analysis. However, the final step is not a 3D analysis, but the creation of a 3D model for viewing. Once generated, these models can be freely viewed, rotated, cut outs performed, and movies generated using special 3D model viewing software, such as the Skyscan CTVol program. The resulting images clearly show the bone destruction in the arthritic animals compared to the control animals (Fig. 6a, b). We have used a similar technique to detect lytic lesions in a mouse model of Paget’s disease of bone (4). One important difference with the arthritis model is, however, that the location of the lesions is more difficult to predict. We, therefore, needed to scan entire mouse legs rather than just a small volume like the proximal tibia. However, even at a low resolution of 17 mm, a single scan can only image part of a mouse leg. The Skyscan 1172 allows the researcher to set up “oversize” scans, in which images from several scans can be combined into a single dataset. Once this has been achieved, reconstruction and generation of a 3D model should be performed as described for the periarticular erosions mentioned previously. Figure 6c, d shows a lytic lesion in a 12-month-old transgenic animal. The overview of the femur in Fig. 6c was produced by stitching two scans together in software.

3.3. In Vivo mCT Analysis

The strength of using in vivo mCT scanners is that changes in bone architecture can be monitored over time in individual animals. However, as each scan involves exposure of the animal to radiation, one should ensure that the cumulative dose does not exceed levels that affect bone metabolism. Also, since the animals need to be anaesthetised during the scan, consideration should be given to the effects of repeated anaesthesia on the animal. The best area to scan is around the knee since the hind limbs can be easily positioned, and this region is not covered with thick

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Fig. 6. Visualising focal bone lesions. (a, b) Show 3D models of mouse hind paws scanned at 18 mm. A is a control animal while b is a scan of a mouse 3 weeks after the induction of collagen-induced arthritis. Joint destruction is indicated by the arrows in (b). (c) Shows an 18-mm scan of the femur of a animal model of Paget’s disease. The lytic lesions in the cortex are clearly visible at this resolution. (d) Shows a cross section indicated by the dotted line in (c), scanned at 5 mm. (e) Shows the same level from a wild-type control mouse. Figure (a, b) by E. Coste, R van ‘t Hof, and S.H. Ralston; (c–e) by A. Daroszewska, R. van ‘t Hof, and S.H. Ralston.

layers of soft tissue. We use a small, circular polystyrene holder and masking tape to position and hold the limbs in place (Fig. 7a). When performing in vivo scans, it is very important to restrain leg movement since this can lead to serious artefacts in the reconstructed images (Fig. 7c).

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Fig. 7. In vivo scanning. For proper and reproducible positioning and to minimise movement, the mouse legs are inserted into a polystyrene ring and the feet taped together (a). (b) Shows a slice from a good 9-mm scan of the tibia and fibula of a 4-month-old mouse. (c) Shows the artefacts resulting from leg movement during the scan. (d) Shows the amount of trabecular detail that can be obtained from in vivo scans of mouse bone.

The scanner parameters should be set up essentially as described for the ex vivo scans (50 kV, 0.5-mm Al filter, 0.6° rotation step), with the exception of the resolution settings. We use a resolution of 9 mm on a Skyscan 1076 for analysis of mouse trabecular bone, and using this setting a scan takes approximately 20 min. For the detection of bone lesions, a resolution of 18 mm is sufficient, and this reduces the scan time to about 8–9 min. Reconstruction and analysis are performed as described for the ex vivo scans. However, the limited resolution of the in vivo scanners can make thresholding for mouse trabecular bone more difficult. One way to get more reliable threshold results is to use special, adaptive thresholding techniques. The wide field of view necessary for scanning of live animals means that the image files at 9-mm resolution are large and that reconstruction times can take several hours per scan.

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4. Notes 1. Analysis of cortical porosity in mice may require higher resolution as many pores are smaller than 5 mm. 2. The NRecon reconstruction software provided by Skyscan requires an additional step. The raw reconstructed image data is in a floating point format, and this is scaled by the software to an 8-bit integer image file. An important step is to choose the correct maximum value from the floating point data and to use the same scaling for all samples in an experiment. 3. For accurate VOI selection, bones should be oriented straight. This can be done by positioning the samples correctly in the sample holder. However, due to the irregular shape of bone samples, especially the tibia, it is very difficult to position all samples exactly the same way. The Skyscan Dataviewer allows the user to rotate the image stack along all three major axes, and this can be used to ensure the correct orientation of all samples. This step is extremely important when measuring 2D shape parameters (such as elongation and minimum and maximum diameter) on single slices of the cortical bone. References 1. Hounsfield, G. N. Computerized transverse axial scanning (tomography): Part1. (1973) Description of system. Br. J. Radiol. 46, 1016–1022. 2. Sasov, A., and Van Dyck, D. (1998) Desktop X-ray microscopy and microtomography. J Microsc.; 191(Pt 2):151–158.

3. Feldkamp, L. A., Davis, L. C., Kress, J. W. (1984) Practical cone-beam algorithm. J. Opt. Soc. Am. A6, 612–619. 4. A. Daroszewska, A., Van ’t Hof, R., Rose, L,. Rose, K., and Ralston, S. (2009) The P392L Mutation of SQSTM1 Causes a Paget’s DiseaseLike Phenotype In Mice. J. Bone Miner. Res. 24 (Suppl 1), S38.

Chapter 28 Bone Measurements by Peripheral Quantitative Computed Tomography in Rodents Jürg A. Gasser and Johannes Willnecker Abstract This chapter provides information for the use of peripheral quantitative computed tomography in small animals, including suggestions for study design, instrument setting, and data interpretation. Key words: In vivo computed tomography, Bone structure, BMD, Rat, Mouse

1. Introduction Peripheral quantitative computed tomography (pQCT) is a powerful technique for non-invasive monitoring of envelopespecific changes in bone mass and cortical geometric parameters in rodents (1). QCT is unique among methods of bone mineral measurement in that it can provide separate estimates of trabecular and cortical bone mineral density as a true volumetric mineral density value [g/cm3] (2). Additionally, QCT can assess the geometric properties of cortical bone with high accuracy (3) and predict some of the mechanical properties of bone with remarkable precision (4–6). If used appropriately, the technique can monitor bone changes in disease states (OVX, immobilisation, inflammation, tumour osteolysis), characterise the effects of therapeutic interventions at various envelopes, and initiate the process of analysing a bone phenotype in genetically modified mice. Through the use of specific algorithms, calculated parameters, such as the axial or polar bone strength index, can reliably predict bone strength for bending or torsion. The technique clearly has its limitations and should, thus, be used in conjunction with other established methods. Also, the prediction of ultimate strength (BSI) calculated by the pQCT on 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_28, © Springer Science+Business Media, LLC 2012

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basis of structural information does not provide information on all aspects of mechanical properties of bones, such as changes in matrix composition leading to brittleness, abnormal collagen structure as observed in woven bone, and the effects of surface-based bone remodelling which creates focal weak spots (stress raisers) in cancellous bone. All these gaps have to be filled in by other methods, such as mechanical testing, backscattered electron imaging, and histopathological investigations. The prototypes of the XCT900 and XCT960 scanners were first developed at the University of Würzburg (7), and brought into commercial clinical and preclinical application by Stratec Medizintechnik GmbH, Germany. Subsequently, dedicated small animal scanners with higher resolution, such as the XCT Research SA for small animals and the XCT3000 Research for larger animals, were developed (Table 1). pQCT measurements can be carried out ex vivo on any excised bone, including vertebral bodies. All pQCT scanners of the XCT series work according to the translation– rotation principle. The photons emitted by the X-ray tube are detected by the 12 semiconductor detectors which have a near100% efficacy for X-rays of around 38.5 keV. The attenuation coefficients at each point of the cross-sectional image are reconstructed from the projected data (filtered back-projection) (8). Daily calibration of the system using a custom-made phantom containing commercial plastics allows calculating density values (mg/cm3) from the attenuation coefficients.

Table 1 Technical data of the XCT series XCT Research SA

XCT Research M

XCT Research SA+

XCT3000 Research

Slice thickness

0.8 mm

0.5 mm

0.7–0.13 mm

1 mm

High voltage

50 kV

50 kV

50 kV

60 kV

X-ray energy

38 keV

38 keV

38 keV

45 keV

Matrix size

Up to 1,000 × 1,000

Up to 1,000 × 1,000

Up to 1,000 × 1,000

Up to 1,000 × 1,000

Resolution

Up to 90 μm

Up to 70 μm

Up to 70 μm

Up to 100 μm

No. of projections

180–720

180–720

180–720

180–720

Object size

Up to 90 mm

Up to 50 mm

Up to 90 mm

Up to 270 mm

Scan length

170 mm

170 mm

170 mm

390 mm

Scatter radiation dose in 1-m distance

0.1 μSv/h

5 μSv/h

0.1–5 μSv/h

0.1 μSv/h

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The true potential of pQCT lies in its ability to non-invasively assess bone and muscle development in vivo, where it can be applied to study the skeletal phenotype in genetically modified animals and to monitor the effectiveness of therapeutic interventions. The present chapter focuses entirely on the use of the XCT Research SA+ pQCT scanner for imaging the skeleton of rats and mice. The chapter focuses on technical aspects, such as the anaesthesia, choice of animal (age), experimental design, choice of the scan site and region of interest, positioning of limbs, instrument setting, data analysis and interpretation.

2. Materials 1. Scanners: XCT Research SA+ scanner (Stratec Medizintechnik GmbH, Germany) (see Note 1) or XCT960A scanner (Stratec Medizintechnik GmbH, Germany). 2. Inhalational anaesthetic apparatus capable of delivering 2.5% isoflurane with 0.8% oxygen and 0.8% air (e.g. Fluovac). 3. Syringes and needles. 4. Ketarom® injectable anaesthetic (1.2 ml ketamine hydrochloride + 0.8 ml rompun 2% + 8 ml NaCl). 5. Device to position animal limbs in scanner.

3. Methods 3.1. Experimental Design and Choice of Animals

Most of the pQCT data reported in the literature is derived from measurements carried out in Sprague-Dawley or Wistar rats (2–6, 9, 10), but given the more frequent use of genetically modified mice in skeletal biology, pQCT measurements are increasingly added to study mice (11, 12). Accordingly, it is now possible to use pQCT to analyse rat or mouse bones, but a number of factors must be taken into consideration to assure the best possible quality of the data.

3.1.1. Group Size and Scanner Precision In Vivo and In Vitro

1. A minimum of ten animals should be studied per group to take account of the coefficient of variation obtained on the XCT960A (rats) and XCT Research SA+(rats and mice) (Table 2). 2. As a rule of thumb, the difference between the means in two groups of animals must be at least three times the value of the CV displayed in Table 2 in order for you to obtain statistically significant differences.

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Table 2 Coefficient of variation for the XCT960A and XCT Research SA+ scanners XCT Research SA+ Rats: XCT960A Parameter (units)

Rat

Mouse

CVi%

CVv%

CVr%

CVr%

CVr%

0.331

0.598

0.869

1.13

0.97

Total bone mineral density (mg/mm )

0.116

0.170

0.273

0.36

0.25

Total bone area (mm2)

0.327

0.700

1.087

1.51

1.11

0.681

1.331

2.330

3.02

3.76

0.444

0.423

0.740

0.77

1.54

Trabecular bone area (mm )

0.542

1.060

1.754

2.25

2.32

Cortical bone mineral content (mg/mm)

0.456

0.576

0.856

0.44

0.79

Cortical bone mineral density (mg/mm3)

0.214

0.319

0.540

0.77

0.39

Cortical bone area (mm )

0.605

0.849

1.353

0.90

1.03

Cortical thickness (circular ring model) (mm)

0.585

0.688

1.152

0.42

0.97

Periosteal perimeter (mm)

0.163

0.350

0.544

0.74

0.55

0.283

0.528

0.867

1.11

1.26

0.877

1.332

2.315

nd

nd

Total bone mineral content (mg/mm) 3

Trabecular bone mineral content (mg/mm) 3

Trabecular bone mineral density (mg/mm ) 2

2

Endocortical perimeter (mm) 4

3

Bone strength index (mm g/cm )

The coefficient of variation in percent was calculated using the formula (CV = STDEV/mean × 100). Instrument precision (CVi%) was determined by repeating ten measurements in the proximal tibia ex vivo, without repositioning of the bone at a distance of 5 mm from the proximal end of the bone. In vivo precision was calculated from ten measurements with (CVr%) or without repositioning of the limb (CVu%)

3. Investigators should determine their own set of parameters (CV values) for the experimental conditions they are using since the precision is dependent on various factors, such as strain of mouse or rat, age of the animals studied, model of instrument, site of interest, method used to hold the limbs, and instrument setting. 3.1.2. Age of Animals

1. For studies in rats, the minimum age should be 6 months unless the focus of the study is on the evaluation of skeletal growth or factors influencing it (13–15). The use of younger animals inevitably confounds disease- or drug- induced effects with those of skeletal growth. Figure 1 clearly demonstrates that, even though there is little change in cross-sectional bone area in ageing 6- and 9-month-old rats, periosteal modelling drifts are triggered by OVX in the younger, but not older animals.

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% change from baseline

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25 20 Sham (age 6 mo)

15

OVX (age 6 mo)

10

Sham (age 9 mo)

5

OVX (age 9 mo)

0 -5 -10

0

4

8

12

16

weeks Fig. 1. Changes in cross-sectional area in the tibia of 6- and 9-month-old rats. All values shown represent means plus SEMs. Note that sham-operated rats still show some growth-related increase in cross-sectional area up to age 7 months. OVX induces pronounced periosteal bone apposition in 6-month-old rats, but this effect is absent in 9-month-old rats.

2. For studies in mice, the minimum age should generally be 4 months (see Note 2). However, please keep in mind that unlike in rats trabecular bone parameters are not maintained at the level of peak bone mass in most mouse strains. This may make it very difficult for you to assess trabecular parameters in skeletally mature mice at a typical measurement site, such as the proximal tibia metaphysis. The situation for trabecular bone measurements in mice is somewhat better in the distal femur metaphysis, but you may find it difficult to position this site. For this reason, you may be forced to use skeletally immature mice (<4 months of age), if your focus is in trabecular bone, leaving you to deal with the difficulties of interpretation of your data due to interference of skeletal growth with your primary response. 3.1.3. Study Design

1. For longitudinal studies, it is important to obtain a baseline value for each animal so that you can relate changes induced by the disease or drug treatment to that value. 2. For longitudinal studies where different treatments are being compared, it is advisable to allocate the animals to the various treatment groups based on their total cross-sectional BMD and their trabecular BMD. Apart from matching the treatment groups for these two parameters, it is also recommended that you ensure that there is homogeneity of variances between the groups (see Note 3). 3. For studies where you plan to carry out measurements on excised bones, such as lumbar vertebral bodies (which you cannot measure in vivo), or where you plan to carry out ex vivo biomechanical tests or histomorphometric analyses, it is important to include a baseline group (necropsy at baseline).

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4. For intervention studies where excised bones are being measured, additional control groups may be required, where bones are collected at baseline (necropsy at baseline) and prior to initiating active treatment. 3.1.4. Analysis of Genetically Modified Mice

There may be situations which prevent you from setting up a “perfect protocol” as outlined under Subheadings 3.1.1–3.1.3. For example, some genetically modified mice may show reduced survival and it may be impossible to study their mature skeletal phenotype or to conduct longitudinal studies. It is also important for investigators to realise that abnormalities of skeletal phenotype may derive from disturbances occurring during intrauterine skeletal morphogenesis or changes in skeletal growth in the post-natal period, leading you to conclude perhaps wrongly that the respective gene, receptor, or protein in question may represent an interesting target for the treatment of osteoporosis.

3.2. Skeletal Regions of Interest

For in vivo monitoring of bone mass, density, and cortical architecture, pQCT measurements are restricted to locations in the appendicular skeleton and tail vertebra, although in the past most measurements were carried out in the hind limbs (2–6, 10).

3.2.1. Preferred Sites to Study Cancellous and Cortical Bone Parameters In Vivo

Various skeletal sites can be conveniently analysed by pQCT scanning in vivo and the benefits and limitations of each of these are discussed below. 1. Proximal tibia metaphysis (PTM): The PTM is an ideal site for pQCT scanning for various reasons (Fig. 2). This site is rich in cancellous bone and reacts with the greatest magnitude of change to interventions, such as OVX (2), immobilisation, and bone anabolic therapies, such as parathyroid hormone (PTH). Another reason for choosing the PTM is the fact that it has been the most popular site for measuring structural and dynamic histomorphometric parameters. Serial scans carried out at various locations throughout the rat PTM showed that a section placed at a distance of 4.5–5 mm from the proximal end of the bone, a position in the secondary spongiosa, is most suited for skeletally mature rats (6 months and older) (Figs. 3 and 4) (3). The ideal “corresponding” site in mice is located 2.5 mm from the proximal end of the tibia (Fig. 2). Not surprisingly, the magnitude of cancellous bone loss in the rat decreases when the section is placed further away from the knee joint because of the decreasing amount of spongy bone found, especially in older animals. 2. Distal femoral metaphysis (DFM): The DFM is another good site to study cancellous and cortical bone even though the local strain pattern, which generates large forces in the direction of the patella, limits the magnitude of cancellous bone loss at this site (2).

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Fig. 2. Regions of interest for pQCT analysis in skeletally mature rats and mice. The distances (in mm) from the joint space for the distal femur metaphysis and the proximal tibia metaphysis are indicated. A pure “cortical” bone site can be easily assessed in the mid-diaphysis of the tibia (horizontal white dotted line).

Fig. 3. Scout scan of the knee joint in a rat. The scout scan is shown on the right of the picture 4.5 mm from the proximal tibia. For comparison, a radiograph of the same region is shown on the left of the picture.

For rats, an ideal slice is placed at 5 mm from the distal end of the femur (3 mm in mice). 3. Distal tibial metaphysis (DTM): The DTM is of limited value for pQCT scanning. Its cross section is small and separating cancellous and cortical bone is in most cases not possible in rats and mice.

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Fig. 4. Example of QCT scan at proximal tibial metaphysis. The scan shown was taken by an XCT960 scanner, 4.5 mm from the knee joint at the proximal tibial metaphysis taken in a 9-month-old rat. In most cases, the software is able to define the ROI automatically on which the scan analysis is run. The dotted line delineates the contour of the muscle. Using a lower threshold, muscle area can be determined in all sections.

4. Tail vertebral bodies (TVBs): Measurements can be carried out in the TVBs which are easy to position. Unfortunately, the magnitude of bone loss in the TVB in response to manipulations, such as ovariectomy, is considerably smaller than observed in PTM, DFM, and most importantly the lumbar vertebral bodies (LVBs) so that this site is only of limited use. The reasons for this “failure” are not clear but may include factors, such as differences in cellularity (fatty marrow as opposed to red marrow), vascularity (metabolism), or protective local strain patterns. 3.2.2. Preferred Sites to Study Cortical Bone Parameters In Vivo

None of the algorithms can possibly separate cancellous and cortical bone in a perfect way. For this reason, we recommend to choose a “pure” cortical bone site in addition to the PTM or DFM. One of the easiest sites to localise repeatedly with high precision in longitudinal studies is the mid-diaphysis of the tibia (MDT). Bone length can be easily determined on a contact X-ray and the position calculated in growing as well as in skeletally mature rats and mice.

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3.3. Quality Assurance

Before initiating your measurements in animals, you must start the quality assurance (QA) procedure by scanning the QA phantom provided by the vendor. This procedure has to be carried out once every day before the machine allows you to collect data. The routine starts by initiating a Scout View scan (SV-scan) (Fig. 3) followed by the collection of data from the phantom on three different density values, all of which have to stay within the tolerated range of <1% of the true value.

3.4. Limb Positioning

Accurate positioning of the limb is one of the crucially important factors in order to guarantee high-quality data in longitudinal studies. For this purpose, we developed a special holder which guarantees exact positioning and does not allow movement of the limb in any direction, as well as prevents its rotation. 1. Place the animal on a plastic tray in the lateral position. 2. Place the mouth and nose of the animal in a hole to which a tube is plugged on to deliver the inhalation anaesthetic [2.5% isoflurane (Forene®, Fig. 5), which we deliver together with 0.8% oxygen and 0.8% air] (see Note 4). 3. Place the animal’s leg into the tube with the foot sticking out at the other end of the holder (Fig. 6). 4. Place the conical plug around the limb to hold the tibial muscle with the iron rod inserted into the lateral slit, thus preventing any rotation of the limb (see Note 5). 5. Placing the foot holder around the ankle and securing it with adhesive tape (Fig. 6) to prevent it from sliding back into the tube.

Fig. 5. Anaesthetic apparatus. The animals are placed into a lateral position under inhalation anaesthesia with 2.5% isoflurane (Forene®). Suction around the nose prevents spillage of fumes to the surrounding air. The overview shows the system with a special “flow-through chamber” in which the next animal can be anaesthetised while measuring the limb of another.

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Fig. 6. Device to position animal limbs in scanner. The custom-made device to position the hind limb in the pQCT scanner is shown. Detailed view of the leg holder is shown on the left panel. This consists of a tube, a conically shaped plug with a rod, and a foot holder. The function of the conical plug is to hold the tibial muscle preventing its rotation. The foot holder is placed around the ankle preventing it from withdrawing the leg into the tube.

6. Give each animal a unique code and keep this designation for all later time points. As soon as the code is typed into the scanner for a second time, the software knows that it should compare the results of the upcoming with an earlier measurement from the same animal. This software-based automatic detection procedure is very efficient at placing your measurements (ROI) at the same location of any previous scan. 3.5. Instrument Setting

The numbers of possibilities for setting your instrument are virtually endless and it is beyond the scope of this guide to discuss all of them. Suggested instrument settings for rats being scanned on the XCT Research SA, XCT Research M, Research SA+, and the XCT3000 Research scanners are summarised in Table 3. In our experience, they work very well and give robust results of high quality for both cortical and cancellous parameters.

3.5.1. Detection of Cortical and Trabecular Compartments

We have chosen to use the same threshold-based contour finding for detection of both the periosteal and endocortical surface (CONTMODE 1, CORTMODE 2, PEELMODE 2) (see Note 6). The CONTMODE 1 parameter determines the separation of soft tissue from bone for delineation of the outer contour (periosteal perimeter). The threshold attenuation coefficient can be freely chosen by the operator. The CORTMODE 2 algorithm separates trabecular from cortical bone at the endocortical surface. All voxels with a lower attenuation coefficient than the selected threshold are eliminated and counted as being part of the cancellous bone compartment. The voxels are then “proofed” by a three-by-three filter to ensure continuity. We recommend using the same threshold to define the inner contour of cortical bone as was used for detection

350 mg/cm

2

610 mg/cm

3

610 mg/cm3

2

2

1

10 mm/s 5 mm/s

1.4–0.25 mm 0.07 × 0.07 × 0.7–0.13 mm3

Research SA+

400

400

2

2

1

10 mm/s 5 mm/s

2 mm 0.1 × 0.1 × 1 mm3

XCT 3000 Research (primates, sheep, dogs)

Threshold determines the outer contour in the CONTMODE part of scan analysis while Threshold2 defines the cortical and cancellous bone compartment in CORTMODE

710 mg/cm

3

2

710 mg/cm3

CORTMODE

Threshold

Threshold2

350 mg/cm3 3

2

2

PEELMODE

1

1

CONTMODE

10 mm/s 5 mm/s

1.0 mm 0.07 × 0.07 × 0.5 mm3

20 mm/s 10 mm/s

1.6 mm 0.09 × 0.09 × 0.8 mm3

XCT Research M (mice)

Scan speed SV-scan Final scan

Collimator (diameter) Voxel size

XCT Research SA (rats)

Table 3 Instrument setting for measurements of the proximal tibia metaphysis with the XCT Research Series of scanners

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of the outer contour (i.e. 610–710 mg/cm3 in rats and 350– 400 mg/cm3 in mice). PEELMODE 2 implements an operatorselected threshold density defining the way subcortical and trabecular bones are separated. In higher species, especially human and non-human primates, where it is possible to distinguish between subcortical and trabecular bones, the PEELMODE 2 threshold is always selected at a lower level than the CORTMODE 2 threshold. However, in rats and mice, no subcortical bone compartment can be identified anatomically and thus all bones are either defined as cortical or trabeculae bones. For this reason, in small rodents, the PEELMODE 2 threshold has to be identical to the CORTMODE 2 threshold. 1. Select the values between 610 and 710 mg/cm3 in rats and 350 and 400 mg/cm3 in mice. You should choose the highest possible threshold for contour finding to minimise the partial volume effect. The ideal threshold is 710 mg/cm3, which corresponds to an attenuation coefficient of 0.94 cm−1 (see Note 7). 2. It is a good idea to run the scan analysis-loop always twice with two to three different thresholds (an ideal and a lower one). The thresholds we routinely chose are 510, 610, and 710 mg/cm3 in skeletally mature rats, and 280, 350, and 400 mg/cm3 for skeletally mature mice. If your experimental conditions result in bone loss and your initially chosen “higher” threshold fails during later data acquisitions, you can chose to work with the data of the “lower”, non-optimal second threshold without having to re-analyse all your earlier time points. 3. Choose the same peeling algorithms, including the thresholds for each animal and each measurement point. If your chosen set-up fails during later time points in your experiment, you must go back and re-analyse all your earlier data points of the entire experiment using lower thresholds. 3.5.2. Special Situations

Younger animals, which are subjected to skeletal growth, often require you to lower the thresholds displayed in Table 1. The same may be true for genetically modified animals with a severely osteopenic phenotype, animals with high rates of bone remodelling, and animals where mineralisation is disturbed, such as vitamin D deficiency and osteogenesis imperfecta.

3.6. Data Analysis and Interpretation

The analytical part of the software generates a great number of parameters, but luckily you do not require all of them. We believe that the 14 parameters listed in Table 4 are sufficient for adequate interpretation of your data. In the following, we attempt to describe such an “analytical” process on a true data set. The 12-week study was performed in 9-month-old virgin Wistar rats. Ten animals per group were ovariectomised and treated

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Table 4 Commonly used pQCT parameters Parameters

Definition

Unit

TOT_CNT

Total bone mineral content

mg

TOT_DEN

Volumetric total bone mineral density

mg/cm3

TOT_A

Total bone mineral area

mm2

CRT_CNT

Cortical bone mineral content

mg

CRT_DEN

Volumetric cortical bone mineral density

mg/cm3

CRT_A

Cortical bone mineral area

mm2

TRAB_CNT

Cancellous bone mineral content

mg

TRAB_DEN

Volumetric cancellous bone mineral density

mg/cm3

TRAB_A

Cancellous bone mineral area

mm2

PERI_C

Periosteal circumference

mm

ENDO_C

Endocortical circumference

mm

CRT_THK

Mean cortical thickness ring model

mm

xBSI

Axial bone strength index (bending strength)

mm4 g/cm3

pBSI

Polar bone strength index (torsional strength)

mm4 g/cm3

Column 1 lists the parameters as denominated by the software and their unit is listed in column 3. Only the most relevant parameters required for interpretation of the data are listed

daily p.o. by 0.3 mg/kg of 17α-ethinylestradiol dissolved in corn oil. Other groups of animals were injected twice weekly subcutaneously with 10 μg/kg alendronate or vehicle (corn oil). A shamoperated group was also measured to determine age-related changes with pQCT measurements being carried out in all groups at baseline, 4, 8, and 12 weeks. Although it is possible to analyse absolute data in well-matched groups of animals, we recommend that you calculate the percent change for each parameter and time point of all animals from their own baseline value. The reasons for this is simple: you find it virtually impossible to adjust the values of all treatment groups for all the 13 parameters that you want to evaluate when allocating the animals to their groups at baseline. When looking at your graphs containing the data, always keep in mind the “rule of thumb” mentioned earlier. In order for any “effect” to be real, the % change should be at least three times bigger than the respective CVr value displayed in Table 2.

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3.7. Interpretation of Data 3.7.1. Definition of Trabecular Density

3.7.2. Estimation of Bone Bending Strength

Trabecular density is defined as the mass of trabecular bone divided by the endosteal volume which includes all tissues within the endosteal area, including bone marrow. It is not the actual material density of trabecular bone. Given a material density of bone of 1,200 mg/cm3 and a trabecular volume of 20%, the apparent trabecular density is 240 mg/cm3 (Fig. 7). Cortical bone is composed of 90–98% of bone material and 2–10% of osteons. Therefore, the measured cortical density is 90–98% of the material density. More information about the different meanings of density can be found in an article by Rauch (32). The bending and torsional strength of an object can be calculated from its cross-sectional geometry and the material properties. The physical parameters are the moment of inertia I and the section modulus R. In addition, the pQCT software displays the SSI, a combination of section modulus and the cortical density (Fig. 8). The BSI, a combination of moment of inertia and cortical density, has been used by Ferretti to predict bone bending strength in rat femurs with high accuracy (33). In repeat measurements (time course), the axial values are affected by different rotation of the limb during the various measurements. In contrast, the polar SSI is

Bone marrow

Bone material density ≅ 1200

Volume of trabecular bone = apparent trabecular 20% density = 240 mg/cm3 Fig. 7. Calculation of trabecular density.

Volume of trabecular bone apparent trabecular =12% density = 144 mg/cm3

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x =distance of a voxel from the axis xmax = maximum distance of a voxel from the axis

x

a = area of a voxel [mm2]

xma

SSI =

Center of gravity

å X 2 *a*CD X max *1200

CD = measured cortical density [mg/cm3] 1200 = normal physiological cortical density

Fig. 8. Calculation of bone strength.

independent of rotation. Therefore, the polar value shows better reproducibility. Furthermore, the cortical density, mean attenuation coefficient, and cortical area are displayed. 3.7.3. Estimation of Fracture Load

An estimation of the load at which bones fracture can be calculated from the SSI using the formula: FB =

4sB ´ SSI , l

where FB is the fracture load, σB is the ultimate strength of bone, and l is the bending test distance. 3.7.4. Partial Volume Effect

When the object size is in the same order of magnitude as the scanning resolution, the partial volume effect must be taken into consideration. This is especially relevant with regard to cortical bone parameters. The CT image is displayed by individual voxels. If a voxel is not entirely filled with bone, the measured density is a mixture of the cortical bone density and the density of the surrounding tissue (Fig. 9). Therefore, the “measured” density is too low and the area too large. To avoid partial volume effect, the cortical thickness must be at least 2.5 times the voxel size. Rittweger suggested an algorithm to correct for the partial volume effect (34).

3.8. Analysis of Scan Data

Commence the analytical process by looking at the periosteal perimeter (PERI_C) or the total bone area (TOT_A). Both parameters describe the same feature of bone size. However, TOT_A, being an area measurement, shows a greater magnitude of change.

3.8.1. Analysis of Bone Size

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Voxel Size

Measured Density: 1200 mg/cm3 » 600 mg/cm3 0 mg/cm2 Fig. 9. Definition of partial volume effect.

Only an increase of 2% or greater in PERI_C or greater than 3% in TOT_A gives you an indication of a disease or drug-induced effect. Such changes are easily obtained during early skeletal growth up to the age of 6 months in rats. Anabolic agents, such as GH (9, 16, 17), vitamin D (18), and high-dose PTH, can induce them. In our example (Fig. 10a), none of the changes measured in PERI_C exceeded the 2% magnitude indicating that observed variations are within the precision of the measurement and therefore not real effects. 3.8.2. Analysis of Bone Mineral Content

Next, look at the total cross-sectional bone mineral content (TOT_CNT) which indicates whether there has been an absolute gain or loss in bone mass caused by the intervention. Together with the previous parameter (PERI_C), we can learn whether this increase is happening entirely at the endosteal envelope (i.e. in the absence of changes in PERI_C) or connected to an increase in bone size (increase in PERI_C). In our example, there was no change in bone size; therefore, TOT_CNT gives the same information as the volumetric cross-sectional BMD (TOT_DEN) displayed in Fig. 10b. Our “rule of thumb” tells us that the sham OP and the alendronate group are stable and not different from each other over time. However, the curve for the ethinylestradiol-treated rats drops off by more than three times the CVr found in Table 2 (>0.8%) indicating that treatment is not 100% effective in preventing bone loss. OVX rats show highly significant changes in TOT_ DEN already at 4 weeks and this parameter can already give meaningful information at a 2-week measurement point. In the absence of any change in bone size, we can now also conclude that bone loss must have occurred entirely on the endosteal envelope (cancellous or endocortical bone resorption). Our analytical process should, therefore, try to distinguish between these two possibilities.

a

change (%)

Periosteal Perimeter (PERI_C) 10 8 6 4 2 0 –2 –4 –6 –8 –10

SHAM OP OVX Ethinylestradiol Alendronate

0

4

8

12

weeks

b

change (%)

Volumetric Cross-Sectional BMD (TOT_DEN) 2 0 –2 –4 –6 –8 –10 –12 –14 –16 –18

SHAM OP OVX Ethinylestradiol Alendronate

0

4

8

12

weeks

c

Mean Cortical Thickness (CRT_THK) 10

change (%)

5 0 SHAM OP

–5

OVX

–10

Ethinylestradiol

–15

Alendronate

–20 –25 –30 0

4

8

12

weeks

d

Endocortical Perimeter (ENDO_C) 20

change (%)

15 SHAM OP OVX Ethinylestradiol Alendronate

10 5 0 –5 0

4

8

12

weeks

Fig. 10. Note that all values shown represent means plus SEMs. (a) Change in periosteal perimeter (% from baseline) after OVX. (b) Change in volumetric cross-sectional BMD (% from baseline) after OVX. (c) Change in mean cortical thickness (% from baseline) after OVX. (d) Change in endocortical perimeter (% from baseline) after OVX. (e) Change in volumetric cancellous BMD (% from baseline) after OVX. (f) Change in volumetric cortical BMD (% from baseline) after OVX.

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change (%)

e

Volumetric Cancellous BMD (TRAB_DEN) 5 0 –5 –10 –15

SHAM OP OVX Ethinylestradiol Alendronate

–20 –25 –30 –35 –40 0

4

8

12

weeks

f

Volumetric Cortical BMD (CRT_DEN)

change (%)

10

SHAM OP OVX Ethinylestradiol Alendronate

5

0

-5 0

4

8

12

weeks

Fig. 10. (continued)

3.8.3. Analysis of Changes in Cortical Bone

The best parameter available to address this question is to look for changes in mean cortical thickness (CRT_THK, Fig. 10c) and the threshold delineated endocortical perimeter (ENDO_C, Fig. 10d) or, alternatively, the cancellous bone area (TRAB_A). In our example, both the sham OP- and alendronate-treated animals are stable over time in these three parameters. The ethinylestradiol-treated animals show changes which just about exceed three times the CVr value for this parameter displayed in Table 2 (>2.6%). This indicates that some cortical thinning through endocortical bone resorption has occurred. Apparently, this resorptive process is very fast and significant cortical thinning is observed in vehicle-treated OVX rats as early as 4 weeks after the operation (Fig. 10c). A decrease in CRT_THK in the absence of changes in bone size (PERI_C) indicates that all of the changes in CRT_THK are the result of endocortical bone resorption. Treatments which are known to increase cortical thickness through endocortical bone apposition are PTH (2, 19, 20) and PGE2 (21, 22). The magnitude of these changes (decrease in ENDO_C) is large and can easily be monitored after 4 weeks or longer.

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3.8.4. Analysis of Changes in Trabecular Bone

Analysis of changes in trabecular bone density is best achieved by using the area-corrected volumetric cancellous BMD (TRAB_ DEN, Fig. 10e) as opposed to cancellous BMC (TRAB_CNT) (see Note 8). In the example shown, TRAB_DEN clearly shows that cancellous bone is lost at even a greater rate than is observed in cortical thinning (CRT_THK, Fig. 10c). Again, alendronate appears to be fully protective while cancellous bone loss in ethinylestradiol-treated animals exceeds the threefold CVr displayed in Table 2, indicating small but significant bone loss from this compartment. Some of the known agents acting to increase cancellous bone in rats are PTH (23, 24), FGF (25), PGE2 (21, 22, 26), vitamin D analogues (27), and GH.

3.8.5. Analysis of Volumetric Cortical Bone Density

Volumetric cortical BMD (CRT_DEN) (Fig. 10f) is the only parameter, which reflects an intrinsic material property of bone. In contrast to TOT_DEN and TRAB_DEN, which in fact are nothing but projected area density measurements similar to those obtained by DXA, CRT_DEN is a true volumetric measurement of density in the cortical compartment. Under most circumstances, this parameter changes very little over time but it may increase in long-term studies with bisphosphonates to indicate increased matrix mineralisation (secondary mineralisation). Conversely, this parameter may decrease at least transiently during long-term treatment with bone anabolic agents, such as PTH, since the newly formed endocortical bone is less densely mineralised and since secondary mineralisation takes some time to catch up (28). Volumetric CRT_DEN may also decrease in cases, where bone mineralisation is disturbed (vitamin D deficiency or osteogenesis imperfecta) (29) or in the presence of increased Haversian remodelling (only exceptionally in rodents). In our example, there is indeed little change over time in CRT_DEN, except for a trend to an increase in the alendronate and OVX group which, perhaps, could be indicative of increased matrix mineral content in bisphosphonate-treated animals.

3.8.6. Summarising the Results

The example outlined above suggests rapid cancellous and endocortical bone loss, the latter resulting in cortical thinning. Cortical thinning is not counteracted by activation of a compensatory periosteal bone formation drift in these skeletally mature animals, in contrast to what would be seen in younger rats (see Fig. 1). The magnitude of bone loss is greater in the cancellous than in the cortical bone compartment. None of the interventions has significant effects on matrix mineralisation as concluded from cortical BMD measurements, but there is at least a trend towards an increase in alendronate-treated rats. It may be worthwhile to follow up such a trend with more sophisticated methods with better discriminatory power for matrix mineralisation, such as backscattered electron imaging (30, 31).

