Skeletal Development and Repair: Methods and Protocols [2nd ed.] 9781071610275, 9781071610282

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Methods in Molecular Biology 2230

Matthew J. Hilton Editor

Skeletal Development and Repair Methods and Protocols Second Edition

METHODS

IN

MOLECULAR BIOLOGY

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

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

For over 35 years, biological scientists have come to rely on the research protocols and methodologies in the critically acclaimed Methods in Molecular Biology series. The series was the first to introduce the step-by-step protocols approach that has become the standard in all biomedical protocol publishing. Each protocol is provided in readily-reproducible step-bystep fashion, opening with an introductory overview, a list of the materials and reagents needed to complete the experiment, and followed by a detailed procedure that is supported with a helpful notes section offering tips and tricks of the trade as well as troubleshooting advice. These hallmark features were introduced by series editor Dr. John Walker and constitute the key ingredient in each and every volume of the Methods in Molecular Biology series. Tested and trusted, comprehensive and reliable, all protocols from the series are indexed in PubMed.

Skeletal Development and Repair Methods and Protocols Second Edition

Edited by

Matthew J. Hilton Department of Orthopaedic Surgery and Cell Biology, Duke Cellular, Developmental, and Genome Laboratories, Duke University School of Medicine, Durham, USA

Editor Matthew J. Hilton Department of Orthopaedic Surgery and Cell Biology Duke Cellular, Developmental, and Genome Laboratories Duke University School of Medicine Durham, USA

ISSN 1064-3745 ISSN 1940-6029 (electronic) Methods in Molecular Biology ISBN 978-1-0716-1027-5 ISBN 978-1-0716-1028-2 (eBook) https://doi.org/10.1007/978-1-0716-1028-2 © Springer Science+Business Media, LLC, part of Springer Nature 2014, 2021 This work is subject to copyright. All rights are reserved by the Publisher, whether the whole or part of the material is concerned, specifically the rights of translation, reprinting, reuse of illustrations, recitation, broadcasting, reproduction on microfilms or in any other physical way, and transmission or information storage and retrieval, electronic adaptation, computer software, or by similar or dissimilar methodology now known or hereafter developed. The use of general descriptive names, registered names, trademarks, service marks, etc. in this publication does not imply, even in the absence of a specific statement, that such names are exempt from the relevant protective laws and regulations and therefore free for general use. The publisher, the authors, and the editors are safe to assume that the advice and information in this book are believed to be true and accurate at the date of publication. Neither the publisher nor the authors or the editors give a warranty, expressed or implied, with respect to the material contained herein or for any errors or omissions that may have been made. The publisher remains neutral with regard to jurisdictional claims in published maps and institutional affiliations. This Humana imprint is published by the registered company Springer Science+Business Media, LLC, part of Springer Nature. The registered company address is: 1 New York Plaza, New York, NY 10004, U.S.A.

Preface This Second Edition of Skeletal Development and Repair: Methods and Protocols in the Methods in Molecular Biology series is designed as a comprehensive laboratory manual for all levels of basic research scientists working in the broad fields of skeletal development and skeletal repair research utilizing mouse models. The protocols highlighted here not only encompass the most current and cutting-edge techniques in skeletal development and repair but also showcase those protocols that have been modified and perfected over the course of several decades of skeletal research. These protocols presented by experts in the field include surgical, transplantation, organ culture, and parabiosis methods that permit analyses of skeletal tissues undergoing repair in vivo and permit analyses of cellular interactions ex vivo. Additionally, these methods include histological, cellular, and molecular techniques developed to study gene and protein expression in whole embryos, whole skeletal tissues, or tissue sections, as well as in vitro primary cell culture protocols designed to assay cellular, metabolic, and gene functions within specific cell populations. By design, most of the described methods utilize the laboratory mouse as the platform for surgical manipulation and/or transplantation as well as the source of tissues and cells for in vitro culture and analyses. The mouse has become the organism of choice for nearly all areas of skeletal research due to the development of numerous transgenic, cre recombinase expressing, and floxed mice available to the research community. The variety of skeletal research protocols contained in this volume will make it an invaluable tool that we hope will find its way into all labs studying skeletal development and repair using mice as their primary model system. Durham, NC, USA

Matthew J. Hilton

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

PART I

MURINE SKELETAL DEVELOPMENT, REPAIR, AND GENETIC MODELS

1 Overview of Skeletal Development . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Tatsuya Kobayashi and Henry M. Kronenberg 2 Overview of Skeletal Repair (Fracture Healing and Its Assessment) . . . . . . . . . . . Elise F. Morgan, Anthony De Giacomo, and Louis C. Gerstenfeld 3 Advantages and Limitations of Cre Mouse Lines Used in Skeletal Research . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Florent Elefteriou and Greig Couasnay

PART II

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SKELETAL REPAIR, PARABIOSIS, TRANSPLANTATIONS, AND ORGAN CULTURES

4 Generation of Closed Transverse Fractures in Small Animals . . . . . . . . . . . . . . . . . Anthony De Giacomo, Elise F. Morgan, and Louis C. Gerstenfeld 5 A Mouse Femoral Ostectomy Model to Assess Bone Graft Substitutes. . . . . . . . . Ryan P. Trombetta, Emma K. Knapp, and Hani A. Awad 6 Surgical Induction of Posttraumatic Osteoarthritis in the Mouse. . . . . . . . . . . . . . Robert D. Maynard, David A. Villani, William G. Schroeder, Douglas J. Adams, and Michael J. Zuscik 7 Parabiosis: Assessing the Effects of Circulating Cells and Factors on the Skeleton . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Benjamin Alman and Gurpreet Baht 8 Murine Limb Bud Organ Cultures for Studying Musculoskeletal Development . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Martin Arostegui and T. Michael Underhill 9 Murine Limb Explant Cultures to Assess Cartilage Development . . . . . . . . . . . . . Manuela Wuelling and Andrea Vortkamp 10 Renal Capsule Transplantation to Assay Angiogenesis in Skeletal Development and Repair . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Anais Julien, Simon Perrin, Rana Abou-Khalil, and Ce´line Colnot

PART III

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63 75 91

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STRUCTURAL, HISTOLOGICAL, AND MOLECULAR ANALYSES ON SKELETAL TISSUES AND TISSUE SECTIONS

MicroCT for Scanning and Analysis of Mouse Bones . . . . . . . . . . . . . . . . . . . . . . . . 169 Yung Kim, Michael D. Brodt, Simon Y. Tang, and Matthew J. Silva

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Four-Point Bending Testing for Mechanical Assessment of Mouse Bone Structural Properties . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Hattie C. Cutcliffe and Louis E. DeFrate Phenotyping Intact Mouse Bones Using Bone CLARITY. . . . . . . . . . . . . . . . . . . . Jennifer B. Treweek, Aidan Beres, Nathan Johnson, and Alon Greenbaum Processing and Sectioning Undecalcified Murine Bone Specimens . . . . . . . . . . . . Thomas B. Bemenderfer, Jonathan S. Harris, Keith W. Condon, Jiliang Li, and Melissa A. Kacena Preparation of Thin Frozen Sections from Nonfixed and Undecalcified Hard Tissues Using Kawamoto’s Film Method (2020) . . . . . . . . . . . . . . . . . . . . . . Tadafumi Kawamoto and Komei Kawamoto Demineralized Murine Skeletal Histology. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Anthony J. Mirando and Matthew J. Hilton Assessment of Osteocytes: Techniques for Studying Morphological and Molecular Changes Associated with Perilacunar/Canalicular Remodeling of the Bone Matrix . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Neha S. Dole, Cristal S. Yee, Charles A. Schurman, Sarah L. Dallas, and Tamara Alliston Cell Lineage Tracing: Colocalization of Cell Lineage Markers with a Fluorescent Reporter . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Yan Jing, Patricia Simmer, and Jian Q. Feng Immunofluorescent Staining of Adult Murine Paraffin-Embedded Skeletal Tissue . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Neta Felsenthal and Elazar Zelzer Detection of Hypoxic Regions in the Bone Microenvironment . . . . . . . . . . . . . . . Wendi Guo and Colleen Wu EdU-Based Assay of Cell Proliferation and Stem Cell Quiescence in Skeletal Tissue Sections. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Marco Angelozzi, Charles R. de Charleroy, and Ve´ronique Lefebvre Whole Mount In Situ Hybridization in Murine Tissues. . . . . . . . . . . . . . . . . . . . . . Deepika Sharma, Matthew J. Hilton, and Courtney M. Karner

PART IV 23

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PRIMARY CELL ISOLATIONS, CULTURES, AND ASSAYS

Bone Marrow Stromal Cell Assays: In Vitro and In Vivo. . . . . . . . . . . . . . . . . . . . . 379 Pamela G. Robey, Sergei A. Kuznetsov, Paolo Bianco, and Mara Riminucci Isolation and Culture of Periosteum-Derived Progenitor Cells from Mice . . . . . 397 Chinedu C. Ude, Girdhar G. Sharma, Jie Shen, and Regis J. O’Keefe Isolation and Culture of Murine Primary Chondrocytes: Costal and Growth Plate Cartilage . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 415 Yihan Liao, Jason T. Long, Christopher J. R. Gallo, Anthony J. Mirando, and Matthew J. Hilton

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Isolation and Culture of Neonatal Mouse Calvarial Osteoblasts. . . . . . . . . . . . . . . Madison L. Doolittle, Cheryl L. Ackert-Bicknell, and Jennifer H. Jonason Mitochondrial Function and Metabolism of Cultured Skeletal Cells . . . . . . . . . . . Li Tian, Clifford J. Rosen, and Anyonya R. Guntur Radiolabeled Amino Acid Uptake Assays in Primary Bone Cells and Bone Explants . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Leyao Shen and Courtney M. Karner RANKL-Based Osteoclastogenic Assay from Murine Bone Marrow Cells. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Zhenqiang Yao, Lianping Xing, and Brendan F. Boyce Hematopoietic Stem Cell Cultures and Assays . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Benjamin J. Frisch

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

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Contributors RANA ABOU-KHALIL • Univ Paris Est Creteil, INSERM, IMRB, Creteil, France CHERYL L. ACKERT-BICKNELL • Department of Orthopaedics and Rehabilitation, Center for Musculoskeletal Research, University of Rochester, Rochester, NY, USA; Department of Orthopedics, University of Colorado Anschutz Medical Campus, Aurora, CO, USA DOUGLAS J. ADAMS • Department of Orthopedics, Anschutz Medical Campus, University of Colorado, Aurora, CO, USA TAMARA ALLISTON • Department of Orthopaedic Surgery, University of California, San Francisco, San Francisco, CA, USA; UC Berkeley/UCSF Graduate Program in Bioengineering, San Francisco, CA, USA BENJAMIN ALMAN • Department of Orthopaedic Surgery, Duke University, Durham, NC, USA MARCO ANGELOZZI • Division of Orthopedic Surgery, Department of Surgery, Children’s Hospital of Philadelphia, Philadelphia, PA, USA MARTIN AROSTEGUI • Biomedical Research Centre, University of British Columbia, Vancouver, BC, Canada; Department of Cellular and Physiological Sciences, University of British Columbia, Vancouver, BC, Canada HANI A. AWAD • Center for Musculoskeletal Research, University of Rochester Medical Center, Rochester, NY, USA; Department of Biomedical Engineering, University of Rochester, Rochester, NY, USA GURPREET BAHT • Department of Orthopaedic Surgery, Duke Molecular Physiology Institute, Duke University, Durham, NC, USA THOMAS B. BEMENDERFER • Department of Orthopaedic Surgery, Indiana University School of Medicine, Indianapolis, IN, USA AIDAN BERES • Joint Department of Biomedical Engineering, North Carolina State University and University of North Carolina at Chapel Hill, Raleigh, NC, USA PAOLO BIANCO • Sapienza University of Rome, Rome, Italy BRENDAN F. BOYCE • Department of Pathology and Laboratory Medicine, University of Rochester Medical Center, Rochester, NY, USA MICHAEL D. BRODT • Department of Orthopaedic Surgery and Musculoskeletal Research Center, Washington University, Saint Louis, MO, USA CE´LINE COLNOT • Univ Paris Est Creteil, INSERM, IMRB, Creteil, France KEITH W. CONDON • Department of Anatomy, Cell Biology and Physiology, Indiana University School of Medicine, Indianapolis, IN, USA GREIG COUASNAY • Department of Orthopedic Surgery, Baylor College of Medicine, Houston, TX, USA HATTIE C. CUTCLIFFE • Department of Orthopaedic Surgery, Duke University, Durham, NC, USA SARAH L. DALLAS • Department of Oral and Craniofacial Sciences, School of Dentistry, University of Missouri Kansas City, Kansas City, MO, USA CHARLES R. DE CHARLEROY • Division of Orthopedic Surgery, Department of Surgery, Children’s Hospital of Philadelphia, Philadelphia, PA, USA LOUIS E. DEFRATE • Department of Orthopaedic Surgery, Duke University, Durham, NC, USA; Department of Biomedical Engineering, Duke University, Durham, NC, USA;