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4. Notes 1. The XCT960A was the first commercially available dedicated animal pQCT scanner. It was replaced by the XCT Research SA and XCT Research M. Meanwhile, both machines are replaced by the XCT Research SA+ scanner which combines the characteristics of both machines using an adjustable collimation, and therefore a variable slice thickness. It provides a higher spatial resolution (70 μm) and a slice thickness of 0.7–0.13 mm to account for the size of bones of small animals, such as mice, rats, dogs, primates, and sheep (Table 1). The rat data presented in this chapter was measured on the XCT960A, whereas all mouse data were obtained using the XCT Research SA+. 2. Less data is available from mice to make clear recommendations with regard to the ideal age range for studies, but in our experience, an age of 4 months or older appears to be ideal for C57BL/6J mice. 3. Most interventions induce bone loss (OVX, immobilisation, inflammation) or bone gain (anabolic agents, like PTH, Sclerostin antibodies, GH, PGE2). The magnitude of the changes you are going to observe for agents who are unable to induce endochondral ossification (de novo trabecular bone formation) is for most cases proportional to the cancellous template available at the beginning of your experiment. In other words, animals with high trabecular BMD show a greater magnitude of bone gain under PTH treatment compared to severely osteopenic animals. Adjustment of groups to match for total cross-sectional BMD also assures that animals are well-matched in terms of biomechanical parameters. 4. Since the entire procedure of scanning normally requires less than 6 min, inhalational anaesthesia is ideal. Rats and mice can be kept under anaesthesia almost indefinitely in case multiple slices are being produced. Alternatively, anaesthesia can be induced by intraperitoneal injection of 40 mg/kg Ketarom (1.2 ml of ketamine hydrochloride + 0.8 ml of Rompun 2% + 8 ml of physiological NaCl solution). 5. The conical plug is available in three lengths and the right size is chosen by the operator depending on the size of the animal. This holder guarantees next to identical placement of the leg from one measurement to the next. 6. Some investigators prefer to define the cancellous bone area as a fixed value of 45% of the total bone area. This mode of analysis may have advantages in larger animals, such as dogs, primates, and men, where the cortical compartment is more difficult to separate from cancellous bone by threshold-based algorithms.

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For rodents, however, the threshold-based separation procedure gives much more accurate information on the changes in cortical thickness and endocortical perimeter. 7. The high threshold of 710 mg/cm3 really works only for rats 9 months of age and older and may fail even in this age group at later measurement points, if the intervention results in substantial cortical bone loss. If your cortical ring does not appear to be closed when running your chosen algorithm in the analytical part of the procedure, your threshold is too high and you have to lower it to avoid incorrect specification of cortical architecture. 8. In the example given, we are using a threshold-based algorithm to separate cancellous from cortical bone. The changes discussed in Subheading 3.8.3 show that the marrow cavity is expanding through a strong endocortical resorption process. This ongoing endocortical erosion lowers volumetric cortical density at the interface to such a degree that it falls below the chosen threshold of 690 mg/cm3 and as a consequence is counted as cancellous bone. Because of the constant increase in TRAB_A, just looking at cancellous BMC (TRAB_CNT) gives the misleading impression of an increase in cancellous bone mass even though rapid removal of cancellous bone structures is ongoing. References 1. Guglielmi, G., Glüer C. C., Majumdar, S., Blunt, B. A., and Genant, H.K. (1995) Current methods and advances in bone densitometry, Eur. Radiol. 5, 129–139. 2. Gasser. J. A. (1997) Quantitative assessment of bone mass and geometry by pQCT in rats in vivo and Site specificity of changes at different skeletal sites, J. Jpn. Soc. Bone Morphom. 7, 107–114. 3. Gasser, J. A. (1995) Assessing bone quantity by pQCT, Bone 17, S145–S154. 4. Ferretti, J. L., Capozza, R. F., and Zanchetta, J. R. (1995) Mechanical validation of a tomographic (pQCT) index for non-invasive estimation of rat femur bending strength, Bone 17, S145–S162. 5. Ferretti, J. L., Capozza, R. F., and Zanchetta, J. R. (1995) Mechanical validation of a noninvasive (pQCT) index of bending strength in rat femurs, Bone 18, 97–102. 6. Ferretti, J. L. (1997) Non-invasive assessment of bone architecture and biomechanical properties in animals and humans employing pQCT technology, J. Jpn. Soc. Bone Morphom. 7, 115–125.

7. Schneider, P. and Börner, W. (1991) Peripheral quantitative computed tomography for bone mineral measurements using a new special QCT-scanner: methodology, normal values, comparison with manifest osteoporosis, Fortschr. Röntgenstr. 154, 292–299. 8. Hermann, G. T. (1980) Image reconstruction from projections: the fundamentals of computerized tomography. Orlando: Academic Press. 9. Banu, M. J., Orhii, Pb., Mejia, W., McCarter, R. J. M., Mosekilde L., Thomsen, J. S., Kalu, D. N. (1999) Analysis of the effects of growth hormone, voluntary exercise and food restriction on diaphyseal bone in female F344 rats. Bone 25, 469–480. 10. Breen, S. A., Millest, A. J., Loveday, B. E., Johnstone, D. and Waterton, J. C. (1996) Regional analysis of bone mineral density in the distal femur and proximal tibia using peripheral computed tomography in the rat in vivo. Calcif. Tissue Int. 58, 449–453. 11. Beamer, W. G., Donahue, L. R., Rosen, C. J. and Baylink, D. J. (1996) Genetic variability in adult bone density among inbred strains of mice. Bone 18, 397–403.

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12. Graichen, H., Lochmüller, E. M., Wolf, E,, Langkabel, B., Stammberger, T., Haubner, M., Renner-Müller, I., Engelmeier, K. H., and Eckstein, F. (1998) A non-destructive technique for a 3-D microstructural phenotypic characterisation of bones in genetically altered mice: preliminary data in growth hormone transgenic animals and normal controls. Anat. Embryol. 199, 239–248. 13. Wronski, T. J., Dann, L. M., Scott, K. S.,and Cintron, M. (1989) Long-term effects of ovariectomy and aging on the rat skeleton. Calcif. Tissue. Int. 45, 360–366. 14. Yamazaki, I., and Yamaguchi, H. (1989) Characteristics of an ovariectomized osteopenic rat model. J. Bone Miner. Res. 4, 13–22 15. Kalu, D. N. (1991) The ovariectomized rat model of postmenopausal bone loss. Bone Miner. 15, 175–192. 16. Andreassen, T. T., Jorgensen, P. H., Flyvbjerg, A., Orskov, A. and Oxlund, H. (1995) Growth hormone stimulates bone formation and strength of cortical bone in aged rats. J. Bone Miner. Res. 10, 1057–1067. 17. Andreassen, T. T. and Oxlund, H. (2000) The influence of combined parathyroid hormone and growth hormone treatment on cortical bone in aged ovariectomized rats. J. Bone Miner. Res. 15, 2266–2275. 18. Weber, K., Goldberg, M., Stangassinger, M. and Erben, R. G. (2001) 1α-hydroxyvitamin D2 is less toxic but not bone selective relative to 1α-hydroxyvitamin D3 in ovariectomized rats. J. Bone Miner. Res. 16, 639–651. 19. Ejersted, C., Andreassen, T. T., Oxlund, H., Jorgensen, P. H., Bak, B,. Haggblad, J., Torring, O. and Nilsson, M. H. (1993) Human parathyroid hormone (1–34) and (1–84) increase the mechanical strength and thickness of cortical bone in rats. J. Bone Miner. Res. 8, 1097–1101. 20. Ejersted, C., Andreassen, T. T., Nilsson, M. H. and Oxlund, H. (1994) Human parathyroid hormone (1–34) increases bone formation and strength of cortical bone in aged rats. Eur. J. Endocrinol. 130, 201–207. 21. Jee, W. S. S., Mori, S., Li, X. J. and Chan, S. (1990) Prostaglandin E2 enhances cortical bone mass and actiavtes intracortical bone remodeling in intact and ovariectomized female rats. Bone 11, 253–266. 22. Jee, W. S. S., Ke, H. Z. and Li, X. J. (1991) Long-term anabolic effects of prostaglandinE2 on tibial diaphyseal bone in male rats. Bone Miner. 15, 33–55. 23. Gunness-Hey, M. and Hock, J. M. (1984) Increased trabecular bone mass in rats treated

24.

25.

26.

27.

28.

29.

30.

31.

32.

33.

34.

with synthetic parathyroid hormone. Metab. Bone & Rel. Dis. 5, 177–181. Gunness-Hey, M., and Hock, J. M. (1993) Anabolic effect of parathyroid hormone on cancellous and cortical bone histology. Bone 14, 277–281. Pun, S., Dearden, R. L., Ratkus, A. M., Liang, H., and Wronski, T. J. (2001) Decreased bone anabolic effect of basic fibroblast growth factor at fatty marrow sites in ovariectomized rats. Bone 28, 220–226. Mori, S., Jee, W. S. S., and Li. X, J. (1992) Production of new trabecular bone in osteopenic ovariectomized rats by prostaglandin E2. Calcif. Tissue Int. 50, 80–87. Erben, R. G., Bromm, S., Stangassinger, M. (1998) Therapeutic efficacy of 1α,25hydroxyvitamin D3 and calcium in osteopenic ovariectomized rats: Evidence for a direct anabolic effect of 1α,25-hydroxyvitamin D3 on bone. Endocrinol. 139, 4319–4328 Kneissel, M., Boyde, A., and Gasser, J. A. (2001) Bone tissue and its mineralization in aged estrogen-depleted rats after long-term intermittent treatment with parathyroid hormone (PTH) analog SDZ PTS 893 or human PTH(1–34), Bone 28, 237–250. Boyde A., Travers. R., Glorieux, F. H., and Jones, S. J. (1999) The mineralisation density of iliac crest bone from children with osteogenesis imperfecta. Calcif. Tissue Int. 64, 185–190. Boyde, A., Jones, S. J., Aerssens, J., and Dequeker, J. (1995) Mineral density quantification of the human cortical illiac crest by backscattered electron image analysis: Variations with age, sex, and degree of osteoarthritis. Bone 16, 619–627. Roschger, P., Plenk, H. Jr., Klaushofer, K. and Eschberger, J. (1995) A new scanning electron microscopy approach for the quantification of bone mineral distribution: Backscattered electron image grey levels correlated to calcium K alpha-line intensities. Scan. Microsc. 9, 75–88. Rauch, F., and Schönau, E. (2001) Changes in Bone Density During Childhood and Adolescence: An Approach Based on Bone’s Biological Organisation. J. Bone Miner. Res. 16, 597–604 Ferretti, J. L., Capozza, R. F., and Zanchetta, J. R. (1996) Mechanical Validation of a Tomographic (pQCT) Index for Noninvasive Estimation of Bending Strength of Rat Femurs. Bone 18, 97–102. Rittweger, J., Michaelis, I., Giehl, M., Wüsecke, P., and Felsenberg, D. (2004) Adjusting for the Partial Volume Effect in Cortical Bone analyses of pQCT. J. Musculoskel. Neuron. Interact. 4, 436–441.

Chapter 29 Quantitative X-ray Imaging of Rodent Bone by Faxitron J.H. Duncan Bassett, Anne van der Spek, Apostolos Gogakos, and Graham R. Williams Abstract This chapter describes the use of digital micro-radiography with the Faxitron machine as a means of imaging and quantitating bone mineral content in mice and rats. Key words: Bone mineral content, Digital X-ray analysis, Faxitron, ImageJ, Mouse, Skeletal phenotype analysis

1. Introduction Over the past 10 years there has been an explosion of interest in skeletal phenotyping of different mouse strains, driven in part by initiatives such as ENU Mutagenesis programmes and the systematic generation of genetically modified mice with targeted inactivation of all genes in the genome. Each of these strains needs to be screened for phenotype abnormalities across a range of physiological systems, including the musculoskeletal system. Detailed characterization of skeletal phenotypes in a high-throughput manner using techniques such as MicroCT and quantitative CT represents a specific challenge. Developmental abnormalities may occur early in utero and during neonatal life (skeletal dysplasias) or in the postnatal period (linear growth and endocrine disorders), and such abnormalities impact on bone mass and mineralization in adulthood. By contrast, phenotypes such as osteoporosis do not manifest until adulthood and may display sexual dimorphism. Thus, analyses need to be performed separately in both sexes and at several ages if abnormalities are to be captured efficiently. To address this issue, we have developed and refined a cost-effective, rapid, and highly sensitive method for the quantitative assessment of bone mineral content from fixed post-mortem specimen using 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_29, © Springer Science+Business Media, LLC 2012

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Faxitron point projection digital micro-radiography machine. In addition to quantitating bone mineral content, the Faxitron can also be used to obtain morphological information and direct measurements of anthropomorphic parameters from digital X-ray images.

2. Materials 2.1. General Reagents/ Materials

1. Dissecting instruments. 2. 20-ml Polystyrene tubes. 3. 70% Ethanol solution.

2.2. Equipment and Computer Software

1. Faxitron MX20 variable kV point projection X-ray source and digital image system (Qados, Cross Technologies plc, Berkshire, UK) (see Note 1). 2. Calibration standards – 1-mm steel wire; 1-mm diameter spectrographically pure aluminium wire [Hollinbrow Precision Products (UK) Ltd., Shropshire, UK]; 1-mm diameter polyester fibre. 3. ImageJ 1.43n software (available free of charge at http://rsb. info.nih.gov/ij/). 4. Adobe Photoshop 7.0.1 or later software (Adobe Systems Inc., San Jose, CA, USA).

3. Methods 3.1. Dissection and Preparation of Skeletal Elements

Bones can be imaged from mice at any age from 4 weeks onwards. For initial screening we usually analyse an upper limb, a lower limb, and the proximal six tail vertebrae following careful dissection post-mortem as described below: 1. Sacrifice the animal using an approved method. 2. Carefully remove the skin from the carcass. 3. To remove the upper limb, cut under the scapula with a scalpel blade to detach from the thorax and cut through the clavicle as close to the sternum as possible. 4. To remove the lower limb, cut vertically medial to the pelvis immediately lateral to the vertebrae thereby avoiding the femoral head. 5. To remove the proximal six tail vertebrae, peel off the skin, cut the tail at the base. Count down inter-vertebral spaces and detach the six most proximal vertebrae.

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6. Place the intact limbs and tail vertebrae into 70% ethanol in a 20-ml polystyrene tube and fix at 4°C for at least 48 h. 7. After fixation, carefully dissect the muscle and soft tissues away from the bones of the upper and lower limbs (see Note 2). Carefully detach the humerus from the scapula and the femur from the acetabulum to leave the intact limb bone exposed. 8. Place the dissected bones in a polystyrene tube containing fresh 70% ethanol and store at 4°C until further use (see Note 3). 3.2. Faxitron Digital X-ray Imaging of Excised Bones

1. Switch on Faxitron machine for at least 15 min prior to use (see Note 4). 2. Remove the polypropylene sample stage prior to the calibration step. 3. Calibrate the Faxitron by running four flat dark field images using time set at 15 s and voltage at 26 kV and save the calibration settings (see Note 5). 4. When you are ready to image the bones, remove them from the 70% ethanol solution, place them on a paper towel to remove the excess ethanol and arrange them, within the imaging area of the polypropylene sample tray along with the steel, aluminium, and polyester calibration standards (see Note 6). 5. Insert sample tray into the ×5 magnification uppermost slot in the Faxitron machine and close the door. 6. Ensure that the Auto Level, Auto Exposure Control, Contrast Assist and Sharpen Assist toggles are deselected. Then select the “start exposure” button to commence imaging of the samples. The image will display on the computer screen. 7. Check that the orientation of samples and standards is correct taking care to make sure that all elements are included in the image. 8. Should any adjustment be necessary, remove the polypropylene sample stage, reposition the samples as necessary, replace the sample stage, and repeat step 6. 9. When you are happy with the image save it as a DICOM file, which is the default format. 10. Switch the Faxitron off using the key when you have completed the analysis session.

3.3. Image Processing

1. Open DICOM image in ImageJ and press “h” to reveal a histogram showing the distribution of grey scale pixels within the image (Fig. 1a). 2. Record the minimum and maximum grey level values. 3. Using the Rectangular Selections Tool within ImageJ, select as large an area as possible of the polyester standard without

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Fig. 1. (a) Original DICOM image showing the recommended organisation of limbs and vertebrae alongside polyester (left ), aluminium (right ), and steel (bottom) calibration standards. The histogram on the right shows the grey scale distribution and location of the three standards relative to mineralised tissue. The large peak on the left represents the background. (b) Pseudo-colour 8-bit TIFF image following stretch processing. The histogram shows the stretched grey scale distribution in relation to the 16-colour bins.

including background and press “h”. Note the value of the modal grey level. Similarly, select as large an area as possible of the steel standard and press “h”, noting the value of its modal grey level. This procedure is not applied to the aluminium standard, which acts as an internal reference for the grey scale stretch procedure described below. 4. Using ImageJ, stretch each of the 2,368 × 2,340 DICOM images (16,383 grey levels) to the modal grey level values of the plastic and steel standards noted above. This stretched image is inverted using ImageJ and then converted to an 8-bit TIFF file (256 grey levels). In the resultant image, the plastic and steel standards are thus assigned grey levels of 0 and 255, respectively.

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3.5. Generating a Montage of Pseudocoloured Images of All Bones

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1. Open the stretched 8-bit grey scale TIFF image in ImageJ. 2. Select the “Image” menu, then select Lookup Tables, 16 Colours (see Note 7). 3. Save the image as a TIFF file and repeat the procedure for all images. To compare images from different groups of mice (e.g. wild-type and mutant), it is desirable to display them side by side in a single montage. This can be achieved using Adobe Photoshop as described below. 1. Open the TIFF files of all pseudo-colour images of the bones you wish to analyse in Adobe Photoshop. 2. Select the first image and adjust the canvas size so it is big enough to accommodate all the individual colour images in two rows – wild-type above mutant. 3. Copy and paste each colour image into the enlarged canvas and position appropriately. 4. Save the montage as a TIFF file with selection of the following settings: Image Compression: NONE; Byte Order: Macintosh.

3.6. Processing and Cleaning Montages for Analysis of Mineralisation

1. Open the montage TIFF file in Adobe Photoshop. 2. Select the Brush Tool and adjust brush size to approximately 33 pixel diameter, select black for the brush colour. 3. Zoom in to at least 200% and use the Brush Tool to black out any soft tissue beyond the edge of the bone being careful not to black out any of the bone itself (see Note 8). 4. Select the Magic Wand Tool and click adjacent to the first bone. A yellow line should now appear around the bone and if the edges are not clearly highlighted brush over to black out any soft tissue remaining. 5. Repeat this process for each bone being analysed (see Note 9). 6. Save the processed and cleaned montage as a new TIFF file. 7. Repeat steps 1–6 to give separate TIFF files containing montages of the bones to be analysed.

3.7. Determination of Relative Mineral Content

1. Open the cleaned, montage file in the ImageJ programme (Fig. 2a). 2. Under Plugins, select Macro, CustomHistogram.txt (see Note 10).

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3. Select all bones to be analysed using the Wand (tracing) tool whilst holding down the Shift key.

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Fig. 2. (a) Black and white representation of pseudo-colour 8-bit TIFF files containing cleaned femur montages. (b) Relative frequency distribution plot showing difference in grey scale distributions between WT and mutant femurs. (c) Cumulative frequency plot showing difference between WT and mutant femurs.

4. Under Plugins, select Macro followed by CustomHistogram. Select the following values in the dialog box: X Min: 0, X Max: 255, Y Max: auto, Bins: 16 and press OK. 5. Select the “List” button in the displayed histogram and copy the “bin start” and “count” columns and paste into an Excel spreadsheet. This provides the number of pixels in each of 16 equal grey level divisions (bins) to give a frequency distribution

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of grey level values for the whole montage. The wild-type and mutant frequency distributions thus comprise summed grey scale values from each bone included in the respective montages and therefore facilitate comparison of their relative grey level distributions (Fig. 2b). 6. Statistical analysis of the difference between the cumulative frequency distribution of the grey levels of wild-type versus mutant montages (Fig. 2c) can be performed using a Kolmogorov–Smirnov test on each of the grey scale images to determine if these differ from one another (1).

4. Notes 1. The radiation source in the Faxitron is shielded by lead lining. Inadvertent radiation exposure is prevented by a trip switch locking mechanism that is activated when the X-ray source is switched on. It is essential that these safety features are tested at installation and checks are included as part of a maintenance contract. 2. Muscle and soft tissues should not be removed at the time of dissection. Complete removal is much easier and results in less risk of damage to bone when this is done after fixation in 70% ethanol for at least 48 h. 3. Samples can be stored in 70% ethanol at 4°C for many months and can be re-used for additional analyses including micro-CT imaging, back-scattered electron scanning electron microscopy, and investigation of biomechanical properties by destructive three-point bend and compression testing. 4. The X-ray source needs to warm up in order to maintain stability. 5. The Faxitron requires to be recalibrated each time it is restarted. 6. Samples should be removed from the 70% ethanol solution for as short a period of time as possible to prevent drying out. 7. This procedure applies a pseudo-colour scheme in which grey levels are divided into 16 equal intervals each represented by a different colour (Fig. 1b), thus greatly aiding visual presentation of digital images. 8. If in doubt regarding the identity of bone or soft tissue, it is better to erase a small bit of bone rather than leaving areas of soft tissue because the markedly lower grey levels of soft tissue would introduce artefacts and skew the mineralisation data. By contrast, erasing a bit of bone is readily compensated and has

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little if any overall effect. Nevertheless, if a mistake is made using the Brush Tool it can be rectified using CTRL + Z. 9. A slightly different method is required to erase soft tissue from the vertebral images. Using the Pencil Tool in Adobe Photoshop, select a diameter of 13 pixels with 100% hardness. Zoom to 300% and draw a line along the outside of individual vertebrae. For the inexperienced eye, it can be difficult to distinguish soft tissue and ligament from bone and training from an experienced operator is necessary. Select the Paint Bucket Tool and fill in the background outside the vertebrae with black. 10. If the CustomHistogram.txt macro is absent from the Plugin menu, it can be downloaded from the ImageJ website (http:// rsb.info.nih.gov/ij/). Reference 1. Demidenko E. Kolmogorov–Smirnov Test for Image Comparison. In: Analysis in Computer Science and its Applications, Lagana A. (Ed.),

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Chapter 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 Abstract Optical imaging is a valuable technique for visualizing and quantifying biological processes in living organisms. Optical imaging can be divided into two main imaging modalities: bioluminescence imaging and fluorescence imaging. This chapter describes the use of these imaging techniques to image tumour cells in mouse models of cancer and to detect early bone metastasis. Key words: Fluorescent imaging, Bioluminescent imaging, Tumour cells, Bone metastasis

1. Introduction 1.1. Optical Imaging

In recent years, optical imaging has emerged as a valuable technique for visualizing and quantifying biological processes in living organisms. The technique involves targeted expression of reporter genes such as luciferase derived from the firefly Photinus pyralis (fLuc) or green fluorescent protein (GFP) derived from the jellyfish Aequorea victoria within cells of interest. Using the light emitted from these reporters, biological processes such as tumour host interactions can be monitored. There have been major technological advances in methods of photon detection due to the development of cooled charged coupled device (CCD) cameras, Peltier cooled detectors, and microplate intensifiers which allow researchers to capture photons emitted from weak sources deep within the tissues of small animals. Optical imaging can be divided into two main imaging modalities: bioluminescence imaging (BLI) and fluorescence imaging (FLI). Both can be used to study cell and tissue-specific promoters and to follow trafficking, differentiation, and the fate of reporter gene-expressing cells, or biological processes such as

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apoptosis, protein–protein interactions, angiogenesis, proteolysis, and gene-transfer (1–3). Moreover, optical imaging and optical reporter systems are very cost-effective, time-efficient, and are particularly well suited for use in small animals as well as for in vitro assays. 1.2. Fluorescence Imaging

The technique of FLI makes use of a fluorescent compound (fluorophore) typically a fluorescent protein or dye, which can be excited by an external light source of a certain wavelength and which emits photons of a specific longer wavelength upon relaxation. The emitted photons are captured for imaging and/or quantification. An important disadvantage of FLI is the fact that it is difficult to detect light emitted from cells that lie deep within tissues since both the excitation light and the emitted light have to pass through the animal. The signal decreases exponentially further the source the fluorescent signal is from the surface of the animal. Another limitation of FLI is the fact that there is a relatively high level of signal-to-noise ratio due to the generation of bright background signals arising from the animal’s intrinsic autofluorescence (4, 5). In order to combat these limitations, however, new classes of red-shifted fluorescent proteins, near-infrared dyes and quantum dots have been developed that provide better deep tissue imaging characteristics (6, 7).

1.3. Bioluminescence Imaging

BLI is based on the detection of light produced by an enzymatic reaction in which a substrate, such as luciferin or coelerentarazin, is oxidized by luciferase which can be expressed by cells in vitro or in vivo. Until recently, BLI was the most commonly used technique for whole body optical imaging due to several important advantages of using firefly luciferase (fLuc) as a reporter over other optical reporters (8). First, there is no background bioluminescent activity, making BLI a highly specific and sensitive imaging modality. Second, the fLuc protein has a relatively short half-life of about 1–1.5 h which makes it ideally suited to kinetic and dynamic analyses of gene expression within short time frames. This provides researchers with a tool to examine circadian and infradian rhythms of gene expression. Although the potential applications of BLI are very wide, in this chapter we focus on its application to the study of skeletal metastases.

1.4. Models of Skeletal Metastases

Animal models of bone metastasis have been useful in the identification of genes that regulate susceptibility to the development and progression of metastasis which in turn represent novel targets for drug development (9–11). In vivo bioluminescent bone metastasis can be induced by intra-cardiac; intra-osseous, or arterial tail vein injection of fLuc positive tumour cells. Direct injection of tumour cells into the systemic circulation of an animal leads to the development of distant metastases throughout the body and the standard way to achieve this is by injecting the cells into the left ventricle of the heart (12). Shortly after intra-cardiac injection of

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Fig. 1. BLI of luciferase expressing tumour cells after intra-osseous injection in vivo. Luciferase-expressing breast cancer cells (MDA-MB231-luc) were transplanted (100,000 cells) into the bone marrow cavity of the femur of nude mice. Tumour progression was followed using bioluminescence at 1 (a), 2 (b), and 3 (c) weeks, respectively (adapted with permission from Kaijzel et al. (1), Fig. 3).

luciferase-expressing human MDA-MB231-Luc breast cancer cells, large numbers of these photon-emitting tumour cells are detectible in the circulation. After a few days, however, only a few light emitting spots can be identified in the skeleton, mimicking micrometastatic cell spread (13). After another couple of days or weeks the size of these bone metastatic tumours increases leading to massive bone destruction and eventual death of the animal. To study local tumour growth and the early process involved in bone metastasis, cancer cells can also be injected directly into the marrow space of long bones (13, 14). Quantification of the bioluminescent signal localized over the site of implantation at different time points allows the researcher to monitor tumour growth in vivo (Fig. 1). This model allows regular monitoring of the development and progression of experimental bone metastases in living animals at a stage which largely preceded osteolysis, with high sensitivity (13, 15).

2. Materials 1. Balb/c Nu/Nu athymic mice (see Note 1). 2. MDA-231 cells engineered to express luciferase (MDA-231-B/ Luc+) (see Note 2). 3. Tissue culture facilities.

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4. Sterile phosphate-buffered saline (PBS). 5. Selection of syringes and needles. 6. Sterile surgical instruments. 7. Sutures. 8. Isoflurane gas anaesthesia system. 9. IVIS Spectrum −90°C cooled CCD camera system with imaging software (Caliper Life Sciences, Inc. Hopkinton, USA) (see Note 3). 10. D-Luciferin sodium salt (Synchem OHG, Germany).

3. Methods 3.1. Induction of Bone Metastases by Intra-cardiac Injection of Cancer Cells

1. Harvest the MDA-231-B/Luc+ cells, pellet and wash several times in sterile PBS (see Note 4). 2. Prepare a suspension of MDA-231-B/Luc+ cells at a concentration of 1 × 106 cells/ml in PBS, by aspirating the cells up and down through a 25-G needle. Use a microscope to make sure that there are no clumps in the single cell suspension (see Note 5). 3. Anaesthetize the mouse using isoflurane gas anaesthesia (see Note 6). 4. Fix the anaesthetized mouse in a supine position on a sterile surface with the head of the animal in a nozzle which supplies isoflurane at a maintenance dose of 2% (see Note 6). 5. Aspirate the cell suspension into a syringe, through a 26-G needle, making sure that no air bubbles are present. 6. Carefully insert the needle through the diaphragm approximately 3 mm to the left of the sternum and aim centrally towards the heart (Fig. 2). 7. Advance the needle into the left ventricle making sure that it is correctly positioned (see Note 7). 8. Slowly inject 100 ml of the cell suspension into the left ventricle over a period of 20–40 s. 9. Monitor the development and progression of metastases weekly by BLI (see Subheading 3.3).

3.2. Induction of Bone Metastases by Direct Intra-Osseous Inocculation of Cancer Cells

1. Prepare a single cell suspension of MDA-231-B/Luc+ cells at a concentration of 1.5 × 107 cells/ml in PBS as described in Subheading 3.1, steps 1 and 2. 2. Anaesthetize the mouse using 5% isoflurane gas anaesthesia (see Note 6).

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Fig. 2. Intra-cardiac injection of cancer cells. (a) The skin is pulled back and stretched tightly over the mouse using the thumb and index finger. (b) The needle is carefully inserted through the diaphragm approximately 3 mm to the left of the sternum and aimed centrally and upwards. The needle is advanced into the left ventricle. A small volume is aspired into the syringe, to make sure that it is correctly positioned. The needle is in the left ventricle if bright red, oxygenated blood flows into the needle. (c) Now, 100 ml of the cell suspension can be injected into the left ventricle over a period of 20–40 s.

3. Fix the anaesthetized mouse in a supine position on a sterile surface with the head of the animal in a nozzle which supplies isoflurane at a maintenance dose of 2% (see Note 6). 4. Aspirate the cell suspension into a syringe, through a 30-G needle, making sure that no air bubbles are present. 5. Make a small incision in the right hind limb to access the tibia (Fig. 3a). 6. Make two holes, 4–5 mm apart, in the cortex of the tibia with a 25-G needle (Fig. 3a). 7. Create space in the bone by flushing out the bone marrow with PBS from the proximal end of the tibial shaft using a 30-G needle. The PBS coming out of the distal hole should be clear. Typically, this is the case after ~50 ml PBS has been flushed through the bone (Fig. 3b). 8. Inject 10 ml into the right tibia using a 30-G needle again from the proximal end of the shaft (Fig. 3b) (see Note 8).

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Fig. 3. Intra-osseous inoculation of cancer cells. (a) A small incision is made in the hind limb of the animal and two holes are drilled 4–5 mm apart from each other using a 25-G needle. (b) Through the hole at the proximal end, the bone marrow is flushed out with PBS and 10 ml of the cell suspension is injected. (c) The incision is closed with a suture.

9. Close the skin incision with a suture (Fig. 3c). 10. Monitor the development and progression of metastases twice a week by BLI. 3.3. Bioluminescence Imaging Using the IVIS Spectrum System

Throughout the entire procedure, mice should be observed for any signs of distress or changes in vitality. 1. Initialize the IVIS Spectrum imaging system. 2. Prepare a solution of sodium luciferin at a concentration of 166 mg/ml in sterile PBS. 3. Inject the animal intraperitoneally with 30 ml of the luciferin solution, this results in a dose of 250 mg/kg for a mouse of ~20 g. 4. Allow the luciferin to distribute through the tissues in for between 5 and 15 min (see Note 9). 5. Anaesthetize the mouse using isoflurane gas anaesthesia as described in Subheading 3.1, steps 3 and 4. 6. Select the field of view depending in the number of animals that will be imaged. 7. When fully anaesthetized, place the animal or animals in a supine position in the imaging chamber on the 37°C movable imaging stage with integrated anaesthesia nozzles constantly supplying 2% isoflurane (see Note 6).

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8. Start the image recording sequence (see Note 10). 9. Turn the mice from a ventral to a dorsal position and repeat the image recording. 10. Return the mice to their cages where they should recover consciousness quickly. 3.4. Quantification of the Bioluminescent Signal

1. Acquire the images as described in Subheading 3.3. 2. Superimpose the bioluminescent data on a grayscale photo of the animal allowing the relative light intensity to be visualized by pseudo-colours (see Note 11). 3. Select the regions of interest to be analyzed (see Note 12). 4. Express the measured values of light emission within the ROI as relative light units (RLU) or as photons per cm2 per second per steradian.

4. Notes 1. It is possible to conduct these studies in normal mice if the tumour cells for inoculation are syngenic to the animal model being used. However, immuno-deficient mice such as Balb/c Nu/Nu athymic nude mice must be used if the injected cells are not syngenic to prevent their destruction by the host immune system. 2. The protocol described here has been optimized for MDA-MD231 Luc+ cells. The protocol described may need to be adapted and optimized if different cell lines are used. 3. Other suitable imaging systems are commercially available, such as the Photon Imager (Biospace Lab, Paris, France) or the NightOWL II LB 983 (Berthold Technologies GmbH & Co. KG, Bad Wildbad, Germany). 4. It is important that the cells are free of culture medium since this can trigger an allergic reaction in the animal. 5. It is of utmost importance that the single cell suspension does not contain any cell clumps or aggregates since these are potentially harmful to the animals. 6. Gas anaesthesia is given in a constant oxygen flow of 0.8 l/min. 5% Isoflurane is used for induction of the anaesthesia, and 2% isoflurane for maintenance. 7. If the needle is correctly positioned you will observe spontaneous and continuous entrance of pulsating, bright red oxygenated blood into the transparent needle hub during the entire procedure.

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8. When inserting the cells into the bone marrow care should be taken to ensure that no cells are deposited on the bone surface or subcutaneous tissues since these will have different growth characteristics to those injected into the marrow space. 9. The luciferase signal is optimal 10–15 min after injection with luciferin (16). It is very important that all the mice within one experiment are injected with luciferin at exactly the same time before the imaging is carried out. 10. When activating the “acquire” button on screen in the “Living Image” software, grayscale images of the mice are recorded first with dimmed light. Photon emission is then integrated throughout a period of 1–5 min and recorded as pseudo-colour images superimposed over the grayscale white light photo. 11. Pseudo-colour maps can be adjusted by changing the scale and threshold values, this is purely graphic and has no effect on the actual measurements. 12. The ROI can either be manually selected or can be automatically selected by the imaging software. References 1. Kaijzel, E. L., van der Pluijm, G., and Löwik, C. W. (2007) Whole-body optical imaging in animal models to assess cancer development and progression. Clin. Cancer Res. 13, 3490–7. 2. Massoud, T. F., Gambhir, S. S. (2003) Molecular imaging in living subjects: seeing fundamental biological processes in a new light. Genes Dev. 17, 545–80. 3. Weissleder, R., Tung, C. H., Mahmood, U., and Bogdanov, A., Jr. (2006) In vivo imaging of tumors with protease-activated near-infrared fluorescent probes. Nat. Biotechnol. 17, 375–8. 4. Ntziachristos, V. (2006) Fluorescence molecular imaging. Annu. Rev. Biomed. Eng. 8, 1–33. 5. Lim, Y. T., Kim, S., Nakayama, A., Stott, N. E., Bawendi, M. G., and Frangioni, J. V. (2003) Selection of quantum dot wavelengths for biomedical assays and imaging. Mol. Imaging 2, 50–64. 6. Giepmans, B. N., Adams, S. R., Ellisman, M. H., and Tsien, R. Y. (2006) The fluorescent toolbox for assessing protein location and function. Science 312, 217–24. 7. Shcherbo, D., Merzlyak, E. M., Chepurnykh, T. V., Fradkov, A. F., Ermakova, G. V., Solovieva, E. A., Lukyanov, K. A., Bogdanova, E. A., Zaraisky, A. G., Lukyanov, S., and Chudakov, D. M. (2007) Bright far-red

8.

9.

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11.

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fluorescent protein for whole-body imaging. Nat. Methods 4, 741–6. Contag, C. H., and Bachmann, M. H. Advances in in vivo bioluminescence imaging of gene expression. Annu. Rev. Biomed. Eng. 2002; 4:235–60. Eccles, S. A., and Welch, D. R. (2007) Metastasis: recent discoveries and novel treatment strategies. Lancet 369, 1742–57. Sharpless, N. E, and Depinho, R. A. (2006) The mighty mouse: genetically engineered mouse models in cancer drug development. Nat. Rev. Drug Discov. 5, 741–54. Steeg, P. S. (2006) Tumor metastasis: mechanistic insights and clinical challenges. Nat. Med. 12, 895–904. Arguello, F., Baggs, R. B., and Frantz, C. N. (1988) A murine model of experimental metastasis to bone and bone marrow. Cancer Res. 48, 6876–81. Wetterwald, A., van der Pluijm, G., Que, I., Sijmons, B., Buijs, J., Karperien, M., Löwik C.W., Gautschi, E., Thalmann, G. N., and Cecchini, M. G. (2002) Optical imaging of cancer metastasis to bone marrow: a mouse model of minimal residual disease. Am. J. Pathol. 160, 1143–53. van der Pluijm, G., Sijmons, B., Vloedgraven, H., Deckers, M., Papapoulos, S., and Löwik, C. (2001) Monitoring metastatic behavior of

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human tumor cells in mice with species-specific polymerase chain reaction: elevated expression of angiogenesis and bone resorption stimulators by breast cancer in bone metastases. J. Bone Miner. Res. 16, 1077–91. 15. Kalikin, L. M., Schneider, A., Thakur, M. A., Fridman, Y., Griffin, L. B., Dunn, R. L., Rosol, T. J., Shah, R. B., Rehemtulla, A., McCauley, L.K., Pienta, K. J. (2003) In vivo visualization of

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metastatic prostate cancer and quantitation of disease progression in immunocompromised mice. Cancer Biol. Ther. 2, 656–60. 16. Paroo, Z., Bollinger, R. A., Braasch, D. A., Richer, E., Corey, D. R., Antich, P. P., Mason, R.P. (2004) Validating bioluminescence imaging as a high-throughput, quantitative modality for assessing tumor burden. Mol. Imaging 3, 117–24.

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Chapter 31 Fourier Transform Infrared Imaging of Bone Eleftherios P. Paschalis Abstract Fourier transform infrared imaging (FTIRI) is a technique that can be used to analyze the material properties of bone using tissue sections. In this chapter I describe the basic principles of FTIR and the methods for capturing and analyzing FTIR images in bone sections. Key words: Fourier transform infrared imaging, Bone, FTIR, Mineral, Matrix

1. Introduction Fourier transform infrared imaging (FTIRI) analysis of bone as originally described by Marcott et al. (1) is a vibrational spectroscopy imaging technique that can be used to analyze the material properties of bone in thin bone tissue sections, with a spatial resolution of about 6.3 μm. By combining a Focal Plane Array (FPA) Mercury Cadmium Telluride (MCT) detector with an FTIR microscope, regions of bone approximately 400 μm2 may be analyzed. It can be combined with bone histomorphometry (2), small angle X-ray scattering (SAXS) analysis, (3) and quantitative backscattered electron imaging (qBEI) (4) to better characterize the material properties of bone in tissue sections. To date, FTIR analysis of bone has been successfully applied to the analysis of normal bone and diseased bone as far as material properties are concerned, as well as in the evaluation of the effects of various therapeutic agents on the material properties of bone (5–8). The variables that are most commonly analyzed by FTIRI are as follows: 1. The mineral to matrix ratio in which the amount of mineral per amount of organic matrix per volume analyzed can be measured (8).

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2. The amount and type of carbonate substitution in the bone mineral apatite lattice (9, 10) (see Note 1). 3. The mineral maturity (chemistry) and crystallinity (size and shape of crystallites) of the bone mineral apatitic crystallites. 4. The ratio of two of the major cross-links in bone (pyridinoline/and dehydrohydroxylysinonorleucine) (8). All of these variables can be reported either as an average over the whole bone section, or within a discreet anatomical region within that section (8) (see Note 2).

2. Materials 2.1. Equipment and Software

1. A Fourier transform infrared (FTIR) spectrometer, equipped with a FPA MCT detector with associated software for image analysis and processing (see Note 3). 2. Plotting software capable of handling matrices of data and reconstructing 2D or 3D images from them (e.g. OriginLab or Sigmaplot).

2.2. Tissue Sections

For FTIRI analysis, thin bone tissue sections are required of between 0.5 and 5 μm in thickness, depending on the species the bone tissue was excised from (see Note 4). Sections must be fixed prior to analysis and embedded in a hard supportive material. Suitable fixatives include 100% ethanol, 70% ethanol, glycerol, formaldehyde, electron microscopy (EM) fixative, and formalin in cacodylate or phosphate-buffered saline (see Note 5). Most of the embedding media have chemical components that spectrally overlap the components of mineralized tissues, therefore it is of utmost importance to select optimal fixation and embedding protocols in order to minimize artifactual effects on mineral and matrix FTIRI spectra. Embedding media such as Araldite, Epon, JB4, LR White, PMMA, and Spurr have all been used for FTIR analysis but comparative studies indicate that embedding in LR White, Spurr, Araldite, and PMMA had the least effect on the spectral parameters most often measured by FTIRI analysis (11).

3. Methods 3.1. Preparation of Section

Once an appropriate section has been obtained, place it between two BaF2 windows in order to keep the section as flat as possible, so that no spectral artefacts are introduced.

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3.2. Decide upon Which Type of Analysis is Required

Two distinct types of FTIRI analysis may be performed; one can either extract individual FTIR spectra and calculate the parameter sought through curve fitting of the spectral band of interest and another that can process all 4,096 acquired spectra at once to reconstruct an image depicting the spatial distribution of the parameter(s) under investigation. The typical spectra of PMMA embedded human bone (in the area of primary mineralized trabeculae), pure PMMA, and pure type I mineralizing collagen are shown in Fig. 1. The peaks referred to in the following discussion are appropriately labelled.

3.3. Select the Area to be Analyzed

Selecting the area to be analyzed depends on the question you want to address. Although FTIRI results can be averaged out across a whole specimen, it is possible to analyze specific areas within the bone sample and relate the spectra obtained for factors such as the degree of mineralization, or levels of bone turnover. In the former case, areas can be selected on the basis of qBEI images (12) (thus enabling the selection of either primary or secondary mineralized areas), while in the latter areas may be selected based

Fig. 1. Examples of FTIR spectra. Typical FTIR spectra of PMMA (a) (commonly used embedding medium for hard tissues), human trabecular bone in a primary mineralized area (b), and pure type I mineralizing collagen (c). As is evident, despite the fact that they are distinct, subtle overlaps occur in the spectral bands of interest (Amide I, v3 CO32−, and v1v3 PO43−), which make the subtraction of PMMA spectral contributions essential.

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on parallel histologically stained sections (4, 8, 13, 14). This can allow one to correlate spectroscopically determined parameters with rates of bone remodelling. An advantage of conducting FTIRI in relation to bone remodelling is that it allows one to examine differences between normal and diseased bone independent of bone turnover (4, 13–21). 3.4. Acquisition of Spectral Images

1. Ensure that the spectrometer is continuously powered to minimize warm-up instabilities and has been purged with dry-air or purified nitrogen gas to minimize the water vapour and atmospheric CO2 spectral interference. 2. Collect background spectral images under identical conditions from the same BaF2 windows at the beginning and end of each experiment to ensure instrument stability. 3. When specific regions of the section are being analyzed ensure that data from a minimum of three areas per specimen (each 400 μm × 400 μm) are collected so that the results can be analyzed statistically.

3.5. Analysis of Individual Spectra

Individual FTIR spectra may be extracted from the images at specific operator-chosen pixels and then baseline-corrected using a rubber-band function (a spectral procedure involving the correction of the convex envelope lying below a spectrum; instead of a linear, a polynomial function is utilized (22)).

3.5.1. Analysis of Mineral to Matrix Ratios

These can be calculated by integrating the area under the peaks at ~900–1,200 cm–1 (v1v3 phosphate band, representative of the phosphate functional groups encountered in the apatite bone mineral crystals) and ~1,585–1,725 cm–1 (Amide I, representative of the carbonyl functional groups present in organic moieties) (9).

3.5.2. Analysis of Carbonate/Phosphate Ratio

The relative amounts of carbonate substitution in the bone mineral apatite crystal lattice can be calculated from the ratio of the integrated areas of the respective raw (non-curve-fitted) peaks. In order to deduce the type of carbonate substitution, the v2CO32− spectral region is curve-fitted using a commercially available program (such as GRAMS 32, Galactic Industries Corporation, Salem, NH, USA), with the input of three sub-bands at 866, 871, and 878 cm–1, which have been assigned in the literature as indicative of “unstable – labile,” Type B, and Type A carbonate-substituted apatites, respectively (9). Type A represents substitution of the OH− group in the apatite crystal lattice by CO32−, Type B to the substitution of the PO43− group, and “labile” to surface and/or “loosely” incorporated CO32− groups. The output of this analysis is typically expressed as peak position (expressing the type of substitution) and relative percent area (indicative of the amount of the particular carbonate species).

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3.5.3. Mineral Maturity and Crystallinity

This can be calculated by analysis of the v1v3 phosphate band (Fig. 1). This broad band has minor underlying contributions from the collagen and PMMA which must be accounted for. This is accomplished through spectral subtraction of pure collagen and PMMA spectra, using their 100% spectral peaks (Amide I, and the peak ~1,730 cm–1, respectively). Following this, the second derivative (seven- or nine-point Savitsky–Golay) spectrum in the v1v3 phosphate band spectral region is calculated. The peak positions in the second derivative spectrum are recorded and input into the curvefitting program. The type of these bands is set to a mixture of Gaussian and Lorentzian bands. The output is again expressed as underlying peak position and relative percent peak area. Once the curvefitting process has reached convergence, the results should be checked for fidelity. To achieve this, the individual underlying bands are added to construct a new composite peak, the second derivative spectrum of this peak calculated, and compared and contrasted against the second derivative spectrum of the original band. Of the resulting underlying bands, the ratio of the ones at ~1,030 and 1,020 cm–1 has been shown to be directly proportional to mineral maturity/crystallinity (9).

3.5.4. Pyridinoline/ Dehydrohydroxylysinonorleucine

For analysis of the pyridinoline/dehydrohydroxylysinonorleucine, the amide I and II spectral regions are baseline-corrected, and water vapour (interfering with the Amide I spectral region) and PMMA (overlapping with the Amide I spectral region) spectrally subtracted. The criteria for proper subtraction are described extensively elsewhere (23). The initial position and type (Gaussian) of underlying bands are determined through second derivative and difference spectroscopy (2). Once the curve-fitting process converges, the output of the analysis is expressed as peak position and relative percent area. The ratio of the relative areas of the peaks at ~1,660 and 1,690 cm–1 is finally calculated; this ratio has been shown to correspond to the pyridinoline/dehydrohydroxylysinonorleucine collagen cross-link ratio (2).

3.6. Analysis of Whole Spectral Images

When whole spectral images have to be analyzed, curve fitting is not a viable option due to the enormous amount of time needed to analyze the 4,096 individual FTIR spectra that comprise an image. Nevertheless, calculation of parameters may proceed through monitoring absorbance heights at specific wavelengths. The correspondence between the two approaches (curvefitting and subband percent area vs. absorbance height at specific wavelength) has been discussed elsewhere (1, 24, 25). The acquired spectra are typically baseline-corrected using a rubber-band function native to all spectroscopic software packages.

3.6.1. Correction for PMMA

The first obstacle that needs to be circumvented prior to analysis of whole spectral images is the correction for the spectral contributions of PMMA which are variable depending on the extent of

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mineralization (see Note 6). The easier (time- and effort-wise) way to do this is to apply a masking procedure, which entails dictating to the instrument-native software (or any commercially available spectroscopic processing software package) to ignore any pixels that the corresponding FTIR spectra show the presence of PMMA (based on the ~1,730 cm–1 characteristic vibrational band). Although this is the quickest way to exclude PMMA from the eventual measurements, it is highly recommended not to be used! The reason for this is that anatomical areas of extreme scientific interest such as osteoid and/or lightly mineralized areas (first few microns of primary mineralized areas) will also be excluded from the measurements. Since bone surfaces are the ones that are active, “shaving” even a few microns for convenience is most likely to result in loss of very important biological activity and its influence on the spectroscopically monitored parameters. Instead, a proportional subtraction (based on the ratio of the PMMA to Amide I in the case of collagen analysis, or PMMA to v1v3 phosphate in the case of mineral analysis, band intensity, or integrated peak area) should be performed (1). 3.6.2. Mineral to Matrix ratio

The mineral to matrix ratio for the whole image is calculated as follows: the integrated peak areas for v1v3 phosphate (~900–1,200 cm–1), Amide I (~1,590–1,700 cm–1) and PMMA (~1,690–1,800 cm–1) are recorded in the instrument native software, and exported as an xyz (where x and y are the coordinates for a specific pixels, and z is the outcome monitored at the same pixel) spreadsheet to a commercially available scientific plotting software such as Sigmaplot or Origin, capable of reconstructing matrices (typically 64 × 64) into 2D or 3D images. The next step is going back to the original spectral image and identifying pixels where there is minimal mineral peak present and PMMA. The ratio of the integrated areas of v1v3 PO43−/PMMA at these pixels is calculated. Then back to the xyz spreadsheet, the area of PMMA is multiplied by this ratio and the resulting column subtracted from the original v1v3 PO43− column. It is this resulting column that represents a corrected one for PMMA values (as any value due to anything else other than a “pure” v1v3 PO43− spectral contribution will be either zero or negative). Following this, the resulting column is divided by the Amide I integrated area values, and the resulting xyz matrix converted back to a 2D or 3D image. In this image, values of zero or negative ones are then excluded from further consideration.

3.6.3. Ratio of Carbonate to Phosphate

The most useful FTIR parameter for eventual imaging applications is the band area ratio of the CO3− v3 contour to the PO43− v1,v3 contour. The main difficulty with the utilization of the CO32− v3 mode for IR imaging is the potential interference from overlapping vibrational modes of both the protein component of the tissue and from the PMMA embedding material. To compensate for contributions from these to the CO32− v3, a double subtraction protocol (using

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pure PMMA and collagen spectra) is employed. Detailed description and validation of this approach has been published elsewhere (10). Briefly, for the quantitative determination of CO32−, the contributions of embedding material (PMMA) and organic matrix (collagen) to the original spectra are subtracted based on spectra of the pure compounds. Thus, the intensity of the C=O stretching mode of PMMA at ~1,720 cm–1 and the amide I frequency at ~1,650 cm–1 (for collagen) are used as standards for spectral subtraction because there are no mineral contributions at these frequencies in the tissue spectra. For each spectrum in the IR imaging data sets, these correction factors are applied globally to the entire spectrum, thereby minimizing contributions from non-mineral components of the tissue to the CO32− v3 region. 3.6.4. The Mineral Maturity/Crystallinity

The mineral maturity/crystallinity is calculated based on the absorbance height ratio at 1,030/1,020 after appropriate correction for PMMA contribution has been completed. The general procedure is the same as described for the mineral maturity and crystallinity (Subheading 3.5.3), only in this instance the absorbance heights at 1,030, 1,020, and 1,730 cm–1 are utilized. The correcting ratio that the absorbance height values of PMMA are multiplied by is the 1,030/1,730 ratio, the resulting values subtracted from the 1,030 column values, and then the 1,030/1,020 ratio calculated (1, 24, 25).

3.6.5. Pyridinoline/ Dehydrohydroxylysinonorleucine Ratio

The ratio of pyridinoline/dehydrohydroxylysinonorleucine collagen cross-links is calculated from the baseline-corrected spectra, utilizing the absorbance heights at 1,660, 1,690, and 1,730 cm–1. For PMMA contribution correction, the 1,660/1,730 absorbance height ratio is used, and once calculated it is multiplied with the PMMA values, and the resulting values subtracted from the 1,660 absorbance height values, after which the correct 1,660/1,690 ratio may be calculated, correlating to the above-mentioned collagen cross-link ratio (2). In case, this ratio is the major outcome sought through the FTIRI analysis, then histologically stained sections (facilitating the easy depiction of bone surface metabolic activity) may be used (2).

4. Notes 1. This has a direct bearing on the chemistry, size and shape, and solubility of the apatite crystals. 2. An important strength of FTIR is that the results can be averaged across the whole section or expressed as a function of the extent of mineralization or tissue age within a specific region of the section.

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3. All FTIR instruments come with dedicated software built in; for example OPUS for Bruker Optics instruments. Commercially available software packages include MatLab, Isys, and Grams 32. 4. Where possible thin sections (0.5–1.0 μm) should be used when mouse bone is being analyzed, whereas for human bone sections of up to 5 μm in thickness can be used. Sections thicker than 5 μm usually saturate the MCT detector, resulting in spectral artefacts. 5. Formalin may interfere with the analysis of collagen cross links. 6. PMMA does not penetrate into heavily mineralized tissue areas, but is present in lightly mineralized zones. References 1. Marcott, C., Reeder, R. C., Paschalis, E. P., Tatakis, D. N., Boskey, A. L,. and Mendelsohn, R. (1998) Infrared microspectroscopic imaging of biomineralized tissues using a mercury-cadmium-telluride focal-plane array detector. Cell Mol. Biol. (Noisy-le-grand) 44, 109–15. 2. Paschalis, E. P., Verdelis, K., Doty, S. B., Boskey, A. L., Mendelsohn, R., and Yamauchi, M. (2001) Spectroscopic characterization of collagen cross-links in bone. J. Bone Miner. Res. 16, 1821–8. 3. Camacho, N. P., Rinnerthaler, S., Paschalis, E. P., Mendelsohn, R., Boskey, A. L., and Fratzl, P. (1999) Complementary information on bone ultrastructure from scanning small angle X-ray scattering and Fourier-transform infrared microspectroscopy. Bone 25, 287–93. 4. Durchschlag, E., Paschalis, E. P., Zoehrer, R., Roschger, P., Fratzl, P., Recker, R., Phipps, R., and Klaushofer, K. (2006) Bone material properties in trabecular bone from human iliac crest biopsies after 3- and 5-year treatment with risedronate. J. Bone Miner. Res. 21, 1581–90. 5. Boskey, A., and Mendelsohn, R. 2005 Infrared analysis of bone in health and disease. J. Biomed Opt. 10, 031102. 6. Boskey, A., and Pleshko Camacho, N. (2007) FT-IR imaging of native and tissue-engineered bone and cartilage. Biomaterials 28, 2465–78. 7. Gourion-Arsiquaud, S., West, P. A., and Boskey, A. L. (2008) Fourier transform-infrared microspectroscopy and microscopic imaging. Methods Mol. Biol. 455, 293–303. 8. Paschalis, E. P. (2009) Fourier transform infrared analysis and bone. Osteoporos Int. 20, 1043–7. 9. Paschalis, E. P., DiCarlo, E., Betts, F., Sherman, P., Mendelsohn, R., and Boskey, A. L. (1996)

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FTIR microspectroscopic analysis of human osteonal bone. Calcif. Tissue Int. 59, 480–7. Ou-Yang, H., Paschalis, E. P., Mayo, W. E., Boskey, A. L., and Mendelsohn, R. 2001 Infrared microscopic imaging of bone: spatial distribution of CO3(2-). J. Bone Miner. Res. 16, 893–900. Aparicio, S., Doty, S. B., Camacho, N. P., Paschalis, E. P., Spevak, L., Mendelsohn, R., and Boskey, A. L. (2002) Optimal methods for processing mineralized tissues for Fourier transform infrared microspectroscopy. Calcif. Tissue Int. 70, 422–9. Blouin, S., Thaler, H. W., Korninger, C., Schmid, R., Hofstaetter, J. G., Zoehrer, R., Phipps, R., Klaushofer, K., Roschger, P., Paschalis, E. P. (2009) Bone matrix quality and plasma homocysteine levels. Bone 44, 959–64. Paschalis, E. P., Boskey, A. L., Kassem, M,. and Eriksen, E. F. (2003) Effect of hormone replacement therapy on bone quality in early postmenopausal women. J. Bone Miner. Res. 18, 955–9. Paschalis, E. P., Glass, E. V., Donley, D. W., and Eriksen, E. F. (2005) Bone mineral and collagen quality in iliac crest biopsies of patients given teriparatide: new results from the fracture prevention trial. J. Clin. Endocrinol. Metab. 90, 4644–9. Boskey, A. L., DiCarlo, E., Paschalis, E., West, P., and Mendelsohn, R. (2005) Comparison of mineral quality and quantity in iliac crest biopsies from high- and low-turnover osteoporosis: an FT-IR microspectroscopic investigation. Osteoporos Int. 16, 2031–8. Paschalis, E. P., Betts, F., DiCarlo, E., Mendelsohn, R., and Boskey, A. L. (1997) FTIR microspectroscopic analysis of human

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iliac crest biopsies from untreated osteoporotic bone. Calcif. Tissue Int. 61, 487–92. Paschalis, E. P., Burr, D. B., Mendelsohn, R., Hock, J. M., and Boskey, A. L. (2003) Bone mineral and collagen quality in humeri of ovariectomized cynomolgus monkeys given rhPTH(1–34) for 18 months. J. Bone Miner. Res. 18, 769–75. Paschalis, E. P., Recker, R., DiCarlo, E., Doty, S. B., Atti, E., and Boskey, A. L. (2003) Distribution of collagen cross-links in normal human trabecular bone. J. Bone Miner. Res. 18, 1942–6. Paschalis, E. P., Shane, E., Lyritis, G., Skarantavos, G., Mendelsohn, R., and Boskey, A. L. (2004) Bone fragility and collagen crosslinks. J. Bone Miner. Res. 19, 2000–4. Roschger, P., Manjubala, I., Zoeger, N., Meirer, F., Simon, R., Li, C., Fratzl-Zelman, N., Misof, B., Paschalis, E., Streli, C., Fratzl, P., and Klaushofer, K. (2009) Bone Material Quality in Transiliac Bone Biopsies of Postmenopausal Osteoporotic Women After 3 Years Strontium Ranelate Treatment. J. Bone Miner. Res. 25, 891–900.

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21. Gourion-Arsiquaud, S., Faibish, D., Myers, E., Spevak, L., Compston, J. Hodsman, A., Shane, E., Recker, R. R., Boskey, E. R., and Boskey, A. L. (2009) Use of FTIR spectroscopic imaging to identify parameters associated with fragility fracture. J. Bone Miner. Res. 24, 1565–71. 22. Lieber, C. A., and Mahadevan-Jansen, A. (2003) Automated Method for Subtraction of Fluorescence from Biological Raman Spectra. Applied Spectroscopy 57, 1363–1367. 23. Dong, A., Huang, P., and Caughey, W. S. (1990) Protein secondary structures in water from second-derivative amide I infrared spectra. Biochem. 29, 3303–3308. 24. Mendelsohn, R., Paschalis, E. P., and Boskey, A. L. (1999) Infrared Spectroscopy, Microscopy, and Microscopic Imaging of Mineralizing Tissues. Spectra-Structure Correlations from Human Iliac Crest Biopsies. J. Biomed. Optics 4, 14–21. 25. Mendelsohn, R., Paschalis, E. P., Sherman, P. J., and Boskey, A. L. (2000) IR Microscopic Imaging of Pathological States and Fracture Healing of Bone. Applied Spectroscopy 54, 1183–1191.

Chapter 32 Raman Microscopy of Bone Simon R. Goodyear and Richard M. Aspden Abstract Raman microscopy is a non-destructive technique requiring minimal sample preparation that can be used to measure the chemical properties of the mineral and collagen parts of bone simultaneously. Modern Raman instruments contain the necessary components and software to acquire the standard information required in most bone studies. The spatial resolution of the technique is about a micron. As it is nondestructive and small samples can be used, it forms a useful part of a bone characterisation toolbox. Key words: Raman spectroscopy, Bone chemistry, Composition

1. Introduction Raman microscopy is a non-destructive technique requiring minimal sample preparation that can be used to measure the chemical properties of the mineral and collagen parts of bone simultaneously. Raman spectroscopy uses laser light and measures the change in wavelength of the light (shift) due to inelastic scattering. The amount of shift is dependent on the chemical bonds present in the scattering object (1) and the spectrum including all shifts provides a “chemical fingerprint” of the sample. The intensity of a spectral peak, or band, is due to laser power and frequency, the polarisability of the bond’s electron cloud, the type of bond causing the band, the number of bonds in the area imaged, and sample absorbance (2). This technique is particularly applicable to biological tissues because it is non-destructive, it is insensitive to water content, requires little or no sample preparation and, when coupled with microscope optics for sample illumination and spectral acquisition, allows investigation at the micron level (3). Comparisons can be

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made between, or within, samples by analysing either individual band parameters (e.g. peak area, position or height) (4) or the spectra as a whole using multivariate techniques such as principal component analysis (PCA) (3, 5). Raman microscopy has been used for a wide variety of bone studies including mineralisation (6–9), ageing (10, 11), the variation of composition within a bone (12), loading (13), and comparison between wild-type (WT) and knockout (KO) mice (14, 15). Excellent explanations of the Raman effect and typical apparatus used in a Raman experiment may be found in refs. 2, 16–20.

2. Materials 1. Raman microscope: a typical Raman instrument includes a source of monochromatic light (usually a laser), optics to focus the light onto the sample (a microscope), a method for moving the sample relative to the laser beam (XYZ motorised stage) more optics to collect the scattered light from the sample (the microscope and a filter to remove elastically scattered light), a method for forming a spectrum from the scattered beam (a diffraction grating) and a way of digitising the spectrum (CCD). Choices available within this assembly include the laser frequency and objective lenses used. In addition, polarisation optics before and after the sample can also be included. We use a Renishaw inVia Raman microscope. A 300 mW, 785 nm laser (Renishaw, Wotton-under-Edge, Gloucestershire, UK) is used for sample illumination and focussed on the sample through a Research Grade Leica DMLM microscope (Leica, Milton Keynes) fitted with a H101 motorised XYZ stage (Prior, Cambridge) and a ×63/0.95 numerical aperture Achroplan water immersion objective (Zeiss, Welwyn Garden City, Hertfordshire, UK). A conventional ×5 lens (Leica, Milton Keynes) is used for visualising the whole specimen and targeting sampling points. 2. Software for processing individual spectra and producing simple maps. This is usually included with the Raman equipment. 3. Optional: a programmable package such as Matlab, with the Curve Fitting Toolbox. This is useful where large numbers of spectra are to be processed, or non-standard techniques employed.

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3. Methods 3.1. Instrument Calibration

At the beginning of each imaging session, the laser is checked for alignment with the optical axis of the microscope and the wave number datum checked against a silicon internal standard and adjusted if necessary. Care needs to be taken to ensure that the silicon peak intensity is approximately the same each time, thus ensuring comparability of spectral intensities. Intensity calibration is not routinely available on Raman microscopes so results are commonly presented as ratios of spectral parameters (see Subheading 3.6).

3.2. Sample

Generally, if the laser beam can be focussed on the sample a spectrum can be collected. Minimal preparation is necessary and as the technique is non-destructive, the sample can subsequently be used for other tests. We typically use fresh cross-sections of mouse femora or tibiae cut using a bone saw, previously used for measuring the speed of sound by ultrasound (see Chapter 35 by Goodyear and Aspden, this volume), submersed in PBS (Fig. 1). Fixed or embedded samples can be used although care must be taken with interpretation of results. See ref. 21 for further details.

3.3. Acquiring Spectra

Position sample and determine the locations for spectral acquisition. For the examples used here, spectra were recorded from random sites around the whole cross-section of each sample using the mapping functionality of the Renishaw software and between 10 and 40 spectra are recorded. The motorised stage is used to move each selected location in turn to the focal point of the microscope.

Fig. 1. The sample arrangement for Raman microscopy. Samples are glued to a microscope slide with superglue and imaged submerged in PBS via a water-dipping lens.

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Although the samples are prepared by making parallel cuts on a bone saw and so are reasonably flat, the microscope’s automatic focus function is used to ensure the laser is focussed on the bone surface before data collection started. Each spectrum recorded is the summation of a number of accumulations of a set exposure time. Values are selected in a preliminary scan to give a reasonable signal-to-noise ratio without an excessive scan time. 3.4. Pre-processing

Cosmic ray artefacts, which cause spikes in the spectrum, are removed manually from each spectrum by linear interpolation between nearby unaffected regions in the Renishaw software. Spectra are then smoothed or de-noised. Smoothing techniques include the Savitsky–Golay algorithm, while de-noising uses a wavelet method (22, 23). The underlying background signal is then removed by subtracting a baseline of varying complexity, e.g. a linear or cubic curve, although we use an iteratively fitted polynomial (24).

3.5. Analysis

Figure 2 shows a typical Raman spectrum from bone where information on mineral and matrix components is presented simultaneously but distinctly, allowing information about each phase to be drawn from a single spectrum. Low bands are mostly from mineral, while the higher energy shifts are from collagen. The band at 961 cm–1 corresponds to the symmetric stretching vibration (v1) of the phosphate ion and is the strongest marker of bone mineral. The phosphate bending vibrations v2 and v4 appear at

Intensity Counts

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HPO42– Carb v1/ Phos v3

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Fig. 2. A Raman spectrum from mouse bone showing the origin of the major bands. Information from the mineral and matrix parts of the bone can be seen clearly. The spectrum has been denoised and the background signal removed.

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438 and 589 cm–1, respectively while the non-symmetric stretch (v3) causes a band at about 1,040 cm–1. A peak due to a superposition of carbonate and phosphate v3 appears at 1,070 cm–1. The high-frequency bands, from the organic phase are the amide III (~1,260 cm–1) and amide I (~1,680 cm–1) bands, which arise largely from the collagen (25), and the CH2 peak at approximately 1,450 cm–1 which is present in both collagenous and non-collagenous organic molecules (14, 26). Ratios of quantities derived from these bands are usually reported, peak height or peak area can be used: we have found that both give equivalent results and so use peak height as these are simpler to derive, and there is precedence in the literature, e.g. refs. 10, 27. Where bands were distinct (phosphate bends and the organic bands) their intensities and areas can be calculated in Excel. Intensities and areas from overlapping bands – the phosphate stretches, carbonate, and HPO42− bands lying between 900 and 1,100 cm–1 – were calculated by fitting mixed Gaussian–Lorentzian functions using the curve fitting toolbox in Matlab. This method was also employed to fit sub-bands to the amide I band. A mineral to matrix ratio can be calculated from the ratio of intensities of any of the mineral (phosphate v1, v2, or v4) bands and either of the matrix bands (amide I or III). We use the phosphate v4 and the amide III peak to minimise the possibility of orientation/polarisation complications (see Subheading 3.7) (28). While the peak at 1,070 cm–1 is formed by contributions from both carbonate and phosphate v3, it has been shown to be a good measure of the carbonate content of bone (29) so the carbonate to phosphate ratio was calculated by dividing the value from the carbonate peak (1,070 cm–1) by the phosphate v4 peak. Similarly, the relative amount of HPO42− was obtained by dividing the amplitude of the peak at 1,003 cm–1 by that of the phosphate v4 peak (25). The ratio of peaks fitted within the amide I band gives a measure of ordered to disordered collagen (14). In FTIR spectroscopy, this ratio has been shown to give a measure of mature, trivalent crosslink bonds and immature, divalent bonds (30); however, this has never been verified independently for Raman and so should not be used for this purpose. The FWHM of the phosphate v1 band, taken together with the degree of substitution values (carbonate to phosphate ratio), gives a measure of mineral crystallinity (10, 31–33). 3.6. Principal Component Analysis

Comparisons between whole spectra can be made visually but more subtle differences can be detected using statistical techniques, typically PCA. PCA assesses intensities at all wavenumbers simultaneously and generates a small number of ordered components or variables that account for the variance within the data. The first few components describe the majority of the variance and so can be used instead of the original variables (wavenumbers in this case) to represent the data. Typically, of the order of ten new variables, the

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modes of variation or principal components, are used to replace the vast number of original variables. Due to the nature of PCA these new variables are orthogonal, or linearly independent. Each mode is assigned a score that describes the spectrum from each sample and the set of scores can be compared statistically between samples. More complete descriptions of PCA can be found in refs. 34, 35. Many authors normalise spectra before comparison to overcome the differences in overall magnitude resulting from different laser power, quality of focus, and other factors such as the density of scattering centres. Most authors use the phosphate v1 band to normalise by refs. 12, 36 although the CH2 band (~1,450 cm–1) (7, 11) and total area under the curve (14) have also been used. We believe this approach should be used with caution because it suggests the normalising band is invariant in all spectra processed and this cannot be known a priori. When the materials being compared are very similar, as in the case of bone, and the laser power is checked using the silicon standard, there is a good argument that suggests using the spectrum without normalising has advantages (8). Relating PCs to Raman spectra can be challenging as the PCs can appear very different from the original spectra and so post-processing steps are sometimes applied, starting with manual rotation and combination (3, 25, 27). This is a labour-intensive procedure and success would rely on the expertise of the manipulator. Others use non-negativity constraints so that PCs are positive (37). The most complex procedure used appears to be Band Target Entropy Minimisation (BTEM), introduced by Chew et al. (38), which uses non-negativity, simulated annealing and entropy minimisation to extract individual constituents from mixtures of materials. 3.7. Polarisation/ Orientation

The laser used for Raman excitation is linearly polarised. In addition, bone contains lamellae which themselves contain oriented collagen fibres and mineral crystallites. Some Raman bands are sensitive to polarisation anyway, even in liquids, and the oriented structures in the sample mean that many others are sensitive to the orientation of the sample with respect to the polarisation of the beam. This has been almost unrecognised in the mineralised tissue literature in spite of many publications investigating bone. Orientation bias becomes an issue when the sampling volume contains fibrils that are all oriented in a common direction, for instance when using a high-magnification objective lens to obtain spectra from within one lamella. Using a ×100 objective to image an osteon, Kazanci et al. (28) showed orientation, and composition information can be extracted from the same spectrum by using bands that are more or less sensitive to polarisation. They suggest using the phosphate v2 or v4 and amide III bands for composition information as they are less affected by polarisation. This phenomenon is less likely to be an issue when low powered objectives are used or where many spectra from random sites are averaged.

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References 1. Banwell, C. N., and McCash, E. M. (1994) Fundamentals of Molecular Spectroscopy. 4th ed., McGraw-Hill, London; New York. 2. Smith, E. (2005) Modern Raman Spectroscopy: A Practical Approach. John Wiley, Chichester. 3. Tarnowski, C. P., Ignelzi, Jr. M.A., and Morris, M. D. (2002) Mineralization of Developing Mouse Calvaria as Revealed by Raman Microspectroscopy. J. Bone Miner. Res. 17, 1118–1126. 4. Callender, A. F., Finney, W. F., Morris, M. D., Sahar, N. D., Kohn, D. H., Kozloff, K. M., and Goldstein, S. A. (2005) Dynamic Mechanical Testing System for Raman Microscopy of Bone Tissue Specimens. Vib. Spectrosc. 38, 101–105. 5. Notingher, I., Jell, G., Notingher, P. L., Bisson, I., Tsigkou, O., Polak, J. M., Stevens, M, M., and Hench, L. L. (2005) Multivariate Analysis of Raman Spectra for in Vitro Non-Invasive Studies of Living Cells. J. Mol. Struct. 744–747, 179–185. 6. Wang, C., Wang, Y., Huffman, N. T., Cui, C., Yao, X., Midura, S., Midura, R. J., and Gorski, J. P. (2009) Confocal Laser Raman Microspectroscopy of Biomineralization Foci in UMR 106 Osteoblastic Cultures Reveals Temporally Synchronized Protein Changes Preceding and Accompanying Mineral Crystal Deposition. J. Biol. Chem. 284, 7100–7113. 7. Penel, G., Delfosse, C., Descamps, M., and Leroy, G. (2005) Composition of Bone and Apatitic Biomaterials as Revealed by Intravital Raman Microspectroscopy. Bone. 36, 893–901. 8. Goodyear, S. R. (2009) Physicochemical Methods for Measuring the Properties of Bone and their Application to Mouse Models of Disease. PhD ed., University of Aberdeen, Aberdeen. 9. Goodyear, S. R., Gibson, I. R., Skakle, J. M., Wells, R. P., and Aspden, R. M. (2009) A Comparison of Cortical and Trabecular Bone from C57 Black 6 Mice using Raman Spectroscopy. Bone. 44, 899–907. 10. Akkus, O., Polyakova-Akkus, A., Adar, F., and Schaffler, M. B. (2003) Aging of Microstructural Compartments in Human Compact Bone. J. Bone Miner. Res. 18, 1012–1019. 11. Ager, J. W., Nalla, R. K., Breeden, K. L., and Ritchie, R. O. (2005) Deep-Ultraviolet Raman Spectroscopy Study of the Effect of Aging on Human Cortical Bone. J. Biomed. Opt. 10, 1–8. 12. Ramasamy, J. G., and Akkus, O. (2007) Local Variations in the Micromechanical Properties of Mouse Femur: The Involvement of Collagen Fiber Orientation and Mineralization. J. Biomech. 40, 910–918.

13. De Carmejane, O., Morris, M. D., Davis, M. K., Stixrude, L., Tecklenburg, M., Rajachar, R. M., and Kohn, D. H. (2005) Bone Chemical Structure Response to Mechanical Stress Studied by High Pressure Raman Spectroscopy. Calcified Tissue Int. 76, 207–213. 14. Dehring. K, A., Crane, N. J., Smukler, A. R., McHugh, J. B., Roessler, B. J., and Morris, M. D. (2006) Identifying Chemical Changes in Subchondral Bone Taken from Murine Knee Joints using Raman Spectroscopy. Appl. Spectrosc. 60, 1134–1141. 15. Goodyear, S. R., Gibson, I. R., Skakle, J. M. S., Wells, R. P. K., and Aspden, R. M. (2007) P49 the Mechanical, Material and Chemical Properties of Cortical Bone from nNOS Null Mice. J. Bone Miner. Res. 22, 1138. 16. Weber, W. H., and Merlin, R. (2000) Raman Scattering in Materials Science. Springer, Berlin: London. 17. Laserna, J. J. (1996) Modern Techniques in Raman Spectroscopy. Wiley, Chichester; New York. 18. Baranska, H. (1987) Laser Raman Spectrometry: Analytical Applications. Horwood, Chichester. 19. Long, D. A. (1977) Raman Spectroscopy. McGraw-Hill, New York. 20. Tanaka, M., and Young, R. J. (2006) Review Polarised Raman Spectroscopy for the Study of Molecular Orientation Distributions in Polymers. J. Mater. Sci. 41, 963–991. 21. Yeni, Y. N., Yerramshetty, J., Akkus, O., Pechey, C., and Les, C. M. (2006) Effect of Fixation and Embedding on Raman Spectroscopic Analysis of Bone Tissue. Calcified Tissue Int. 78, 363–371. 22. Cai, T. T., Zhang, D., and Ben-Amotz, D. (2001) Enhanced Chemical Classification of Raman Images using Multiresolution Wavelet Transformation. Appl. Spectrosc. 55, 1124–1130. 23. Barclay, V. J., Bonner, R. F., and Hamilton, I. P. (1997) Application of Wavelet Transforms to Experimental Spectra: Smoothing, Denoising, and Data Set Compression. Anal. Chem. 69, 78–90. 24. Lieber, C. A., and Mahadevan-Jansen, A. (2003) Automated Method for Subtraction of Fluorescence from Biological Raman Spectra. Appl. Spectrosc. 57, 1363–1367. 25. Timlin, J. A., Carden, A., and Morris, M. D. (1999) Chemical Microstructure of Cortical Bone Probed by Raman Transects. Appl. Spectrosc. 53, 1429–1435. 26. Morris, M. D., and Finney, W. F. (2004) Recent Developments in Raman and Infrared

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S.R. Goodyear and R.M. Aspden Spectroscopy and Imaging of Bone Tissue. Spectroscopy. 18, 155–159. Carden, A., Rajachar, R. M., Morris, M. D., and Kohn, D. H. (2003) Ultrastructural Changes Accompanying the Mechanical Deformation of Bone Tissue: A Raman Imaging Study. Calcified Tissue Int. 72, 166–175. Kazanci, M., Roschger, P., Paschalis, E. P., Klaushofer, K., and Fratzl, P. (2006) Bone Osteonal Tissues by Raman Spectral Mapping: Orientation-Composition. J. Struct. Biol. 156, 489–496. Awonusi, A., Morris, M. D., and Tecklenburg, M. M. J. (2007) Carbonate Assignment and Calibration in the Raman Spectrum of Apatite. Calcified Tissue Int. 81, 46–52. Paschalis, E. P., Verdelis, K., Doty, S. B., Boskey, A. L., Mendelsohn, R., and Yamauchi, M. (2001) Spectroscopic Characterization of Collagen Cross-Links in Bone. J. Bone Miner. Res. 16, 1821–1828. Wopenka, B., and Pasteris, J. D. (2005) A Mineralogical Perspective on the Apatite in Bone. Mat. Sci. Eng. C. 25, 131–143. Freeman, J. J., Wopenka, B., Silva, M. J., and Pasteris, J. D. (2001) Raman Spectroscopic Detection of Changes in Bioapatite in Mouse Femora as a Function of Age and in Vitro Fluoride Treatment. Calcified Tissue Int. 68, 156–162.