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Contributors

Department of Mechanical Engineering and Materials Science, Duke University, Durham, NC, USA ANTHONY DE GIACOMO • Department of Orthopedic Surgery, Woodland Hills Medical Center, Woodland Hills, CA, USA; Boston University School of Medicine, Boston, MA, USA NEHA S. DOLE • Department of Orthopaedic Surgery, University of California, San Francisco, San Francisco, CA, USA MADISON L. DOOLITTLE • Department of Orthopaedics and Rehabilitation, Center for Musculoskeletal Research, University of Rochester, Rochester, NY, USA FLORENT ELEFTERIOU • Department of Molecular and Human Genetics, Baylor College of Medicine, Houston, TX, USA; Department of Orthopedic Surgery, Baylor College of Medicine, Houston, TX, USA NETA FELSENTHAL • Department of Molecular Genetics, Weizmann Institute of Science, Rehovot, Israel JIAN Q. FENG • Department of Biomedical Sciences, Texas A&M University College of Dentistry, Dallas, TX, USA BENJAMIN J. FRISCH • Department of Pathology and Laboratory Medicine, University of Rochester School of Medicine and Dentistry, Rochester, NY, USA; Center for Musculoskeletal Research, University of Rochester School of Medicine and Dentistry, Rochester, NY, USA; James P. Wilmot Cancer Institute, University of Rochester School of Medicine and Dentistry, Rochester, NY, USA CHRISTOPHER J. R. GALLO • Department of Orthopaedic Surgery, Duke Cellular, Developmental, and Genome Laboratories, Durham, NC, USA LOUIS C. GERSTENFELD • Department of Orthopaedic Surgery, Orthopaedic Research Laboratory, Boston University School of Medicine, Boston, MA, USA ALON GREENBAUM • Joint Department of Biomedical Engineering, North Carolina State University and University of North Carolina at Chapel Hill, Raleigh, NC, USA; Comparative Medicine Institute, North Carolina State University, Raleigh, NC, USA; Bioinformatics Research Center, North Carolina State University, Raleigh, NC, USA ANYONYA R. GUNTUR • Center for Molecular Medicine, Maine Medical Center Research Institute, Scarborough, ME, USA; Tufts University School of Medicine, Tufts University, Boston, MA, USA; Graduate School of Biomedical Sciences and Engineering, University of Maine, Orono, ME, USA WENDI GUO • Department of Orthopaedic Surgery, Duke University School of Medicine, Durham, NC, USA; Department of Pharmacology and Cancer Biology, Duke University School of Medicine, Durham, NC, USA JONATHAN S. HARRIS • Department of Orthopaedic Surgery, Indiana University School of Medicine, Indianapolis, IN, USA MATTHEW J. HILTON • Department of Orthopaedic Surgery and Cell Biology, Duke Cellular, Developmental, and Genome Laboratories, Duke University School of Medicine, Durham, USA YAN JING • Department of Orthodontics, Texas A&M University College of Dentistry, Dallas, TX, USA NATHAN JOHNSON • Joint Department of Biomedical Engineering, North Carolina State University and University of North Carolina at Chapel Hill, Raleigh, NC, USA JENNIFER H. JONASON • Department of Orthopaedics and Rehabilitation, Center for Musculoskeletal Research, University of Rochester, Rochester, NY, USA ANAIS JULIEN • Univ Paris Est Creteil, INSERM, IMRB, Creteil, France

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MELISSA A. KACENA • Department of Orthopaedic Surgery, Indiana University School of Medicine, Indianapolis, IN, USA; Department of Anatomy, Cell Biology and Physiology, Indiana University School of Medicine, Indianapolis, IN, USA; Richard L. Roudebush VA Medical Center, Indianapolis, IN, USA COURTNEY M. KARNER • Department of Orthopaedic Surgery, Duke Orthopaedic, Cellular, Developmental and Genome Laboratories, Duke University School of Medicine, Durham, NC, USA; Department of Cell Biology, Duke University School of Medicine, Durham, NC, USA KOMEI KAWAMOTO • Radioisotope Research Institute, School of Dental Medicine, Tsurumi University, Yokohama, Japan TADAFUMI KAWAMOTO • Radioisotope Research Institute, School of Dental Medicine, Tsurumi University, Yokohama, Japan YUNG KIM • Department of Orthopaedic Surgery and Musculoskeletal Research Center, Washington University, Saint Louis, MO, USA EMMA K. KNAPP • Center for Musculoskeletal Research, University of Rochester Medical Center, Rochester, NY, USA TATSUYA KOBAYASHI • Massachusetts General Hospital, Harvard University, Boston, MA, USA HENRY M. KRONENBERG • Massachusetts General Hospital, Harvard University, Boston, MA, USA SERGEI A. KUZNETSOV • Skeletal Biology Section, National Institute of Dental and Craniofacial Research, National Institutes of Health, Department of Health and Human Services, Bethesda, MD, USA VE´RONIQUE LEFEBVRE • Division of Orthopedic Surgery, Department of Surgery, Children’s Hospital of Philadelphia, Philadelphia, PA, USA YIHAN LIAO • Departments of Orthopaedic Surgery, Duke Cellular, Developmental, and Genome Laboratories, Durham, NC, USA; Pharmacology and Cancer Biology, Duke Cellular, Developmental, and Genome Laboratories, Durham, NC, USA JILIANG LI • Department of Biology, Indiana University Purdue University Indianapolis, Indianapolis, IN, USA JASON T. LONG • Departments of Orthopaedic Surgery, Duke Cellular, Developmental, and Genome Laboratories, Durham, NC, USA; Cell Biology, Duke Cellular, Developmental, and Genome Laboratories, Duke University School of Medicine, Durham, NC, USA ROBERT D. MAYNARD • Department of Orthopedics, Anschutz Medical Campus, University of Colorado, Aurora, CO, USA ANTHONY J. MIRANDO • Department of Orthopaedic Surgery, Duke University School of Medicine, Duke Cellular, Developmental, and Genome Laboratories, Durham, NC, USA; Department of Cell Biology, Duke University School of Medicine, Duke Cellular, Developmental, and Genome Laboratories, Durham, NC, USA ELISE F. MORGAN • Boston University School of Medicine, Boston, MA, USA; Department of Mechanical Engineering, College of Engineering, Boston University, Boston, MA, USA REGIS J. O’KEEFE • Department of Orthopaedic Surgery, Washington University School of Medicine, St. Louis, MO, USA SIMON PERRIN • Univ Paris Est Creteil, INSERM, IMRB, Creteil, France MARA RIMINUCCI • Sapienza University of Rome, Rome, Italy PAMELA G. ROBEY • Skeletal Biology Section, National Institute of Dental and Craniofacial Research, National Institutes of Health, Department of Health and Human Services, Bethesda, MD, USA

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Contributors

CLIFFORD J. ROSEN • Center for Molecular Medicine, Maine Medical Center Research Institute, Scarborough, ME, USA; Center for Clinical and Translational Research, Maine Medical Center Research Institute, Scarborough, ME, USA; Tufts University School of Medicine, Tufts University, Boston, MA, USA; Graduate School of Biomedical Sciences and Engineering, University of Maine, Orono, ME, USA WILLIAM G. SCHROEDER • Department of Orthopedics, Anschutz Medical Campus, University of Colorado, Aurora, CO, USA CHARLES A. SCHURMAN • Department of Orthopaedic Surgery, University of California, San Francisco, San Francisco, CA, USA; UC Berkeley/UCSF Graduate Program in Bioengineering, San Francisco, CA, USA DEEPIKA SHARMA • Department of Orthopaedic Surgery, Duke Orthopaedic, Cellular, Developmental and Genome Laboratories, Duke University School of Medicine, Durham, NC, USA GIRDHAR G. SHARMA • Department of Orthopaedic Surgery, Washington University School of Medicine, St. Louis, MO, USA JIE SHEN • Department of Orthopaedic Surgery, Washington University School of Medicine, St. Louis, MO, USA LEYAO SHEN • Department of Orthopaedic Surgery, Duke University School of Medicine, Duke University, Durham, NC, USA MATTHEW J. SILVA • Department of Orthopaedic Surgery and Musculoskeletal Research Center, Washington University, Saint Louis, MO, USA PATRICIA SIMMER • Department of Biomedical Sciences, Texas A&M University College of Dentistry, Dallas, TX, USA SIMON Y. TANG • Department of Orthopaedic Surgery and Musculoskeletal Research Center, Washington University, Saint Louis, MO, USA LI TIAN • Center for Molecular Medicine, Maine Medical Center Research Institute, Scarborough, ME, USA JENNIFER B. TREWEEK • Department of Biomedical Engineering, University of Southern California, Los Angeles, CA, USA RYAN P. TROMBETTA • Orthopedic Trauma Department, US Army Institute for Surgical Research, San Antonio, TX, USA CHINEDU C. UDE • Department of Orthopaedic Surgery, Washington University School of Medicine, St. Louis, MO, USA T. MICHAEL UNDERHILL • Biomedical Research Centre, University of British Columbia, Vancouver, BC, Canada; Department of Cellular and Physiological Sciences, University of British Columbia, Vancouver, BC, Canada; School of Biomedical Engineering, University of British Columbia, Vancouver, BC, Canada DAVID A. VILLANI • Cell Biology, Stems Cells and Development Program, Anschutz Medical Campus, University of Colorado, Aurora, CO, USA ANDREA VORTKAMP • Developmental Biology, Centre for Medical Biology, University Duisburg-Essen, Essen, Germany COLLEEN WU • Department of Orthopaedic Surgery, Duke University School of Medicine, Durham, NC, USA; Department of Pharmacology and Cancer Biology, Duke University School of Medicine, Durham, NC, USA; Department of Cell Biology, Duke University School of Medicine, Durham, NC, USA MANUELA WUELLING • Developmental Biology, Centre for Medical Biology, University Duisburg-Essen, Essen, Germany

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LIANPING XING • Department of Pathology and Laboratory Medicine, University of Rochester Medical Center, Rochester, NY, USA ZHENQIANG YAO • Department of Pathology and Laboratory Medicine, University of Rochester Medical Center, Rochester, NY, USA CRISTAL S. YEE • Department of Orthopaedic Surgery, University of California, San Francisco, San Francisco, CA, USA ELAZAR ZELZER • Department of Molecular Genetics, Weizmann Institute of Science, Rehovot, Israel MICHAEL J. ZUSCIK • Department of Orthopedics, Anschutz Medical Campus, University of Colorado, Aurora, CO, USA; Center for Musculoskeletal Research, University of Rochester Medical Center, Rochester, NY, USA

Part I Murine Skeletal Development, Repair, and Genetic Models

Chapter 1 Overview of Skeletal Development Tatsuya Kobayashi and Henry M. Kronenberg Abstract Development of cartilage and bone, the core components of the mouse skeletal system, depends on coordinated proliferation and differentiation of skeletogenic cells, including chondrocytes and osteoblasts. These cells differentiate from common progenitor cells originating in the mesoderm and neural crest. Multiple signaling pathways and transcription factors tightly regulate differentiation and proliferation of skeletal cells. In this chapter, we overview the process of mouse skeletal development and discuss major regulators of skeletal cells at each developmental stage. Key words Bone development, Chondrocyte, Osteoblast, Mouse, Mesenchymal condensation, Method, Skeletal development, Transcription factor, Signaling

Mice became the most popular animal model for studying vertebral skeletal development for several reasons. Their small body size and rapid reproductive cycle facilitate experimentation. Their genetic manipulability and skeletal biology that shares many features with human skeletal biology set them apart from other model organisms, such as rats that currently have limited genetic options and zebrafish that do not have a remodeling skeleton. The remarkable advances in understanding skeletal development in the last two decades mainly owe to technological breakthroughs in mouse genetics and molecular and cellular biology. Mice will continuously be indispensable primary experimental models for skeletal biology until a better research model is developed in response to the increasing demand for research directly relevant to human physiology. In this chapter, we wish to provide a concise overview of mouse skeletal development. We trace the process of mouse skeletal development mainly at the cellular and molecular levels, touching upon major signaling systems and transcription factors that regulate the process. We also briefly discuss applications and limitations of available analytical methods for studying skeletal development in mice.

Matthew J. Hilton (ed.), Skeletal Development and Repair: Methods and Protocols, Methods in Molecular Biology, vol. 2230, https://doi.org/10.1007/978-1-0716-1028-2_1, © Springer Science+Business Media, LLC, part of Springer Nature 2021

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Origin of Skeletogenic Cells The mouse skeletal system is comprised of multiple tissue types including, for example, bone, cartilage, dentin, muscle, tendon, and ligaments. Distinct cell types generate these diverse tissues through serial differentiation steps. They all derive from two distinct embryological origins: mesoderm and neural crest [1]. These embryonic tissues produce mesenchymal cells that further differentiate into specific cell types composing the mouse skeleton. We focus on development of bone and cartilage that are the major components of skeletons. Bone and cartilage provide the framework of the skeletal system while interacting with other types of tissues during development [2]. Cartilage and bone have distinct functions, yet these tissues closely interact during development. Cartilage is formed by chondrocytes that produce abundant extracellular matrix mainly composed of collagens (particularly type II, IX, and XI collagens) and proteoglycans (such as aggrecan). The unique alignment of collagen fibers and proteoglycans allows the tissue to hold a large amount of water, thus providing cartilage with substantial elasticity [3]. This feature is necessary for the function of permanent cartilage such as that in joints and ribs. Cartilage also serves as a temporary template for formation of new bone (the growth plate). Unlike permanent cartilage, growth plate cartilage is continuously being replaced by bone. Therefore, tightly coordinated proliferation and differentiation of growth plate chondrocytes are necessary for longitudinal growth of long bones [4]. Mineralized bones are formed by osteoblasts through chondrocyte-dependent (endochondral bone formation) and chondrocyte-independent (intramembranous bone formation) mechanisms. The former process, mainly used in bones that show longitudinal growth, requires formation of cartilage templates (“growth plates” postnatally) comprised of chondrocytes. In contrast, osteoblasts directly differentiate from their progenitor cells in intramembranous bones such as calvariae. Both osteoblasts and chondrocytes differentiate from common progenitors, mesenchymal cells. Mesenchymal cells originating from the neural crest migrate rostrally and form bone, cartilage, and dentin in the head. Some bones in the craniofacial region, such as calvariae, form through intramembranous bone formation that does not require cartilage templates. Unlike bones in the head, bones in vertebral and appendicular skeletons are formed by mesenchymal cells originating from the paraxial mesoderm and lateral plate mesoderm, respectively. These bones grow through endochondral bone formation, in which cartilage template formation precedes development of mineralized bone.

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Mesenchymal Condensation After migration, mesenchymal cells condense, temporarily stop proliferating, and form clusters that exclude blood vessels (mesenchymal condensation) [5, 6]. These cells then differentiate into chondrocytes and perichondrial cells in endochondral bones, or into osteoblast progenitor cells in intramembranous bones. In mouse limb buds, mesenchymal condensation occurs around E11 in forelimbs. The mechanism regulating this process is not fully understood, but signaling pathways triggered by transforming growth factor ß, Bone morphogenetic proteins (BMP) and fibroblast growth factors (FGF) play critical roles [5]. The transcription factor, Sox9, is essential for mesenchymal condensation both in neural crest-derived and mesoderm-derived condensations [7–9].

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Chondrocyte Differentiation and Endochondral Bone Formation In endochondral bone formation, condensed mesenchymal cells further differentiate into two different cell types. Chondrocytes differentiate from the central part of the condensation, and perichondrial cells differentiate from cells of the outer layer. Upon differentiation into chondrocytes, cells start proliferating again to form a cartilage template for future bone. BMP signaling appears to play a critical role in this process [10, 11]. Chondrocytes further proliferate relatively uniformly, until cells in the center of the cartilage differentiate into hypertrophic chondrocytes. This initial hypertrophic differentiation occurs around E13-14 in mouse tibiae. The mechanism regulating the initial hypertrophy is not clear. However, since parathyroid hormone–related peptide (PTHrP, encoded by Pthlh) overactivity delays this process [12, 13], PTHrP signaling may play a role in determining the timing of the initial hypertrophy. Once hypertrophic chondrocytes appear, chondrocytes in the cartilage start forming a polarized structure comprised of primarily three different chondrocyte layers. The periarticular layer near the end of the cartilage is comprised of round, non–column-forming chondrocytes with a moderate proliferation rate. Some periarticular chondrocytes form the joint surface, while others differentiate into flat, column-forming proliferating chondrocytes that proliferate vigorously. Columnar chondrocytes then differentiate into postmitotic hypertrophic chondrocytes. Upon differentiation into hypertrophic chondrocytes, cells rapidly increase their cellular volumes. This increase in cell volume, likely mediated by an increase in fluid uptake [14] is a major engine of growth plate lengthening, the major process that drives bone lengthening. From this perspective the orchestrated proliferation of chondrocytes can be viewed as a

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way of generating sufficient hypertrophic chondrocytes to lengthen bone sufficiently. The flat columns of proliferating chondrocytes serve both to direct the lengthening along the axis of long bones, as well as to produce matrix that also contributes to the expansion of the growth plate. The complex regulation of the stages of chondrocyte differentiation are regulated by signals expressed by perichondrial cells (FGFs and BMPs, for example) as well as by chondrocytes themselves. The mechanism for the “ballooning” of hypertrophic chondrocytes is not known, but Insulin-like growth factor 1 (IGF1) signaling [14] and extracellular signal-regulated kinase (ERK) signaling [15] appear to positively and negatively regulate the increase in cell size, respectively. In addition to contributing to bone lengthening, hypertrophic chondrocytes regulate the surrounding matrix through mineralization, regulate the differentiation of adjacent perichondrial cells into osteoblasts, regulate the proliferation of their precursors (flat proliferating chondrocytes), and attract blood vessels and osteoblast precursors to the primary spongiosa. Thus, they can be considered the master cell of the growth plate. Hypertrophic chondrocytes and prehypertrophic chondrocytes express Indian hedgehog (Ihh), a critical multifunctional signaling molecule that accomplishes many of these roles of hypertrophic chondrocytes. Ihh stimulates expression of PTHrP by periarticular chondrocytes, proliferation and differentiation of periarticular chondrocytes, proliferation of flat, columnar chondrocytes, and induction of osteoblast differentiation in endochondral bones [16]. Ihh stimulation of PTHrP expression in periarticular chondrocytes and perichondrial cells prevents premature hypertrophic differentiation and thus suppresses expression of Ihh itself. This PTHrP-Ihh feedback loop coordinates chondrocyte differentiation to maintain the growth plate structure while bones achieve dramatic increases in size during development. Hypertrophic chondrocytes further differentiate to express alkaline phosphatase and direct the mineralization of surrounding matrix. Late hypertrophic chondrocytes then express osteopontin (Spp1), matrix metallopeptidase 13 (Mmp13; collagenase 3), vascular endothelial growth factor (VEGF), and RANK ligand (Rankl). Mineralized hypertrophic chondrocytes are subsequently resorbed by chondro/osteoclasts, leaving a remnant of extracellular cartilage matrix upon which invading blood vessels and osteoblasts attach to form the primary spongiosa. Several recent studies suggest that a subset of hypertrophic chondrocytes can differentiate into cells of the osteoblast lineage [17–19].

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Mechanisms for Regulating Growth Plate Chondrocytes Regulation of proliferation, differentiation, and survival of growth plate chondrocytes has been intensively studied. With regard to signaling molecules, chondrocyte proliferation is usually negatively regulated by FGF signaling [20] and positively regulated by insulinlike growth factor 1 (IGF-1) signaling [21]. Mitogen-activated protein kinase (MAPK) pathways differentially regulate chondrocyte proliferation and differentiation in a pathway-specific manner [15, 22]. Appropriate dosage of beta-catenin, the critical mediator of canonical Wnt signaling, is essential for normal chondrocyte differentiation and proliferation [23]. The phosphoinositide-3kinase (PI3K)/Akt pathway positively regulates proliferation and growth [24, 25], while Type C natriuretic peptide (CNP) signaling regulates chondrogenesis by antagonizing FGF signaling [26]. Chondrocyte differentiation is regulated at multiple steps. Differentiation of periarticular to columnar chondrocytes is positively regulated by Ihh, which also regulates PTHrP expression. PTHrP signaling is necessary to prevent premature hypertrophic differentiation of columnar proliferating chondrocytes. Other signaling systems including those triggered by FGFs and BMPs are known to regulate hypertrophic differentiation [27]. Several transcription factors are known to regulate chondrocyte proliferation and differentiation [28, 29]. For example, Runx2, a transcription factor essential for osteoblast differentiation promotes hypertrophic differentiation [30, 31]. Myocyte enhancer factor 2 (Mef2) family transcription factors, critical for muscle development, and their inhibitory regulator, histone deacetylase 4 (Hdac4), play a critical role in hypertrophic differentiation [32, 33]. Regulators of the Mef2–Hdac4 interaction thus play important roles. For example, salt-inducible kinase family kinase 3 (Sik3) [34] and Protein phosphatase 2A [35] change the phosphorylation status of HDAC4 to regulate hypertrophic differentiation. Regulators of chondrocyte survival include Hif1a, which was shown to be indispensable for chondrocyte survival in the hypoxic environment of cartilage and joints [36]. The master transcription factor, Sox9, and associated Sox transcription factors are essential for chondrocyte survival and proliferation [37]. Sox9 likely regulates highly expressed housekeeping genes in addition to chondrocyte-specific genes by directly and indirectly binding to regulatory elements in the genome [38]. MicroRNAs, a relatively newly understood class of regulatory mechanisms, appear to play important roles in skeletogenesis [39]; chondrocytic miRNAs are necessary for normal proliferation, differentiation, and survival [40, 41], while miRNA deficiency in bone results in increased bone mass [42, 43].