33. Penel, G., Leroy, G., Rey, C., and Bres, E. (1998) MicroRaman Spectral Study of the PO4 and CO3 Vibrational Modes in Synthetic and Biological Apatites. Calcified Tissue Int. 63, 475–481. 34. Chatfield, C., and Collins, A. J. (1989) Principal Component Analysis, in Introduction to Multivariate Analysis pp 57–81, Chapman and Hall. 35. Hair, J. F. (1998) Multivariate Data Analysis. Prentice Hall, Upper Saddle River, N.J. 36. Kirchner, M. T., Edwards, H. G. M., Lucy, D., and Pollard, A. M. (1997) Ancient and Modern Specimens of Human Teeth: A Fourier Transform Raman Spectroscopic Study. Journal of Raman Spectroscopy. 28, 171–178. 37. Gentleman, E., Swain, R. J., Evans, N. D., Boonrungsiman, S., Jell, G., Ball, M. D., Shean, T. A. V., Oyen, M. L., Porter, A., and Stevens, M. M. (2009) Comparative Materials Differences Revealed in Engineered Bone as a Function of Cell-Specific Differentiation. Nat. Mater. 8, 763–770. 38. Chew, W., Widjaja, E., and Garland, M. (2002) Band-Target Entropy Minimization (BTEM): An Advanced Method for Recovering Unknown Pure Component Spectra. Application to the FTIR Spectra of Unstable Organometallic Mixtures. Organometallics. 21, 1982–1990.

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Chapter 33 The Calvarial Injection Assay Robert J. van ‘t Hof Abstract This chapter describes the calvarial injection method, whereby the effect of a substance on bone is tested by subcutaneous injection over the calvarium of a mouse. This assay allows testing of the effect of substances on both bone resorption and bone formation in a relatively simple in vivo model. The analysis is carried out by histological means, usually in glycolmethacrylate-embedded tissue, allowing for histochemical analysis and for a variety of different histological staining methods which are also described in detail. Key words: Calvarial injection, TRAP, Bone formation, Bone resorption, GMA embedding

1. Introduction There are several assays available to study the effects of cytokines, drugs, and hormones on bone cells in vitro. However, as the complex interactions between cells are disrupted, these in vitro assays do not always reflect what happens in vivo. The calvarial injection method, originally described by Boyce et al. (1), is valuable for studying the effects of substances on bone metabolism in vivo. In this assay, the substance to be tested is injected subcutaneously over the calvarium of a mouse. At the end of the assay, the animal is euthanised, the calvarium dissected, and analysed by microscopy. Although the assay was originally used to study the effects of cytokines on osteoclast formation and activity (1, 2), it has also been used to study the effects of drugs on bone formation (3).

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2. Materials 2.1. Injection

1. Recombinant murine IL-1a (5 mg/ml; CN Biosciences (UK) Ltd., Nottingham, UK). 2. Hamilton syringe (Luer-lock type; Anachem Ltd, Luton, UK).

2.2. Tissue Processing

1. Histocryl (glycolmethacrylate (GMA), TAAB). 2. Resin mix: Add 1.5 g catalyst (benzoyl peroxide, comes with the Histocryl) to 100 ml Histocryl; keep at 4 C. 3. Accelerator mix: 5 ml PEG 400, 5 ml dibutyl phthalate, 240 ml Histocryl accelerator. 4. Embedding mix: Add 175 ml accelerator mix to 1 ml of resin mix (at 4°C) and use immediately.

2.3. TRAcP/Von Kossa/ Light Green Stain

1. 1.5% (w/v) silver nitrate in dH2O. 2. 0.1% (w/v) hydroquinone. 3. 1% (w/v) Light Green in dH2O. 4. All the reagents for the TRAcP stain are described in Chapter 12, this volume.

2.4. Goldner’s Trichrome Stain

1. Weigert’s haematoxylin: (a) Solution A: Dissolve 10 g haematoxylin in 1,000 ml absolute alcohol. Ripen for at least 4 weeks before use. (b) Solution B: Dissolve 11.6 g ferric chloride (hydrated) in 1,000 ml distilled water and add 10 ml of 2% hydrochloric acid. Immediately before use, mix equal parts of A and B. Do not keep working solution pre-made. 2. Ponceau de xylidine/acid fuchsin: 1.5 g Ponceau de xylidine, 0.5 g acid fuchsin, 2 ml acetic acid (concentrated), 98 ml distilled water. 3. Azophloxine (working solution): 0.5 g azophloxine, 0.6 ml acetic acid (concentrated), 99.4 ml distilled water. 4. Ponceau de xylidine/acid fuchsin/azophloxine (working solution): 12 ml Ponceau de xylidine/acid fuchsin, 8 ml azophloxine, 80 ml 0.2% acetic acid; reuse the working solution. 5. Phosphomolybdic acid/Orange G: 6 g phosphomolybdic acid, 4 g Orange G, 1,000 ml distilled water. 6. Light Green: 2 g Light Green, 2 ml acetic acid (concentrated), 1,000 ml distilled water.

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3. Methods 3.1. Injection Protocol (Resorption)

1. Inject the mice over the calvarial bones with 10 ml recombinant murine IL-1a (5 mg/ml) or vehicle (sterile saline) using a 50-ml Hamilton syringe. Perform injections three times per day for three consecutive days (see Notes 1–3). 2. Euthanise the mice 4 days after the last injection. 3. Dissect out the calvarial bones and fix in 4% buffered formalin/saline (pH 7.4) for 1 h. 4. Rinse the calvaria in PBS and store in 70% alcohol. 5. Embed the undecalcified calvarial bones in GMA (see Notes 4 and 5) and cut 3-mm sections on a microtome (Jung, Heidelberg, Germany) using a glass knife (see Subheading 3.2 for embedding procedure). 6. Stain sections with von Kossa and TRAcP, followed by counter staining with Light Green. Alternatively, especially when one is interested in the effects on osteoblasts, the sections can be stained with Goldner’s trichrome (see Subheadings 3.3 and 3.4 for staining protocols).

3.2. Tissue Processing

Cut out a strip of calvarial tissue from the centre of the calvarium as illustrated in Fig. 1. The following steps are most easily performed using a tissue processor, but can also be performed manually. All the steps are performed at 4°C (see Note 6).

Fig. 1. A strip of tissue is cut out of the fixed calvarium and embedded for processing as indicated in this figure.

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1. Transfer tissue strips to 96% ethanol for 1 h. 2. 1 h in 100% ethanol. 3. 1 h in a 1:1 mix of 100% ethanol and resin mix. 4. 1 h in resin mix. 5. 72 h in resin mix. 6. Transfer tissue to a mould placed in a crushed-ice slush, fill with embedding mix, seal with a stub, and leave to polymerise for 1 h. 3.3. TRAcP/Von Kossa/ Light Green Staining of Mouse Calvariae

This method stains osteoclasts bright red, the mineralised bone black, and the remaining tissue green (Fig. 2). The Von Kossa stain should be performed first because the TRAcP staining solution (which is acidic) dissolves much of the mineral from the section resulting in an unsatisfactory Von Kossa stain. 1. Immerse sections in 1.5% silver nitrate (made up when required and filtered just before use) for 40 s. 2. Wash three times in water. 3. Develop the stain in 0.1% hydroquinone for 25–30 s (maximum). Check using a microscope at this point; mineralised bone should be black, not brown. If the bone looks brown, rinse in water and repeat the procedure. 4. Thoroughly rinse sections in running tap water for 10 min. Hydroquinone inhibits TRAcP staining and this step ensures that all the hydroquinone is washed off. 5. Perform TRAcP stain as described in various other chapters in this volume (for example, Chapter 12). Slides should be lying flat in plastic slide boxes with damp tissue lining the bottom. Boxes should then be covered to avoid drying of the staining solution.

Fig. 2. Calvarial sections from neonatal mice treated with saline (a) or IL-1a (b). Unlike the control section (a), numerous TRAcP-stained osteoclasts (red, arrows) are visible on the bone surface in (b) and extensive bone resorption is evident.

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6. Incubate at 37°C for 1.5 h (check staining after 1 h). 7. Rinse off the TRAcP staining solution with dH2O. 8. Counter stain with 1% Light Green for 30–60 s. Wash off with dH2O. 9. Air dry. 10. Mount with aqueous mounting medium (e.g. Apathy’s). 11. Store in cardboard slide trays and cover to prevent fading. 3.4. Goldner’s Trichrome

This stain results in bright green-stained calcified bone and good contrast of the cells. Although the osteoclasts do not stand out as well as with the TRAcP stain, this stain allows easy identification of osteoblasts. It is essential not to let the sections dry at any time during the staining protocol, as this leads to cracks in the mineralised bone. 1. Keep the sections in distilled water for at least 1 h (to prevent bubbling below section; if this still persists, keep the slides in water for longer). 2. Stain sections in Weigert’s haematoxylin for 20 min (see Note 7). 3. Wash in water. 4. Differentiate with 0.5% acid alcohol. 5. Wash in water for 20 min. 6. Stain sections in Ponceau/acid fuchsin/azophloxine for 5 min. 7. Rinse in 1% acetic acid for 10 s. 8. Stain sections in phosphomolybdic acid/Orange G for 20 min. 9. Repeat step 7. 10. Stain sections in 0.2% Light Green for 5 min. 11. Rinse in water. 12. Blot dry. 13. Rinse in 100% alcohol. 14. Immerse the sections in xylene. 15. Wipe off xylene around section before mounting in DPX. This method stains cell nuclei blue/black, mineralised bone/ muscle green, and osteoid/collagen red (Fig. 3).

3.5. Analysis of Results

Although many qualitative conclusions can be drawn about the effects of test substances by simple microscopical observation, we usually perform a quantitative analysis using computer-assisted histomorphometry. Parameters of interest are numbers of osteoclasts and osteoblasts per bone surface, mineralised bone width and bone formation, and resorption surfaces. We use software developed

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Fig. 3. Calvarial section, stained with Goldner’s trichrome, from a neonatal mouse treated with BMP-2. Large, activated osteoblasts (blue–black nuclei) and osteoid/collagen (red ) are clearly visible above the bone surface (green). Original magnification, ×200.

using the Aphelion ActiveX image analysis toolkit from ADCIS (ADCIS SA, Hérouville-Saint-Clair, France) and a Zeiss Axioskop microscope fitted with a colour camera. The program prompts the user to select and focus a field, grabs the image, and identifies the part of the image that contains tissue. The bone is identified using colour thresholding and the bone surface, volume, and width are calculated (see Note 8). Then, the resorption and formation surfaces are drawn onto the image by the user and finally the user is prompted to enter the number of osteoclasts (see Note 9) and osteoblasts present in this field. We usually measure at least ten fields from a representative area of a section (using a 20× objective lens, see Note 10), three sections at different levels (at least 100 mm apart) per animal, and at least six animals per treatment group (see Note 11).

4. Notes 1. Mice aged from several days up to several months can be used for this assay. Neonates require smaller amounts of injection material (useful when using an expensive drug) and have the advantage that they are easier to handle. 2. The injection schedule needs to be optimised for each substance tested in this assay. One of the most important variables influencing this is the biological half-life of the substance tested. For example, when testing the effects of mevastatin, we used a regime of two injections (5 mg/kg) per day for 5 days and euthanised the animals 1 or 7 days after the last injection.

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3. It is essential that all injection solutions and syringes are sterile. Otherwise, the effects of a test drug could easily be masked by a localised immune response to the injection, which invariably produces some localised bone loss in the calvarium. 4. Embedding in standard MMA plastic is not an alternative, as the TRAcP stain does not work well on material embedded in this plastic. Adapted MMA embedding protocols have been developed that do allow TRAcP staining (Chapter 19, this volume); however, for calvaria, we find GMA easier to section and stain. An alternative could be the embedding of decalcified calvaria in wax. However, the authors do not know of any stains for this material that allow easy distinction between bone and the other tissues using simple colour thresholding; consequently, the semi-automated analysis of these sections is much more difficult. 5. We have found that the manufacturer’s protocol, which uses only the histocryl accelerator, often leads to brittle blocks that are difficult to cut. Our variation, which uses an accelerator mix, produces blocks that are easier to cut. 6. It is essential to perform the embedding at low temperature, especially the polymerisation step. This step is best performed in a crushed-ice slush that optimally cools the polymerising block. 7. Celestine Blue can be used as an alternative to haematoxylin if nuclei are not stained particularly well. Prepare the Celestine Blue as follows: 2.5 g Celestine Blue B, 25 g ferric ammonium sulphate, 70 ml glycerin, 500 ml dH2O. Dissolve the ferric ammonium sulphate in cold distilled water and stir well. Add Celestine Blue to this solution, and then boil the mixture for a few minutes. After cooling, filter the stain and add the glycerine. Use the same staining time for Celestine Blue as for haematoxylin, i.e. 20 min. 8. In many programs, the calvarial width is determined by having the user draw lines across the mineralised bone at multiple sites. This method is fairly time consuming and not very reproducible due to operator variability and bias. We use a mathematical method, whereby the calvarial bone is modelled as a rectangle and the width is calculated from the perimeter and the surface area of the bone according to the following formula: Width =

Perimeter − perimeter 2 − (16 × area) 4

.

To make this method work properly, all holes within the bone binary image should be closed (using an Image Holefill operator) and the outline should be smoothed by a Binary Close operator (or by an Image Dilate, followed by an Image Erode

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operator of the same size). The operators mentioned above are available in all image analysis packages that are currently on the market. 9. It can be difficult to get an accurate number of osteoclasts, as these cells are often present in clusters and not well-separated visually. Furthermore, as osteoclasts are such large, irregularly shaped cells, what appears to be several osteoclasts close together in a section may actually be parts of the same osteoclast. For this reason, it is good practice to analyse several histological sections, separated by at least 100 mm (see Subheading 3.5). 10. The number of fields to measure per sample depends on the field of view of the camera. Therefore, when using a large chip, high-resolution camera like the QImaging Retiga-4000 or Diagnostic Instruments Insight 1400, 4–5 fields are usually sufficient. 11. To avoid possible artefacts introduced by the dissection procedure, do not take histomorphometric measurements at the calvarial ends. References 1. Boyce, B. F., Aufdemorte, T. B., Garrett, I. R., Yates, A. J., and Mundy, G. R. (1989) Effects of interleukin-1 on bone turnover in normal mice Endocrinology 125, 1142–1150. 2. van ‘t Hof, R. J., Armour, K. J., Smith, L. M., Armour, K. E., Wei, X. Q., 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. 3. Mundy, G., Garrett, R., Harris, S., Chan, J., Chen, D., Rossini, G., Boyce, B., Zhao, M., and Gutierrez, G. (1999) Stimulation of bone formation in vitro and in rodents by statins Science 286, 1946–1949.

Chapter 34 Ovariectomy/Orchidectomy in Rodents Aymen I. Idris Abstract This chapter describes the surgical procedures for ovariectomy and orchidectomy in mice and rats. In addition to providing technical details of the surgical techniques, details of anaesthesia and perioperative care are also included. Key words: Surgery, Ovariectomy, Orchidectomy, Oestrogen, Androgen, Rodents, Mouse, Rat, Bone loss

1. Introduction Sex hormones play a vital role in regulating attainment of peak bone mass and in protecting against bone loss with increasing age. Oestrogen protects against bone loss in both men and women, mainly by suppressing bone resorption (1). Conversely, oestrogen deficiency predisposes to osteoporosis by causing accelerated bone loss and relative uncoupling between bone resorption and bone formation. Although oestrogen is mainly thought to act on cells of the osteoclast lineage (2), there is experimental evidence to suggest that oestrogen stimulates bone formation in rodents (3) and some evidence to suggest that high-dose oestrogen stimulates bone formation in humans (4). It has been suggested that the uncoupling between bone resorption and bone formation that occurs in response to oestrogen deficiency might be due in part to a protective effect of oestrogen on osteocyte and osteoblast apoptosis (5). The mechanisms by which oestrogen deficiency affects bone cell activity are incompletely understood, but multiple mechanisms are likely to be operative, including regulation of local production of RANKL and OPG; cytokines, such as IL-1 TNF and M-CSF; and direct effects on osteoblasts and osteoclasts (6). The most commonly used animal model of oestrogen deficiency is ovariectomy in rodents. 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_34, © Springer Science+Business Media, LLC 2012

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This results in fairly rapid bone loss which can be detected over a time frame of 2–4 weeks in mice (7) and 2–3 months in rats (8). This results in a rapid elevation in bone turnover with increases in bone resorption and bone formation with relative uncoupling between these processes. Androgen deficiency in men is associated with an increased risk of osteoporosis, although it remains unclear if this is a direct effect of testosterone or a result of local oestrogen deficiency in bone as the result of aromatisation of gonadal androgens (9). Whatever the underlying mechanism, orchidectomy (ORX) in male mice results in significant loss of bone mineral density and trabecular number due to excessive bone resorption (10). Increased cortical porosity has also been reported to occur following ORX in rats, but cortical bone loss is significantly less than trabecular bone loss (11, 12). In the present chapter, I describe the surgical technique for ovariectomy and orchidectomy in mice and rats.

2. Materials Trade names are occasionally used for identification purposes only and do not imply endorsement. 2.1. Surgical Instruments

1. Sterile instruments (scissors, tooth, and blunt forceps). 2. Metal clips and applying forceps. 3. Swab. 4. Sterile syringes (1 and 5 ml). 5. Hypodermic needles (25 gauge). 6. Infrared lamp. 7. Electric clipper. 8. Beaker.

2.2. Anaesthetic Agents

1. Isoflurane. 2. Ketamine hydrochloride (Vetalar V™) and medetomidine hydrochloride (Dormitor™) cocktail or Ketamine hydrochloride (Vetalar V™) and Xylazine (Rompun™) cocktail. 3. Atipamezole hydrochloride (Antisedan™).

3. Methods 3.1. Animal Husbandry

1. House animals in pathogen-free rooms maintained at constant temperature, with 12-h light/12-h dark cycles. 2. Provide free access to water and standard, pelleted commercial diet.

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3. Allow animals an adaptation/acclimatisation period of at least 3 days prior operating if transported from different facility. 3.2. Anaesthesia

1. Induce anaesthesia by intraperitoneal injection of a cocktail of Vetalar V™ (females, 75 mg/kg; males, 50 mg/kg) in combination with Dormitor™ (females, 1 mg/kg; males, 0.5 mg/kg) (see Note 1). 2. Check the depth of anaesthesia by monitoring respiratory rate (anaesthetised animals show reduced respiratory rate) or simply testing the animal response to gentle pressure on the hind paws.

3.3. Preoperative Care

1. After the onset of anaesthesia, place animals under infrared lamp to prevent heat loss. 2. Using an electric clipper, shave fur bilaterally over the lumbar spine (Ovx) or ventral side of the scrotum (Orx) to expose skin (see Note 2). 3. Swab the shaved skin with 70% (v/v) ethanol followed by sterile PBS. 4. Ensure that all experimental protocols are approved and conducted in accordance with regulation provided by regulatory ethics committee. 5. Sterilise and disinfect all surgical instruments and hard surfaces with 70% ethanol prior to use.

3.4. Operative Technique for Ovariectomy

1. Place the anaesthetised animal on the operating table with its back exposed and its tail towards you (Fig. 1a). 2. Make a single midline dorsal incision (0.5 cm for mice and 2 cm for rat) penetrating the skin using small scissors (see Fig. 1a). Incision should be made in the lower back, directly below the bottom of the rib cage (see Note 3). 3. Gently free subcutaneous connective tissue from the underlying muscle on each side using blunt forceps (see Fig. 1b). 4. Locate ovary under the thin muscle layer and make a small incision (less than 1 cm) on each side to gain entry to the peritoneal cavity (see Fig. 1b). 5. Hold securely the edge of the incision with tooth forceps and retract the ovarian fat pad surrounding ovaries with blunt forceps to expose oviduct (see Fig. 1c). 6. Identify and replace ovaries back into the abdominal cavity for sham operations. 7. Perform a single ligature around the oviduct (0.5 cm for mice and 2 cm for rats from ovary) to prevent bleeding following removal of ovary (see Fig. 1d).

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Fig. 1. Illustration of ovariectomy in the mouse. The ovary is indicated by the arrow.

8. Remove ovary by gently severing the oviduct, using sterile, small scissors (see Fig. 1e). 9. Replace uterus and remaining part of the oviduct back into the abdominal cavity. 10. Suture the muscle layer (see Note 4). 11. Turn the animal over so that it is still laid on its ventral surface but its tail is pointing away from you, and repeat the procedure described in steps 3–10 for the other ovary if using one skin incision (see Fig. 1f and Note 2). 12. Close the skin incision using metal clips (see Note 5). 3.5. Operative Technique for Orchidectomy

1. Place the anaesthetised animal on the operating table on its back with the tail towards you. 2. Make a single incision on the ventral side of the scrotum (0.5 cm for mice and 1.5 cm for rat) penetrating the skin using sterile scalpel (see Fig. 2a, b). 3. Localise the testicular fat pad on the left side and pull it through the incision using blunt forceps (see Fig. 2c). 4. Cut the cremaster muscles and locate the testicular fat pad and gently pull it through the incision using sterile, blunt forceps.

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Fig. 2. Illustration of orchidectomy in the rat.

5. Expose testicular content by gently freeing testicular fat pad with sterile, blunt forceps (see Fig. 2d). 6. Gently expose the cauda epididymis, caput epididymis, vas deferens, and testicular blood vessels while holding the testicular sack with sterile tooth forceps. 7. Perform a single ligature around the blood vessels to prevent bleeding following removal of testis. 8. Gently severe cauda epididymis and caput epididymis from the testis (see Fig. 2e). 9. Remove testis by gently severing blood vessels with small scissors (see Fig. 2f ). 10. Replace the remaining content of testicular sac back using blunt forceps. 11. Repeat steps 1–9 for the other testis. 12. Close the skin with metal clips (see Note 5). 3.6. Post-operative Care

1. Reverse anaesthesia using an intraperitoneal injection of atipamezole hydrochloride (Antisedan, 1 mg/kg for both males and females).

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2. House animals individually and keep under close observation for approximately 2–4 h until they fully recover from anaesthesia. 3. Following recovery period (approximately 24 h after surgery), the animals can be grouped together as normal. 4. Delay administration of experimental treatments for at least 24 h after surgery.

4. Notes 1. Consult your local veterinary official and local guidelines for preferred mode of anaesthesia and dose recommendations. We use intraperitoneal injection in preference to inhaled anaesthesia since it allows greater freedom of movement of the anaesthetised animal during operation and because induction and recovery are quicker. The anaesthetic should be prepared in as small a volume as possible (0.1 ml for mice and 0.2 ml for rats). 2. Shaving is optional when operating on mice. 3. The procedure can also be carried out by making two incisions to separately reach each ovary/testis. 4. If operating in mice, suturing the muscle layer is optional. 5. The use of metal clips to close skin incision is essential in rodents to prevent reopening of the incision.

Acknowledgement The author would like to thank Dr. Antonia Sophocleous PhD for providing the images for Figs. 1 and 2 and for critically reviewing the manuscript. References 1. Khosla, S., Melton, L. J. III, and Riggs, B. L. (2001) Estrogens and bone health in men. Calcif. Tissue Int. 69, 189–192. 2. Khosla, S., Melton, L. J. III, Atkinson, E. J., O’Fallon, W. M., Klee, G. G., and Riggs, B. L. (1998) Relationship of serum sex steroid levels and bone turnover markers with bone mineral density in men and women: a key role for bioavailable estrogen. J. Clin. Endocrinol. Metab. 83, 2266–2274.

3. Samuels, A., Perry, M. J., and Tobias, J. H. (1999) High-dose estrogen induces de novo medullary bone formation in female mice. J. Bone Miner. Res. 14, 178–186. 4. Tobias, J. H., and Compston, J. E. (1999) Does estrogen stimulate osteoblast function in postmenopausal women? Bone 24, 121–124. 5. Weinstein, R. S., and Manolagas, S. C. (2000) Apoptosis and osteoporosis. Am. J. Med. 108, 153–164.

34 6. Raisz, L. G. (2005) Pathogenesis of osteoporosis: concepts, conflicts, and prospects. J. Clin. Invest. 115, 3318–3325. 7. Idris, A. I., Van’t Hof, R. J., Greig, I. R., Ridge, S. A., Baker, D., Ross, R. A., and Ralston, S. H. (2005) Regulation of bone mass, bone loss and osteoclast activity by cannabinoid receptors. Nat. Med. 11, 774–779. 8. Rissanen, J. P., Suominen, M. I., Peng, Z., Morko, J., Rasi, S., Risteli, J., and Halleen, J. M. (2008) Short-term changes in serum PINP predict long-term changes in trabecular bone in the rat ovariectomy model. Calcif. Tissue Int. 82, 155–161.

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9. Khosla, S., Melton, L. J. III, and Riggs, B. L. (2001) Estrogens and bone health in men. Calcif. Tissue Int. 69, 189–192. 10. Gunness, M., and Orwoll, E. (1995) Early induction of alterations in cancellous and cortical bone histology after orchiectomy in mature rats. J. Bone Miner. Res. 10, 1735–1744. 11. Danielsen, C. C., Mosekilde, L., and Andreassen, T. T. (1992) Long-term effect of orchidectomy on cortical bone from rat femur: bone mass and mechanical properties. Calcif. Tissue Int. 50, 169–174. 12. Orwoll, E. S. (1996) Androgens as anabolic agents for bone. Trends Endocrinol. Metab. 7, 77–84.

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Part VII Mechanical Loading Techniques

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Chapter 35 Mechanical Properties of Bone Ex Vivo Simon R. Goodyear and Richard M. Aspden Abstract The primary functions of bone are to do with support and protection – mechanical functions. The aim of this chapter is to set out some of the methods that can be used to measure these properties in cortical and cancelleous bone from large (e.g. human or bovine) and small (e.g. mouse) animals. The difference between properties of the sample (intrinsic properties) and properties of the material (extrinsic properties) is introduced and techniques for measuring them suggested. The addition of other tests to give a complete characterisation of a bone sample is presented. Key words: Mechanical testing, Bone, Material properties, Mechanical properties

1. Introduction The primary function of bone is to form the skeleton, which provides support for the body and protection for vital organs. These are primarily mechanical functions. To fulfil these, the bone matrix has to have the right combination of stiffness and strength to enable it to withstand the forces imposed upon it. These forces may be repetitive and moderate, such as those generated during walking, or high and transient, such as inflicted by a blow on the head. The structure and composition of bone can adapt over time to try to match the mechanical properties of the bone to the prevailing demands being placed on it. How to measure some of these mechanical properties is the aim of this chapter. There are two types of bone which are considered in this chapter: cortical or dense bone and cancellous or trabecular bone. Cortical bone is a solid, compact material which forms the diaphysis of long bones, and a shell around the metaphysis and the vertebrae. Cancellous bone has an open, porous structure comprising rods or plates. Hence, it is less dense than cortical bone and also less stiff and strong. It makes up the centre of the vertebrae and 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_35, © Springer Science+Business Media, LLC 2012

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metaphysis. However, the two materials in combination form structures which are strong and light and their properties in situ may differ considerably from those measured from tests on isolated samples (1, 2). In cancellous bone, the distinction between material and structure is not always easy to define and treating it as a cellular material has met with a considerable degree of success (3, 4). Mechanical testing is one test in an armamentarium of methods that can be used to characterise bone, as illustrated in Fig. 1, which shows a scheme we have developed for testing mouse bones. Together, the methods measure the material, mechanical, and geometrical properties to describe the factors that determine a

Fig. 1. A suggested flowchart for characterising mouse bones. The order of tests is determined by whether preparation irreversibly changes the bone and also the size of sample required for the test. For example, mCT is positioned first as it is non-destructive but requires an intact piece of bone, whereas although Raman is also non-destructive it can be used on small samples, and so is at the bottom of the chart. The decision point half way through assesses whether there is a phenotype by testing both geometrical and mechanical properties. If neither shows an effect of treatment, it is likely the factor does not affect bone.

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bone’s competence in fulfilling its primary function, outlined above. In this chapter, methods to measure the intrinsic and extrinsic mechanical properties of bone (the shaded methods in Fig. 1) are described. Although conceptually relatively simple, the mechanical properties of these tissues are not easy to measure. If only a comparison is required between equivalent sites in different patient groups, then careful use of the same technique on all samples will provide consistent relative values. If, however, absolute values are required, then great care must be taken as bone is not isotropic or homogeneous, water content is important, and its properties depend on the rate at which it is deformed. In addition, sample preparation can be difficult. Unlike engineering materials, which can be cut and machined to predetermined sizes for testing and for which there are officially recognised standards, bone samples are generally limited in size and shape by the site from which they are taken. The size of the sample being tested may affect the measurements being made. This chapter describes methods of preparation and testing that can be applied to cortical and cancellous bone from human, bovine, or equine sources and goes on to present a method for testing whole bones in small animals, such as that required to measure bone properties in genetically modified mice. These are so small that preparing and testing isolated samples of cortical or cancellous bone becomes very difficult and tests are generally performed on intact bones. Differences in approach that are required because of the different natures of the materials are noted. The descriptions are restricted to methods that can reasonably easily be applied by anyone having access to a limited range of laboratory equipment, such as a materials testing machine, a balance of suitable precision, an ultrasound generator, an oven, and furnace. More sophisticated techniques for mechanical and material characterisation include micro- and nano-indentation, infrared spectroscopy [Fourier transform infrared (FTIR) or Raman], backscattered electron microscopy, and atomic force microscopy. Researchers requiring these techniques or more details of variations are referred to the book by Cowin (5) and to the original papers.

2. Materials 1. Phosphate-buffered saline (PBS): Used to keep specimens moist during preparation and testing (see Note 1). 2. Washing up liquid: For reducing surface tension. 3. Tweezers, scalpel, and scissors: For removing soft tissue and manipulating samples.

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4. Hacksaw: For cutting large bone samples to manageable-sized pieces prior to precision cutting. 5. Mineralogical saw [Accutom 2 (Struers Ltd., Rotherham) or Isomet (Buehler, Lake Bluff, IL, USA)] fitted with either a diamond or an aluminium oxide cut-off wheel: For precision cutting of bone samples (see Note 2). 6. Lathe or milling machine, as might be found in any engineering workshop: To machine bone samples to desired shape. 7. Coring bits, 5- and 9-mm internal diameter and break-off tools: Custom made by Bolton Surgical Ltd. (Sheffield, UK). 8. Materials testing machine (e.g. Instron, High Wycombe, Bucks or Electroforce test Instruments, Bose, Minnesota): For mechanical testing of samples. 9. Balance (Sartorius Gmbh, Göttingen, Germany) with specific gravity determination kit Sartorius 6080/60801 Specific Gravity Determination Kit: For density measurement by Archimedes’ principle. 10. Ultrasound generator (Panametrics Pulser–Receiver model 5052PR (Panametrics Inc, Waltham, MA, USA) with a V211BA transducer) and oscilloscope (Hitachi V-665A, Tokyo, Japan): For measuring the speed of sound. 11. Oven (Hybaid, Teddington) and furnace (Carbolite, Sheffield): For determining the relative composition. 12. Software for calculation of biomechanical variables: Origin (OriginLab Corp., Northampton, MA, USA), Matlab (the MathWorks Inc., Natick, MA, USA), or MathCad (Mathsoft Engineering and Education Inc., Cambridge, MA, USA).

3. Methods 3.1. General Considerations for the Preparation of Bone Samples

3.2. Preparing Prisms of Cortical Bone

All human tissues must be handled with due regard to health and safety because of the risk of infection. In addition to gloves and laboratory coat, a mask and eye shield are required since there is a risk of aerosols being generated during cutting, drilling, or machining. Samples must be kept moist at all times during the preparation procedure. Bones must be dissected free of soft tissues, taking care not to create notches in the bone that is going to be tested, since this weakens it. General methods for the preparation of samples for testing are described in the following sections. 1. Cut the bone into manageable-sized pieces using a hacksaw or a junior hacksaw. 2. Precision cut the bone samples to the desired size using a mineralogical saw rotating at 600–800 rpm. For compression and

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Fig. 2. Tensile test specimens are traditionally produced with a waist to ensure that failure consistently occurs in the centre and not near the grips.

bending tests, the samples should be cut to give rectangular parallelepiped-shaped specimens. For tensile testing, the specimens should be milled to give a waisted configuration (Fig. 2). 3. If the samples are large enough, the samples should have cylindrical symmetry, but for thin specimens the waisting can be done in two dimensions. This is to ensure that fracture occurs consistently in the central part of the specimen away from the gripped ends. Water should be run over the sample at all times during sawing and machining. 3.3. Preparing Cancellous Bone Cores

1. Obtain the bone core by drilling through the bone sample with a coring bit (Fig. 3). To remove the core, it either has to be drilled right through a piece of bone or broken off the underlying bone. Providing the core is not too deep; this can be done using the break-off tool shown in Fig. 3. This is made to the same specification as the coring bit, except it does not require teeth and half of its circumference has been removed. This is inserted in place of the coring bit and, by levering on the end of it, the base of the core may be broken from the underlying tissue (see Note 3). 2. Trim the ends of the cores plane and parallel using a mineralogical saw. If the cores have been obtained from the articular surface of joints, they will have a cartilage and subchondral bone at one end which needs to be removed using the saw.

3.4. Preparation of Mouse Bones

1. Remove the hind limb from the animal by cutting through the pelvis with scissors. Take care not to damage the head and neck of the femur. Strip the fur from the leg. If not immediately

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Fig. 3. Coring bits and matching “break-off” devices. Unless cores can be drilled right through a piece of bone, they need to be removed from the underlying bone to which they are attached. Inserting the break-off device after drilling the core and levering it sideways generally successfully snaps the core at its base after which it may be removed using forceps.

required for testing, samples should now be immersed in PBS and stored below −20°C. 2. Before testing, samples are defrosted and the soft tissue removed. First, the foot is removed by cutting though the ankle joint and then the soft tissue stripped from the bone using a scalpel. The femur is then separated from the tibia and pelvis, where present, and each bone placed in a separate vial of fresh PBS. The fibula is also removed from the tibia at this stage. If testing is to proceed in the next 24 h, bones are stored in the fridge or returned to the freezer if a longer interval is expected. 3. In mice, density and speed of sound measurements are made in cortical bone and so the metaphysis are removed to separate cortical from predominantly trabecular bone. The broken end of each bone section from the bending test (see below) is wrapped in PBS-soaked tissue paper and lightly clamped in a mineralogical saw and the metaphysis cut-off. After removing the distal metaphysis from the femur, a ~1.5-mm section is cut off for ultrasonic testing and the remaining cortical bone used for density determination. In the tibia, the proximal metaphysis is removed first, with the ultrasound sample being taken from the adjacent diaphysis. During cutting, the sample is irrigated with distilled water to remove cutting debris and keep it cool.

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3.5. General Considerations for Mechanical Testing

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Before testing, it is important to decide what parameters are going to be measured. Most testing machines apply a deformation to a sample, which normally increases linearly with time. This deformation is commonly referred to as displacement. The operator can choose the rate at which the displacement occurs and the properties measured depend on this rate as bone is viscoelastic. Choosing when to stop the test is also important depending on what is to be done next with the specimen. In tension or bending, the machine can be set to stop automatically on fracture, as the load drops rapidly at this point. In compression, failure is not so easy to determine and various methods can be used. The applied load and displacement are recorded throughout the test. The load– displacement curve represents the extrinsic properties of the specimen under test and is, therefore, used to measure the properties of whole bones (Fig. 4a). These properties are important for understanding why bones fracture. For prepared samples of

Fig. 4. A load–displacement curve (a) measures the extrinsic properties of a specimen, e.g. an individual bone. The main parameters are stiffness, work to failure (shaded ), and ultimate load and displacement. The stress–strain curve (b) is similar but, because it is normalised for the sample dimensions, measures the intrinsic properties of the material being tested. These are the elastic or Young’s modulus (E ), the yield stress and strain, the ultimate stress and strain, and the energies to yield and failure.

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known dimensions, what are commonly required are the intrinsic properties of the bone material itself. To obtain these, the load is divided by the cross-sectional area of the specimen to produce the stress, and the displacement is divided by the original length to give the strain. The stress–strain curve is then very similar to the load–displacement curve, but refers now to the material not the structure (Fig. 4b). 3.6. Repeating Tests on the Same Specimens

It has been found that if a test is repeated several times on the same specimen within a short period then the resulting load– displacement plots do not immediately coincide but appear to converge towards a stable curve. Because of this, many investigators apply up to 5 or 6 loading cycles to a specimen, often at a reduced load, before performing the actual test: a process called preconditioning. This is a point of some contention because it is not understood what happens during this process. It can be argued that what is finally being measured is not the natural property of the specimen but one that has somehow been modified by a series of cyclically applied loads. Does repeatability necessarily imply accuracy? We always apply a single test to a specimen.

3.7. Types of Mechanical Testing

There are four main types of test: tension, compression, bending, and torsion. 1. Tension and compression testing. These are most commonly done to measure the intrinsic properties of bone matrix. These tests use machined samples of known dimensions. Tension is generally used for cortical bone, and compression is more common for cancellous bone because of the difficulty obtaining and mounting suitably sized specimens for tensile testing. Bending and torsion can be applied in this way, but the interpretation of the results is more difficult. They are probably more commonly applied to whole bones. In this section, descriptions are given of tension and compression tests for machined specimens. 2. Bending. This is most commonly applied to whole long bones from rodents because of the difficulties of machining tensile or compressive specimens from such small bones. It is best for measuring extrinsic properties of the whole bone and, therefore, is very useful for studies comparing the effects of different drug therapies or genetic modifications. It is not easy to estimate intrinsic properties from this test for a variety of reasons, mainly related to the size and shape of the bone. More details are given by Turner and Burr (6). There are two bending configurations, called three-point and four-point loading (Fig. 5). In three-point loading, there is a significant shear stress generated at the mid-point of the beam. For this reason, four-point loading is sometimes preferred because in a uniform specimen shear is minimised and a pure bending moment is applied.

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Fig. 5. Whole bones may be tested in either (a) three-point or (b) four-point loading configuration. Force is applied through the upper plate and the resulting displacement recorded. The span is the distance between the lower supports.