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Development of Joints, Articular Cartilage, Tendons, and Ligaments Joint formation starts with appearance of the interzone, a cluster of flat, condensed cells, in the cartilage anlage. These early joint progenitor cells express specific markers, such as Wnt9a and Gdf5, signaling by which coordinately regulates normal joint development [44]. Joint progenitor cells further differentiate into different types of cells in synovial joints, including synoviocytes and articular chondrocytes. Articular cartilage exhibits a structure distinct from that of growth plate cartilage. It is covered by the superficial layer comprised of flat cells expressing Prg4. This unique cell population serves as “progenitor cells” for articular chondrocytes [45]. This study demonstrated that articular cartilage slowly increases its size through appositional growth. The mechanism by which the size and structure of articular cartilage are defined in adult bone has not been determined. Evidence suggests that regulation of joint progenitor cells during embryonic joint morphogenesis is a major determinant [46]. Progenitors of tendons and ligaments are first detected by expression of Scx [47], a transcription factor required for normal tendon and ligament development. Other transcription factors, including Mkx and Egr1, follow Scx. TGF-β and FGF signaling plays critical roles in inducing their differentiation [48]. Tendons connect bone and muscle. Transition zones between tendon and bone (the enthesis), or tendon and muscle (the myotendinous junction, or MTJ) subsequently develop to show unique histological and cellular phenotypes.

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Osteoblast Differentiation and Bone Development Initial bone formation in endochondral bones starts in the perichondrium adjacent to the hypertrophic zone of the growth plate. Ihh produced by hypertrophic chondrocytes signals to perichondrial cells to induce osteoblast differentiation [49]. Ihh signaling is essential for osteoblast differentiation of endochondral bones, while hedgehog signaling is not essential for osteoblasts of intramembranous bones. Wnt/beta-catenin signaling is also necessary for osteoblast differentiation [50]. In endochondral bones, the hedgehog signaling appears to induce Wnt ligand expression to initiate osteoblast differentiation [51]. These signaling molecules ultimately regulate gene expression through modulating expression and/or functions of transcription factors. Runx2 was the first transcription factor demonstrated to be essential for osteoblast differentiation [52, 53]. Another essential transcription factor for osteoblast differentiation is Sp7 (Osterix), which acts at a level genetically downstream of Runx2 [54]. The precise mechanisms by which Runx2 and Sp7 regulate osteoblast differentiation are

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unknown. Osteoblasts further differentiate into mature osteoblasts that vigorously produce bone matrix proteins including type I collagen and osteocalcin. ATF4, an important mediator of the endoplasmic reticulum stress pathway, regulates osteocalcin and type I collagen expression in osteoblasts [55].

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The Life of Osteoblasts We are gaining better understanding of the origin, life, and fate of individual osteoblasts mainly owing to the advancement of genetic tools. For example, lineage tracing of genetically labeled osteoblasts revealed that different subsets of cells in the osteoblast lineage follow different fates [56, 57]. Both marrow and cortical osteoblasts appear to originate from perichondrial cells of developing bone. A subset of these cells that expresses a collagen II transgene represent perichondrial chondro-osteoblast progenitors [58]. These perichondrial cells share some common properties with the pericytes surrounding bone marrow blood vessels that have been shown in adult bone to also have the potential to form osteoblasts and osteocytes in vivo [59]. While these mesenchymal cells appear to be the primary source of osteoblasts during normal development and likely serve as osteoprogenitors in postnatal stages, other cell types in other settings may contribute to the osteoblast lineage. Recent studies have provided genetic evidence that hypertrophic and even earlier resting stage chondrocytes may transdifferentiate into osteoblastic cells in vivo [17–19, 60]. Vascular endothelial cells that, in the setting of inflammatory stimuli and activation of BMP signaling in the disease, fibrodysplasia ossificans progressiva, can become chondrocytes and osteoblasts in vivo [61]. Some circulating cells of hematopoietic origin express markers of the osteoblast lineage and can calcify matrix after implantation into nude mice [62]. Thus, the identification of precursors of osteoblasts during development and postnatal life remains an important research agenda. Many osteoblasts that lay down mineralized bone matrix die, but some osteoblasts further differentiate into osteocytes and bone lining cells. Osteocytes embedded in the mineralized bone matrix are connected to each other to form a fine network, and thus may sense and mediate effects of mechanical stress on osteoblasts to regulate bone homeostasis. Recently, osteocytes have been recognized also as an important regulator of bone metabolism and systemic mineral homeostasis [63]. From the point of view of bone development, it is noteworthy that osteocytes are the major source of sclerostin (encoded by Sost) that inhibits Wnt signaling, and thus represents a mechanism whereby osteocytes can suppress osteoblast development. Recently it was shown that Sost expression is regulated by the SIK/HDAC/MEF2 axis and that PTH,

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through PKA targeting of SIK, can suppress Sost expression in osteocytes in a fashion analogous to the way that PTHrP targets SIK3 to suppress chondrocyte hypertrophy in the growth plate [64, 65]. This finding is particularly intriguing that two very different cell types in the skeleton, although diverged from common progenitors, use a common regulatory mechanism to control two very different processes, growth plate development and osteoblast homeostasis. In addition to osteoblasts and osteocytes, genetic evidence demonstrates that cells of the osteoblast lineage give rise to other types of stromal cells, such as Cxcl12-abundnat reticular (CAR) cells [66, 67] that play a critical role in supporting hematopoietic stem cells [68]. CAR cells are considered to constitute a cell population that mostly overlaps with those expressing Leptin receptor (Lepr) and may include osteoprogenitor cells in adult bone [69, 70].

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Skeletal Stem Cells After birth, the skeleton continues to grow and, throughout adult life, remodeling of normal bone and repair of injured bone necessitates regulated new bone formation. The idea that skeletal precursors in cartilage, periosteum, marrow, and perhaps elsewhere might act as stem cells to provide new cells to bone throughout life is a plausible hypothesis. Through development of analytical tools and technologies for study of bone cells, such as genetic labeling and flow cytometry, several studies have identified markers for distinct and overlapping stem/progenitor populations of the chondrocyte and osteoblast lineages, such as Sox9 [67], Cd105 /Cd200+ [71], Acta2 [72], Lepr [70], Grem1 [73], Gli1 [74], Pthlh [60], and Ctsk/Cd200+ [75]. Currently, the relationships among osteoprogenitor cell populations identified in these different studies, if any, is uncertain and the distinct roles of these cells in mediating postnatal skeletal homeostasis and response to injury are under study.

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Bone Remodeling and Osteoclastogenesis Bones grow while maintaining their shapes and functions through continuous modeling and remodeling. Osteoclasts initiate bone remodeling by resorbing bone matrix. Osteoclasts differentiate from monocytic precursors of the hematopoietic lineage. Multiple transcription factors, such as PU.1, MITF, Fos, NFkb, and NFATc1, as well as signaling molecules, including M-CSF and RANKL, are known to be essential for osteoclastogenesis [76, 77]. RANKL is produced by hypertrophic chondrocytes and by cells of the osteoblast lineage, including osteocytes, and is crucial

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for the needed communication among these cell types during bone development [78]. Defects in osteoclast functions usually result in osteopetrosis, and increases in osteoclast activity leads to osteopenia. Osteoclasts and osteoblasts interact directly and indirectly, and thus osteoclasts play important roles during bone development. However, because osteoclasts and bone remodeling have been investigated mainly in the context of postdevelopmental stages, we will not further review the vast literature related to osteoclasts and bone remodeling in this chapter.

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Experimental Methods for Studying Mouse Skeletal Development Skeletal development has been studied at the organ, tissue, cell, and molecular levels. Technological advances in molecular and cell biology, and mouse genetics greatly influenced the way mammalian skeletal development is investigated. Most importantly, genetic manipulation in mice has significantly advanced our understanding of mammalian skeletal development at the cellular and molecular levels in vivo. Because of the difficulty in reconstructing bone and cartilage tissues in vitro, investigations using reverse mouse genetics will continue to be a valuable tool for understanding the interactions of cells and tissues of bone. In addition, several technological breakthroughs combined with mouse genetics to lead to remarkable discoveries in skeletal biology. Among them, mouse genome manipulation with the CR ISPR/Cas9 technology, advanced cellular analysis using flow cytometric analysis, and various single cell technologies are particularly noteworthy. However, there is still large room for technical improvement toward thorough understanding of skeletal development. We would like to conclude this chapter with a brief discussion of a future agenda for research methodologies in skeletal biology. In addition to general limitations of available analytical methods, the unique properties of skeletal tissues still pose idiosyncratic problems in investigation of mouse skeletal development. For example, the following are a few technical and biological problems: 1. Limited direct manipulability in vivo. Other than through genetic means, it is difficult to manipulate developing mouse skeletal tissues. Although there are a few reports, such as intrauterine surgical manipulation [79], direct, physical manipulation is technically demanding. 2. Difficulty in dynamic analysis in vivo. Analysis of bone development heavily depends on static histological sections, while skeletal development is a dynamic process. Development of methods that allow for longitudinal observation of bone development at the cellular level is highly desired.

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3. Difficulty in reconstituting the in vivo environment in vitro. In vitro systems have a variety of advantages over in vivo systems in manipulability and reproducibility. However, reconstituting the process of bone development in vitro has not been very successful. 4. Difficulty in applying biochemical and molecular analyses. Primarily for the reason stated above, existing biochemical or molecular biological methods are difficult to directly apply to in vivo samples. For example, gene expression analysis using intact bones mostly depends on in situ hybridization, but in situ hybridization is prone to technical variability, and lacks sensitivity and quantitative accuracy. 5. Biological differences between rodents and humans. The structure and mechanical properties of the bones of rodents are different from those of humans. Cortical bone in humans contains Haversian systems that are absent from the much smaller rodent bones, for example. Nevertheless, it is remarkable how much the structure and signaling systems, as well as effects of mutations and drugs are usually similar between rodents and humans. Particularly for studies of conditions related with mechanical stresses, such as osteoarthritis, mice may not be an ideal model of human diseases. Humanization of the mouse skeleton is currently not possible. In summary, research in mouse bone development has been remarkably advanced due to the development of the genetic manipulation technology and understanding of the mouse genome. As our desire for deeper understanding of bone development increases, we encounter limitations of currently available analytical methods and model systems. Therefore, research in this field will greatly benefit from future technological breakthroughs such as direct micromanipulation of developing bones and highresolution, real-time imaging, and development of novel model systems.

Acknowledgments This work is supported by NIH grants, DK011794 to H.K. and AR056645 to T.K. References 1. Olsen BR, Reginato AM, Wang W (2000) Bone development. Annu Rev Cell Dev Biol 16:191–220 2. Sharir A, Stern T, Rot C, Shahar R, Zelzer E (2011) Muscle force regulates bone shaping

for optimal load-bearing capacity during embryogenesis. Development 138 (15):3247–3259 3. Mow VC, Holmes MH, Lai WM (1984) Fluid transport and mechanical properties of

Skeletal Development articular cartilage: a review. J Biomech 17 (5):377–394 4. Kronenberg HM (2003) Developmental regulation of the growth plate. Nature 423 (6937):332–336 5. Goldring MB, Tsuchimochi K, Ijiri K (2006) The control of chondrogenesis. J Cell Biochem 97(1):33–44 6. Hall BK, Miyake T (2000) All for one and one for all: condensations and the initiation of skeletal development. BioEssays 22(2):138–147 7. Akiyama H, Chaboissier MC, Martin JF, Schedl A, de Crombrugghe B (2002) The transcription factor Sox9 has essential roles in successive steps of the chondrocyte differentiation pathway and is required for expression of Sox5 and Sox6. Genes Dev 16(21):2813–2828 8. Bi W, Deng JM, Zhang Z, Behringer RR, de Crombrugghe B (1999) Sox9 is required for cartilage formation. Nat Genet 22(1):85–89 9. Mori-Akiyama Y, Akiyama H, Rowitch DH, de Crombrugghe B (2003) Sox9 is required for determination of the chondrogenic cell lineage in the cranial neural crest. Proc Natl Acad Sci U S A 100(16):9360–9365 10. Niswander L (2002) Interplay between the molecular signals that control vertebrate limb development. Int J Dev Biol 46(7):877–881 11. Yoon BS, Ovchinnikov DA, Yoshii I, Mishina Y, Behringer RR, Lyons KM (2005) Bmpr1a and Bmpr1b have overlapping functions and are essential for chondrogenesis in vivo. Proc Natl Acad Sci U S A 102 (14):5062–5067 12. Schipani E, Lanske B, Hunzelman J, Luz A, Kovacs CS, Lee K et al (1997) Targeted expression of constitutively active receptors for parathyroid hormone and parathyroid hormonerelated peptide delays endochondral bone formation and rescues mice that lack parathyroid hormone-related peptide. Proc Natl Acad Sci U S A 94(25):13689–13694 13. Weir EC, Philbrick WM, Amling M, Neff LA, Baron R, Broadus AE (1996) Targeted overexpression of parathyroid hormone-related peptide in chondrocytes causes chondrodysplasia and delayed endochondral bone formation. Proc Natl Acad Sci U S A 93 (19):10240–10245 14. Cooper KL, Oh S, Sung Y, Dasari RR, Kirschner MW, Tabin CJ (2013) Multiple phases of chondrocyte enlargement underlie differences in skeletal proportions. Nature 495 (7441):375–378 15. Murakami S, Balmes G, McKinney S, Zhang Z, Givol D, de Crombrugghe B (2004) Constitutive activation of MEK1 in chondrocytes causes Stat1-independent achondroplasia-like

13

dwarfism and rescues the Fgfr3-deficient mouse phenotype. Genes Dev 18(3):290–305 16. Kronenberg HM (2006) PTHrP and skeletal development. Ann N Y Acad Sci 1068:1–13 17. Yang L, Tsang KY, Tang HC, Chan D, Cheah KS (2014) Hypertrophic chondrocytes can become osteoblasts and osteocytes in endochondral bone formation. Proc Natl Acad Sci U S A 111(33):12097–12102 18. Park J, Gebhardt M, Golovchenko S, PerezBranguli F, Hattori T, Hartmann C et al (2015) Dual pathways to endochondral osteoblasts: a novel chondrocyte-derived osteoprogenitor cell identified in hypertrophic cartilage. Biol Open 4(5):608–621 19. Zhou X, von der Mark K, Henry S, Norton W, Adams H, de Crombrugghe B (2014) Chondrocytes transdifferentiate into osteoblasts in endochondral bone during development, postnatal growth and fracture healing in mice. PLoS Genet 10(12):e1004820 20. Ornitz DM, Marie PJ (2015) Fibroblast growth factor signaling in skeletal development and disease. Genes Dev 29(14):1463–1486 21. Wang Y, Cheng Z, Elalieh HZ, Nakamura E, Nguyen MT, Mackem S et al (2011) IGF-1R signaling in chondrocytes modulates growth plate development by interacting with the PTHrP/Ihh pathway. J Bone Miner Res 26 (7):1437–1446 22. Zhang R, Murakami S, Coustry F, Wang Y, de Crombrugghe B (2006) Constitutive activation of MKK6 in chondrocytes of transgenic mice inhibits proliferation and delays endochondral bone formation. Proc Natl Acad Sci U S A 103(2):365–370 23. Akiyama H, Lyons JP, Mori-Akiyama Y, Yang X, Zhang R, Zhang Z et al (2004) Interactions between Sox9 and beta-catenin control chondrocyte differentiation. Genes Dev 18 (9):1072–1087 24. Ford-Hutchinson AF, Ali Z, Lines SE, Hallgrimsson B, Boyd SK, Jirik FR (2007) Inactivation of Pten in osteochondroprogenitor cells leads to epiphyseal growth plate abnormalities and skeletal overgrowth. J Bone Miner Res 22(8):1245–1259 25. Yang G, Sun Q, Teng Y, Li F, Weng T, Yang X (2008) PTEN deficiency causes dyschondroplasia in mice by enhanced hypoxia-inducible factor 1alpha signaling and endoplasmic reticulum stress. Development 135(21):3587–3597 26. Yasoda A, Komatsu Y, Chusho H, Miyazawa T, Ozasa A, Miura M et al (2004) Overexpression of CNP in chondrocytes rescues achondroplasia through a MAPK-dependent pathway. Nat Med 10(1):80–86

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Tatsuya Kobayashi and Henry M. Kronenberg

27. Goldring MB (2012) Chondrogenesis, chondrocyte differentiation, and articular cartilage metabolism in health and osteoarthritis. Ther Adv Musculoskelet Dis 4(4):269–285 28. Liu CF, Samsa WE, Zhou G, Lefebvre V (2017) Transcriptional control of chondrocyte specification and differentiation. Semin Cell Dev Biol 62:34–49 29. Sun MM, Beier F (2014) Chondrocyte hypertrophy in skeletal development, growth, and disease. Birth Defects Res C Embryo Today 102(1):74–82 30. Enomoto H, Enomoto-Iwamoto M, Iwamoto M, Nomura S, Himeno M, Kitamura Y et al (2000) Cbfa1 is a positive regulatory factor in chondrocyte maturation. J Biol Chem 275(12):8695–8702 31. Takeda S, Bonnamy JP, Owen MJ, Ducy P, Karsenty G (2001) Continuous expression of Cbfa1 in nonhypertrophic chondrocytes uncovers its ability to induce hypertrophic chondrocyte differentiation and partially rescues Cbfa1-deficient mice. Genes Dev 15 (4):467–481 32. Arnold MA, Kim Y, Czubryt MP, Phan D, McAnally J, Qi X et al (2007) MEF2C transcription factor controls chondrocyte hypertrophy and bone development. Dev Cell 12 (3):377–389 33. Vega RB, Matsuda K, Oh J, Barbosa AC, Yang X, Meadows E et al (2004) Histone deacetylase 4 controls chondrocyte hypertrophy during skeletogenesis. Cell 119(4):555–566 34. Sasagawa S, Takemori H, Uebi T, Ikegami D, Hiramatsu K, Ikegawa S et al (2012) SIK3 is essential for chondrocyte hypertrophy during skeletal development in mice. Development 139(6):1153–1163 35. Kozhemyakina E, Cohen T, Yao TP, Lassar AB (2009) Parathyroid hormone-related peptide represses chondrocyte hypertrophy through a protein phosphatase 2A/histone deacetylase 4/MEF2 pathway. Mol Cell Biol 29 (21):5751–5762 36. Rankin EB, Giaccia AJ, Schipani E (2011) A central role for hypoxic signaling in cartilage, bone, and hematopoiesis. Curr Osteoporos Rep 9(2):46–52 37. Henry SP, Liang S, Akdemir KC, de Crombrugghe B (2012) The postnatal role of Sox9 in cartilage. J Bone Miner Res 27 (12):2511–2525 38. Ohba S, He X, Hojo H, McMahon AP (2015) Distinct transcriptional programs underlie Sox9 regulation of the mammalian chondrocyte. Cell Rep 12(2):229–243

39. Harfe BD, McManus MT, Mansfield JH, Hornstein E, Tabin CJ (2005) The RNaseIII enzyme Dicer is required for morphogenesis but not patterning of the vertebrate limb. Proc Natl Acad Sci U S A 102 (31):10898–10903 40. Kobayashi T, Lu J, Cobb BS, Rodda SJ, McMahon AP, Schipani E et al (2008) Dicerdependent pathways regulate chondrocyte proliferation and differentiation. Proc Natl Acad Sci U S A 105(6):1949–1954 41. Kobayashi T, Papaioannou G, Mirzamohammadi F, Kozhemyakina E, Zhang M, Blelloch R et al (2015) Early postnatal ablation of the microRNA-processing enzyme, Drosha, causes chondrocyte death and impairs the structural integrity of the articular cartilage. Osteoarthr Cartil 23 (7):1214–1220 42. Raaijmakers MH, Mukherjee S, Guo S, Zhang S, Kobayashi T, Schoonmaker JA et al (2010) Bone progenitor dysfunction induces myelodysplasia and secondary leukaemia. Nature 464(7290):852–857 43. Gaur T, Hussain S, Mudhasani R, Parulkar I, Colby JL, Frederick D et al (2010) Dicer inactivation in osteoprogenitor cells compromises fetal survival and bone formation, while excision in differentiated osteoblasts increases bone mass in the adult mouse. Dev Biol 340 (1):10–21 44. Salva JE, Merrill AE (2017) Signaling networks in joint development. Dev Dyn 246 (4):262–274 45. Kozhemyakina E, Zhang M, Ionescu A, Ayturk UM, Ono N, Kobayashi A et al (2015) Identification of a Prg4-expressing articular cartilage progenitor cell population in mice. Arthritis Rheumatol 67(5):1261–1273 46. Kobayashi T, Kozlova A (2018) Lin28a overexpression reveals the role of Erk signaling in articular cartilage development. Development 145(15):dev162594 47. Schweitzer R, Chyung JH, Murtaugh LC, Brent AE, Rosen V, Olson EN et al (2001) Analysis of the tendon cell fate using Scleraxis, a specific marker for tendons and ligaments. Development 128(19):3855–3866 48. Asahara H, Inui M, Lotz MK (2017) Tendons and ligaments: connecting developmental biology to musculoskeletal disease pathogenesis. J Bone Miner Res 32(9):1773–1782 49. Long F, Chung UI, Ohba S, McMahon J, Kronenberg HM, McMahon AP (2004) Ihh signaling is directly required for the osteoblast lineage in the endochondral skeleton. Development 131(6):1309–1318

Skeletal Development 50. Long F (2011) Building strong bones: molecular regulation of the osteoblast lineage. Nat Rev Mol Cell Biol 13(1):27–38 51. Hu H, Hilton MJ, Tu X, Yu K, Ornitz DM, Long F (2005) Sequential roles of Hedgehog and Wnt signaling in osteoblast development. Development 132(1):49–60 52. Komori T, Yagi H, Nomura S, Yamaguchi A, Sasaki K, Deguchi K et al (1997) Targeted disruption of Cbfa1 results in a complete lack of bone formation owing to maturational arrest of osteoblasts. Cell 89(5):755–764 53. Otto F, Thornell AP, Crompton T, Denzel A, Gilmour KC, Rosewell IR et al (1997) Cbfa1, a candidate gene for cleidocranial dysplasia syndrome, is essential for osteoblast differentiation and bone development. Cell 89(5):765–771 54. Nakashima K, Zhou X, Kunkel G, Zhang Z, Deng JM, Behringer RR et al (2002) The novel zinc finger-containing transcription factor osterix is required for osteoblast differentiation and bone formation. Cell 108(1):17–29 55. Yang X, Matsuda K, Bialek P, Jacquot S, Masuoka HC, Schinke T et al (2004) ATF4 is a substrate of RSK2 and an essential regulator of osteoblast biology; implication for CoffinLowry Syndrome. Cell 117(3):387–398 56. Park D, Spencer JA, Koh BI, Kobayashi T, Fujisaki J, Clemens TL et al (2012) Endogenous bone marrow MSCs are dynamic, faterestricted participants in bone maintenance and regeneration. Cell Stem Cell 10 (3):259–272 57. Maes C, Kobayashi T, Selig MK, Torrekens S, Roth SI, Mackem S et al (2010) Osteoblast precursors, but not mature osteoblasts, move into developing and fractured bones along with invading blood vessels. Dev Cell 19 (2):329–344 58. Colnot C, Lu C, Hu D, Helms JA (2004) Distinguishing the contributions of the perichondrium, cartilage, and vascular endothelium to skeletal development. Dev Biol 269 (1):55–69 59. Mendez-Ferrer S, Michurina TV, Ferraro F, Mazloom AR, Macarthur BD, Lira SA et al (2010) Mesenchymal and haematopoietic stem cells form a unique bone marrow niche. Nature 466(7308):829–834 60. Mizuhashi K, Ono W, Matsushita Y, Sakagami N, Takahashi A, Saunders TL et al (2018) Resting zone of the growth plate houses a unique class of skeletal stem cells. Nature 563(7730):254–258 61. Medici D, Shore EM, Lounev VY, Kaplan FS, Kalluri R, Olsen BR (2010) Conversion of