However, in whole bones, the non-uniform cross section means that these assumptions do not apply. In addition, it is difficult to load all four points identically and test results are subject to large errors. Three-point and four-point loading jigs can be bought or made to fit most materials testing machines. It is important to be able to adjust the span, the distance between the lower supports. For testing whole bones, this should be maximised to reduce shear stresses but be constant for all specimens to be compared. For machined specimens, testing at different spans enables extrapolation to be made for an infinite span to give a shear stress-free modulus (7). 3.8. Tension Testing

1. Prepare the specimen as described in Subheading 3.2 or 3.3 ensuring that it is at least 15–25-mm long and 5 mm across (see Note 4). 2. Keep the sample moist by lightly wrapping in cling film or moist tissue or gauze. 3. Fix the specimen in the testing machine: Most testing machines have grips for holding specimens in tension. These can be tightened on to the broader ends of the specimen and are

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adequate for testing cortical bone (see Note 5). If you are testing cancellous bone cores, set the ends into cups using dental or bone cement and the cups are then held in the testing machine grips. 4. Set up the machine to apply the desired strain rate, and program in any predetermined limits on load, displacement or fracture detection, and the rate of data recording. It is advisable to set the machine to stop as soon as fracture occurs because at this point the load falls to zero. 5. Start the testing machine. 6. After the test, transfer data from the instrument to a spreadsheet for further analysis if the instrument does not already have appropriate software. 3.9. Analysing Data from Tension Testing

1. If intrinsic properties are required and not already calculated, convert load values to stress by dividing by the original crosssectional area. (No account is taken of changes in cross-sectional area which occur during the test.) 2. Convert displacement values to strains by dividing by the unstretched distance between the points of attachment or reference marks of the displacement transducer (see Note 5). 3. Plot stress as a function of strain: The elastic or Young’s, modulus, yield strength and strain, ultimate strength and strain, and energies to yield and failure may then be calculated. 4. Calculate the elastic modulus by estimating the gradient of the linear part of the curve. For linear relationships, this is often calculated by the instrument software. If not, this can be found crudely by hand calculation, better by fitting a straight line to the data or for non-linear relationships by differentiating the curve using software, such as Origin, Matlab, or MathCad. 5. Calculate the yield point by estimating when the elastic modulus starts to reduce. This is not always easy to define, but one way is to construct a line parallel with the linear part of the curve but offset along the strain axis by a small amount. The point at which this intersects the curve is defined as the offset yield point and the stress and strain at this point are the yield stress and strain. 6. Calculate the failure stress and strain from the point at which fracture occurs. 7. Integrate the curve to either the yield point or failure point to find the area beneath to estimate the energy needed to produce yield or failure, respectively, per unit volume of material. The energy to failure is often referred to as the modulus of toughness.

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3.10. Compression Testing

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1. Prepare a cylinder or rectangular parallelepiped of bone as described above. Try to ensure that the length of the specimen is about twice the diameter (see Note 6). 2. Place the sample on the lower anvil of the testing machine. As in the case of tensile testing, a universal joint must be used in the loading path (see Note 4). Alternatively, a steel plate with small indentation containing a ball bearing between the specimen and the upper loading anvil can be used; this allows for any slight non-parallelism between the two faces. 3. Accurate measurements of strain require the use of a displacement transducer, as for tension, but for most comparative studies the displacement of the cross head is probably adequate (see Note 5). 4. Lower the cross head of the machine until it is just in contact with the specimen. 5. Set the strain rate and any predetermined load or displacement limits as before. 6. Start the test, and stop the test when the chosen criterion is met (see Note 7). 7. Calculate stress and strain and plot stress as a function of strain as described above. Where testing is done between two anvils, there is often a region near the origin, where the slope of the curve starts small and increases rapidly before becoming approximately linear, resulting in a J-shaped curve. This toe region is an artefact of the method and is commonly found in this sort of testing both of natural and synthetic polymeric materials. Because it can be difficult to determine a straight portion of the curve or to fit a straight line to it, it is best to differentiate the curve to find the modulus as a function of strain. The elastic modulus can then be taken to be the peak value of this curve. 8. The yield stress may be defined as the stress at which the modulus starts to decrease. We used a reduction of 3% from its peak value to provide a defined reference point (8). 9. Energy to yield is found by integration of the area under the stress–strain curve up to the yield stress. Strains are ill-defined in this testing protocol because of the toe region. Embedding the ends of longer specimens and gripping as for a tensile test can eliminate this problem. Failure cannot usually be defined in the same way as for tensile testing because progressive crushing of the material often occurs rather than a fracture.

3.11. Bending

The method described here applies to testing whole bones, from a mouse for instance, but with small alterations could be used for regular sections cut from larger bones.

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1. Set the bending jig in the materials testing machine. 2. Adjust the spacing of the lower supports to be as close to the ends of the diaphysis as possible and centred on the moving anvil. 3. Place the bone in a stable position on the supports. For femur, the anterior side is usually upwards and for tibia the fibula insertion point is normally at the top. 4. Lower the cross head of the instrument so that the upper loading anvil is just in contact with the bone surface. 5. Set the displacement rate, any load or displacement limits, and the data collection rate. 6. Start the test. 7. The machine can be set to stop as soon as fracture occurs because at this point the load falls very rapidly, though not always to zero. 8. Load is plotted as a function of displacement and the stiffness, yield load and failure load, and energy to fracture may be found using the same criteria as used for stress–strain curves in the tensile and compression tests (see Note 8). 3.12. The Elastic Modulus of Cortical Bone

The elastic or Young’s modulus of cortical bone can be calculated from density and speed of sound measurements described below using equation (9): E = rc 2 . This provides a direct measure of the intrinsic or material stiffness of the bone and can be applied to small samples, such as mouse bone and suitably machined specimen from larger mammals.

3.13. Density of Bone by Archimedes’ Principle

This technique can be applied to both cortical and cancellous bones and is particularly applicable to hollow, porous, or irregularly shaped specimen. With mouse bone, it is difficult to obtain repeatable results because the bone’s weight is small and comparable with surface tension, evaporation, and excess water effects, which must all be controlled. 1. The density of the bone is measured by comparing the weight in air with the weight when suspended in a fluid, typically distilled water. We use a five-figure balance (Sartorius Gmbh, Göttingen, Germany) for all measurements. 2. The true weight of each bone is measured first by weighing wet in sealed containers to eliminate any loss of weight due to evaporation. Prior to weighing, excess water is removed from the bone surface by wiping on damp tissue and the marrow canal emptied of fluid by passing air through it using a syringe

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and needle. Bones are then placed in a pre-weighed microcentrifuge tube containing PBS and weighed. 3. Each bone is then weighed suspended in distilled water using a Sartorius 6080/60801 Specific Gravity Determination Kit. The bone is suspended by a fine wire that passes through the medullary canal. Larger specimens can be suspended in a basket. Buoyancy effects due to trapped air are eliminated by filling the medullary canal with fluid using a syringe and needle. A droplet of detergent (washing-up detergent is adequate!) is added to the fluid to reduce surface tension effects. 4. The fluid density is calculated by measuring the upthrust exerted on a glass bob provided as part of the specific gravity kit when suspended in the fluid and using the equation: U = V rf , where U is the upthrust (weight in air – weight in fluid), V the volume of the body immersed, and rf the fluid density. 5. Bone density is calculated from the formula: r=

rf m , m − ms

where r f is the fluid density, m the true weight or weight in air, and ms the weight suspended in fluid. All weights are taken in triplicate and averaged. Where one of the measurements is clearly different to the other two, it is discarded and a repeat measurement made. 3.14. Speed of Sound Measured Using Ultrasound

Thin sections of cortical bone were prepared for ultrasound testing as described in Subheading 3.4, step 3. 1. Measure the length of the bone section accurately using a digital micrometer, for example. 2. Measure the time taken for ultrasound pulses to traverse the sample. We use pulse-echo mode, so the same transducer is used to generate and receive ultrasound pulses and a second, identical but unconnected transducer used as a reflecting surface at the other bone surface, although any glass or metal block would serve just as well as long as the impedance is different to that of the bone. The bone surface must be coupled to the transducers ultrasonically using a fluid, such as water or ultrasound gel. In our arrangement, pulses of ultrasound, of 10 MHz frequency, are generated using a Panametrics Pulser– Receiver model 5052PR (Panametrics Inc, Waltham, MA, USA) with a V211BA transducer in pulse-echo mode. Ultrasound echoes are displayed on an oscilloscope (Hitachi V-665A, Tokyo, Japan) and the time between echoes measured using the oscilloscope’s internal callipers.

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3. The speed of sound is then calculated using the formula: c=

2d , t

where d is the section thickness and t the time measured between echoes (9). 4. The bone sections are subsequently rinsed in PBS to remove any adhering ultrasound gel. 3.15. Ashing

After density measurements have been made, the bone can be dehydrated and ashed to determine the gross composition in terms of water, organic, and mineral content. 1. Crucibles are prepared by washing and drying in a furnace (Carbolite, Sheffield) at 600°C for 24 h and allowed to cool in a desiccator and then weighed. This ensures that they are free from contamination and moisture. 2. Remove excess water from the bone surface and medullary canal, as for density measurement, and add to the pre-weighed crucibles to measure wet weight. 3. Heat at 100°C for 24 h in an oven (Hybaid, Teddington), allow to cool (in a desiccator), and reweigh to determine the dry weight. 4. Then, heat at 600°C for a further 24 h in the furnace and, after cooling, weigh again to determine the ash weight. The bones are cooled in a desiccator to prevent any rehydration. 5. Water content is found from wet weight – dry weight, the organic content from dry weight – ash weight and mineral content as the ash weight (10). Measurements are then expressed as a percentage of the wet (or sometimes dry) weight to allow comparison between bones of different sizes.

4. Notes 1. Storage of samples: Samples should be kept moist with PBS prior to mechanical testing; formalin and other fixatives must never be used as this affects the mechanical properties. If the samples are not being tested immediately (within 2–4 h), the specimens can be stored frozen at −20°C. They must be wrapped in tissue or gauze dampened with PBS and vacuum sealed into plastic freezer bags. A vacuum bag sealer can be obtained for this purpose. Freezing is reported to have no effect on the mechanical properties of bone (11, 12). 2. Instruments for precision cutting: We use an Accutom 2 mineralogical saw (Struers Ltd., Rotherham) and a 125-mm

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aluminium oxide wheel, but the most common saw cited in the bone literature is the Isomet (Buehler, Lake Bluff, IL, USA). Whatever saw is used, faster wheel speeds and slower specimen feed result in a better surface finish. Where samples of a predetermined size are required, it is important to know how much of the sample is lost during the cutting procedure. Any saw removes an amount of material corresponding to the width of the blade, termed the kerf. For an aluminium oxide wheel, the kerf is about 0.5 mm compared with about 0.3 mm for a diamond blade. When the kerf is known and the machine is fitted with a micrometer or other precision positioning device, it is possible to obtain samples of the desired size by adjusting the cutting position appropriately. It is possible to cut specimens 100–150-mm thick using an aluminium oxide wheel. Care must be taken to keep specimen feed speeds, i.e. the rate at which the specimen is moved past the cutting wheel, fairly low, as aluminium oxide wheels are brittle and shatter easily and diamond wheels are thin and are easily bent. However, aluminium oxide wheels cost about £10, in contrast to about £200–300 for a diamond cutter, depending on size. 3. Obtaining bone cores: When coring right through a block of tissue, the sample is commonly left in the coring bit. Occasionally, this can happen, too, if a core snaps off while being drilled. In order to be able to remove this core, we have had the bits made with a hole from end to end so that a close fitting plunger may be inserted to push out the core. However, this can damage the top of the core if it is tightly wedged into the drill bit. For this reason, we prefer not to drill right through and to use the break-off tool to snap the core from its base after the coring bit has been removed. 4. Tension testing: The specimen must be large enough to get consistent results; Keaveny’s methods start with a specimen 40-mm long, which highlights the difficulties that may sometimes be encountered obtaining suitable specimens (13). If the grips are not exactly in line or the specimen is not precisely machined, then applying a tensile load will also result in a degree of bending. This adds uncertainty to the test data which is different for each specimen and makes accurate comparison between specimens impossible. A universal joint must be placed in the path of the load to ensure that only tensile forces are transmitted to the specimen. These are available from the instrument manufacturers. 5. Measuring strain during tension tests: An estimate of the strain can be calculated from the displacement of the cross head recorded by the machine. However, for accurate studies, a strain gauge or displacement transducer must be applied to the central, often waisted, part of the specimen. This can be done using either a clip-on device or video recording methods.

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6. Shape of samples for compression testing: It is desirable for the length of the specimen to be approximately twice the diameter. It is possible to conduct tests on samples, where these dimensions are roughly equal without too much of a problem and in many cases this may be unavoidable due to anatomical considerations. Boundary conditions, e.g. friction between the sample and the loading platens, can become a problem if the sample is not longer than it its width. 7. Setting limits for compression testing: Care needs to be taken when setting applied loads and limits into certain testing machines. Conventionally, engineers use positive numbers to denote tensile loads and negative numbers for compressive loads. The most commonly used test is tension and here there is generally no problem as everything is positive. However, setting a limit for a compressive load often needs to include a negative sign. It sounds trivial but is easily overlooked, and many students have stood watching the machine crush their specimen wondering why the machine has not stopped! Only after hitting the emergency stop button have they realised that they omitted the negative sign in front of the load limit they had set. There is often no clear end point for failure in compression, unlike tension, and determining when to stop the test has to be decided beforehand. In our studies, we have decided that any reduction in stiffness, that is a reduction in the slope of the load–displacement curve, is a sign of the beginning of failure. As we usually wish to do further analysis on the specimens and minimise the damage caused by the test, we watch the load–displacement curve (which is plotted on the computer monitor during the test) carefully during the test, and stop the test as soon as it becomes obvious that the slope of the curve is decreasing. 8. Calculating intrinsic properties of bone from bending tests: Equations have been derived to calculate intrinsic properties from these data but are fraught with difficulties when testing whole bones because of the asymmetric cross section of the bone. It is not recommended to use these without expert advice. References 1. Aspden, R. M. (1990) Constraining the lateral dimensions of uniaxially loaded materials increases the calculated strength and stiffness: application to muscle and bone. Journal of Materials Science: Materials in Medicine 1, 100–104. 2. Bryce, R., Aspden, R. M., Wytch, R., and Langrana, N. A. (1995) Stiffening effects of

cortical bone on vertebral cancellous bone in situ. Spine 20, 999–1003. 3. Gibson, L. J. (1985) The mechanical behaviour of cancellous bone. Journal of Biomechanics 18, 317–328. 4. Gibson, L. J., and Ashby, M. F. (1988) Cellular solids, Pergamon Press, Oxford.

35 5. Cowin, S.C. (2001) Bone mechanics handbook, CRC Press, Boca Raton. 6. Turner, C. H., and Burr, D. B. (2001) Experimental techniques for bone mechanics, in Bone mechanics handbook (Cowin S.C. ed.) CRC Press, Boca Raton, pp. 7-1-7-35. 7. Spatz, H. -Ch., O’Leary, E. J., and Vincent, J. F. V. (1996) Young’s moduli and shear moduli in cortical bone. Proceedings of the Royal Society B: Biological Sciences 263, 287–294. 8. Li, B., and Aspden, R. M. (1997) Composition and mechanical properties of cancellous bone from the femoral head of patients with osteoporosis or osteoarthritis. Journal of Bone and Mineral Research 12, 641–651. 9. Lees, S., Heeley, J. D., and Cleary, P. F. (1979) A study of some properties of a sample of bovine cortical bone using ultrasound. Calcified Tissue International 29, 107–117. 10. Mkukuma, L. D., Imrie, C. T., Skakle, J. M. S., Hukins, D. W. L., and Aspden, R. M.

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(2005) Thermal stability and structure of cancellous bone mineral from the femoral head of patients with osteoarthritis or osteoporosis. Annals of the Rheumatic Diseases 64, 222–225. 11. Nazarian, A., Hermannsson, B. J., Muller, J., Zurakowski, D., and Snyder, B. D. (2009) Effects of tissue preservation on murine bone mechanical properties. Journal of Biomechanics 42, 82–86. 12. van Haaren, E. H., van der Zwaard, B. C., van der Veen, A. J., Heyligers, I. C., Wuisman, P. I., and Smit, T. H. (2008) Effect of long-term preservation on the mechanical properties of cortical bone in goats. Acta Orthopaedica 79, 708–716. 13. Keaveny, T. M., Guo, E., Wachtel, E. F., McMahon, T. A., and Hayes, W. C. (1994) Trabecular bone exhibits fully linear elastic behavior and yields at low strains. Journal of Biomechanics 27, 1127–1136.

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Chapter 36 Mechanical Stimulation of Bone Cells Using Fluid Flow Carmen Huesa and Astrid D. Bakker Abstract This chapter describes several methods suitable for mechanically stimulating monolayers of bone cells by fluid shear stress (FSS) in vitro. Fluid flow is generated by pumping culture medium through two parallel plates, one of which contains a monolayer of cells. Methods for measuring nitric oxide production by bone cells in response to FSS are also described. Key words: Mechanical stimulation, Fluid flow, Shear stress, Cell culture

1. Introduction The skeleton provides mechanical support, in order to withstand the force of gravity, and supports muscle forces, allowing movement. This function is secured by the constant adaptation of bone to its mechanical loading environment, which results in bones that combine a proper resistance to fracture with a minimal use of material. Loading-induced deformations (strains) in bone matrix are believed to be too small to directly stimulate cells within bone; therefore, it has been hypothesised that fluid flow over the network of osteocytes embedded within the bone is the main signal-generating factor (1, 2) and that osteocytes are the main cell type involved in producing a response (3–5). The flow of extracellular tissue fluid as a result of mechanical loading of bone has been shown experimentally by Knothe Tate et al. (6, 7). Importantly, (static) loading regimes that do not elicit fluid flow through the lacuno-canalicular network do not seem to be osteogenic (8), whereas the most osteogenic-loading regimes are those that are dynamic, elicit high strain rates, and thereby presumably result in strong flow of interstitial fluid in bone (9).

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_36, © Springer Science+Business Media, LLC 2012

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How fluid flow results in a cellular response is not clear. Fluid flow produces fluid shear stress (FSS) over the osteocyte membrane that could stimulate the osteocytes, similar to the mechanism of stimulation of endothelial cells (6, 10, 11). It has been predicted that fluid-induced shear stresses over the cell extensions of osteocytes in vivo will be in the order of 0.8–3 Pa (12). However, the accuracy of these calculations is currently under debate and it has not been determined which cellular features are most important for the function of the osteocyte as mechanosensor. Currently, there are three different models/hypotheses that try to explain how osteocytes sense the fluid flow-derived mechanical stimulus, involving different anatomical features of the cell: (a) osteocyte cell extensions, (b) cell body, and (c) cilium. You et al. (13) have described a mathematical model in which the attachments of the osteocyte cell extensions to the extracellular matrix are described as tethering elements, protein filaments of unknown composition, believed to attach the cytoskeleton to the bone matrix. The mathematical model predicted that fluid flowinduced drag forces on the tethering elements will amplify the strains on the bone matrix by a factor of 10 (11). Recently, tethering elements have been suggested to be integrins (14). It has been suggested that the same FSSs acting on the cell processes would also act on the cell body (15), although this is refuted by Anderson et al. (16). Most experimentation with fluid flow is conducted with parallel plate flow chambers, where fluid flows over the cell body, rather than through canaliculi. Such flow systems indeed induce cellular responses, supporting the notion that the cell body is able to detect mechanical stimuli. Whether this is also the case in vivo remains unresolved. In a multicellular organism, cilia are the equivalents of bacterial flagella. Primitive flagella are thought to have worked as signal receivers and sensors of the surrounding environment. Growing evidence suggests that cilia might be undertaking a similar role in mammalian cells by acting as a mechanosensory organelle (17). Cilia have recently been identified in osteoblasts and osteocyte-like cells in vitro (18) and their possible role in mechanosensing is currently under investigation. The paragraphs above demonstrate that a lot remains to be investigated in mechanosensing at the cellular level. It is important to remember that bone cells cultured in a monolayer on a flat surface may not respond to a controlled mechanical stimulus in the same way as they might do in vivo, in a three-dimensional environment. Application of fluid flow to monolayers of cells on a flat substrate results in maximum displacements at the apical surface of the cell body (19). In contrast, in vivo, fluid flow will likely only occur over the cell extensions (16). Keeping in mind these limitations, we also want to emphasise the many insights that have been gained using in vitro systems for application of mechanical stimuli

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to cells and in particular systems that allow stimulation of cultured bone cells using a controlled FSS. This chapter, therefore, describes in vitro methods to study the effects of fluid flow on bone cells. Although osteocytes are the likely mechanosensors of bone, primary osteocytes are not easy to obtain in high numbers and osteoblasts are often used instead of osteocytes, even though they are less sensitive to a mechanical stimulus than osteocytes (3). Primary osteoblasts can be obtained as described in various chapters in this volume. The use of wellcharacterised mechanosensitive cell lines (e.g. MC3T3-E1 osteoblasts or MLO-Y4 osteocytes) also has its place in such studies and is described elsewhere (see Chapter 6, this volume).

2. Materials 2.1. Tissue Culture Media and Solutions

1. Alpha modified Eagle’s medium (a-MEM; 22571, Gibco, Paisley, UK), supplemented with 10% heat-inactivated foetal calf serum (FCS), 2 mmol/l L-glutamine, 100 IU/ml penicillin, and 50 mg/ml streptomycin (culture medium) for the maintenance of osteoblast-like cell lines, such as MC3T3-E1. If using other cell types, change the contents of the medium according to the requirements of that specific cell type. 2. a-MEM with 2% FCS, 2 mmol/l L-glutamine, 100 IU/ml penicillin, and 50 mg/ml streptomycin (flow medium) to be used for the fluid flow experiments. 3. Dulbecco’s modified Eagle’s medium (D-MEM) without phenol red (GIBCO, Paisley, UK). 4. Hanks’ balanced salt solution (HBSS) without calcium and magnesium. 5. Sterile phosphate-buffered saline (PBS), pH 7.4. 6. Trypsin–tetrasodium ethylenediamine tetraacetic acid (EDTA) solution consisting of 0.25% trypsin and 1 mM EDTA 4 Na. 7. 70% alcohol. 8. 0.15 mg/ml rat tail collagen type I in glacial acetic acid. 9. Cell permeable DAR-4M AM chromophore Nottingham, UK) for live imaging of nitric oxide.

(Axxora,

10. Griess reagent for determination of nitric oxide production. Solution A: 2% sulfanilamide and 5% phosphate in water. Solution B: 0.2% naphthylethylene diamine HCl in water. Store the solutions separately in a refrigerator and protected from light. 11. 0.1 M NaNO2 in water (stock solution) for quantification of nitric oxide production.

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12. ATP Bioluminescence assay kit HSII (Roche, UK) and Prostaglandin E2 EIA system (Amersham). 13. TaqMan® Gene Expression Cells-to-CT™ Kit (Applied Biosystems). 2.2. Instruments

1. 25- and 75-cm2 tissue culture flasks. 2. 15- or 25-ml polypropylene centrifuge tubes, 50-ml centrifuge tubes (Falcon), and 94 × 16-mm cell star Petri dishes. 3. Pasteur pipette. 4. 5-, 10-, and 50-ml syringes and 27-G½ needles. 5. Medical 3-way Connecta® stopcock (BD Medical, Oxford, UK) to be used as a pressure-release valve (Fig. 1a). 6. Bubble trap: We designed our own out of autoclavable materials (Delrin, Cole-Parmer, USA) with two quick release fittings

Fig. 1. Adaptation of peristaltic pump to produce oscillating flow. (A) Displacement of the syringe (D), (B) rotating head, (C) lever connected to A, (D) 5-ml syringes set up back to back. (E) Reference 0 point. (F) Radius of the pump (rp ) in mm that provides the displacement of the syringe.

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on either side, limiting the reservoir to 1–2 ml of medium (Fig. 1b). 7. Platinum-cured silicone tubing to connect the flow chamber to the pump and/or medium reservoir. Silicone tubing is flexible and cell friendly, and is available from a wide variety of stores. We use MasterFlex® (Cole-Parmer, USA). 8. Quick release fittings for tubing (Cole-Parmer, USA). 2.3. Pumps

An array of pumps is available in a range of prices. For simple fluid flow experiments, a peristaltic pump is a cheap option that is easy to keep sterile and is durable.

2.3.1. Peristaltic Pump to Create Pulsatile Flow

Most peristaltic pumps work well. We use Masterflex L/S with easy load head (Cole-Parmer, USA). The revolving speed and the number of pins in the pump head of a peristaltic pump determine the frequency of the pulse. The flow rate is determined by the speed at which the pump head revolves and the inner diameter of the tubing used.

2.3.2. Peristaltic Pump Modified to Create Oscillating Flow

We have built a pump head that fits two syringes that are coupled back to back in order to produce an oscillating flow (Fig. 2). The advantage of this system is that it causes minimal changes in hydrostatic pressure. The size of the syringe can be changed as well as the diameter of the radius in order to vary the flow rates and thereby the shear stresses exerted on the cells (see Note 4).

Fig. 2. Aberdeen live imaging fluid flow chamber. (a) Top. (b) Side. (c) Front.

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2.3.3. Syringe Pump to Create Oscillating Flow

Several models of syringe pumps are commercially available (e.g. Harvard Apparatus, Holliston, MA). The advantage of using commercially available pumps is that they are pre-programmed and therefore the flow rates do not have to manually calculated.

2.3.4. Micro-Annular Gear Pump

For more demanding experiments, a computer-controlled microannular gear pump (HNP Mikrosysteme GmbH, Parchim, Germany) can be used. This pump allows the creation of a defined pulsatile fluid flow, oscillating fluid flow, or steady fluid flow. The disadvantages of this machine are the relatively high cost and the sensitivity for macromolecular proteins in the medium (use of serum should be avoided).

2.4. Flow Chamber

Many flow chambers are available, both commercially or custom made. Below, we offer four examples, but other designs are also possible. All chambers have in common that they provide a controlled laminar flow over the cells because the medium is forced to flow between two parallel plates. The shear stress induced by a specific flow is mainly determined by the flow rate of the culture medium and the width of the channel and the height difference between the two parallel plates. For details about the required dimensions of the chamber, see Note 2.

2.4.1. mSlide Chamber (Ibidi® GmbH, Munich, Germany)

The mSlide chambers are extremely suitable for live imaging during fluid flow, but due to the limited dimensions of the chambers they are less suited for experiments where the goal is to isolate total RNA from mechanically stimulated bone cells or quantification of production of secreted substances. The mSlide chamber is available in a variety of dimensions and with various substrate coatings. In all cases, the plastic microscope slide that forms the bottom side of the chamber is a polymer that allows for high-quality imaging. On top of the slide are two reservoirs connected by a straight channel in which cells are seeded. The dimensions of this channel affect the magnitude of the shear stress exerted on the cells (see Note 2). We generally use collagen IV-coated slides for bone cells.

2.4.2. FlexFlow™ Shear Stress Device (Flexcell, Hillsborough, NC)

Flexcell chambers can be used for live imaging, but also allow the culture of a large number of cells, which makes them suitable for RNA and protein extraction. The Flexcell flow chamber consists of a polycarbonate plate, with a vacuum channel on one face. Laminar parallel flow is provided in a flow channel formed by a rectangularsize gasket and a glass slide with adherent cell monolayer. The main characteristics of this chamber are that it is sealed by a vacuum and it is suitable for an upright microscope. Flexcell® also offers a variation on this chamber, the Streamer® Shear Stress Device that allows the stimulation of cells on 6 matrix-coated glass slides simultaneously. The streamer is suitable for experiments that require large amounts of protein lysate and/or RNA for further analysis and a protocol for its use is given in Chapter 6, this volume.

36 2.4.3. Aberdeen Live Imaging Fluid Flow Chamber

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The Aberdeen Live Imaging Fluid Flow (ALIFF) chamber is custom made and suits an upright microscope. It is similar to the FlexFlow™ chamber, but requires screws rather than vacuum to stay closed and is hence less prone to leakage. The bottom of the ALIFF chamber is custom made from microscope slides or tissue culture flasks to fit the 25 × 49-mm hold at the bottom of the chamber. The bottom slide is separated from the slide containing monolayer of cells with a 0.75-mm thick Press-to-Seal™ silicone sheet (Invitrogen, UK). The design of the chamber is shown in Fig. 3.

Fig. 3. (a) Picture of a fluid flow set-up for quantification of signalling molecule production by bone cells. (b) Schematic drawing of the fluid flow set-up depicted in (a). (c) Filling the Amsterdam flow chamber with cells. (d) Placing of the glass slide with a monolayer of cells, facing down, in the Amsterdam chamber. Note that the polycarbonate plate is on the bottom during assembly while it is on top during the fluid flow experiments. (1) Peristaltic pump, (2) Amsterdam flow chamber, (3) glass reservoir, (4) inlet for CO2, (5) flow meter, (6) glass slide containing cells, (7) polycarbonate plate, and (8) aluminium lid of the Amsterdam flow chamber.

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Fig. 4. Culture of cells for ALIFF chamber. (a) Press-to-Seal™ silicone sheet (Invitrogen, UK) was cut to dimensions specified in Fig. 3 to limit the area of cell growth to 56 × 13 mm. (b) Spacers to lift glass slide from Petri dish floor to avoid adherence of the glass slides to the Petri dish. (c) Area of cell growth.

2.4.4. Amsterdam Fluid Flow Chamber

The Amsterdam flow chamber is custom made and contains a glass slide (25 × 65 mm) containing bone cells that serves as the bottom plate. The top plate is made in polycarbonate and contains the medium inlet and outlet, a bubble trap, a rectangular flow area, and gasket (Fig. 4d). The glass slide is secured to the top plate using an aluminium lid and screws. The advantage of this chamber is that it is easy to use and durable; the disadvantage is that it cannot be used for live imaging.

2.5. Incubator

37°C with 5% CO2 or, alternatively, an air box with a high-throughput blower and a thermostatic regulator.

2.6. Microscope

Fluorescence microscope (i.e. Zeiss Axioskop with an MRm camera and AxioVision software) or deconvolution microscope (API DeltaVision, CA, USA) able to excite at 530–550 nm, and read emission at 580 nm.

3. Methods 3.1. Preparation of Cells

Normal techniques for working under sterile conditions (use of sterile media and instruments and working in a flow cabinet) should be adhered to. 1. Wash osteoblasts in T75 flasks with PBS before adding 2 ml of trypsin–EDTA. 2. Incubate cells in trypsin–EDTA for 3–5 min. Make sure that all cells are loose by tapping the flask lightly on the side.

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3. Add 8 ml of culture medium to the cells and transfer to a 15-ml Falcon tube. Reduce quantities by half when using a T25 flask. 4. Centrifuge the cell suspension at 1,000 × g for 5 min. 5. Discard supernatant, resuspend cells in 1 ml culture medium, and count the cells. 3.2. Seeding of Cells

Methods are given for the various types of flow chambers discussed in Subheading 2.4.

3.2.1. mSlide Chambers

1. Seed between 2 × 103 and 8 × 103 cells/cm2 for live imaging. The total number of cells needed per chamber depends on the surface area of the channel. Dilute the total number of cells needed per chamber in the exact volume of medium needed to fill the channel of the chamber (e.g. a mSlideI chamber fits 100 ml in the channel). 2. Pipet the cell suspension into one side of the channel, so it fills the channel. Add the exact volume of medium to the mSlide chamber as specified by manufacturer. 3. Incubate for 1 h at 37°C + 5% CO2 to allow the cells to attach. 4. Fill up the reservoirs on the side of the chamber with culture medium. 5. Incubate overnight at 37°C with 5% CO2 before starting the fluid flow experiment.

3.2.2. Seeding of Cells on Glass Slides (FlexFlow™ Chamber/ALIFF Chamber/ Amsterdam Fluid Flow Chamber)

1. Choose glass slides that fit the dimensions of the fluid flow chamber used. Sterilise the slides with 70% ethanol. Dry the slides thoroughly and keep sterile. 2. Place a glass slide in a Petri dish. Two sterile spacers made of silicone can be placed between the glass slide and the bottom of the Petri dish to prevent adherence of the glass slides to the Petri dish, but make sure that the glass slide is level. 3. Pipet 1 ml of rat tail collagen type I per 1.4 cm2 and incubate for an hour at room temperature. Wash three times in sterile PBS to remove acid and unbound collagen. Glass slides can be stored for several days in PBS. Before seeding the cells, the glass slides can be sterilised again by exposure to UV light for 30 min. Let the glass slides air dry before seeding the cells. 4. Release the cells from the T75 flask using trypsin as described above in Subheading 3.1, step 2. We recommend to seed between 1.3 × 104 and 3 × 104 cells/cm2 for isolation of cDNA, isolation of protein, or quantification of secreted molecules by mechanically stimulated cells in the medium. Suspend the cells in medium, and use 50 ml of medium per cm2 glass slide.

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Fig. 5. Experimental fluid flow circuits. (a) Live imaging set up for pulsatile fluid flow. (b) Live imaging set up for oscillating fluid flow. Description of parts (1) pump/motor, (2) inlet, (3) outlet, (4) reservoir, (5) parallel plate chamber, (6) bubble trap, (7) valve, and (8) flow sensor.

5. Seed cells only on the exact area of the glass slide that is exposed to the laminar fluid flow when placed inside the flow chamber. An easy way to achieve this is by drawing the circumference of the glass slide on a piece of paper and marking the area that should be seeded with cells on this drawing. The sheet of paper can then be placed underneath the Petri dish as a visual guide. Trace the outline of the marked area while slowly releasing the cell suspension from a pipet tip onto the glass slide. Then, gently fill up the rest of the marked area with the remainder of the cell suspension. The goal is to end up with a rectangular “drop” of medium containing the cells on the glass slide that matches the area that is exposed to a laminar fluid flow. Alternatively, cut a press-to-seal silicon sheet to fit the desired dimensions and carefully adhere to glass slide before seeding the cells (see Fig. 5). 6. Let the cells adhere for 1–1.5 h at 37°C with 5% CO2. If the medium volume used to seed the cells is relatively small, 2–3 ml of culture medium (without cells) can be added to the sides of the Petri dish to prevent evaporation. 7. Fill the Petri dish with enough medium to submerge the cell monolayer. 8. Incubate overnight at 37°C with 5% CO2. 3.3. Assembly of the Flow Apparatus

Make sure that all materials are sterilized before use. Use 70% ethanol, Virkon, UV light, or autoclave materials (not suitable for polycarbonate, but suitable for polyoxymethylene plastic also known as Delrin, DuPont, USA).

3.3.1. Pulsatile Flow, Online Monitoring of Cell Responses

Here, we describe how to apply a pulsatile flow to cultured bone cells using the apparatus set up as shown in Fig. 6a. It consists of a peristaltic pump, a pressure valve, a mSlide chamber, and a bubble

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Fig. 6. Parallel plate fluid flow chamber with steady flow. Diagram based on Frangos et al. (26). The flow movement happens through gravity. The speed of flow is determined by the distance between the top and the bottom reservoir. (1) Pump/motor, (2) inlet, (3) outlet, (4) reservoir, and (5) parallel plate chamber, where cells are plated. The distance between the top and the bottom reservoir (8) determines the pressure at which the cells would be in the chamber, but there is no drop of pressure across the chamber.

trap connected with T14 silicone tubing. The amount of medium required to fill this system is 10 ml. 1. Make fresh flow medium before assembling the fluid flow system and keep warm. 2. The flow chamber should be already filled with media, as described in Subheading 3.2.1, step 4. 3. Prepare inlet tubing. This consists of silicone tubing, a pressure valve (make sure that it stays closed), and tubing with the proper connector to the chosen chamber. This tubing should be long enough to connect later to the chamber and the bubble trap and be placed into the peristaltic pump. 4. Prepare the outlet tubing, which consists of tubing connected to an opened bubble trap with the appropriate fittings. Connect the tubing to the chamber and the bubble trap before starting to fill up the system. 5. With a 50-ml syringe, completely fill the inlet tubing. 6. Connect the filled inlet tubing to the chamber inlet avoiding bubble formation. If tubing were to be filled while connected to the mSlide chambers, the air would push the fluid out of the

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chamber and introduce air bubbles in the chamber. Once this happens, it is very difficult to fill up the chamber with medium. 7. Continue to fill up the system (chamber, tubing, and bubble trap). Then, completely seal the bubble trap. 8. With care, remove the syringe from the inlet tubing and put in a quick-release CPC coupling insert to attach the tubing to the bubble trap, creating a closed loop. 9. Lock the chamber onto the microscope stage (preferably keep warm by using a heated stage). Set up the pump at the same height as the microscope stage. Place tubing connected to the inlet of the chamber into peristaltic pump head (do not lock the pump head yet). Open the pressure valve. Lock the pump head. Let the cells rest for 15 min before the start of the experiment. 10. Turn on the pump. Adjust the flow rate by changing the speed of the peristaltic pump. It is not recommended to run an experiment for more than 2 h using this system. 3.3.2. Oscillating Fluid Flow, Online Monitoring of Cell Responses

The system for oscillating flow does not require the use of a bubble trap or a pressure-release valve. It requires a peristaltic pump with an adapted pump head to fit two syringes back to back (Fig. 6b). Be aware that creating an oscillating flow might induce turbulent flow at high flow rates. 1. Prepare fresh flow medium before assembling the fluid flow system and keep warm. 2. Prepare the pump to produce the specific amount of shear required (see Note 4). 3. Change medium in chamber by tilting the chamber to one side and emptying the bottom reservoir, and then filling the top reservoir. Repeat twice. Leave chamber flat and fill both reservoirs with medium. 4. Connect 50 cm of small-diameter silicone tubing to a 5-ml syringe filled with medium. Fill the tubing completely with medium using the syringe. Keep the syringe connected. 5. Repeat step 4. 6. Fit the syringes in the pump head syringe holder. Be careful not to introduce bubbles in the tubing. Excess medium comes out from the tubing, so prepare a beaker for collection. 7. Connect one piece of tubing to the inlet of the chamber, and the other piece of tubing is connected to the outlet of the chamber. Avoid trapping air bubbles in the system. 8. Lock the chamber onto the microscope stage (preferably keep warm). Set up the pump at the same height as the microscope stage. 9. Turn on the pump. Adjust the flow rate by changing the speed of the peristaltic pump (see Note 4).

36 3.3.3. Pulsatile Fluid Flow, Isolation of RNA or Protein

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The following set-up is suitable for simultaneously stimulating a greater number (up to 5 × 105) of cells to a pulsatile fluid flow. In combination with the relatively low amount of medium required in the flow loop (in the example below, we use 12 ml) and the possibility of sampling medium during the experiment, this set-up is extremely useful for measuring the secretion of molecules by the cells. The system consists of a peristaltic pump, the Amsterdam flow chamber, and a medium reservoir connected with silicone tubing (1.6-mm inner diameter). Although we generally use a custom-made glass reservoir, a simple 50-ml Falcon tube can also serve as a medium reservoir, as described below. 1. Make fresh flow medium before assembling the fluid flow system and keep warm. Make approximately 15 ml per glass slide. 2. Make four holes (6-mm diameter) in a lid of a 50-ml Falcon tube. Sterilise the lid using 70% alcohol. 3. Heat the incubator to 37°C after thoroughly cleaning with 70% alcohol. 4. Fix a sterile 50-ml Falcon tube with holes in the lid in an upright position within the incubator. Push one end of 70 cm of silicone tubing through one of the holes in the lid until it almost reaches the bottom of the Falcon tube, and then fix the rest of that tubing in the pump head. 5. Push one end of a second 40-cm-long piece of silicone tubing through one of the empty holes in the lid. This end does not need to reach the bottom. 6. Connect the system providing humidified 5% CO2 in air to the Falcon tube via one of the remaining two empty holes in the lid. 7. Fill the Falcon tube with 9.5 ml of flow medium using a 10-ml syringe and a needle, and use the pump at a low speed in order to completely fill the 70-cm silicone tube attached to the pump. 8. Inside a laminar flow hood, attach one end of 15 cm of silicone tubing to the entrance port of the polycarbonate top plate of the Amsterdam flow chamber (Fig. 4), and the other end to the exit port. Fill the bubble traps, tubing, and rectangular flow area completely with 2.5 ml of flow medium using a 5-ml syringe and a 27-G½ needle. 9. Put the glass slide containing the cells, with the cells facing down, on top of the medium in the rectangular flow area. Make very sure that no air is trapped underneath the glass slide. Position the metal lid over the glass slide and fix the glass slide in place using screws. Turn over the chamber, so the metal lid and glass slide form the bottom of the chamber while the polycarbonate part serves as the top.

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10. Bring the camber to the incubator, remove the 15-cm silicone tube from the inlet of the chamber (squeeze the end shut to prevent medium from leaking out of the chamber), and connect the medium-filled tube on the pump to the inlet of the chamber. Avoid trapping air bubbles. 11. Remove the 15-cm silicone tube from the outlet of the chamber and connect the outlet to the Falcon tube using the 40-cm silicone tube described in step 5. Lay down the flow chamber at equal height as the pump on a level surface. Make sure that the glass slide serves as the bottom. If the assembly of the system is done gently, only a minimal amount in mechanical stimulation of the glass slide will have occurred. We advise to start the experiments shortly after assembly of the system as only a very limited amount of medium is available to the rather large amount of cells positioned within the flow chamber. 12. Turn on the pump. Adjust the flow rate by changing the speed of the peristaltic pump. Medium samples can be taken from the Falcon tube during the experiment (e.g. by using a Pasteur pipette). 13. As static control for the mechanically stimulated cells, a glass slide with the same number of cells attached is placed into a Petri dish containing 12 ml fresh flow medium and incubated for the same amount of time. 3.3.4. Steady Fluid Flow

The apparatus shown in Fig. 7 can be used for subjecting cells to a steady flow of fluid and allows sampling of medium during the flow experiment. However, the relatively large volumes of medium required to run this system dilute the secreted molecules produced by the stimulated bone cells, and therefore large quantities of cells should be used. The flow loop consists of two reservoirs, situated one above the other with a flow chamber (e.g. the Amsterdam chamber) between them. The continuous pumping of the culture medium from the lower to the upper reservoir, at rates greater than those through the chamber, maintains the flow loop. 1. Make fresh culture medium before assembling the fluid flow system and keep warm. 2. Fill the chamber as described in steps 7 and 8 of Subheading 3.3.3. 3. Add medium to the top reservoir (10–20 ml), filling the lower reservoir as well and the tubing to connect to the chamber. Once the tubing is filled with medium, connect the tubing to the chamber and let the medium flow to the lower reservoir. 4. Adjust the FSS by changing the column height in the loop. Recirculation minimises the amount of flow medium required, allowing easy quantification of secreted molecules, as they are present in medium samples in high concentrations.

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Fig. 7. Fluorescence recorded from single cells caused by production of nitric oxide (NO) from images taken during NO imaging experiment on wild-type pOBs. Each line indicates to the fluorescence of one cell plotted against time. (a) Representation of all cells with fluorescence in a field of view. Some show the signal to be saturated before changes after flow can be measured. Others show no response. Half of the cells in this figure show a response to fluid flow. This response is monitored by a steeper NO synthesis rate immediately after flow. (b) Comparison of NO increase rates before and during PFF of the cells represented in red and blue colours in (a). Linear curves were fitted as f = y0 + ax. The correlation values were all r2 = 0.99. The curves after PFF are steeper, indicating that NO production is increased.

3.4. Analysis of the Cellular Response to Fluid Flow

Mechanical forces on the osteocyte trigger a series of chemical responses. An important early response to mechanical loading is the influx of calcium ions through ion channels in the plasma membrane and the release of calcium from internal stores (20, 21). The rise in intracellular calcium concentration activates many

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downstream signalling cascades, such as protein kinase C and phospholipase A2, and is necessary for activation of calcium/calmodulindependent proteins, such as nitric oxide synthase. The activation of phospholipase A2 results, among others, in the activation of arachidonic acid production and PGE2 release (22). Genes whose expression in osteocytes is modified by mechanical loading include c-fos, RANKL, osteoprotegerin, and MEPE (23). We commonly use the Prostaglandin E2 EIA system from Amersham which allows determination of prostaglandin E2 concentrations between 1 and 6,400 pg/ml in a 96-well plate format. For determining ATP concentrations in the medium, we found the ATP Bioluminescence assay kit HSII from Roche to perform best. Quantification of nitric oxide production is described below. 3.4.1. Online Monitoring of NO Production

1. Prepare and warm up medium: D-MEM without phenol red supplemented with 1% FCS, 2 mmol/l L-glutamine, 100 IU/ ml penicillin, and 50 mg/ml streptomycin. 2. Incubate cells in chambers with 10 mM DAR–4 M AM chromophore in flow medium for 1–2 h at room temperature. Introduce chromophore in chamber in the same manner as in step 3 in Subheading 3.3.2, very slow to avoid stimulation of the cells that could be caused by the movement of fluid over the cell monolayer. 3. After incubation, remove extracellular chromophore by slowly washing the cell monolayer twice in medium taking care not to stimulate the cells with the movement of the fluid over the cell monolayer. 4. Set up a flow system as described in Subheading 3.3.1. Mount the flow chamber on an inverted fluorescent microscope equipped with a rhodamine filter. 5. Start image capture. Exactly 1 min later, start the flow. Stop the image capture after 5 min. Images are recorded using a rhodamine filter (530–550 nm of excitation) at a rate of one image every 4 s. 6. Images are analysed using ImageJ. Each intensity in a gray spectrum is assigned a number between 0 and 255. Consequently, each pixel within a selected region of interest (ROI) is allocated a number. In this way, it is possible to obtain the gray value of an ROI (i.e. a cell) by expressing it as the average of the values in the ROI. 7. The first image is subtracted from the whole stack to eliminate background and innate fluorescence. The perimeter of a cell is selected (for cells visible for most of the stack). The mean signal intensity within that ROI is measured, and plotted as a function of time. This is done for each field of view and a curve is fitted for all the cells in a field of view. An example is given in Fig. 7. For information on how to quantify exact amounts of NO produced by the cells, see ref. 24.

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Samples of culture medium can be taken at any time point after initiation of the flow of fluid for analysis of secreted products by the mechanically stimulated cells. For rapid responses, such as NO production (and ATP release), medium can be sampled at 2–5 min after initiation of the mechanical stimulus. If multiple factors are analysed, make sure that separate aliquots of the samples are frozen to avoid repeated freezing and thawing cycles. Many methods are available to determine NO production in medium samples. Most of these methods are based on determining NO2− concentrations, since NO is a gas that quickly reacts with the culture medium to form NO2− (the half-life of NO is 14 s). A cheap and easy way is using Griess reagent, although this method is not sensitive enough to detect low amounts of NO produced. 1. Dilute the 0.1 M NaNO2 stock solution in culture medium (without serum) to make a standard curve of the following concentrations: 100, 50, 25, 10, 5, 2.5, and 1.25 mM. 2. Pipet 75 ml standard, blank, or unknown sample in each well of a 96-well plate. 3. Mix 4 ml Griess reagent part A with 4 ml Griess reagent part B immediately prior to use. Add 75 ml Griess reagent per well. 4. Incubate at room temperature for 15 min on a microplate shaker. 5. Measure the absorbance at 540 (up to 570) nm using a plate reader.

3.4.3 Cell Lysate/PCR

It is possible to obtain RNA from mSlide chambers by using TaqMan® Gene Expression Cells-to-CT™ Kit (Applied Biosystems). For all other chambers, use standard methods of RNA and protein extraction (see, for example, Chapter 17, this volume).

4. Notes 1. For most fluid flow experiments, it is recommended to eliminate the serum from the medium, unless experiments are running for over 12 h. In that case, serum is necessary for cell survival. Do use a source of large proteins in the medium, such as BSA or low serum (2% or less), because this is needed to keep the glycocalyx intact, a structure essential for the flow response. In studies designed to measure loading-induced PGE2 production, it is also advisable to add a small amount of serum to the medium to ensure that arachidonic acids, necessary for PGE2 production, are not rate limiting.

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2. The volumetric rate of flow (Q) required to produce a desired FSS (t wall ) at the cells may be obtained from the equation below: t wall =

6 mQ , bh 2

where b and h are the width and height, respectively, of the parallel plate flow chamber and m is the fluid viscosity. The FSS is, thus, strongly related to the dimensions of the parallel plate chamber used (25). 3. Reynolds (Re) number determines whether a fluid flow is laminar. The transition from laminar to turbulent flow starts at 1,000–8,000 Re. The Reynolds number can be calculated as Qr/(mb), where Q is the volumetric rate of flow, r is the fluid density, b is the width of the chamber, and m is the fluid viscosity. Although the Reynolds number is meant to be used for steady flow, it is valid for most fluid flow applications, where applied flow frequencies generally not exceed 10 Hz. On average, the majority of the surface of a parallel plate chamber is exposed to a homogenous wall shear stress as long as the width of the chamber is at least 20-fold bigger than the height. 4. An adaptor was designed for the peristaltic pump to enable it to drive syringes coupled back to back to produce oscillating flow (Fig. 2). The pistons of the syringes fit into a holder, A, which is constrained to move only back and forth as shown by the arrow. The rotating head drives this holder via an arm whose position from the centre of rotation can be adjusted to govern the maximum displacement, d. If the frequency or rotation is f, then the volume of fluid expressed at time t is given by: V (t ) = Asd × sin(2pft ) and the volumetric flow rate, Q, is the time derivative of this: Q=

dV = 2p fAs d × cos(2p ft ), dt

where t wall is the shear stress at the wall of the chamber, Q is the volumetric fluid flow, m is the fluid viscosity, b is the width of the chamber, h is the height of the chamber, rp is the radius in oscillating pump, As is the cross-sectional area of the syringe in the oscillating pump, and f is the frequency. 5. With the set-ups described in this chapter, we generally do not encounter problems with cell death or “shearing of cells”. Whether cells are removed by application of FSS can be easily assessed by visual inspection of the cultures before and after application of fluid flow. Also the total amount of protein or DNA

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in the monolayer of cells can be measured after application of flow, and this number can be compared to cells kept under static culture conditions. In addition, a number of cell viability assays is available, such as the Caspase-Glo 3/7 Assay (Promega, Madison, WI, USA). 6. When you are unsure what type of flow regime to use (steady, pulsatile, or oscillating), remember that it is unknown exactly which type of flow exists in vivo. It is, therefore, important to realise the limitations of each system and being clear in reporting results on the precise method used in the experimental design. Each flow regime produces different responses (8), and dynamic flow (pulsatile or oscillating) generally produces more responses than steady flow. If desired, the flow regime can be monitored by the introduction of a flow sensor to the system. This can be useful in order to monitor the exact flow rates (and thus shear stress) applied to the cells, and the waveform of the stimulus, especially when using peristaltic pumps, when the waveform is not always sinusoidal. References 1. Piekarski, K., and Munro, M. (1977) Transport mechanism operating between blood supply and osteocytes in long bones. Nature. 269, 80–82. 2. Cowin, S. C., and Weinbaum, S. (1998) Strain amplification in the bone mechanosensory system. Am. J. Med. Sci. 316, 184–188. 3. Klein-Nulend, J., van der Plas, A., Semeins, C.M., Ajubi, N. E., Frangos, J. A., Nijweide, P. J., and Burger, E. H. (1995) Sensitivity of osteocytes to biomechanical stress in vitro. FASEB J. 9, 441–445. 4. Klein-Nulend, J., Roelofsen, J., Sterck, J. G., Semeins, C. M., and Burger, E. H. (1995) Mechanical loading stimulates the release of transforming growth factor-beta activity by cultured mouse calvariae and periosteal cells. J. Cell Physiol. 163, 115–119. 5. Turner, C. H., Forwood, M. R., and Otter, M. W. (1994) Mechanotransduction in bone: do bone cells act as sensors of fluid flow? FASEB J. 8, 875–878. 6. Knothe Tate, M. L., Knothe, U., and Niederer, P. (1998) Experimental elucidation of mechanical load-induced fluid flow and its potential role in bone metabolism and functional adaptation. Am. J. Med. Sci. 316, 189–195. 7. Knothe Tate, M. L., Steck, R., Forwood, M. R., and Niederer, P. (2000) In vivo demonstration of load-induced fluid flow in the rat tibia and its potential implications for processes associated with functional adaptation. J. Exp. Biol. 203, 2737–2745.

8. Jacobs, C. R., Yellowley, C. E., Davis, B. R., Zhou, Z., Cimbala, J. M., and Donahue, H. J. (1998) Differential effect of steady versus oscillating flow on bone cells. J. Biomech. 31, 969–976. 9. Turner, C. H., Owan, I., and Takano, Y. (1995) Mechanotransduction in bone: role of strain rate. Am. J. Physiol. 269, E438–442. 10. Weinbaum, S., Guo, P., and You, L. (2001) A new view of mechanotransduction and strain amplification in cells with microvilli and cell processes. Biorheology. 38, 119–142. 11. Han, Y., Cowin, S. C., Schaffler, M. B., and Weinbaum, S. (2004) Mechanotransduction and strain amplification in osteocyte cell processes. Proc. Natl. Acad. Sci. USA 101, 16689–16694. 12. Weinbaum, S., Cowin, S. C., and Zeng, Y. (1994) A model for the excitation of osteocytes by mechanical loading-induced bone fluid shear stresses. J. Biomech. 27, 339–360. 13. You, L. D., Weinbaum, S., Cowin, S. C., and Schaffler, M. B. (2004) Ultrastructure of the osteocyte process and its pericellular matrix. Anat Rec. A. Discov. Mol. Cell Evol. Biol. 278, 505–513. 14. Wang, Y., McNamara, L.M., Schaffler, M. B., and Weinbaum, S. (2007) A model for the role of integrins in flow induced mechanotransduction in osteocytes. Proc. Natl. Acad. Sci. USA 104, 15941–15946. 15. Bonewald, L. F. (2007) Osteocytes as dynamic multifunctional cells. Ann. N Y Acad. Sci. 1116, 281–290.

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16. Anderson, E. J., Kaliyamoorthy, S., Iwan, J., Alexander, D., and Knothe Tate, M. L. (2005) Nano-microscale models of periosteocytic flow show differences in stresses imparted to cell body and processes. Ann. Biomed. Eng. 33, 52–62. 17. Whitfield, J. F. (2003) Primary cilium--is it an osteocyte’s strain-sensing flowmeter? J. Cell Biochem. 89, 233–237. 18. Xiao, Z., Zhang, S., Mahlios, J., Zhou, G., Magenheimer, B. S., Guo, D., Dallas, S. L., Maser, R., Calvet, J. P., Bonewald, L., and Quarles, L. D. (2006) Cilia-like structures and polycystin-1 in osteoblasts/osteocytes and associated abnormalities in skeletogenesis and Runx2 expression. J. Biol. Chem. 281, 30884–30895. 19. McGarry, J. G., Klein-Nulend, J., Mullender, M. G., and Prendergast, P. J. (2005) A comparison of strain and fluid shear stress in stimulating bone cell responses--a computational and experimental study. FASEB. J. 19, 482–484. 20. Hung, C. T., Allen, F. D., Pollack, S. R., and Brighton, C. T. (1996) Intracellular Ca2+ stores and extracellular Ca2+ are required in the real-time Ca2+ response of bone cells experiencing fluid flow. J. Biomech. 29, 1411–1417.

21. Hung, C. T., Pollack, S. R., Reilly, T. M., and Brighton, C. T. (1995) Real-time calcium response of cultured bone cells to fluid flow. Clin. Orthop. Relat. Res. 256–269. 22. Ajubi, N. E., Klein-Nulend, J., Alblas, M. J., Burger, E. H., and Nijweide, P. J. (1999) Signal transduction pathways involved in fluid flowinduced PGE2 production by cultured osteocytes. Am. J. Physiol. 276, E171–178. 23. Kulkarni, R. N., Bakker, A. D., Everts, V., and Klein-Nulend, J. (2010) Inhibition of Osteoclastogenesis by Mechanically Loaded Osteocytes: Involvement of MEPE. Calcif. Tissue Int. 87, 461–8. 24. Vatsa, A., Mizuno, D., Smit, T.H., Schmidt, C.F., MacKintosh, F.C., and Klein-Nulend, J. (2006) Bio imaging of intracellular NO production in single bone cells after mechanical stimulation. J. Bone Miner. Res. 21, 1722–1728. 25. Bacabac, R.G., Smit, T.H., Cowin, S.C., Van Loon, J.J., Nieuwstadt, F.T., Heethaar, R., and Klein-Nulend, J. (2005) Dynamic shear stress in parallel-plate flow chambers. J. Biomech. 38, 159–167. 26. Frangos, J.A., McIntire, L.V., and Eskin, S.G. (1988) Shear stress induced stimulation of mammalian cell metabolism. Biotechnol. Bioeng. 32, 1053–1060.

Chapter 37 Using Cell and Organ Culture Models to Analyze Responses of Bone Cells to Mechanical Stimulation Andrew A. Pitsillides and Simon C.F. Rawlinson Abstract Bone cells of the osteoblastic lineage are responsive to the local mechanical environment. Through integration of a number of possible loading-induced regulatory stimuli, osteocyte, osteoblast, and osteoclast behaviour is organized to fashion a skeletal element of sufficient strength and toughness to resist fracture and crack propagation. Early pre-osteogenic responses had been determined in vivo and this led to the development of bone organ culture models to elucidate other pre-osteogenic responses where osteocytes and osteoblasts retain the natural orientation, connections and attachments to their native extracellular matrix. The application of physiological mechanical loads to bone in these organ culture models generates the regulatory stimuli. As a consequence, these experiments can be used to illustrate the distinctive mechanisms by which osteocytes and osteoblasts respond to mechanical loads and also differences in these responses, suggesting co-ordinated and cooperatively between cell populations. Organ explant cultures are awkward to maintain, and have a limited life, but length of culture times are improving. Monolayer cultures are much easier to maintain and permit the application of a particular mechanical stimulation to be studied in isolation; mainly direct mechanical strain or fluid shear strains. These allow for the response of a single cell type to the applied mechanical stimulation to be monitored precisely. The techniques that can be used to apply mechanical strain to bone and bone cells have not advanced greatly since the first edition. The output from such experiments has, however, increased substantially and their importance is now more broadly accepted. This suggests a growing use of these approaches and an increasing awareness of the importance of the mechanical environment in controlling normal bone cell behaviour. We expand the text to include additions and modifications made to the straining apparatus and update the research cited to support this growing role of cell and organ culture models to analyze responses of bone cells to mechanical stimulation. Key words: Bone, Mechanical load, Mechanical strain, Fluid shear

1. Introduction As bone’s primary function is mechanical, it is not surprising that almost all studies using intact bone concern its morphology. Such histomorphometric studies are used to provide insights into how bone responds, as an organ, to mechanical loading. Despite this, 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_37, © Springer Science+Business Media, LLC 2012

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the cellular basis for “sensing” mechanical stimuli or “communicating” their influence to coordinate any loading-induced changes that they engender is not known. Studies in intact bone are, however, rarely used to establish the direct links with any changes in bone cell biochemistry. It is also evident that most studies aimed at defining these mechanisms currently use bone cells grown in vitro, and that this has produced rapid advances in our understanding of the factors that might be involved in regulating bone cell responses to loading-induced stimuli. It is clear that such in vitro studies facilitate the mechanistic deciphering and constitute a useful initial approach. However, it is also evident that they generally take little regard of the influence that might be provided by cell–cell and cell–matrix interactions within a bone’s complex environment, architecture and morphology (1). It is, therefore, imperative to undertake studies aimed at bridging any gap between the cell biology of bone’s response to loading on the one hand, and the morphological approach to this same problem on the other. Here, we concentrate on techniques by which the cellular responses to mechanical stimulation can be examined in cell monolayer and in organ culture models. We provide details and discuss a wide range of devices that can be used to generate precise, measurable mechanical strains in these preparations and give details of those devices which we have used extensively in both organ and cell culture. We hope that this provides a basis upon which new investigators into this field can decide how to approach their own particular set of questions. In our opinion, these organ and cell culture models for studying bone responses to mechanical stimuli should continue to grow as a key aspect of any investigation into bone cell biology, because they take into account the essential load-bearing role of skeletal tissues. Thus, it may be considered appropriate that even those studies examining the effects on non-mechanical factors (e.g. PTH, LRP5, and Wnt signalling, oestrogen, and sclerostin) should ideally also be conducted in an environment in which skeletal tissues and/or cells derived from these tissues are concomitantly exposed to their normal or pathophysiological range of mechanical stimuli. As a corollary, it is therefore tempting to suggest that any such studies conducted in the absence of mechanical stimulation are for practical purposes examining responses to these non-mechanical factors in an environment effectively mimicking unloading. We first address some of the limiting aspects of in vitro culture. This is because they should be considered during the interpretation of results derived from experiments investigating the cellular responses to mechanical stimuli, and extrapolation for in vivo relevance. We then describe model cell culture systems for investigating the response of isolated bone cells to mechanical strain, and finally we describe methods for loading bone explants in culture.

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1.1. Limitations of In Vitro Organ and Cell Culture Strain Application Models

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Organ culture, the maintenance of tissue explants in vitro, is an attempt to bridge the gap between cell culture and in vivo models. Many tissues including cartilage, tendon, and bone have been studied in organ culture (2–6). The major advantage offered by organ culture is that it maintains an intact extracellular matrix (ECM). These extracellular matrices are the product of resident cells, are structurally diverse, and are also uniquely specialized according both to tissue type and local requirements. For example, mineralization is important for bone rigidity and type I collagen endows bone with mechanical toughness (7, 8). Retention of the ECM is important as it maintains normal cell–matrix attachment sites and spatial relationship between cells in a tissue. By retaining the tissue’s architectural organization, it is also likely that the relationship between distinct mechanical sequelae of loading, such as strain, fluid shear stress, and streaming potentials will also be conserved. Both organ and monolayer cell cultures allow the responses to be investigated without the complication of systemic factors. Yet at the same time it is possible, indeed probable, that some of these responses to mechanical stimulus will be complicated by the consequence of tissue or cell isolation and maintenance in culture. Connective tissues, including bone, play major structural roles; for example, the skeleton must support body weight, facilitate movement, and protect internal organs. In such tissues, the ECM and not the resident cells, fulfils such roles. Thus, organ preservation in culture may be considered essential in experiments that are aimed at establishing the basis of the tissue’s response to loadbearing. Bones undergo an adaptive response to dynamic loading in vivo, with very low cycle numbers capable of inducing changes in bone architecture and mass (9–11). Indeed, an osteogenic/antiresorptive regimen of 36 loading cycles at 0.5 Hz is sufficient to produce a maximal response (9). The brief duration (72 s) of this mechanical stimulation, required to activate a full osteogenic response in vivo, makes the study of the early post-loading, “preosteogenic” events feasible. Thus, similar loading regimens have been used in organ culture and “strain models” in cell culture to investigate loading-induced responses (5, 6, 12). Since strain, fluid shear stress, and streaming potentials have all been reported to influence bone cell metabolism, it is important to appreciate that the ECM may modify both the signals to which resident cells respond, as well as the specific cellular reaction which such signals generate (9–11, 13–15). Many bone ECM molecules contain integrin-binding Arg– Gly–Asp (RGD) sequences and in vivo changes in integrin expression profoundly effect bone metabolism (16–19). This is important, as any lack of integrin-binding sites in cell monolayer culture may also lead to changes in integrin expression. ECM molecules are known to influence cell behaviour. Indeed, osteoblasts that bind preferentially to fibronectin in vitro, also exhibit a faster rate of

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proliferation on this substrate than when seeded onto poly-L-lysine (20, 21). Differential integrin expression has been shown in osteoclasts seeded on bone sialoprotein-coated glass compared with plastic (22). It is, therefore, pertinent, at least in the short term, that organ culture maintains normal cell–ECM attachment sites, and retains cell–cell associations and their three-dimensional relationships. By contrast, cell cultures on a foreign substrate are usually two-dimensional monolayers in which there is little control of such relationships. Another essential feature of bone explant culture is that it retains the relative positions of osteocytes, osteoblasts, and osteoclasts. These relative positions might be important for maintaining signalling gradients; for example, all bone cells produce nitric oxide and its release may establish local concentration gradients that regulate behaviour (23). It is highly unlikely that any corresponding gradients are produced in cell culture models. Further, osteocytes and osteoblasts can communicate via gap junctions found at the termini of cell processes stretching through the bone canaliculi (24). Despite the fact that these processes and junctions are also elaborated by cultured cells (25), any positional information that they may confer will be lost as their expression patterns are altered as a consequence of their novel two-dimensional organization. These arguments may be particularly pertinent for the terminally differentiated osteocytes, which are normally embedded in the bone ECM and are difficult to isolate and grow in cell culture. Easy to use osteocyte-like cell lines are now being exploited more widely, but it is pertinent to highlight that they have many differences as well as many similarities to their in vivo counterparts, and that there remains few well-established methods for isolating primary osteocytes for cell culture; these include immunomagnetic separation methods (26) (see Chapter 4) and FACS based on DMP1-driven GFP expression (27) (see Chapter 20). It may also be relevant that as well as maintaining the relative position of bone cell types, organ culture models also conserve their normal cell ratios. Thus, organ culture models can be used to examine potential cellular cross talk, which may occur during the tissue’s response to mechanical stimuli. By contrast, cell culture can only partially achieve this through the use of co-culture systems in which cell ratios and positional relationships will only approximate to those in vivo. 1.2. The Effects of Loading: Mechanical Strain, Fluid Shear, and Streaming Potentials

Retention of structural integrity in organ culture allows the response to applied loads that are capable of generating physiological levels of mechanical strain to be investigated. Ideally, cell culture would complement this by allowing for particular responses to specific single component, uniaxial strains, applied to large numbers of uniform cells, to be determined. However, logistical problems and the fact that substrates stretched along one axis, will inevitably contract at 90° to this principal strain axis (subject to Poisson’s

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ratio), means that such “ideals” are not readily achievable. As this ratio is fixed for any single material, but can vary between different materials, direct comparison between studies in which substrates differ should only be made with care. Currently, such considerations are unavoidable, but it is possible that in the future they will represent developments that will add new strengths to in vitro studies; that is, added scope to readily compare bone cell responses to applied strains across a range of substrates, with differing material properties, to better understand their responses. The identification of cell substratum stiffness as a regulator of specific differentiation may have paved the way for such investigations (28). Nonetheless, with appropriate loading, cells in explants can probably experience physiological levels of strain at their given location. It should be stressed, however, that despite precise loading regimens, cells within explants will experience a range of strain magnitudes in either compression or tension and these location-specific variations should ideally be taken into account during experimental design and interpretation. New methods for accurately mapping strain magnitude and direction across loaded bone surfaces will likely make such variation less of a hindrance to interpretation (29). In fact, it is possible that these strain mapping strategies may promote major advances in our understanding of how the bone cell responses to applied strains orchestrate appropriate, three-dimensionally co-ordinated, changes in bone cell behaviour. Another consideration involves the fact that the lacuna-canalicular network of bone is filled with tissue fluid, and that whilst this network is retained in organ culture models, its contents are replaced by culture media. When this fluid flows over, or through the ECM, fluid shear forces are generated along with solute transportation (30). Application of shear forces in vitro has been shown to induce a response in endothelial cells, chondrocytes, and bone cells (13, 14, 31, 32). In bone, fluid shear forces are generated as a result of a mechanical load-induced shift in tissue fluid, and thus mechanical strain cannot be investigated independently of fluid shear in vivo or in organ culture models. It is possible to investigate the independent effects of fluid shear application in cell culture systems (see Chapter 36, this volume). Nevertheless, it is also clear that the ECM modifies fluid shear rate and magnitude in a way that it is practically impossible to reproduce currently in cell culture systems. A further complication is introduced by the contribution of the flow of charged tissue fluid over the surface of bone’s charged ECM, which results in electrical, streaming potential, currents. Media bathing lacuna-canalicular networks in bone explants contain many charged molecules, including amino acids and proteins. Thus, both the media and the surface of the tissue show a net charge, and any potential difference created at this site of contact results in an electrostatically charged layer around the tissue. In intact bone, cyclical mechanical loading creates fluid flow and establishes

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a streaming potential. Removal of bone from an animal isolates it from the load it normally experiences and results in fluid shifts. It is therefore important to acknowledge the disturbance to these streaming potentials that are an unavoidable consequence in both organ and monolayer cultures. Cell cultures have as their main advantage that they are universally established; easy and reproducible (see various chapters in this volume). However, recent work has shown that isolated osteoblasts, like fibroblasts (33) and osteoclasts (34), retain features that are characteristic of the source of their derivation even when they are maintained in culture (35). It has yet to be determined whether these differing source-specific phenotypic characteristics of osteoblasts exert any major influence to control likely consequences of applied mechanical strains in vitro, or indeed in vivo. The relative ease with which the media, cell number, percentage confluence and differentiation state can all be defined and controlled, are all advantages that make it possible to reproduce cell culture experiments readily in different laboratories. Similar standardization is not yet achieved in organ cultures, but should be achievable in the future by agreed and standardized use of the same culture media, explants of similar size, origin, age, sex, hormonal status, etc. In contrast with cell cultures, it is almost impossible to control cell content, tissue architecture or sample heterogeneity in organ cultures, and the responses within any one cell type is likely to be subject to paracrine controlling influences. Although recent developments have been made (see Subheading 3.5), another current drawback of organ culture models is the limited time that explants can retain viability ex vivo. Finally, loadable organ cultures are clearly not appropriate for the application of all techniques. For instance, mechanically stimulated increases in intracellular free calcium are impossible to monitor in organ culture, but can be observed relatively easily in cell monolayer culture. It is clear that the selection of the culture system must be considered carefully and bear in mind the specific question that the experiment is intended to address, as well as the outcome that it is proposed will be measured. 1.3. In Vitro Cell Culture Loading Models

Before describing any specific methods, it must be recognized that the history of cell-straining techniques has, for the most part, relied upon custom-built devices that remained relatively unique to individual investigators. To some extent this may have been due to a desire for progressive augmentation in the system’s efficiency in delivering a specific component of the mechanical loading environment. This is clearly a desirable objective. Nonetheless, it does not negate the contribution to our understanding made by systems in which the precise nature of the stimulus to which cells are exposed, remain largely undefined or even to some extent variable and uncontrolled. It is however, surprising in our opinion that few,

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if any, of these studies have attempted to relate zonal variations in the applied stimulus, to any differences in the cell’s biochemical response at the level of an individual cell. To do this, the investigator would simply need to measure some response, at the level of an individual cell, and relate this to the magnitude of the mechanical stimulus to which that cell was exposed. In an attempt to delineate the defining character of straining techniques, it is important that a number of principles are considered: 1. Mechanical strain (often tensional) is applied by substrate deformation. 2. Known strain magnitude is generated in the substrate onto which the cells are seeded. 3. The substrate is able to support cell growth and differentiation. 4. Cells must attach to the substrate and not detach as a consequence of deformation. 5. The substrate should have perfect elastic properties. 6. Adequate access for observation, extraction and other processing is possible. 7. During experiments, the apparatus should be housed in a controlled gaseous and thermostatic environment. 8. Techniques should aim to subject all cells to equal levels of mechanical strain, or, as a minimum, ensure that their levels are accurately measured or estimated and described. 9. Controls are available that ideally mimic every other variable but the applied strain stimulus (36). The extrinsic mechanical stimulus to which bone cells respond may be a change in tension, compression, gravitation, vibration or hydrostatic pressure. Herein, we outline different techniques that have been developed to study the response of cultured cells to tension, by stretching (or bending) the substrate onto which the cells have adhered. Thereafter, we describe in detail one such four-point straining device that we have used extensively. 1.4. Biaxial Straining

In “dish-deforming” cell culture models, the imposed biaxial substrate deformation provides a non-homogenous strain on cells. Commonly, dishes are intermittently deformed over a template to produce a 5% change in surface area (37, 38), such that cells near the centre of the dish would be strained in excess of those at the periphery. Several systems have been developed (single and multiple dish versions) that control the input strains by varying the curvature of the template. In these systems, estimates of the template’s arc length of spherical distension provide a basis for calculating average strains (39). Numerous other platen-driven devices for applying biaxial strain have been developed for cells grown in culture dishes with a

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flexible membrane, instead of rigid tissue culture plastic, bottoms. Examples of such techniques include the following: upward or downward “tenting” of membranes using vertically pulsating ball-ended prongs (40, 41), electrocylinder-driven pulsations of spherical watchglass sectors to indent culture dishes (42), and upward indentation of Petri dishes with flat-ended circular pistons (43). In 1985, flexible-bottomed, circular cell culture plates were specifically designed that could be interfaced with a computercontrolled vacuum manifold system (44). Vacuum application to the culture well undersurface stretched it downward in a manner that could be controlled by varying vacuum magnitude, waveform, frequency and number. Another differential pressure, flexible-substrate system uses positive, solenoid valve-mediated, pressure to deform circular, peripherally clamped, 100 μm polyurethane/urea membranes (45). A common problem with such devices is that the deformation can result in non-uniform strains of the substrate. It is clear that these will also be subject to variations in the strain applied to cells at various positions across their diameter. This is an important consideration, as it has been predicted that biaxial straining is at least twice as potent as uniaxial strains (46). Thus, unless one was going to investigate a particular response in a single cell of known location and strain, then it would be necessary to appreciate that any results may represent an aggregate of total inhomogeneous cellular responses to the applied range of strains. One might also consider the effect of “puddling”; that is, as the substrate is drawn downwards by the vacuum, the overlying medium will collect at the lowest point. The effect of this fluid movement has not been tested. These effects have recently been addressed and uniaxial tensile strains are now achievable in these systems using “Arctangle Loading Posts”. In addition, three-dimensional cultures are amenable to such computer controlled mechanical stimulation (47). Recent reports have demonstrated differential activation of AP-1, Erg-1 induction and NF-kappa B nuclear translocation in osteoblastic ROS 17/2.8 cells by direct strain compared with simulated weightlessness (48, 49). Strain-induced gene expression responses reflect those seen in distraction gap osteogenesis (50). One concern with these experiments is the relatively high levels of applied strain compared with known in vivo strain levels, and Columbo et al. make a very valid case for performing pressurestrain calibration tests for each particular waveform as a “control” in such experiments (51). Another method that aims to apply uniform biaxial strain involves using a square elastic substrate that is stretched on four sides, over the rim of a platen. As increasing amounts of the original (flat) membrane slip radially, outward over an axially advancing platen rim (its area increasing as it does), the portion of the membrane remaining directly over the platen theoretically experiences

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homogeneous biaxial strain. Brighton et al. found that when straining cultured osteoblasts in this way, they could detect a physiological response at strains as low as 300 με (52). 1.5. Uniaxial Straining

These experiments purport to investigate the effect of uniaxial strains, however, it is important to note that they nonetheless are subject to the restrictions imposed by Poisson’s ratio (a substrate stretched along one axis will inevitably contract at 90° to this principal strain). Most of these devices have relied upon four-point bending, but a three-point bending technique has also been designed by Hasegawa et al. (38). Whilst this successfully achieves the application of uniaxial strains, their magnitude over the plate’s surface will be unequal.

1.5.1. Stretching of Substrates

Uniaxial straining devices have been developed in which a strip of silicone (see Note 1) is placed in a trough and anchored at one end, a magnet attached at the strip’s free end and another magnet placed outside the culture system is then operated by hand or via a motor-driven cam to stretch the substrate. Other methods using silicone as a substrate require film spools to create the length change (46). Using derivatives of such systems, in which cells are seeded onto pre-stretched membranes, it is possible to observe cells during their response to compression as well as tension. It may be vital in the future to make greater use of these compression devices as responses to applied tension and compression are both physiological relevant. Cell culture chambers attached with silicone sealant to the surface of polycarbonate sheets have also been used to subject adherent cells to mechanical strain (53). These systems, based on a design originally described by Murray and Rushton (54), use strips of polycarbonate which are adapted into cell culture chambers, by attaching lids removed from 4-well slides with silicone sealant. Cell straining is achieved using a device consisting of two platens that run on dry linear bearings, driven by a servo-controlled pneumatic ram. The actuator and compressor are housed outside the incubator and the former is controlled from a computer supplied with feedback from a linear variable displacement transducer. The polycarbonate strips, with adherent wells and cells, are clamped across the two platens. Controls consist of strips clamped across “static” platens as well as strips that are clamped across a moving platen that will generate medium perturbation equivalent to that experienced by strips subjected to strain stimuli (53). This device is capable (determined by appropriate strain gauge measurements) of applying controlled cyclical strains between 100 and 200,000 με, at strain rates from 100 to 1,000,000 με/s and in this system, strains have been applied as a ramped square wave pattern. Similar systems for applying uniaxial strain using a polyurethane substrate have also been developed by Grabner et al. (55),

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in which a motor-driven linear stage was used for the application of a cyclical tension. These devices appear to represent a key to addressing questions that relate to the differential effects of mechanical strain and fluid flow. 1.5.2. Four-Point Bending of Substrates

Four-point bending of a plate requires a pair of lateral forces acting outside two fulcra. This produces an even curve at both the tension (convex) and compression (concave) surfaces, and the curve makes up a segment of a circle, between the fulcra. Several four-point bending systems have been designed, and in the most simple terms these differ in the manner by which loads are applied, and the culture substrates compounds that are used. One such system uses rectangular culture plates (polycarbonate or glass) suspended within a common 40 mm× 40 mm silicone rubber well (56). By allowing both sides of the substrate to act as the surface onto which cells can be seeded, the device elegantly permits cellular responses to either tensile or compressive strains to be examined. It also includes scope for direct strain gauge readout to monitor the parameters of strain to which cells are exposed. Another powerful feature of this device is that it permits the use of a range of culture substrate materials, with different Young’s moduli, thus facilitating application of a broad range of strain magnitudes (see Subheading 3.1).