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vascular endothelial cells into multipotent stem-like cells. Nat Med 16(12):1400–1406 62. Fadini GP, Albiero M, Menegazzo L, Boscaro E, Vigili de Kreutzenberg S, Agostini C et al (2011) Widespread increase in myeloid calcifying cells contributes to ectopic vascular calcification in type 2 diabetes. Circ Res 108 (9):1112–1121 63. Bonewald LF (2017) The role of the osteocyte in bone and nonbone disease. Endocrinol Metab Clin N Am 46(1):1–18 64. Wein MN, Liang Y, Goransson O, Sundberg TB, Wang J, Williams EA et al (2016) SIKs control osteocyte responses to parathyroid hormone. Nat Commun 7:13176 65. Nishimori S, Wein MN, Sakamoto K, Foretz M, Berdeaux R, Kronenberg HM (2018) PTHrP targets salt-induced kinases to regulate chondrocyte differentiation. J Bone Miner Res 33(Suppl1):S74 66. Greenbaum A, Hsu YM, Day RB, Schuettpelz LG, Christopher MJ, Borgerding JN et al (2013) CXCL12 in early mesenchymal progenitors is required for haematopoietic stemcell maintenance. Nature 495(7440):227–230 67. Ono N, Ono W, Nagasawa T, Kronenberg HM (2014) A subset of chondrogenic cells provides early mesenchymal progenitors in growing bones. Nat Cell Biol 16(12):1157–1167 68. Anthony BA, Link DC (2014) Regulation of hematopoietic stem cells by bone marrow stromal cells. Trends Immunol 35(1):32–37 69. Seike M, Omatsu Y, Watanabe H, Kondoh G, Nagasawa T (2018) Stem cell niche-specific Ebf3 maintains the bone marrow cavity. Genes Dev 32(5–6):359–372 70. Zhou BO, Yue R, Murphy MM, Peyer JG, Morrison SJ (2014) Leptin-receptor-expressing mesenchymal stromal cells represent the main source of bone formed by adult bone marrow. Cell Stem Cell 15(2):154–168 71. Chan CK, Seo EY, Chen JY, Lo D, McArdle A, Sinha R et al (2015) Identification and specification of the mouse skeletal stem cell. Cell 160 (1–2):285–298 72. Grcevic D, Pejda S, Matthews BG, Repic D, Wang L, Li H et al (2012) In vivo fate mapping identifies mesenchymal progenitor cells. Stem Cells 30(2):187–196 73. Worthley DL, Churchill M, Compton JT, Tailor Y, Rao M, Si Y et al (2015) Gremlin 1 identifies a skeletal stem cell with bone, cartilage, and reticular stromal potential. Cell 160 (1–2):269–284 74. Shi Y, He G, Lee WC, McKenzie JA, Silva MJ, Long F (2017) Gli1 identifies osteogenic

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progenitors for bone formation and fracture repair. Nat Commun 8(1):2043 75. Debnath S, Yallowitz AR, McCormick J, Lalani S, Zhang T, Xu R et al (2018) Discovery of a periosteal stem cell mediating intramembranous bone formation. Nature 562 (7725):133–139 76. Asagiri M, Takayanagi H (2007) The molecular understanding of osteoclast differentiation. Bone 40(2):251–264

77. Kobayashi T, Kronenberg HM (2014) Overview of skeletal development. Methods Mol Biol 1130:3–12 78. Xiong J, Onal M, Jilka RL, Weinstein RS, Manolagas SC, O’Brien CA (2011) Matrixembedded cells control osteoclast formation. Nat Med 17(10):1235–1241 79. Ngo-Muller V, Muneoka K (2010) In utero and exo utero surgery on rodent embryos. Methods Enzymol 476:205–226

Chapter 2 Overview of Skeletal Repair (Fracture Healing and Its Assessment) Elise F. Morgan, Anthony De Giacomo, and Louis C. Gerstenfeld Abstract The study of postnatal skeletal repair is of immense clinical interest. Optimal repair of skeletal tissue is necessary in all varieties of elective and reparative orthopedic surgical treatments. However, the repair of fractures is unique in this context in that fractures are one of the most common traumas that humans experience and are the end-point manifestation of osteoporosis, the most common chronic disease of aging. In the first part of this introduction the basic biology of fracture healing is presented. The second part discusses the primary methodological approaches that are used to examine repair of skeletal hard tissue and specific considerations for choosing among and implementing these approaches. Key words Fracture healing, Radiography, Micro-computerized tomography, Histomorphometry

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Introduction: Overview of Fracture Healing

1.1 Bone Repair Recapitulates Embryological Skeletal Development

Fracture healing and bone repair are unique in that they are postnatal processes that mirror many of the ontological events that take place during embryological development of the skeleton (reviewed by [1–5]). Indeed, many of the genes that are preferentially expressed in embryonic stem cells and the morphogenetic pathways that are active during embryonic skeletal development are also expressed in fracture callus and skeletal repair tissues [6, 7]. It is generally believed that the recapitulation of these ontological processes during fracture healing facilitates the regeneration of damaged skeletal tissues to their preinjury structure and biomechanical function. In this regard, the interplay among regenerative processes in a number of different tissues—vascular, hematopoietic, and skeletal—is essential for the unimpeded repair of injured bones. Furthermore, the appropriate temporal differentiation of the various stem cell populations that form the different tissues and make up skeletal organs is dependent on the proper temporal spatial orchestration of specific paracrine, autocrine, and systemic signaling pathways [7, 8].

Matthew J. Hilton (ed.), Skeletal Development and Repair: Methods and Protocols, Methods in Molecular Biology, vol. 2230, https://doi.org/10.1007/978-1-0716-1028-2_2, © Springer Science+Business Media, LLC, part of Springer Nature 2021

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1.2 Fracture Healing Cascade

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The cascade of events that is commonly described for fracture healing involves formation of a blood clot at the site of injury; an inflammatory phase in which specific cell types involved primarily in innate immune response participate; callus generation in which skeletal stem cells are recruited and differentiate into chondrocytes; primary bone formation in which stem cells are recruited to form bone; and secondary bone remodeling involving osteoclasts in which first mineralized cartilage and primary bone are resorbed followed by prolonged coupled remodeling. While these processes take place in a consecutive temporal manner, they overlap significantly and represent a continuum of changing cell populations and signaling processes within the regenerating tissue. The disruption to the normal bone microenvironments that is caused by the fracture leads to the interactions of cell populations from the medullary space, periosteum, and enveloping muscular tissues. The signaling and cellular contributions from these different tissues and their microenvironments are unique and contribute to the heterogeneous nature of tissue formation at the fracture site [9]. Fracture healing and skeletal tissue repair broadly encompasses an initial anabolic phase that is characterized by de novo recruitment and differentiation of skeletal stem cells that form a cartilaginous callus and, subsequently, of those that form the nascent blood vessels that will feed the new bone. This anabolic phase is followed by a very prolonged catabolic period encompassing resorption of the cartilaginous callus with its replacement by primary bone. Finally, the phase of coupled remodeling takes place, during which the marrow space and hematopoietic tissues are reestablished and regeneration of the original structural features of the injured skeletal organ is achieved. A temporal overview of the biological and histological events of fracture healing, the known cell types that are prevalent at each stage of fracture healing and the stages at which specific signaling molecules are produced are presented in Fig. 1.

Assessing Skeletal Repair

2.1 Using an Integrated Approach to Assessing Tissue Repair

All skeletal healing can be defined both functionally (i.e., by the injured skeletal tissue’s regain of its original structure and biomechanical properties) and in terms of the biological processes that facilitate the regain in function. Because of this complexity, an integrated approach should be taken to asses skeletal tissue repair at the level of the whole organ (via biomechanical and microcomputed tomography (μCT) assessments), at tissue and cellular levels (via histological assessments and some methods of contrastenhanced μCT), and molecular levels (via immunohistological, in situ hybridization, and other assessments of mRNA and protein expression) to define most clearly the mechanisms that promote

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Fig. 1 Summary of the multiple stages of fracture healing. Summary of the stages of fracture repair and the timing of the development of these stages as seen in C57B6 strain of mouse are denoted at the top of the figure. Histological sections are presented below for each stage. All histological specimens are from sagittal sections of mouse tibia transverse fractures and were stained with H&E or Safranin-O and Fast Green, micrographic images are at
200 magnification. Section for the initial injury was taken from the fracture site 24 h postinjury (far left). Sections depicting the initial periosteal response and endochondral formation are from 7 days postinjury (left middle). Arrows denote blood vessels (BV) of the vascular in-growth from the peripheral areas of the periosteum. Sections depicting the period of primary bone and cartilage tissue resorption are from 14 days postinjury (middle right). Sections depicting the period of secondary bone formation are from 21 days postinjury (far right). Insert depicts
400 images of an osteoclast (chondroclast) resorbing an area of calcified cartilage. The major cell types associated with each stage and the relative time frames of the anabolic and catabolic stages of fracture healing are depicted by yellow and orange triangular overlays. The associated molecular processes and regulators next presented in the second boxed area. At the bottom of the figure, the levels of expression of various marker mRNAs for various molecular processes that have been examined in our laboratories and are denoted by three line widths. The levels of expression are by percent over baseline for each and are not comparable between individual mRNAs. Data for expression levels for the proinflammatory cytokines and the ECM mRNAs were from Kon et al. (2001) [39] and Gerstenfeld et al. (2003) [4]

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healing. A description of the use of these techniques presented below provides examples of how biological and functional characteristics are relatable to each other during the normal processes of fracture healing. Many of the methods described here in the context of fracture healing are also detailed in the various chapters of this book. Within this introduction a specific review is provided to illustrate how whole-organ and molecular assessments can be used in conjunction with the many other tissue- and cell-based assays that are outlined in greater technical detail within the other chapters. When considering use of multiple assessments, it is important to recognize that performing several assessments on the same sample can be advantageous from the standpoint of cost savings and statistical power. For example, following μCT imaging with mechanical testing allows direct examination of relationships between mechanical behavior and both structure and mineralization. With this particular sequential approach, one can also use the measured mechanical properties to provide an important calibration or validation of finite element models created from the μCT data [10]. Alternatively if the tissue is first fixed, such as with formalin, μCT imaging can be followed by histological assessments. This sequence allows one to relate cellular composition to structure and mineral content. Use of multiple approaches may also be used as a means of validating biological and structural findings. As an example, qRT-PCR analyses of mRNA expression of markers of chondrogenesis or osteogenesis, or osteoclastogenesis may be used to confirm histological and histomorphometric data of cartilage, bone compositions as well as to define the remodeling and developmental progression of these tissues. By assessing bone healing at multiple levels (organ, tissue, cellular, and molecular) relationships and interactions between the various mechanisms that work at each levels during fracture healing can then be developed. A summary of the most common measurements that are made in the assessment of skeletal repair tissues is presented in Table 1. 2.2 Selection of a Model to Assess Skeletal Repair

The model that one chooses to use in assessing skeletal tissue repair should be carefully considered in relationship to the research questions that are being asked. In this context four considerations come into play: 1. If one is using a surgical model as a means of extrapolating developmental and regenerative characteristics about skeletal tissues, the nature of the type of skeletal formation process that one wishes to examine (intramembranous bone formation versus endochondral bone formation) is important to consider. 2. The nature of both the bone (cortical versus intermedullary) and the surrounding soft tissue compartments that are affected and contribute to the repair process should be considered.

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Table 1 Metrics of bone repair A. Radiographic data

Units

Faxitron radiography Areal BMD Projected bone area

Gray level μm2

Computed tomography Total callus volume Mineralized callus volume Mineralized callus volume fraction Callus mineral content Average tissue mineral density Standard deviation of tissue mineral density

mm3 mm3 – mg HA/cm3a mg HA/cm3a mg HA/cm3a

B. Biomechanical data (Torsion) Ultimate torque Torsional stiffness Torsional rigidity Twist to failure Toughness (work to failure)

Units N-mm N-mm/deg N-mm2/deg deg N-deg

C. Histomorphometric datab Callus diameter (CDm)c Mean value for measurements made, in two orthogonal planes, of the diameter of the midpoint of the fracture callus.

Units Mm

Total Callus Area (CAr) Mean value for measurements of the total callus areas inclusive of all tissues both within and outside the original bone cortices

mm2

Area of Cartilage (Cg/Ar) Mean value for measurements of the total cartilage in the callus. May alternatively be expressed as the percent of total callus volume that is cartilage. %Cg

mm2

Area of Total Osseous Tissues TOT/Ar Mean value for measurements of total callus area that is osseous tissue. (Includes preexiting cortical bone, new woven bone and surfaces lined by osteoblasts). May alternatively be expressed as the percent of total callus volume that is osseous tissues. %TOT Area of Void (V/Ar) Mean value for measurements of total callus area that includes the marrow cavity, hematopoietic elements and empty unstained space. May alternatively be expressed as the percent of total callus volume that is the void. %V Area of Fibrous Tissue FT/Ar Mean value for measurements of traced areas of fibrous tissue within the callus. May alternatively be expressed as the percent of total callus volume that is fibrous tissue (%FT) Osteoclast Volume Density (Oc/Ar) Mean value for measurements of tartrate resistant acid phosphatase stained cells calculated as the number of osteoclasts per unit area of callus. Osteoblast Bone Surface Density (Ob/Ar) Number of osteoblasts lining a bone or mineralized cartilage surface as calculated per unit surface area of new trabecular bone

mm2

mm2

mm2

(#/mm2) #/mm2 (continued)

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Table 1 (continued) Number of Vessels per Callus Area

(#/mm2)

HA ¼ hydroxyapatite: While area, diameter, and cartilage values are not listed in this table and are not commonly used measurements in metabolic bone studies, they are defined and their nomenclature is as in the original agreed standard b Relational determinations to biomechanical testing: CDm and CAr measurements provide comparative values to X-ray, qCT, and biomechanical determinations. Diameters can be used in calculations of moments of inertia. C:TOT measurements can be used in material assessments in relationship to stiffness and strength determinations. Vd Since the external callus tissue initially is devoid of a hematopoietic containing marrow space the assessment and the progression of Vd area gives total measurements of rates of resorption and subsequent remodeling of the external callus and may be related to OcD c Values in parenthesis () denotes nomenclature as is the agreed standard2 a

3. The skeletal organ and its developmental background should be considered. At its simplest level this would divide surgical models into those that assess bone repair in appendicular, axial, and cranial tissues or in different soft tissue elements. 4. If one is assessing a therapeutic modality the surgical model that is chosen should most closely approximate the orthopedic application and therapeutic modality that is being assessed. It is also important to note that, due to the effects of systemic interactions and the heterogeneity in cellular composition, in vivo models cannot be used to fully dissect the molecular mechanisms of the various biological processes that effect repair. It is therefore ideal that in vivo studies should be complimented with in vitro methods of cell or organ culture that are presented in this book. A fracture or any surgical repair model may be tracked temporally and isolated spatially. In the case of a fractured long bone, the injury induces one round of endochondral bone formation in which callus cells differentiate in a synchronous manner that temporally phenocopies the spatial/temporal variation of the cell zones from the top to the bottom of the growth plate. This round is followed by a prolonged period of coupled remodeling. As such fractures, represent an ideal biological process to examine in a postnatal context many cellular and molecular mechanisms that underlie both the endochondral bone formation that takes place during skeletal tissue development and the coupled remodeling that takes place during skeletal tissue homeostasis.

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Whole Organ Assessments The two central, functional attributes of skeletal tissues are their ability to regulate apatite mineral deposition and resorption and to assemble and model the microstructure of the mineralized tissue to meet the biomechanical needs of the animal. These unique functional attributes make radiographic approaches particularly useful in examining skeletal tissue repair since these approaches focus on the mineralized tissues within the callus.

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3.1

Plain-Film X-Ray

This assessment is the most common clinical tool to assess hard tissue repair, although this assessment is limited by its relatively qualitative nature [11–13]. Figure 2 presents a series of plain-film X-ray assessments made across a time course of fracture healing. This type of study provides a first approximation of the progression of tissue repair (Fig. 2a). For these studies, an X-ray device with a high-energy beam and capability of high resolution, such as a Faxitron® cabinet X-ray system, is ideal. The best resolution is obtained if the bone is removed from the animal and cleaned of a large amount of surrounding soft tissue. Fixation devices should be left in place to maintain the integrity of the construct up until the tissue construct is stable. At least two separate anatomical orientations should be used to make such measurements of callus dimensions and bone bridging since the tissues that form during repair can be irregular in shape and cortices of most bones are not true cylindrical structures. This is illustrated in Fig. 2b which is used for cortical bridging and callus formation scoring. A scoring metric has been developed and applied clinically to assess the progression of healing first in tibia (Radiological Union Score of Tibia [RUST]) [14] and now subsequently in other appendicular bones [15]. This scoring system is based on assessment of X-ray images from two separate anatomical orientations and counting the number of cortices showing both the development of callus bridging across the fracture gap [14, 16]. This approach is summarized in (Fig. 2b). Use of this tool provides a quantitative comparative measure of the progression of bone healing between human and animal studies [16, 17]. This tool has high intraclass correlation coefficients (ICCs) between raters of the radiographs and the scores were positively correlated with callus bone mineral density and regain of strength [17, 18].

3.2

μCT

Given that the size, shape, and composition of the bone repair tissues change over the time course of healing, μCT can provide important, qualitative and quantitative assessment of these changes so as to provide nondestructive, and even noninvasive, evaluation of repair progress. We and others have developed “standard” methods of μCT evaluation of mineralized tissues in fracture healing as well as contrast-enhanced μCT methods for examining contributions of non-mineralized tissues and vascular elements.

3.2.1 μCT Assessment of Mineralized Tissues in the Callus

Many of the technical considerations for μCT studies of intact bones [19] also apply to studies of fracture healing. These considerations include scanning parameters (voltage, current, integration time, and resolution) and methods of image processing (noise filtering, thresholding and defining volumes of interest for analysis). Typically, image resolutions of 16 μm/voxel and 12 μm/voxel are sufficient for rat and mouse calluses, respectively. A Gaussian

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Fig. 2 Radiographic Series for assessment of fracture healing. (a) Time series of evolution of mouse femur fracture callus structure and mineral content.

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filter is also commonly used. These resolutions and filter are standard for commercially available desktop μCT systems. The boundaries of the callus must be defined in μCT assessments, if one is interested in quantifying callus size and the fraction or percentage of the callus occupied by mineralized tissue, just as described below for histomorphometric assessments of callus tissues. Identification, or segmentation, of the callus can be achieved by defining the outer and inner boundaries of the callus on transverse cross-sectional images distributed along the length of the callus (Fig. 3). Although we often use an inner boundary that excludes the cortex and medullary space, this boundary can be omitted for complete analysis of the extent of bridging (since some bridging of the cortex might be present) and if healing is so advanced that the boundary between the cortex and mineralized callus tissue cannot be reliably identified. Upon defining the callus or some portion of the callus as the volume of interest, one needs to choose one or more thresholds for specific quantification and visualization of the mineralized portions of the callus. A threshold is a gray value above which a voxel in the μCT scan will be considered to contain mineralized tissue. If a simple categorization of mineralized vs. unmineralized tissue is desired, then one threshold is sufficient. If three or more categories of the relative extent of mineralization are desired, or if a scaffold or implant material is present, then more than one threshold may be used. At present, there is no standard method for choosing a threshold. We recommend that regardless of how many thresholds are used, a threshold should be defined as a percentage of the average gray value of the preexisting cortex or implant material [20]. Although commercial μCT systems have software algorithms for quantifying trabecular structure, this type of analysis is not appropriate for repair tissues, because of the wide range of mineralization that is present in these tissues renders the measures of “trabecular” thickness, “trabecular” number, connectivity, etc. extremely sensitive to the choice of threshold. A number of outcome measures can be quantified using μCT (Table 1). Some of these measures describe callus size and quantity of mineralized tissue (total volume, mineralized volume, mineralized volume fraction, and mineral content), while others describe the mineralization (average and standard deviation of the tissue mineral density) or overall structure (moment of inertia). ä Fig. 2 (continued) (b) Anterior and posterior views of mouse femur fracture callus at 10 days postfracture. The images include numeric scoring for each cortices based on the modified Radiological Union or Score of Tibia system. Numbers in white denote each score while numbers in black denote the overall bridging or healing score. All images were produced with a Faxitron® device set at 40 s and 30 kV

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Fig. 3 (a) Definition of the outer (green) and inner (red) callus boundaries on a transverse cross section of micro-computed tomography scan of a fracture callus at 14 days postfracture. (b) The 3D volume of interest that is defined by the area enclosed by these two boundaries on each transverse cross section along the length of the callus is shown rendered in entirety on the left and in a longitudinal cut-away view on the right. Reprinted from Morgan et al. (2009) [20] 3.2.2 Contrast-Enhanced μCT Imaging of Cartilaginous Tissues in the Callus

Formation of the cartilage tissues is a key phase of many skeletal tissue studies. During skeletal repair, cartilage tissues provide initial stability at the surgery or fracture site and serves as a template for subsequent formation of mineralized tissue. In order to provide nondestructive assessment of the soft callus with μCT, a contrast agent is required to increase the X-ray attenuation of these tissues. We have used a cationic, iodinated contrast agent [21] for this purpose. On account of the large, fixed negative charge in cartilage, the contrast agent, via electrostatic attraction, preferentially accumulates in regions of cartilage within the callus. These regions incur the largest increase in attenuation from pre- to postincubation images. The attenuation of the noncartilaginous soft tissues is moderately increased, allowing clear delineation of the callus boundaries, while the attenuation of bone tissue is unchanged. The basic experimental approach in this contrast-enhanced μCT (CECT) method is to perform μCT scans both before and after incubation of the callus in the contrast agent (Fig. 4a). Analysis of the preincubation images, postincubation images, and images formed by subtracting the former from the latter enables discrimination among cartilage, noncartilaginous soft tissue, and mineralized tissue in the callus. The respective locations of these different tissues within the callus can be nondestructively visualized and quantified in both 2-D and 3-D (Fig. 4). Measurements of callus area and cartilage area made with CECT compare well to those made using histomorphometry (Fig. 4c–e).