1.5.3. Alternative Methods for Applying Strain

Magnets are used to provide a high degree of control in devices capable of stretching a substrate, and such magnetostrictive actuators have been used to generate up to 22,000 με on particular substrates. The electromagnetic elements placed at either end of the substrate are attached to clamps mounted on a guide plate that ensures unidirectional travel. It is an aspect of these devices that they generate powerful electromagnetic fields that may directly influence bone cell behaviour, and shielding from these fields should always be used. As an alternative, piezoelectric extension may be used to provide the displacement. This allows accurate control, however, like magnetostrictive actuators, generates high electrical fields. Such actuators have been used in vitro to apply strains of controlled magnitude (200–40,000 με), frequency (up to 100 Hz) and waveform, which adequately cover the range experienced by bone cells in vivo. The cells are mechanically strained by moving a plunger, connected to the centre of an actuator, both ends of which are inserted into grooves on an acrylic resin frame, thus generating maximum actuator displacement. Cultured cells are seeded on, or in, a collagen gel block between plungers made of a non-conductive acrylic resin material, and the gel block anchored by stainless-steel wire meshes (lattice size; 0.4 mm × 0.4 mm) fixed to the plungerends (57).

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1.5.4. Methods to Examine the Effects of Weightlessness

Prolonged exposure to near weightlessness for astronauts in orbiting space stations and extended space travel missions may help provide major insights into the control of bone mass. The mechanism for bone loss in these situations needs to be thoroughly investigated as it may inform our understanding of the processes involved (58–60). In order to avoid the need to train astronauts in specialized cell culture techniques, earth-bound simulated microgravity devices based on the clinostat are increasingly being used. These include: the random postitioning machine (RPM) (61), the rotating wall vessel bioreactor (RWV) devised by NASA and the commercially made variants available from Synthecon Inc., Houston, Texas, USA. In these devices, cells are cultured in slowly rotating vessels filled with culture medium and are therefore maintained in constant “free-fall” (shear forces would be generated on the cell surfaces, but these are thought to be insignificant). As a consequence, the cultured cells never settle or flatten on any substrate. Investigations using such devices have demonstrated that microgravity leads to the disruption of collagen I/integrin signalling (62), reduced alkaline phosphatase, RUNX2, osteomodulin and PTHR1 gene expression in bone cells (61). Induced microgravity has also been shown to alter the flux of human mesenchymal stem cells through differentiation pathways, favouring adipogenesis over osteoblastogenesis in a manner that is dependent upon the disruption of the cell cytoskeleton (63). It is clear that these investigations might also lead to identification of mechanically responsive genes (64) and strategies for reducing bone loss.

1.5.5. Alternative Substrates

There have been advances in development of rigid bioglass, glass– ceramic implants for the stimulation of local bone formation and fracture healing (65–68). It is envisaged that future studies will need to explore bone cell responses to mechanical stimuli grown on these novel compounds. Indeed, it will be important to establish the consequences of mechanical loading of the bioglass substrate itself.

2. Materials 2.1. Tissue Culture Media

For canine cancellous bone cores : MEM + Hank’s salts and 25 mM HEPES supplemented with 2.0 mM L-glutamine, 0.1% bovine serum albumin, 100 IU/ml penicillin, and 100 μg/ml streptomycin (Gibco). Perform all cultures at 37°C in an air incubator. For rat ulnae or rat calvarial organ cultures: DMEM plus 10% charcoal-dextran extracted foetal calf serum, supplemented with 2.0 mM L-glutamine (Gibco) and 100 IU/ml penicillin, and 100 μg/ml streptomycin (Gibco).

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For chick tibiotarsus: Fitton–Jackson’s modification of BGJb medium containing 2 mM L-glutamine, 100 IU/ml penicillin, 100 μg/ml streptomycin, 50 μg/ml L-ascorbic acid, and 2% heatinactivated foetal calf serum.

3. Methods 3.1. An Example of Four-Point Bending of Monolayers of Cells on Plastic Strips

A four-point bending system that is able to engender low-strain levels has been used in which tensional strain is applied to plastic strips containing adherent monolayer bone cell cultures in a custom-designed loading apparatus (Fig. 1). This four-point bending system was designed to deliver strain levels of several hundred to several thousand με, which are within the physiological range recorded for bone cells in vivo (see Note 2) (5, 69, 70). It should be noted that this system is also capable of delivering compressive strain to cells seeded on the strips lower surface. 1. Passage cells onto pre-sterilized plastic strips for at least 24 h (this time depends upon initial seeding density), and maintain cells for 24 h and throughout the experimental application of mechanical strain in a humidified atmosphere of 95% air–5% CO2 in serum-free medium (see Note 1). 2. Transfer the strips (n = 5 for each variable) to the loading apparatus under sterile conditions. To do this, separate the loading apparatus into its two component parts: a base for the strained strips and an upper portion with the cam, platen, and flow chambers. Remove strips from the dishes in which they have been pre-incubated and position into individual chambers with the requisite volume of medium (10 ml) added. 3. Reconstruct the whole apparatus, place back into the humidified incubator, and allow for equilibration. Keep disturbance of the medium minimal at all times. 4. Apply a load generating mechanical strain of 3,400 με at 1 Hz for 600 cycles to cells adherent on the strips (see Note 2). 5. Subject similar strips with cells attached to cyclic perturbation of the medium (flow controls) without applied loads, and subject others only to identical changes of the medium without any mechanical perturbation to serve as “static” controls (see Note 3). 6. Following strain application, remove strips from the loading apparatus and return to dishes for various times post-straining, depending on the response to be investigated.

3.2. In Vitro Organ Culture Loading Models

Culturing bone as a tissue allows for the preservation of normal cell–matrix attachments as well as cell–cell attachments between resident cells. Preservation of the structural, load-bearing, mineralized

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Flow (control) Fulcrum

Rotating cam

Spring

Strain Fig. 1. Schematic diagram of the custom-built apparatus used for straining cells by subjecting plastic strips to four-point bending. As the eccentric cam rotates it presses the loading platen onto the vertical edges of the plastic strips and causes their deformation (to the dotted position) in an arc around the paired fulcra underlying the strips (white). The platen is returned to the unloaded position by the spring. The rotation of the cam also acts to rock the chamber into which strips to be subjected to flow “control” are placed. For dose– response experiments, peak strain magnitudes can be altered by vertical displacement of the base unit by using a range of washers (with different thickness, placed between base and upper units, see Fig. 2), and frequency altered with rheostat control of cam revolution rate. This unfortunately affects the waveform produced. However, a novel device has recently been designed, in which the platen’s vertical displacement is governed by a screw mechanism that is computer controlled. This offers greater flexibility in the waveforms that can be generated, such that on/off dwell times and the “on” as well as “off” strain rates can be varied independently (see Fig. 3, Stromberg et al., personal communication). In addition, this model utilizes commercially available cell culture strips (of equivalent size to microscope slides) which do not possess the end walls.

compartment means that loading of segments in vitro, can be controlled so that the extent of bending produced, engenders levels of strain (measured directly in loaded segments using attached strain gauges) identical to those measured at the same site during normal physiological activity in vivo (see Note 2). Providing disruption is kept to a minimum, it is also likely that in vitro loading of bone segments will reproduce other associated phenomena of in vivo loading (fluid shear/streaming potentials). On the contrary, such isolation will result in many unavoidable changes, including loss of blood flow, a potential restriction upon the ability of nutrients reaching cells, and changes in the relationship between

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Strain (me)

Strain/Washer curve 4000 3500 3000 2500 2000 1500 1000 500 0 0

0.5

1

1.5

2

Washer Thickness (mm) Fig. 2. Peak strain magnitude in plastic strips can be altered by changing the difference in height between the lower and upper base units in the four-point loading device.

5000 4000 3000 2000 1000 0

Fig. 3. Waveform of the loading generated strains in two strain gauges attached to one strip in the loading apparatus (Stromberg et al., personal communication).

bone and its marrow. Nonetheless, this approach allows attempts at bridging the gap between the morphological disciplines and those of the cell biologist, in determining the mechanisms of bone’s response to loading. The in vivo loadable avian ulna model developed by Lanyon and Rubin (9, 71) determined that dynamic mechanical loads applied to a functionally isolated portion of bone could result in an increase in bone formation. The extent of new bone produced, was dependent on the magnitude and the rate at which the strains were engendered (72). Further studies indicated that only 36 cycles (over a period of 72 s) per day of applied load were sufficient to produce a maximal formative response (9). That all the information needed to produce new bone was contained in only a short period of time provided the rationale for generating organ culture models to investigate other early loading-related responses that might contribute to controlling these ensuing osteogenic responses. The in vitro loadable models which have been developed include adult canine cancellous bone cores, rat cortical ulnar bones,

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rat parietal bones, and embryonic chick tibiotarsi. All of these have used the aforementioned in vivo studies as their basis and have, as a consequence, used similar strain regimes as the mechanical stimulus. Further models include a loadable bovine cancellous bone core system (73) and a human bone core model (74). Each system requires that attendant soft tissues and marrow (where possible) are removed, to leave exclusively bone tissue with resident bone cells and associated internal vasculature. In order to generate results that are likely to have in vivo relevance, it is imperative that these organ culture systems are calibrated. Thus, prior to loading bone in vitro, the amount of force required to generate the levels of strain that are engendered by normal activity in vivo should be defined (see Note 2). In this section, we describe several models of loadable bone organ cultures that have been defined in this way. 3.2.1. Loading of Adult Canine Cancellous Bone Cores (see Note 5)

Preparation of bone cores: 1. Euthanize animal with an overdose of barbiturate, flex the stifle, and make an incision longitudinally in the skin. 2. Deflect the kneecap to expose the trochlea groove of the distal femur. Using a trephine with a non-retractable pointed insert (slightly proud of the cutting edge) make a circular groove in the articular cartilage surface. Remove the pointed insert and take a full depth cartilage and bone plug from the epiphysis of both left and right limbs. 3. Place each core into a cutting rig and trim to a uniform length (1 cm). 4. Introduce the core into a syringe barrel with the nozzle portion and end removed. Flush sterile, warm PBS through the core to remove marrow from between the bony trabeculae. 5. Store cores individually in culture medium in sterile bottles at 37°C until all cores required for an experiment have been collected. 6. Perform cultures at 37°C in an air incubator. Loading of bone cores: 1. Within separate flexible collars of silicone tubing (see Note 4), support each of the bone cores between a pair of milled Perspex supports, which have been drilled to allow the passage of culture medium through them and the core. Make sure the end face of each support is cut to be of equal diameter to the core. To create a seal between the supports and to surround the core, the internal bore of the silicone tubing should be equal to the core diameter. Hold the control channels in place on a backboard with clips. 2. Place cores into the loading apparatus (Fig. 4). To do this, mount the lower Perspex support onto a rigid, non-flexible,

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Fig. 4. Loading apparatus for bone cores.

bar attached to the backboard. Connect the upper support to a pneumatically operated actuator, fixed to the backboard, which is operated by an air compressor unit. Introduce recirculating medium into the reservoir syringe above each core and draw through the silicone tubing, by a peristaltic pump, back to the reservoir syringe above the core (one lane per core). Controls comprise cores housed similarly in silicone tubing and Perspex supports, which are attached to the backboard with clips but remain unloaded. 3. Prior to loading, pre-incubate all cores for 4–5 h in a recirculating system, in which culture medium is delivered at a flow rate of 0.3 ml/min. Following this pre-incubation period, convert the recirculating system to single-passage perfusate mode. Refill all reservoirs and replenish continually for the remainder of the experiment for any “real-time” analyses. Alternatively, retain the recirculating system as appropriate. 4. To load bone cores, regulate air pressure to deliver the same force that generates a bulk strain of 5,000 με in separate cores, at a loading frequency of 1 Hz.

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3.3. Rat Ulnae Organ Cultures (see Notes 5 and 6)

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The culture medium used in these experiments depends upon the conditions employed during the application of loading (see below). Preparation of bones for loading: 1. Dissect ulnae and clear of attendant soft tissue. Cut ulnae to equal length and remove marrow. 2. Pre-incubate on hand-milled polytetrafluoroethylene (PTFE) supports at the air–medium interface for 6 h in 12-well culture plates in a humidified atmosphere of 95% air 5% CO2 at 37°C. 3. Place into one of two different loading devices (weight-lifting, or pneumatic actuator; see below). Following loading, remove bones from the loading apparatus and return to PTFE supports in 12-well plates for various times post-loading, depending on the response to be investigated. 4. The culture medium used in these experiments depends upon the conditions employed during the application of loading. Loading of bones in weight lifting model: 1. Ulnae are loaded in individual chambers with medium recirculating around the cortical shaft. This loading apparatus is maintained in an air incubator, so during the loading period, the culture medium is supplemented with 25 mM HEPES buffer. 2. Each bone is held vertically between two cups. The lower cup is connected to an eccentric cam, whilst the upper cup is connected to a weight-carrying platform that rests on a ledge. As the cam rotates, the bone is raised and lowered. On the up-stroke, the bone acts as the sole support for the weight-carrying platform and on the down stroke the weight returns to rest upon the ledge. The amount of weight on the platform can be altered, thus permitting different levels of mechanical strain to be engendered. (12, 75, 76). 3. Following loading, remove bones from the loading apparatus and return to 12-well plates for various times post-loading, depending on the response to be investigated. Controls comprise ulnar segments, treated identically, that do not lift weights. Loading of bones in the pneumatic model: 1. This rat ulna organ culture model maintains the tissue in a fixed volume of medium (see Fig. 5). The device consists of a milled polycarbonate block containing ten chambers, in which five are customized to permit loading and five serve as controls; loads are applied by pneumatic actuators. 2. The bone shafts are removed from the pre-incubation medium and introduced into the holding cups of the loading apparatus, either in loading or control chambers each containing 4 ml of culture medium.

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FIXED NYLON CAP

BONE CULTURE MEDIUM NYLON CAP (MOVABLE)

AIR PISTON

BASE Fig. 5. Diagram of organ culture loading apparatus, showing the bone segment held between nylon caps immersed in culture medium. Oscillating air pressure in the pneumatically operated cylinders applies a dynamic load to the tissues.

3. After 5-min equilibration time, bone explants are loaded axially to generate mechanical strains levels on the lateral mid-shaft, similar to those generated in vivo. Control bone shafts are not loaded. 4. After the loading period, bone shafts are transferred back to 12-well plates and cultured for various periods of time, depending on the response to be investigated (6, 77, 78). 3.4. Loadable Rat Calvaria Organ Cultures (see Notes 5 and 6)

Essentially the method used is identical to that described under Subheading 3.3, but an adaptation is made in the caps that hold the tissue within the apparatus (6). The caps retain bones to limit uncontrolled translation. For the ulnar bone shafts these nylon caps are fashioned into cups, whilst for the parietal bone (calvaria) explants, the caps have a shallow, narrow rebate engineered to accommodate the thin, plate-like architecture of these bones. 1. Cut the parietal bones into rectangular explants and culture as described in Subheading 3.3. 2. To apply loads, first locate the rostral and caudal ends of the bones into the “cap” rebates, thus allowing axial loading without scope for slip. 3. Perform loading using the pneumatic actuator device (see Subheading 3.3 and Notes 6 and 7). 4. After the loading period, bone shafts are transferred back to 12-well plates and cultured for various periods of time, depending on the response to be investigated.

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3.5. Loading of the Chick Tibiotarsus (see Notes 5, 6, and 8)

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Preparation of bone segments: 1. Euthanize 18-day-old embryonic White Leghorn chicks using an approved method and remove the tibiotarsi. 2. Remove adherent soft tissues and fibula, and both cartilaginous ends to leave a 9-mm bone shaft segment. 3. Aspirate the marrow, but leave the periosteum intact. 4. Wash briefly in phosphate-buffered saline. 5. Culture bone segments at 37°C in a humidified 5% CO2 incubator at the air-medium interface on PTFE supports for 5 h. Loading of bone segments: 1. Hold the embryonic tibial bone shafts at either end by polypropylene caps, in chambers milled into a Perspex block. 2. Both “weight lifting” and “pneumatic” actuator devices (79, 80) have been used to load these bones (see Subheading 3.3). Each device is calibrated to generate the required strain levels (see Note 2). For weight-bearing induced loading, an eccentric cam (rotating at 1 Hz) is employed to raise and lower an L-shaped cradle holding the weights. The force is transferred to the bones via a pivot positioned at the right angle of the L-shaped cradle, and thus, when the cradle is lowered, the tibia supports the weight of the cradle. As in other models, the level of engendered strain can be altered by changing the amount of weight that is “lifted”.

3.6. Longer-Term Perfusion Loading Models

Bone tissue in the cancellous bone core model, first developed in the mid-late 1980s, had only a limited life-span for experimentation and investigated only early responses following physiological mechanical loading (81–83). A more recently described long-term model system has been designed in an attempt to overcome the limited viability preserved in previous organ explant models (74). It has been hoped that this system will allow us to make significant advances in our understanding of loading-related bone formation. The system involves perfusion of trabecular bone cores including their marrow and has been purported to extend tissue lifetime to 72 days. Preliminary studies using the incorporation of fluorescent labels into newly mineralized surfaces, to provide a direct measurement of bone formation rate, have reported that these cultured bone cores appear to retain their in vivo rates of bone formation for up to 20 days (73). The loadability of such explants in these systems suggest that it is possible that advances may be made in our understanding of the means by which the cells “sense” mechanical stimuli and “communicate” this influence to coordinate loadinginduced changes in bone remodelling. The Zetos system was introduced (84), with culture conditions that allow for loading-related changes in bone architecture (85) and stiffness (86) to be detected over a 3-week loading period.

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A more recent addition to the systems for maintaining bone in long-term organ culture has been devised by Chan et al. (87). The stated purpose of this system is to provide scope for the investigation of osteocyte–osteoblast connectivity and interactions. It is based on what might appear to be at first glance an overcomplicated approach, in which the cultured trabecular bone explants are first denuded of resident osteoblasts and then reseeded with primary osteoblasts from an alterative, genetically distinct, source of bone. The advantage that this may provide is that it will allow changes in the behaviour of the “tagged” reseeded, genetically modified osteoblasts to be monitored and an opportunity to localize the responses and molecular pathways involved in the response to mechanical stimulation.

4. Notes 1. A vital consideration for cell culture experiments is that both the substrate and culture wells should be made from biocompatible materials. Indeed, it has been found that “biocompatibility” of material positively correlates with the amount of non-collagenous matrix protein produced by cultured bone cell monolayers (88). Medical grade silicone, with its high biocompatibility, is, therefore, a rational choice of substrate. Aclar-33C (Allied Chemical Co.) is a clear, biocompatible material that may be sectioned for electron microscopy, transmits UV light with little attenuation or scatter and also useful in such devices 2. Load–strain relationships have to be determined in any new model of in vitro strain application by prior calibration. Thus, for example, plastic strips identical to those that will be used experimentally must previously have had strain gauges attached to them and loaded in a controlled manner. During load application to these substrates in vitro (and indeed loading of bone segments in organ culture), the correct force must be delivered to produce the required level (physiological or if required nonphysiological) of mechanical strain. Strain gauges cannot be attached to samples that will be used for experimental investigation; therefore it is necessary before each new device is used to establish the relationship between applied load, which can be monitored and adjusted directly, and the resultant strain, which cannot. To calculate the relationship between strain and load for plastic strips (and bone explants; see Figs. 2 and 3 (11)), single element micro-miniature strain gauges can be used. Bond gauges to the test substance (or bone surface) with cyanoacrylate adhesive, ensuring that that the pre-wired strain gauge will detect strains along the substrate’s principal strain axis.

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Connect gauges to a strain gauge conditioner and amplifier, in a quarter bridge configuration, and record the output voltage on a personal computer equipped with an analog-to-digital (A/D board) converter. Convert to strain values using a 1,000-με calibration shunt resistance built into the conditioner units. Use the recorded strain data files to assess peak strain magnitude and strain rate and determine frequency of a loading cycle. We highlighted that current systems for applying mechanical strain are for the most part unique to individual laboratories. This makes a detailed description of a generic device impossible. For this reason, we have given a broad account of some of the many, and various, “models” that have been used to apply mechanical strain to isolated bone cells in vitro. We have also tried to give some insights into their strengths and weaknesses, the materials used and the methods employed, and some of the considerations that should be taken into account when deciding which model might be chosen. This choice clearly depends on the hypothesis that each investigator has selected to address, and is one that must take into account which mechanical consequence of loading (strain, flow, streaming potential) is being examined. Whilst each of these can be accurately defined, each also exhibits many components that can vary. For example, magnitude, frequency and rate of change can be varied during mechanical strain application, and it is vital that each of these is considered prior to selecting a “model” in which bone cell responses to are to be examined. Perhaps the most obvious and pertinent of these deliberations is whether the applied stimulus falls within the physiological range. This can only be reflected upon, if the variable in question has been measured directly in vivo. 3. Mechanical strain application results in the exposure of cells to both tensional strains, via bending of their substrate, and perturbation of their medium as a consequence of the cyclical displacement of these strips vertically through their medium. As a precaution, devices are therefore designed with flow “controls”, which are included to allow identification of both flow-related and non flow-related responses in the cells that have been exposed to mechanical strain. Examination of the cellular responses to such flow stimulation (medium perturbation) has made it clear that this often includes components of the cell’s response to strain application, but that it also often induces a range of responses that are particular and specific to the application of flow itself (70). Thus, it is vital that controls are included that have been subjected to similar “preparatory” changes in the medium, but that are not subjected to any form of further medium perturbation or substrate deformation

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(i.e. “static” controls). An essential caveat, however, in the flow “controls” included, is that the precise nature of the flow stimulus to which the cells are exposed remains, unlike in other studies, ill defined. Many other cell culture variables may also modify the cellular response to strain application. These are likely to include the following: the effects of changing the medium, serum deprivation, cell density, growth rate, differentiation status, etc. Although most devices allow the investigator to sample medium at various times after strain application, the investigations are often limited by the fact that large volumes of medium are required in such studies and by the fact it is very difficult to make direct microscopic observations in vitro. 4. When rigid Perspex is used to surround the core, “barreling” of the core is prevented and no loading-related responses are measurable. When silicone tubing is used instead, “barreling” of the core is allowed when loads are applied and changes in biochemical activity in the loaded compared with non-loaded cores are detected. 5. Cell viability should be tested in preliminary experiments by measuring extracellular lactate dehydrogenase activity and the ability to produce cAMP in response to PTH. The methods can be found elsewhere in this volume. 6. In these experiments, medium can be supplemented with exogenous factors, such as, enzyme inhibitors/activators, and their effects on the response to loading investigated. It is also possible using these systems to measure the accumulation of soluble metabolites in the medium conditioning each individual culture (6, 77, 78, 89). An important strength of such models is that they allow detailed investigation of the specific changes associated with individual cells within bone tissue. Thus, appropriate treatment (for example, chilling tissue and preparing cryostat sections) of individual bone segments, at various times after loading, allows for the analysis of specific components of the strain response to be examined at the individual cell level by in situ analyses. 7. In our studies examining the response of calvarial bones in vitro, we first applied a 100-με loading episode. This equates to strains that are higher than those determined in vivo, since the loading apparatus was not sensitive enough to lower the strains to physiological levels. In later experiments, loads that generated 1,000 με were applied. Although, this level of strain is approximately 30 times greater than physiological, the bone did not fracture, reflecting the high safety factor in such bones (6). 8. This model was initially established to ascertain whether embryonic bone responded similarly to adult bone. In some experiments, the periosteum was removed to elucidate the contribution from the periosteal cells (79).

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Acknowledgements We are grateful to Arthritis Research UK, the Biotechnology and Biological Sciences Research Council, and The Wellcome Trust for their contribution to the work done in the laboratories of AAP. We would also like to thank Dr. Gul Zaman for his constructive and critical comments and Victoria Das-Gupta and Dominic Simon for their contributions to the original edition. We are also grateful to Prof. Lance Lanyon. References 1. Bradbeer, J. N. (1992) Cell biology of bone remodelling. In Recent Advances in Endrocrinology and Metabolism (Edwards, C. R. E., and Lincoln, D. W., eds) Chruchill Livingstone, PP. 95–113. 2. Bonassar, L. J., Grodzinsky, A. J., Srinivasan, A., Davila, S. G., and Trippel, S. B. (2000) Mechanical and physicochemical regulation of the action of insulin-like growth factor-I on articular cartilage. Arch. Biochem. Biophys. 379, 57–63. 3. Bayliss, M. T., Howat, S., Davidson, C., and Dudhia, J. (2000) The organization of aggrecan in human articular cartilage. Evidence for age-related changes in the rate of aggregation of newly synthesized molecules. J. Biol. Chem. 275, 6321–6327. 4. Packer, D. L., Dombi, G. W., Yu, P. Y., Zidel, P., and Sullivan, W. G. (1994) An in vitro model of fibroblast activity and adhesion formation during flexor tendon healing. J. Hand Surg. Am. 19, 769–776. 5. Zaman, G., Pitsillides, A. A., Rawlinson, S. C. F., Suswillo, R. F., Mosley, J. R., Cheng, M. Z., Platts, L. A., Hukkanen, M., Polak, J. M., and Lanyon, L. E. (1999) Mechanical strain stimulates nitric oxide production by rapid activation of endothelial nitric oxide synthase in osteocytes. J. Bone Miner. Res. 14, 1123–1131. 6. Rawlinson, S. C. F., Mosley, J. R., Suswillo, R. F., Pitsillides, A. A., and Lanyon, L. E. (1995) Calvarial and limb bone cells in organ and monolayer culture do not show the same early responses to dynamic mechanical strain. J. Bone Miner. Res. 10, 1225–1232. 7. Currey, J. D. (1979) Mechanical properties of bone tissues with greatly differing functions. J. Biomech. 12, 313–319. 8. Riggs, C. M., Lanyon, L. E., and Boyde, A. (1993) Functional associations between collagen fibre orientation and locomotor strain direction in cortical bone of the equine radius. Anat. Embryol. (Berl) 187, 231–238.

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Chapter 38 In Vivo Mechanical Loading Roberto Lopes de Souza and Leanne Saxon Abstract The skeleton fulfils its mechanical functions through structural organisation and material properties of individual bones. Both cortical and trabecular morphology and mass can be (re)modelled in response to changes in mechanical strains engendered by load-bearing. To address this, animal models that enable the application of specific loads to individual bones have been developed. These are useful in defining how loading modulates (re)modelling and allow examination of the mechanisms that coordinate these events. This chapter describes how to apply mechanical loading to murine bones through points of articulation, which allows changes in endosteal, periosteal as well as trabecular bone to be revealed by double fluorochrome labelling and computed tomography, respectively. Key words: Mouse, Mechanical loading, Cortical bone, Cancellous bone, Adaptation

1. Introduction 1.1. MechanoAdaptation

When bone is mechanically loaded it will deform. The degree of deformation at any one site can be quantified as the change in length divided by the original length and this is termed strain. One microstrain (με), therefore, is equivalent to a strain of 0.0001%. Strain can be positive (tensile) or negative (compressive) and because it is a ratio, it is dimensionless. Strain is considered the controlling stimulus for bone’s adaptive response to mechanical loading (1, 2). In 1983 Frost first postulated that bone adapts its structure to maintain strains within a safe window to protect against fracture, while optimising bone mass and flexibility. He termed the threshold strain required to trigger an adaptive response as the “minimum effective strain” (1). He also likened this control mechanism to thermostat and thus coined the term “mechanostat” (3).

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_38, © Springer Science+Business Media, LLC 2012

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Frost’s theory has since evolved to incorporate the idea that cells not only sense the local strain, but also integrate strain information throughout the bone to form an overall appreciation of strain distribution (4). The response of bone therefore represents a coordinated effort of cells acting in unison, most likely via the network of osteocytes (5). In addition, other parameters besides strain magnitude, such as the dynamic nature of the mechanical loading stimulus, strain distribution; loading cycle number; strain rate and loading frequency are known to regulate the scale of adaptive responses to loading (6). Strains need to be dynamic and applied at a rapid strain rate to stimulate an osteogenic response, while static loads will fail to initiate an osteogenic response (4, 7). Unusual, novel strains, within the range experienced during everyday activities, also initiate bone modelling to improve the structure and physical properties of the tissue (6, 8–10). The number of daily loading cycles that are required to induce the maximum adaptive response is surprisingly small. Bone formation in response to artificial bending has been found to plateau at 36 cycles/day (9, 11). Moreover, a single 5 min, 300 cycle period of loading employing known osteogenic parameters is sufficient to transform bone from a quiescent state to a state of active bone formation within 5 days (12). Strain rate, or time to peak strain, is also a major determinant of the adaptive osteogenic/antiresorptive responses to mechanical load. High-strain-rates are more osteogenic than low or moderatestrain rates (13, 14). Loads applied at a high frequency, between 1 and 20 Hz, are also known to increase strain-induced adaptive responses (15, 16). Loads applied at higher frequency >20 Hz and very low in magnitude may also stimulate bone formation (17–19). This pattern of loading is believed to replicate the strains generated by muscles to maintain a normal posture (20). The introduction of rest periods in between loading cycles, such as one cycle every 10 or 15 s will also significantly increase the osteogenic response to high strains (21) (see Note 3). This is based on the notion that rest periods enhance canalicular fluid flow and stimulation of osteocytes. So too will breaks over a long periods of time, such as a month off in between 2 months of loading (22). In summary, relatively large strains alone are not sufficient to activate bone formation, and that a few, unusual loads applied at a high strain rate, with rest periods in between will also stimulate new bone formation (18). 1.2. Loading Models

Bones respond to changes in load-induced mechanical strains by altering (re)modelling activities to ensure appropriate cortical and trabecular bone morphology and mass. Determining a relationship between exercise and bone has been difficult in humans because both the dose and the response are relatively ill defined. Nevertheless, many human exercise studies show local site-specific changes in

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bone architecture (23–25). In tennis players and baseball players, a pitcher for example, humeral hypertrophy occurs in the playing arm in which stimulatory loading is experienced (23, 25, 26). Clinical studies have shown local increases in bone mass in response to specifically designed exercises (27, 28). To overcome the difficulty of defining the mechanical stimulus that drives bone’s adaptive response, researchers have used animal models instead. For example, animals have been trained to run on a treadmill or to perform jumping exercises. These models provide a means by which the duration, magnitude, and frequency of loading can be controlled and the resultant changes in cellular behaviour, bone strength and bone formation response can be directly measured using histomorphometry, imaging equipment such as micro Computed Tomography (microCT) and peripheral quantitative analysis (pQCT), ultimate yield of fracture, or ash weight. Early surgical models, such as applying loads through wires inserted into rabbit tibia (7) or pins through the rat caudal vertebrae (29), or osteotomy of sheep ulnae (4, 30, 31), have been superseded by non-surgical models because they were often associated with inflammation, uncontrollable or ill-defined loads and complications post-surgery. Non-surgical models such as the rat and mouse tibia four-point bending model obviously do not need any surgical intervention, but the loads are applied in a non-physiological distribution and apply pressure directly onto the periosteum, resulting in local trauma and the production of woven bone. Torrance et al. (32) developed the rat ulna axial loading method, which presents several advantages: (1) it does not apply direct pressure on the periosteum; (2) a normal range of activity is permitted between the loading periods; and (3) the loading variables can be precisely controlled (32). Non-surgical mouse models for the study of bone’s adaptive response to mechanical loading are highly desirable, because of the great potential afforded by the study of transgenic mice and the large number of inbreed strains of mouse with differing bone phenotypes. In addition, like the rat, the mouse is regarded as an appropriate model for the study of human skeletal disease and in particular postmenopausal osteoporosis (33). Furthermore, unlike the rat, the mouse achieves skeletal maturity in terms of peak bone mass and strength at the comparatively early age of 22 weeks (34, 35). This facilitates the study of bone adaptation in the absence of the confounding affects of high growth rates seen in the rat loading models (11, 32). A mouse ulna and tibia loading model has been designed by our laboratory that applies quantifiable, controlled axial loads to the bone positioned in between custom-designed, padded upper and lower loading cups using a hydraulic or an electrical device, such as Dartec or Bose. The tibia model is advantageous over the ulna

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Fig. 1. Diagrammatic representation of the flexed mouse right hind-limb in position in the loading apparatus, showing the relative position of the bones, their relationship with the upper and lower loading cups (not drawn to scale).

model in that it allows the quantification of changes in both cortical and cancellous bone compartments. The methods for this in vivo murine loading model are described in this chapter (Fig. 1). 1.3. Assessment of Adaptive Modelling and Remodelling In Vivo

The adaptive response to mechanical loading can be assessed using a number of methods which are not be discussed in detail, as this information is covered elsewhere in this volume. If the duration of the loading regimen is sufficient (i.e. 3 days/week for 2 weeks), changes in bone geometry and cancellous bone architecture can be measured by microCT (Fig. 2). If the duration is shorter (i.e. 1–3 days) the loading response is best measured by histology and dynamic histomorphomety. By giving two short courses of flurochrome labels (i.e. calcein or alizarin), which binds to calcium at sites of new bone formation, bone formation can be visualised under the microscope in calcified sections of the bone (Fig. 3). Keep in mind that mechanical strain will vary along the length of the bone and so will the response to loading (see Note 2). Finite element modelling can be used to estimate the strain distribution along the diaphysis and in the cancellous bone and predict sites of greatest bone formation (36). Static histomorphometry is also useful

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Fig. 2. MicroCT images of the cancellous bone in a loaded (right) and non-loaded (left) proximal tibia of a female mouse. Two weeks of in vivo loading induced a significant increase in BV/TV mostly due an increase in trabecular thickness.

for assessing the response to loading in cancellous and cortical bone, that is, staining bone sections and quantifying the presence of osteoblasts, osteoclasts, osteoid, and eroded surface relative to how much bone is present. For cancellous bone, the percent bone volume fraction (BV/TV), surface of bone in the bone tissue volume (BS/TV), trabecular thickness (Tb.Th (μm)), and trabecular separation (Tb.Sp (μm)) can be quantified. 1.4. Determination of the In Vivo Mechanical Environment of Bone

The mechanical environment of bone can be sampled by measuring surface bone strain in vivo or ex vivo. This technique involves bonding electrical strain gauges to the bone’s surface that provide in vivo peak strain rates and magnitude data. It has been used in a large variety of vertebrates (37) and is considered the gold- standard for this purpose (38). It has been validated ex vivo by showing excellent correlation (within 2%) between strain recordings obtained simultaneously from strain gauges and optical extensometers (39, 40). Strain gauge data can also be used with bone imaging data to create a finite element model that allows the strain and stress distribution throughout the entire bone to be calculated. Strain gauging methodology is described below as it is critical to perform before any artificial bone loading experiment. During loading, a known force (in Newtons) must be applied to produce a level of strain that will induce an osteogenic response. To achieve this, strain gauges are attached to a representative group of tibiae or ulnae (n = 4–6). These are subsequently positioned in the loading apparatus, and the load magnitudes required to engender peak surface strains of between 500 and 3,000 με at the medial surface (37% of bone length from proximal end) are determined ex vivo. Our laboratory typically calibrates using data from the medial surface because the highest strains are recorded from this surface and is flatter and more uniform than the lateral surface, facilitating consistently reproducible strain measurements.

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Fig. 3. Histomorphometric evaluation of new bone formation at endosteal and periosteal diaphyseal locations. (a) An image of a transverse section though the diaphysis of the loaded mouse tibia, highlighting the calcein labels incorporated into the mineralising surfaces, obtained using a laser scanning confocal microscopy. (b) Periosteal and endosteal new bone formation.

2. Materials 2.1. Measurement of Bone Strains In Vivo

1. Saline: 0.9% NaCl solution. 2. Foam board. 3. Multimeter. 4. Strain gauge wire (38 TDQ wire from Phoenix Wire Co) (see Note 1). 5. Pins.

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6. 70% Ethanol. 7. Chloroform or Degreaser spray (Vishay, PA, USA). 8. Strain gauges. 9. Solder (361A-20R, Vishay, PA, USA). 10. Light Microscope. 11. Iron Razors. 12. Cotton swabs. 13. Scalpel blades. 14. Tweezers. 15. Glue Catalyst-C (Vishay, PA, USA). 16. Soldering M-Flux (Vishay, PA, USA). 17. M-Bond Adhesive Glue (Vishay, PA, USA). 18. CSM-1 degreaser (Vishay, PA, USA). 19. Amplifier (2100 Amplifier System; Vishay, PA, USA). 20. Dartec Machine. 21. Vicryl Suture 3.0 (Kirkton, Scotland)

3. Methods 3.1. Strain Gauge Preparation

1. Strain gauges are assembled into an implantable unit before the day of strain gauging. Using Vishay strain gauges (06-015-DJ120), tape the gauge down flat onto a dissecting board with Scotch tape and ensure that it is visible under a microscope. 2. Trim three sides of the backing using a single edged razor blade. 3. Lightly scratch each terminal using a pin (this will help the solder stay on the terminal) and apply a small drop of soldering M-Flux AR to each terminal. Then apply a small bead of solder to each terminal using a fine-tipped soldering iron. 4. Prepare the strain gauge lead wires by skinning the tips of two short (~10 cm in length) 38 TDQ wires that have been wrapped together. Dip the ends in flux and apply a small bead of solder to each end. 5. Re-melt the solder beads on the gauge terminals and insert the skinned strain gauge lead wires into the hot beads to establish electrical continuity. Allow the beads to cool and test the resistance of the gauge using a multimeter (should be 120.0 ± 0.3 Ω). If the gauge has deviated from this range the strain gauge has been damaged and should not be used. 6. Coat the gauge with a light spray of polyurethane (we use Clear Lacquer) to waterproof the gauge. 7. Trim the last side of the strain gauge that is stuck down by the Scotch tape.

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3.2. Prepare the Bones Ex Vivo

1. After euthanising the mouse, extract the limb with the bone you wish to attach the strain gauge to, leaving all the muscle and skin attached. Store in 70% alcohol. 2. The day before strain gauging, transfer the limb to 0.9% NaCl solution and incubate overnight at room temperature. 3. Securely pin the excised limb to a styrofoam board. If strain gauging the ulna, have the lateral side facing up. If strain gauging the tibia, expose the medial and/or lateral side. 4. Using a sharp scalpel blade, slice through the muscle to expose the underlying bone. Using a cotton tip, rub the bone to get a nice shine on the bone surface. Spray a clean swab with CSM-1 degreaser or dip in alcohol and quickly wipe down the centre of the bone to degrease.

3.3. Prepare the Mouse In Vivo

1. Pre-medicate the mice (0.5 mg/kg buprenorphine) by subcutaneous injection (Vetergestic, Animal Care Ltd., York, UK), and induce anaesthesia with oxygen and halothane (Rhone Merieux, Ltd., Essex, UK). Clip the fur on the limb to be gauged, wash with dilute povidone iodine, and finally rinse with 70% ethanol. 2. Make a skin incision on the lateral or medial aspect of mouse tibia. Undermine the skin and retract the fibrous attachment of the medial and lateral muscle masses. Periosteum is removed from the implantation site and the bone is gently scraped with scalpel blade to ensure removal of any tissues remaining attached to the bone. Swab with CSM-1 degreaser or alcohol soaked cotton tips to degrease the bone surface. A dry, clean site is necessary to ensure a good bond between the gauge backing film and the bone.

3.4. Attach the Gauge

1. When strain gauging, it is important that the strain gauge is positioned at the same site for each bone. This site can be determined by measuring bone length from radiographs and positioning the gauge at a calculated length along the bone (for example 50% of bone length) or by using anatomical landmarks. The site chosen must allow the gauge to be attached evenly and correspond to a region where significant osteogenic changes occur in response to loading. 2. If attaching ex vivo, use a pencil to mark the bone where the centre of the gauge should be attached. Brush some Catalyst-C onto a soft surface (we use the foil wrapper of an opened scalpel blade) and immediately dip the back of the gauge into the film of catalyst. Squeeze a small dot of M-Bond 200 Adhesive glue onto the same surface and quickly dip in the back of the gauge. 3. Lay the gauge in the correct position on the bone (the gauge should run parallel to the length of the bone) and gently push

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down on the solder bumps on the gauge terminals one at a time. Hold for ~20 s, and then prop up the wire with the tweezers so that the weight of the wire does not peel the gauge up. 4. If strain gauging ex vivo, wait 1.5–2.0 min and apply another thin coating of polyurethane spray to the bone. The purpose of this coating is to seal the places on the gauge terminals where you pushed with the tweezers. 5. If strain gauging ex vivo, store the bones in a vial containing saline to keep them hydrated. 6. If strain gauging in vivo, close the primary skin incision by an interrupted braised Vicryl Suture. The lead wires are fixed to the skin by interrupted suture at the lateral face of the tibia and on the dorsal neck. 7. As soon as the mice recovered from anaesthesia, data collection should begin. 3.5. Measure Strains

1. Connect the stain gauge wires to a shielded cable that leads to an amplifier that converts the change in resistance of each gauge to a change in voltage, which has a known relationship with microstrain. Connect the amplifier to the Dartec machine that will display the microstrain reading. Other amplifiers can be used; however, in the following we give only the methods specific to the 2100 Amplifier System in conjunction with the Dartec HC10 machine. 2. Before turning on the amplifier, check EXCIT toggle switches are turned off and the CAL switches are in the centre (OFF) position. Turn on the 2100 Amplifier System. The red pilot lamp should light up. Attach the 3-OUTPUT lead wires from the back of the amplifier to the power board of the Dartec machine. 3. Select a Channel from which you will connect the strain gauge to. Ensure that the Dartec is programmed to read this Channel (check software under Workshop → Configuration → Define Channels). 4. Connect the wires of a shielded cable to the input pin of the selected Channel. 5. Turn the Channel selector to “AC”, the meter should read between 9 and 11 on the scale. 6. Turn the Channel selector to “DC”, the meter should read close to 10 on the scale. 7. Turn the selector to the Channel you will use. For strain gauging we use ~1 V to minimise error due to self-heating. If the voltage needs to be adjusted, use a small screwdriver to read the desired BRIDGE VOLTS on the Power Supply Meter.

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8. Adjust the GAIN for the Channel. The gain needs to be high enough to detect the lowest output voltage, if it is too low you will only detect background noise. 9. Adjust the Amplifier Balance for the Channel with the EXIT switch off. Using a small screwdriver, adjust the AMP BAL until both OUTPUT lamps are off. 10. Turn on the Dartec HC10 machine, follow steps 2–7 and 12– 13 below. For step 5, have one “Feedback” window showing the microstrain (με) output from the Channel the strain gauge is attached. 11. Attach the two strain gauge wires to the shielded cable using a soldering iron. 12. Turn on the EXCIT switch for the selected Channel and adjust the balance to extinguish the OUTPUT lamps. In doing so you are balancing the Wheatstone bridge. 13. To check the Channel is calibrated correctly, turn the CAL switch to position A to show +1,000 με and to position B to show −1,000 με. Adjust these values if need be using the BALANCE. 3.5.1. For In Vivo Measurements

1. Record strain measurements during locomotion at different paces, over 5-s periods and a jump from a 30 cm height. 2. After recording, euthanise mice and perform a post-mortem to confirm robustness of union between gauges and bone surface.

3.5.2. For Ex Vivo Measurements

1. Position the bone with the gauge attached in-between the custom-built loading cups. Use the Cycle Generator or the Ramp Program to apply a range of loads (N) and record the corresponding strain (με) as shown in the Feedback Window. 2. Ensure that you get at least three repeatable measures for each load and there is a linear increase in microstrain with load.

3.6. Application of Bone Loads In Vivo

3.6.1. Setting up the Dartec

The following methods are specifi c to the Dartec HC10. This machine is one of many materials testing units available on the market today, other machines can be purchased from Endura TEC (BOSE Corp, USA), Instron Lt (USA) and Lloyd Instruments (AMTEK Inc, UK). 1. To use the Dartec HC10 you must be trained as an authorised user; if you have not been trained contact Zwick/Roell (www. zwick.com). If working in the UK, you must be working under an approved institutional project licence and have a personal licence from the Home Office (http://scienceandresearch. homeoffice.gov.uk/animal-research/) that allows you to conduct mechanical loading and anaesthetic procedures on rodents.

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Similar arrangements exist in other countries and should be strictly followed. 2. Turn on the Dartec machine by switching the button on the tower unit. 3. On the computer, open the software “Workshop 96” and select the window “Toolkit 96”. Ensure that the software shows a yellow light next to “Isolator”. The status display window will indicate the software is in SET UP mode. 4. Check the following are selected: under “Options → Toolbar and Monitor Line” and under “Settings → Auto Load Default Screen”. 5. The feedback can be displayed by selecting “Tools → Status” or by clicking the “Feedback” button from the toolbar. Under “Settings → Feedback Channel” select Actuator 1 and then “Clone” to open another window. In one window, go to “Monitor → Upper Peak” (the smallest recorded peak value) and in the other window select “Lower Peak” (the maximum recorded peak value). For both windows, select “Rest Interval” = 10 s. 6. On the top toolbar, click on “Pump Start” to start the hydraulic pump and select “Main Pressure” to apply hydraulic pressure to the actuator. 7. Open the “Systems Configuration” window. Open the “Offset/ Gains” window and zero the load cell with no sample mounted. Check that the “Offset Number” for the load cell is not greater than the capacity of the load cell. If it is, the load cell is damaged. 8. Under “Cycle Generator → Waveform” enter in the desired loading protocol. (See Notes 2 and 3 for a typical loading program and factors that need to be considered.) “Actuator” needs to be Actuator 1. “Control mode” is the mode it will cycle in (i.e. Load Cell). “Wave type” is the pattern of loading and “Number of Cycles” is the number of impacts you want to apply to the bone. Enter in the low and high magnitudes of load you wish to apply in “Level A and B”, respectively. Note the loads are negative indicating a compressive load. Enter in “Hold Time A and Hold Time B”, for the time to be spent at Level A and B (s) and the “Fall Time and Rise Time”, or strain rate (N/s). Depending on which waveform selected you may also need to enter in “Frequency” (number of cycles/s). Other variables such as “Amplitude, Mean, Fall Rate and Rise Rate” will be calculated automatically. Click on “Send”, “Read” to save. 9. Under “Cycle Generator Panel → Define Option”, select “Enable”. Under “Select Options Mode” select “Peak Control” and enter in the maximum and minimum loads (N) to be applied to the bone (same as values entered for Level A and B). This allows the upper and lower levels to be controlled and maintained. Click “Send, Read” to save.

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10. See Note 4 on Occupational Health and Safety for setting limits to the machine to prevent damage to the load cell and for checking the accuracy of the load cell. 11. Carefully screw in the upper and lower loading cups to the end of the actuator and load cell respectively. (Ideally screw in the lower cup when the machine is turned off.) Use the button ++ on the Manual Control panel to move the actuator up if needs be. The lower loading cup should protrude through a stage that the mouse will rest on during loading. Ensure that the stage does not touch the load cell or the lower cup. 12. In the “Offset/Gains” window ensure the Actuator is Actuator 1 and the Channel is Load Cell. The P (Proportional), I (Integral) and D (Derivative) gains need to be entered in the text boxes. The P value is less for stiffer samples and greater for more elastic samples. The I value is generally ~1/3 of the P value and the D value is ~1/10 of the P value. When entered, click on “Send Gains” and “Read Gains”. The Dartec is a close-loop system, which means the input signal is adjusted by the PID values to achieve the desired output. Therefore these values are critical for ensuring the waveform is not over or undershooting the desired load. For mouse bone, we use P = 12, I = 4 and D = 1.2. 13. To determine if the PID gains are correct, place a bone sample in between the loading cups (see Subheading 3.7 Mechanical Loading of Tibia/Ulna) and from “Tools” open “Oscilloscope”; this allows high speed capture displays from the actuator. For “Actuator” select Actuator 1, the “Capture Period” is the duration you want to capture the waveform and the “Feedback Channel” is the channel you wish to monitor (i.e. Load Cell). To change any parameters turn the “Scope” off, conversely when all parameters are set, enable the “Scope”. It is best to run a square wave to check the gains are correct. The PID gains can be changed from the “System” window until the waveform is read back as a perfect square wave. 14. When all parameters are set, induce anaesthesia to the animal using either an injectable or inhaled anaesthetic. It is preferable to use an inhaled anaesthetic (i.e. Halothane or Isoflurane) because it provides more control over the dosage and the animals recover faster. 3.7. Mechanical Loading of the Tibia/ Ulna

1. If loading the tibia, position the mouse’s right ankle in the lower cup and bend the knee so the tibia is vertical. Alternatively if loading the ulna, place the mouse’s right elbow in the lower cup. 2. Use the “Manual Control” window to move the actuator down (− = slow, –– = fast) until a force of approximately −2 N is applied to the knee or −0.2 N to the dorsal surface of the volar flexed knee.

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3. When the bone is positioned correctly, click on “Start” to begin loading. The mode will change from SET UP to LOAD CELL. 4. The “Cycle Generator” window shows you the number of cycles completed and the “Feedback” window indicates what loads are being applied to the bone. When the loading cycles are complete, click the “Global Setup” button in the top righthand corner so that the actuator is in SETUP mode. (This will help prevent damage to the load cell.) The “Manual Control” window will now display “mm”, not “N”. 5. Using the “Manual Control” panel move the actuator up (+ = slow, ++ = fast) and remove the mouse from the machine. 6. Weigh the mouse and observe, if it has recovered. When the animal is no longer in decubitus recombinant, they have fully recovered. 7. When all the mice have been loaded click “Pump Stop” and remove the upper loading cup. Turn off the machine at the main tower unit and close the software.

4. Notes 1. Strain gauges have a number of advantages; they are universal, simple to use, have low mass, show high stability over time, demonstrate excellent linearity over a large strain range, and are relatively cheap. However, they also have their disadvantages; the change in resistance is very low; therefore, an amplifier is needed to measure strain, they are for single use only, they need protection against temperature and moisture, and their correct positioning on bone is critical. 2. The typical protocol we use for mechanical loading of the mouse tibia is as follows: Actuator: Actuator 1 Control mode: 20 N load cell. Wave type: Trapezoid. No. cycles: 40. Level A: −0.5 N

Level B: −13 N

Amplitude: 5 N

Mean: −7 N

Time period

Frequency: not relevant for trapezoid waveform

Hold time A: 10 s or 14.9 s

Hold time B: 0.05 s

Fall time A: 0.025 s

Fall time B: 0.025 s

Fall rate: 400 N/s

Fall rise: 400 N/s

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3. It is important to consider how long each loading session will be, how many days/week you will apply loading, the frequency of the loading waveform, the type of waveform, the strain rate, and whether you will introduce a rest period in between loading cycles. Although no data are available on how bone responds to 3 alternate days versus 5 continuous days of axial loading a week, rest days allow the animal to recover from anaesthetic and stress to the joints. As reviewed earlier in this chapter, rest periods in between loading cycles (for example 10 or 14.9 s rest) are likely to enhance the osteogenic response to loading. 4. The following occupational Health and Safety instructions should be complied with the Dartec machine: ●

In case of emergency, use the red emergency stop button on the left-hand side of the Dartec machine.

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Reduce the pressure of the pump to avoid unnecessarily large forces being applied by the machine.

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Check the load cell is calibrated correctly by placing a known weight directly on the load cell (one that will not exceed the load cells capacity), and check that the load output in Newtons is correct. Use the following equation to calculate the expected output of a known weight: Kilogram (kg) × 9.807 = X N Therefore, if a 2 kg weight is placed on the load cell the output should read –19.614 N. This is the maximum compressive load you should place on a 20 N load cell (remember that compressive loads are negative numbers and tensile loads are positive numbers). If a heavier weight is used it will permanently damage the load cell.

●

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Under “Tools → Limits” set the error trips to stop the machine if it is overdriven. Error trips detect a difference between the required load and the actual load applied to the bone. To program error trips, select Mode → i.e. Load Cell and “Lower Limit” – enter in 1 N over the load cell capacity (i.e. 21 N), select an “Action” to take place if the limit is exceeded (i.e. Stop Generator) and under “Enable Limits” select “Active” “Send” and “Read”. Should one or more limit detectors be activated, the “Controller Status” display will change to indicate either DETECTED or ACTIVATED. These limits are crucial to prevent damage to the load cell. Do not place fingers in between the loading cups when the machine is running and ideally screw in the bottom-loading cup when the machine is turned off.

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INDEX A

B

Acidosis..... ........................................................................ 40 Adaptive modelling ................................................. 626–627 Adaptive remodelling .............................................. 626–627 Adenoviral vectors ........................................... 208, 213–216 Adipocytic differentiation............................................ 85, 91 Agarose gel electrophoresis...................................... 256–257 Alamar blue assay ............................................................ 218 ALIFF flow chamber............................................... 581–584 Alizarin red staining for mineralisation ......................... 13, 33, 36–39 in live cell imaging ..................................................... 439 Alkaline phosphatase enzyme activity assay ................................................... 92 in immunohistochemistry .................. 307–308, 315–316 staining solution ............................................ 85, 92, 308 staining kit ............................................................... 8, 37 Anaglyph images ..................................... 384–385, 417–418 Anaesthesia........ 87, 308, 485, 510, 513, 547, 549–550, 628, 632 Antibodies 23C6 (αvβ3), vitronectin receptor ..................... 149, 164 E11...... .................................................................. 70, 79 F4/80.... ..................................................................... 185 LM609 (αvβ3) .................................................. 135, 139 MoAb 121F............................................... 135, 139, 141 MAb OB 37.11 ........................................................... 51 MAb OB 7.3 ......................................................... 45, 51 SOX–9 ......................................................................... 86 STRO–1 .......................................................... 83, 88, 89 type II collagen ............................................................ 86 Antigen retrieval methods ................................. 94, 328–329 Apoptosis................................................................. 335–336 DNA laddering.......................................... 336, 342–344 nick translation assay ................................. 336–337, 342 caspase assay .............................................. 338, 345–346 toluidine blue staining ............................................... 340 Archimedes principle....................................... 560, 568–569 L-Ascorbic acid ........................................................... 12, 15 L-Ascorbic acid-2-phosphate........................................ 7, 13 Ashing....... .............................................................. 568, 570 Autofluorescence ..................................................... 333, 402 Autostainer ...................................................................... 323 Avian. See Chick

Backscatter electron (BSE) imaging. See SEM BCA protein assay ........................................................... 227 Bending strength by pQCT ........................................................... 490–491 by mechanical testing......................................... 565–566 Bioluminescence imaging (BLI) .............. 507–508, 512–513 Blood gas analyser ..................................................... 40, 113 Bone formation assay. See Bone nodule assay Bone histomorphometry. See Histomorphometry Bone marrow stem cells ............................................... 83–99 Bone marrow stromal cells................................................. 83 Bone metastases....................................................... 511–514 Bone nodule assay human osteoblasts.................................................. 12–13 rodent osteoblasts .................................................. 35–40 role of ascorbate ..................................................... 16–17 role of β-glycerophosphate .......................................... 38 role of pH .............................................................. 38–40 quantification............................................................... 37 Bone resorption assay. See Resorption pit assay Bone saw. See Cutting bone Bone slices. See Dentine slices

C Calcein labeling ....................................................... 291–293 Calvarial injection.................................................... 537–544 Calvarial bone, whole mount staining ............................. 455 Carbon dioxide concentration ......................... 338, 345–346 Caspase assay ........................................................... 345–346 Celestine Blue stain ......................................................... 543 Cell death. See Apoptosis Cell lines 2T3....... ............................................................. 433, 442 293 LTC .................................................................... 210 HEK293 .................................................................... 210 MDA-231B/Luc+ ..................................................... 509 MLO-A5............................................................... 67–81 MLO-Y4 ............................................................... 67–81 MG63............................................................................6 RAW 264.7 ............................................................... 197 SAOS-2 .........................................................................6 TE85.................................................................................6

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, © Springer Science+Business Media, LLC 2012

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BONE RESEARCH PROTOCOLS 638 Index Cement line stain ............................................ 283, 288, 297 Chick osteoblast. See Osteoblast, chick Chick osteoclast. See Osteoclast, chick Chick osteocyte. See Osteocyte, chick Chondrocyte ...................................................83, 93–95, 319 Chondrogenic differentiation ...................................... 89, 91 Chromatin immunoprecipitation (ChIP) assay ....... 241–246 Coating for SEM ............................................................ 376 Collagen production Sircol red assay............................................................. 37 Sirius red staining ........................................................ 36 Collagenase types ................................... 7, 14, 21, 26, 33, 45 Collagen coating of plastics ...................................... 71–72, 78–79 coating of glass slides ................................................. 581 gel for osteoclast culture .................................... 180–181 staining assay (see Sirius Red stain) Collagen production Sircol red assay............................................................. 37 Sirius red staining ........................................................ 36 Collagen type II staining ............................................. 94–95 Collagenase types ....................... 7, 14, 21, 26, 33, 45, 57, 84 Compression testing ........................................................ 565 Confocal laser scanning microscopy ........................ 401–423 colocalisation of signal ............................................... 415 principles and applications................................. 401–406 probes for osteoclasts ................................................. 408 Cortical bone analysis by pQCT ...................................... 484, 486–488 histomorphometry ............................................. 298–299 Critical point drying ........................................................ 375 CrossLaps for Culture kit, CTX-I .................................. 171 Cryogenic mill ................................................................. 250 CryoJane® Tape –Transfer System .................. 306, 310, 316 Cryopreservation and retrieval expanded macrophages .............................................. 167 primary osteoblasts ...................................................... 14 osteocyte cell lines ....................................................... 70 Cryosectioning ................................................................ 310 Crystal Violet stain ................................................ 70, 74–75 Cutting bone using aluminium oxide wheel .................................... 569 using band saw........................................................... 373 using milling tool ....................................... 373–374, 468 using low-speed diamond saw ................... 105, 374, 569

Dentine disks alternative mineralised substrates for ................. 112, 163 labelling with bisphosphonate ........................... 409–410 polishing ............................................................ 150, 184 preparation................................. 105–106, 123, 150, 163, 178, 190–191, 397 sources of dentine .............................................. 112, 397 DEPC water ............................................................ 251–264 Diffusion chambers ..............................12, 16, 17, 96–97, 99 DIG labelling, 311–312. See also Probes Dissection chick calvaria ......................................................... 46–48 human bone chips...................................................... 8–9 mouse calvaria.........................................24, 58, 437, 539 mouse long bone .............................22, 60, 308–309, 500 mouse vertebrae ......................................................... 500 rat calvaria ................................................................... 34 rat long bone .......................................................... 34–35 DMP-1 GFP mice .................................................... 56, 433 DNA laddering................................................ 336, 342–344 DNA quantification ................................................ 255–256 DNA extraction from cell cultures ....................................................... 252 from dried/embedded bone ............................... 252–253 with PicoGreen® ........................................... 85, 268 from fresh or frozen bone .................................. 251–252 Dual luciferase reporter assay .................................. 234–237 Dynamic histomorphometry. See Histomorphometry

D

F

Dartec loading machine........................................... 630–634 Data storage .............................................433, 452, 463–464 Decalcification with Decalcifying Solution-Lite ................................ 325 with EDTA ....................................................... 309, 339 with EDTA/glutaraldehyde for TEM ....................... 353 with EDTA on ultra-thin sections............................. 362

Faxitron..... ...................................................... 393, 499–506 File compression .............................................................. 450 Firefly luciferase ...................................................... 235–237 Fixation general considerations................................................ 328 glutaraldehyde for TRAP .......................................... 108 immersion fixation for TEM ............................. 355–356

E E11 antibody ....................................................... 70, 79, 451 Electrophoretic Mobility Shift Assay (EMSA) ......................................... 225, 238–241 Electron microscopy. See Transmission electron microscopy & Scanning electron microscopy Electrophoresis DNA.......................................................................... 343 Protein. ...................................................................... 229 RNA.... ...................................................................... 256 Embedding in epoxy resin ............................................................. 354 in glycol methacrylate ........................................ 538–540 in MMA ............................................................ 283–285 in OCT...................................................................... 310 Endosteal bone formation ............................................... 628

BONE RESEARCH PROTOCOLS 639 Index osmium/ferrocyanide for TEM ................................. 352 osmium tetroxide for TEM ....................................... 352 paraformaldehyde ...................................... 112, 323, 435 paraformaldehyde/glutaraldehyde mixture for IHC ......................................................... 323 paraformaldehyde/glutaraldehyde mixture for TEM ........................................................ 352 paraformaldehyde/glutaraldehyde mixture for SEM ......................................................... 372 perfusion fixation ....................................... 309, 354–355 picric formalin ........................................................... 338 Fluid flow................................................................. 573–592 flow chambers ALIFF ......................................................... 577, 579 Amsterdam .......................................................... 580 Flexcell® .............................................................. 578 Flexcell® Streamer® ............................................... 69 μSlide Ibidi®........................................................ 578 circuits.............................................72–74, 579, 582, 583 flow rate calculation ................................................... 590 oscillating flow ................................................... 576, 584 pulsatile flow ..................................................... 577–578, 585–586 shear stress calculation ............................................... 590 steady flow ......................................................... 583, 586 streaming potentials ........................................... 596–597 Fluid shear stress. See Fluid flow Fluorescent activated cell sorting (FACS) ................... 61–62 Fluorescent bisphosphonates ................................... 409–410 Fluorescence in vivo imaging................................... 507–508 Fluorescent probes for osteoclasts ........................... 408–409 Fluorescent probes for live cell imaging................... 432–452 Focal drift.. .............................................................. 432–433 Formation parameters. See Histomorphometry Four point bending .......................................... 602, 604–605 Fourier transform infrared imaging (FTIR) ............ 517–525 embedding materials.................................................. 518 fixation of samples ..................................................... 518 Fracture load .................................................................... 491 Freeze drying ........................................................... 373–375 Furnace...... ...................................................................... 558

G Gel shift assays ................................................................ 237 Gene expression. See qRT/PCR β-Glycerophosphate role in mineralising cultures ......................................... 38 Glycol methacrylate. See Resin Gold coating for SEM..................................................... 376 Gold-covered coverslips preparation of .................................................... 124–125 use in kinetic studies .................................. 133, 137–138 Goldner’s Masson Trichrome Stain .................353, 358–359, 538, 541

H HEK293 cells .................................................................. 210 Histology. See Fixation; Embedding; Staining Histomorphometry.................................................. 279–303 cortical bone ...................................................... 298–299 dynamic parameters ................................... 291–293, 299 mineralisation .................................................... 294–295 remodelling parameters ..................................... 297–298 resorption parameters ........................ 293–294, 296–297 structural parameters ......................... 290–291, 296–297 Homogenisers.................................................................. 258 Human bone marrow .................................................. 87–88 Human osteoblasts. See Osteoblasts Human osteoclasts. See Osteoclasts Hypoxia..... ................................................................ 68, 109

I Image analysis for microCT ...................................................... 467–472 for resorption pit assay ....................................... 182–183 for time lapse series............................................ 450–452 for X-ray imaging .............................................. 501–505 Image analysis software ............ 182, 373, 409, 447, 541–542 Immunomagnetic separation CD 14-positive cells .................................................. 166 osteoclasts .......................................... 129–131, 153–154 osteocytes..................................................................... 49 STRO-1 positive cells ................................................. 90 Immunohistochemistry antibody considerations ..................................... 329–330 antigen retrieval ........................................... 94, 328–329 using alkaline phosphatase ......................... 307–308, 315 using avidin-biotin............................................. 325–327 using β-galactosidase ................................. 123, 136–137 using peroxidase .......................................... 86, 325–327, 329–330, 332 Immunofluorescence amplification of signal with tyramide ........................ 332 autofluorescence......................................................... 333 staining method ......................................................... 154 Immunoprecipitation ............................................... 227–228 In situ hybridization (ISH)...................................... 305–320 probe preparation............................................... 310–312 tissue preparation ............................................... 308–310 Instron loading machine....... ................................... 560, 632 Intra-cardiac injection ............................................. 510–511 Intra-osseous inoculation......................................... 510–512

L 293 LTC cells .................................................................. 210 Lactate dehydrogenase (LDH) assay ............... 336, 340–341 Lead nitrate stain..................................................... 354, 360 Lentiviral vectors ..............................205, 207–208, 212–213

BONE RESEARCH PROTOCOLS 640 Index Live cell imaging ..................................................... 425–457 general considerations........................................ 425–433 generating movies ...................................................... 450 postacquisition image analysis ........................... 447–449 Loading of bones by perfusion (Zetos system) ............................... 611–612 in vitro.................................................................. 607–611 in vivo................................................................... 621–636 Loading machines ........................................................... 630 Lyso-ID® Green ............................................................. 408 Lysotracker dyes .............................................................. 408

M MAb OB7.3 ................................................................ 45, 51 MACS. See immunomagnetic separation MATra..................................................................... 234, 236 M-CSF....... ..................................................... 109, 163, 172 MDA-231 B/Luc+ cells .................................................. 509 MG63 cell line ....................................................................6 Magic Red stain .............................................................. 409 Magnetic separation. See Immunomagnetic separation McNeal’s stain ......................................................... 286–287 Mechanical loading. See Mechanical stimulation; Mechanical testing Mechanical stimulation. See also Fluid flow in vitro.................................................................. 593–617 in vivo................................................................... 621–636 Mechanical testing .................................................. 555–571 failure... ...................................................................... 561 flow chart ................................................................... 556 yield stress.................................................................. 551 strain.... ...................................................................... 551 Young’s modulus ................................................ 551, 566 Mechano-adaptation ............................................... 623–624 Metamorph software ....................................................... 447 Metastases in bone imaging by bioluminescence ...................................... 509 imaging by μCT......................................................... 473 Methylene blue stain ....................................................... 353 Methylmethacrylate (MMA) embedding................ 283–285 Microcomputed tomography (μCT) ....................... 461–476 ex vivo.. .............................................................. 468–473 focal lesions................................................................ 473 general considerations........................................ 461–463 in vivo... ............................................................. 473–475 resolution ........................................................... 465–466 sample holders ................................................... 468, 473 signal to noise ratio .................................................... 467 Micromilling of bone sections ......................... 285–286, 371 Mineralisation. See also Bone nodule assay Mineral maturity ............................................. 369, 521, 523 assessment by TEM ..................................... 39, 361–363 assessment by SEM ........................................... 369, 393

Mineralising osteoblasts for live cell imaging ........................................... 438–441 MLO-A5..................................................................... 67–81 culture conditions ........................................................ 77 mineralisation ........................................................ 77–78 MLO-Y4.........................................................................67–81 culture conditions .................................................. 70–71 dendrite length analysis ......................................... 74–76 transfection ............................................................ 76–77 Mouse bone marrow isolation ......................................... 179 Mouse osteoblast. See Osteoblast Mouse osteoclast. See Osteoclast Mouse osteocyte. See Osteocyte Mouse spleen cell isolation .............................................. 184

N Nanodrop......................................................................... 255 Nick translation assay ...................................... 336–337, 342 Nitric oxide measurement with DAR-4 .............................................................. 588 with Griess method ................................................... 589 Nude mice. ........................................................................ 87

O Oestrogen... ............................................................. 545–546 Oil red O stain....................................................... 87, 95, 97 Optical in vivo imaging ........................................... 507–514 Orchidectomy .......................................................... 545–549 Osteoblast chick isolation and culture (Obmix) ................................ 48 human...................................................................... 3–18 characterisation .................................................. 8, 11 isolation and culture........................................... 8–14 sources of bone for ................................................. 15 mouse characterisation ................................................ 27–28 for cocultures ................................................... 26–27 isolation and culture......................................... 22–27 rat characterisation ...................................................... 36 isolation and culture......................................... 31–35 Osteoclast chick antigenic profile ........................................... 134–135 characterisation ............................................ 131–139 culture .................................................................. 130 effect of pH on formation and activity .............. 109–112 F-actin staining ................................................. 148, 154 generation from human peripheral blood ...................... 164–166 from mouse bone marrow ............................ 106–116 from rabbit bone marrow ..................................... 153

BONE RESEARCH PROTOCOLS 641 Index from RAW 264.7 cells ................................. 187–202 in mouse cocultures...................................... 177–181 human analysis by quantitative vitronectin receptor staining .................................... 169–170 characterisation .................................................... 162 generation from buffy coats ......................... 164–165 generation from CD14+ cells....................... 165–166 generation from expanded macrophages .......................................... 168–169 generation from venous blood.............................. 164 isolation from chick bone ........................................... 125–131 from rabbit bone .......................................... 145–157 from rat bone ............................................... 107–108 immunomagnetic purification ................... 129, 153–154 immunostaining ................................................. 412–413 motility assay ..................................................... 137–138 mouse characterisation ............................................ 194–197 culture .......................................................... 106–108 culture on collagen ............................................... 180 from RAW 264.7 cells ................................. 187–202 serum purification ........................................ 192–194 survival ................................................................. 200 yield ..................................................................... 198 Percoll fractionation........................................... 127–128 purification by pronase-EDTA .......................... 153, 181 rabbit characterisation ............................................ 154–155 culture .......................................................... 150–151 immunomagnetic purification ..................... 153–154 plating density ..................................................... 151 rat culture .......................................................... 107–108 serum fractionation .................................... 152, 192–194 transfection ........................................................ 205–221 vitronectin receptor staining ...................................... 154 Osteocyte cell lines. (see also MLO-A5 and MLO-Y4) .......... 67–81 characterisation .......................................... 62–64, 67–69 chick characterisation .......................................... 46, 47, 51 isolation and culture......................................... 48–50 mouse from DMP1-GFP mice................................... 55–66 characterisation ................................................ 63–65 isolation and culture......................................... 58–62 resin casts in SEM ............................................. 389–391 Osteogenic differentiation ..................................... 85, 91–92 Osteogenic strain ............................................. 622, 625, 634 Osteoid staining. See Goldner’s trichrome Osteologic™ glass or dishes .................................... 138, 407 Osteosarcoma cell lines.................................................... 4–6

Osteosense ............................................................... 410, 680 Ovariectomy ............................................................ 545–548

P pCO2 measurement ................................................. 113–114 pH importance in resorption assays ......................... 109–112 importance in mineralisation assays ....................... 38–40 measurement with blood gas analyser ......................... 40, 112–114 pO2 measurement .................................................... 113–114 Paraformaldehyde fixative preparation .................... 112, 435 Parietal bone. See calvarial bone Periosteal bone formation ................................................ 495 Peripheral blood mononuclear cells (PBMC’s) isolation ......................................... 160, 164–167 Peripheral quantitative computed tomography (pQCT) imaging ................................... 477–498 cortical bone ...................................................... 477, 482 leg holders ......................................................... 485–486 paramaters ................................................................. 489 quality assurance ........................................................ 485 trabecular bone .................................................. 490–495 Phalloidin conjugates. See also F-actin staining ............... 409 Phototoxicity ................................................................... 429 Pit assay. See Resorption pit assay Primer design .......................................................... 266–267 Principal component analysis .................................. 531–532 Probe design ............................................................ 266–267 Promoter assay, dual luciferase ................................. 235–237 Pronase-EDTA solution

Q Quantitative computed tomography. See pQCT Quantitative reverse transcription polymerase chain reaction (qRT/PCR) assay and analysis ............................................... 261–275 delta-delta Ct method ....................................... 270–271 exogenous control ...................................................... 267 generating standards .................................................. 268 primer/probe design .......................................... 266–276 Quantitative X-ray imaging..................................... 499–506

R Rabbit osteoclast. See Osteoclast Raman microscopy .................................................. 527–532 RANKL ................................................................. 109, 163, 189, 198, 218 Rat osteoblast. See Osteoblast Rat osteoclast. See Osteoclast RAW 264.7 cells, culture ....................................................................... 197 use in osteoclast generation ............................... 191–192

BONE RESEARCH PROTOCOLS 642 Index Renilla luciferase...................................................... 235–237 Resorption, measurement of calcium ............................................ 171 measurement of collagen breakdown, CTX-I............ 171 Resorption pit assay analysis by confocal microscopy ................................. 418 analysis by reticle ....................................................... 138 analysis by SEM ................................................ 381–383 biochemical assays ..................................................... 171 counting stained pits with reflected light ............................................... 108–109, 115 counting stained pits with transmitted light ....................................................... 155, 171 importance of pH .............................................. 109–112 measuring resorbed area by dot counting system ........................................ 115 by automated image analysis ........................ 182–183 method ............................... 107–109, 138–139, 155, 171 Reverse transcription (RT) .............................................. 265 Reynolds number............................................................. 590 RNA extraction ..................................................... 62, 249–259 from cultured cells ....................................... 254–255 from fresh bone............................................ 253–254 from frozen bone ................................................. 254 quantification..................................................... 255–256

S Safranin O stain .............................................. 86, 89, 93–94 SaOS-2 cell line...................................................................6 Scanning electron microscopy (SEM) ..................... 365–399 backscattered electron imaging of resin embedded bone ....................................................... 387–389 coating of samples...................................................... 376 correlation with other imaging methods ............ 391–394 drying of samples ............................................... 374–376 general principles ............................................... 365–367 making casts of resin embedded bone................ 389–390 optimising image display ................................... 383–387 removal of organic material ............................... 377–381 SDS polyacrylamid gel ............................................ 228–230 Sectioning cryostat .............................................................. 310, 336 epoxy embedded bone................................................ 357 MMA embedded bone .............................................. 280 Serum gradient for purification of osteoclasts .....................152, 192–195, 199, 200 Shear stress. See also Fluid flow calculation.................................................................. 590 Shipping bone samples .................................................... 394 SOX-9 staining ........................................................... 94–95 Spectral imaging ...................................................... 418–419 Speed of sound ................................................ 558, 567–568

Stains/staining methods Alcian blue............................................................. 86, 97 Celestine Blue............................................................ 543 DAPI ........................................................ 123, 150, 154 Diff-Quick ................................................................ 122 Fast green .................................................................... 94 Goldner’s Masson trichrome ..................................... 353 Haematoxylin QS ...................................................... 327 Lead nitrate for TEM........................................ 354–360 Light Green ................................................... 86, 95, 541 Mayer’s haematoxylin ................................................ 282 Methylene blue .................................................. 353, 359 Oil Red O.............................................................. 87, 95 Safranin O ............................................................. 89, 93 Sirius red...................................................................... 97 Sytox Green ....................................................... 154, 408 Toluidine blue ............................................ 108, 282, 287 TO-PRO-3 ....................................................... 209, 408 Uranyl acetate for TEM .................................... 354, 359 Von Kossa ............................. 85, 287, 354, 359, 540–541 Weigert’s haematoxylin.................................. 86, 93, 538 Stiffness. See also Mechanical testing ............................... 566 Strain biaxial ................................................................ 599–601 definition ................................................................... 621 parameters ................................................................. 622 magnitude .................................................................. 597 measurements ............................................................ 625 uniaxial .............................................................. 601–602 Strain gauges ....................................612–613, 625, 627–630 Streaming potentials. See Fluid flow STRO-1. See Antibodies Structural parameters of bone.......................................... 290

T 2T3 cell line ..................................................................... 433 Tartrate-resistant acid phosphatase (TRAcP) concentration of tartrate ............................................ 185 fast-garnet method .................................................... 122 fast red method .................................................. 287–288 kit .............................................................. 106, 181–182 pararosanilin method ........................................ 149–150, 154–155, 179 quantitative colorimetric assay ........................... 163, 170 staining of GMA-embedded tissue ................... 540–541 TE85 cell line .............................................................. 6, 236 Tension testing ........................................................ 563–564 Tergazyme treatment for SEM imaging .......................... 378 Three point loading ................................................. 562–563 Tibia, in vivo loading model ............................................ 630 Toluidine blue stain ......................................... 106, 282, 287 Transfection ............................................................. 205–221 by Amaxa nucleofector ...................................... 211–212 by MATra magnetic system ............................... 234, 236

BONE RESEARCH PROTOCOLS 643 Index with adenovirus.................................................. 216–217 with lentivirus ............................................................ 213 with siRNA ............................................................... 207 Transmission electron microscopy ........................... 351–363 choice of buffer .......................................................... 361 decalcification ............................................................ 356 embedding ................................................................. 357 fixation............................................................... 354–355 sectioning................................................................... 357 staining of semi-thin sections ............................ 357–359 staining of ultra-thin sections ............................ 359–360

U Ultrasound ...................................................... 558, 567–568 Uniaxial straining .................................................... 603–605 Uranyl nitrate stain .................................................. 354, 359

V Vit C. See L-ascorbic acid Vitronectin receptor. See Antibodies Von Kossa stain .......................... 85, 287, 354, 359, 540–541 Von Kossa/McNeal stain ......................................... 286–287

W Weigert’s haematoxylin........................................ 86, 93, 538 Weightlessness ................................................................. 603 Western blotting ...................................................... 228–230 Wheat germ agglutinin ................................................... 408

X XCT scanners. See pQCT X-ray imaging.......................................................... 409–506