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Fig. 4 (a) CECT method: Labeling of tissues in the far right image is as follows: callus boundary ¼ green outline, cartilage ¼ red/pink; cortex ¼ purple; other mineralized tissue ¼ gray. (b) Comparison of (left) a histological section (bright orange-red ¼ cartilage) and (right) CECT cross section (blue ¼ cartilage; red ¼ mineralized cartilage; gray ¼ bone) of a murine fracture callus (postoperative day 10). (c) 3-D rendering of a callus imaged with CECT (red ¼ cortex; blue ¼ cartilage; yellow ¼ noncartilaginous soft callus). Comparison of histomorphometric and CECT measurements of (d) cartilage area and (e) total callus area. Each symbol represents a different callus (n ¼ 4 measurements per callus, each corresponding to one quadrant of the cross section) 3.2.3 Contrast-Enhanced μCT Imaging of Vessel Structure in the Callus

A different contrast agent that is perfused at the time of euthanasia allows for μCT assessment of the vasculature during bone repair. In this method, a contrast agent such as a mixture of lead chromate and silicone rubber (Microfil MV-122; Carver, MA), is injected through the left ventricle of the heart and allowed to perfuse with drainage into the body cavity by cutting the vena cave. After perfusion the contrast agent is allowed to polymerize [22, 23]. The choice of contrast agent and imaging procedure depends on the

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Fig. 5 (a) 3D renderings of the mineralized tissue (yellow) without (top row) and with (bottom row) the vasculature (red) in calluses from a murine model of distraction osteogenesis (DO). “FA” denotes femoral artery. (b) Vessel volume in the gap periphery vs. gap center for the DO time course and unoperated controls (*p < 0.05) Reprinted from Rundle (2006) [46]

type of CT scanner available and the type of vasculature to be quantified [24]. For imaging of only intermediate- to large-sized vessels or when using synchrotron μCT, a very highly attenuating contrast material, such as bismuth [25], could be used and discrimination between the contrast-enhanced vascular casts and the surrounding mineralized tissue can be achieved purely with a threshold [26–28]. However, for analyses of small vascular elements using a desktop μCT system, only a μCT scan performed after decalcification of the host bone is likely to allow clear discrimination between vessel and mineralized tissue. The disadvantages of performing the μCT scan after decalcification are twofold. First, decalcification results in large changes in shape and size of the callus, and the original anatomic positions of the vessels are thus lost. Second, the spatial relationship between vascular elements and mineralized tissue cannot be determined. An extension of the aforementioned method is to perform μCT scans both before and after decalcification. The postdecalcification images are registered to and then subtracted from the predecalcification images to yield data on the vessels (in their original anatomic position), mineralized tissue, and the respective locations of these two tissues (Fig. 5). 3.3 Mechanical Approaches

In the laboratory setting, the mechanical properties of a healing bone are also commonly assessed by mechanical tests that load the bone in torsion or in three-point bending. The choice of the type of test is dictated by technical as well as physiological considerations. Tension and compression tests are not commonly used, because variability in the alignment of the fracture and in the asymmetry of the callus will lead the applied tensile or compressive displacements to induce variable amounts of bending and shear within the callus. Bending and torsion are logical choices when studying fracture

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Fig. 6 Representative torque–twist curve for a mouse tibia 21 days postfracture. The curve is annotated to show definitions of basic biomechanical parameters. Torsional rigidity is computed by multiplying the torsional stiffness by the gage length. Analogous definitions hold for bending tests

healing in long bones, because these bones experience bending and torsional moments in vivo. However, whereas torsion tests subject every cross section of the callus to the same torque, three-point bending creates a nonuniform bending moment throughout the callus. As a result, failure of the callus during a three-point bend test does not necessarily occur at the weakest cross section of the callus. Regardless of the type of mechanical test, the outcome measures that can be obtained are the strength, stiffness, rigidity, and toughness of the healing bone (Fig. 6). For torsion tests, an additional parameter, twist to failure, can be used as a measure of the ductility of the callus. Although strength, a measure of the force or moment that causes failure, can only be measured once for a given callus, it is possible to obtain more than one measure of stiffness and rigidity. Multistage testing protocols have been reported that apply nondestructive loads to the callus in planes or in loading modes that are different from those used for the stage of the test in which the callus is loaded to failure. With these protocols, it is possible to quantify the bending stiffness in multiple planes [29] or the torsional as well as compressive stiffness [30]. The mechanical properties illustrated in Fig. 7 are structural, rather than material, properties. Material properties describe the

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Fig. 7 Prediction of torsional strength by BV, TMD, and σTMD. The beta weight (β) and partial correlation coefficient (rpartial) are given for each μCT measure to indicate its relative contribution. Data from Kakar et al. (2007) [42]

intrinsic mechanical behavior of a particular type of material (tissue), such as woven bone, fibrocartilage, or granulation tissue. The structural properties of a fracture callus depend on the material properties of the individual callus tissues as well as the spatial arrangement of the tissues and the overall geometry of the callus. While it is possible to use measurements of callus geometry together with those of structural properties to gain some insight into callus tissue material properties [31] true measurement of these material properties requires direct testing of individual callus tissues [32, 33]. Although there is no substitute for mechanical testing to assess the extent of healing at intermediate-to-late time points, it is of biological and translational interest to identify relationships between the mechanical properties of the callus and nondestructively obtained measurements of callus composition and structure. In a large, composite dataset of murine calluses at multiple time points postfracture [20], torsional strength was best predicted (as determined by stepwise regression) by the combination of average tissue mineral density, mineralized callus volume, and the standard deviation of mineral density (r2 ¼ 0.62, p < 0.001) (Fig. 7, Table 1). Torsional rigidity was best predicted by the combination of average tissue mineral density, callus mineral content, mineralized volume fraction, and the standard deviation of mineral density (r2 ¼ 0.70, p < 0.001). Changes in the calluses over time were characterized primarily by an increase in average tissue

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mineral density, while variability among calluses at a given time point was seen primarily in the measures that quantify the absolute and relative amounts of mineralized tissue in the callus, that is, mineralized callus volume, callus mineral content, and mineralized volume fraction. Overall, these results illustrate how the mechanical properties of the callus depend on measures of both the quantity and mineral density of the hard tissue.

4

Tissue Level Histological Approaches Of the recommendations and conventions defined by Parfitt et al. [34] for general histological assessments of skeletal tissue, we have put forth our perspective on the aspects of these assessments that would be most appropriate for bone repair [35]. Extensive details of histological methods including in situ hybridization and immunohistochemistry are discussed in other chapters in this book and will not be discussed in detail here. Rather, the present focus is on general recommendations as to how to approach the use of histological and histomorphometric approaches in the analysis of fracture callus tissues. It is important to note that repair tissues are very heterogeneous, being composed of cartilage, bone, and fibrous and hematopoietic tissues, which are changing throughout the repair process. Therefore, the greatest challenge confronting any histological assessment of a bone reparative process is how to sample this heterogeneous tissue so that any quantitative measurements are representative of the repair tissue that is formed. In this regard two sets of issues should be considered in any histological assessment of skeletal repair tissues. The first is related to the anatomical plane and sampling of the repair tissue. While fracture calluses may be examined in a longitudinal plane such approaches provide primarily a qualitative overview of the tissue heterogeneity. This, in part, is due to difficultly of reproducibly positioning a bone during embedding such that a uniform longitudinal plane is always sectioned between individual specimens. This problem largely is related to the fact that long bones are not perfectly cylindrical as illustrated by comparisons of the anterior and posterior X-ray images in Fig. 2b and the MicroCT reconstructions presented in Fig. 3b. Sampling of tissue compositions is further complicated by the fact that endochondral bone formation, and cartilage resorption and bone remodeling that occurs during tissue repair arises also in a nonuniform manner. This is underscored by Fig. 8, which presents the immense structural heterogeneity in representative longitudinal and transverse sections of fracture calluses at 14 days postinjury. To address this challenge, transverse sections can be collected at fixed increments along the long axis of the callus. This sectioning approach provides an optimal means for both observing the variability of the tissue formed in

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Fig. 8 Histological approach to assess fracture. (a) Demonstration of the tissue heterogeneity and morphological irregularity of both the in longitudinal and transverse dimensions. Three transverse slices taken from center at three increments from the center to edge of the callus showing how the cartilage content varies. (b) Demonstration of pseudo coloring of Safranin-O/Fast Green–stained sections segmenting cartilage from voids and bone areas

all three planes of the callus and obtaining accurate cross-sectional diameters and area measurements at precisely defined anatomical positions within the callus and in relationship to the fracture site. We have used serial sections to reconstruct in three dimensions the tissue compositions of whole calluses [36] and to obtain

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measurements of the mechanical properties of callus tissues in conjunction with histological assessment [32]. The second practical aspect to be considered in the application of histomorphometric techniques to bone healing studies is to identify appropriate histological stains to assess the callus. It is important to recognize that a tissue stain should not be used to determine tissue phenotype, but rather to improve and enhance visualization of that tissue by distinguishing it, by color, from a different histologically contiguous tissue. Multiple stains can be used to discriminate cartilage from bone and have been used in studies assessing fracture healing. In our studies we have used Safranin-O/Fast Green staining, which has been widely used and shown to be effective in measurements of cartilage thickness in studies of osteoarthritis [37] and to quantify the amount of cartilage tissue repair of joint surfaces [38]. Other studies also have used combinations of Alcian Blue with hematoxylin and eosin in order to obtain differential staining of cartilage and bone [39, 40]. In general, a staining protocol should provide consistent staining outcome that will enhance histological tissue discrimination of cartilage from bone and non-osseous tissue and can be used with semiautomated measurement techniques.

5

Molecular Approaches The assessment of the expression of mRNAs in conjunction with immunohistochemical approaches provides a means of identifying the underlying morphogenetic signaling and regulatory processes that regulate bone repair. In this final section three separate approaches to assay mRNAs (in situ hybridization, individual candidate mRNA assays and large-scale microarray) will be discussed.

5.1 In Situ Hybridization

At the single mRNA level in situ hybridization is used to identify the nature of specific cell populations and their anatomical localization within the callus surrounding muscle and marrow spaces that are expressing a given mRNA. Such approaches can be used both to identity and to validate simple histological assessments of the cell type (chondrocytes, osteoblasts, etc.) and the differentiated states of these cells [36]. This approach can also be used in conjunction with immunohistological methods to place in anatomical context the cells expressing a given regulatory factor or morphogenetic protein relative to those expressing receptors that make them responsive to these signals.

5.2 Individual mRNA Assessments

Isolation of total RNAs from the entire callus allows one to look at expression of individual mRNAs within the total cell population of the callus tissue. This approach can be used to provide an extremely sensitive temporal road map of the differentiation and development

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of the tissues and cell types of the callus as well as the expression or any regulatory factors that one is interested, which may be functional in fracture healing. It is critical that calluses be isolated quickly and in as reproducible manner devoid of surrounding muscle as possible with manual dissection methods. Tissues should be dissected as exactly as possible from the point the callus initially rises from cortical surface and since it is physically impossible to separate the original bone and marrow elements from the external callus we isolate the mRNAs from the intact tissue specimen. At very early time points before the callus has condensed (day 5 for mouse and day 7 for rat) a margin of muscle may be needed to ensure that the early regenerative cells are isolated, but some caution should be taken to interpret the anatomical localization of the cells that are expressing a given mRNA species. Confirmatory analysis using in situ hybridization or immunohistochemistry is recommended. It is important to note that there are some differences of opinion on how to assess replicates. For our studies in mouse, we generally use three biological replicates, each representing a pool of total RNAs from calluses of three animals. Reference mRNAs are made from unfractured bones (three replicates pooled from groups of three animals) isolated from the mid-diaphyseal region of these bones. This approach incorporates aspects of obtaining reproducibility from assaying replicates and takes into account biological variability since the replicates represent experimental repeats of multiple pooled animal samples. We have taken this approach since mRNA yields from single mouse calluses are very small and insufficient to carry out assays for large numbers of genes or for use in microarray studies. In general this approach has provided very sensitive means of seeing reproducible differences in temporal profiles of expressed genes under differing experimental conditions [39, 41, 42]. For studies performed in rats, individual calluses provide sufficient yields of total mRNA for running multiple mRNA assays [38]. 5.3 MicroArray Approaches

Sequencing of the entire genomes of multiple species has provided the means by which the transcriptome of fracture healing can be assessed by microarray analysis. Given the expense of such studies these approaches should only be carried out in conjunction with a core facility that has technical staff, instrumentation, and a track record of executing this type of study. The setup of a microarray study involves a number of crucial steps, the first and most important is to have sufficient replicates per experimental group (treatment condition or animal genotype, and time point). In planning a study, sufficient RNA is needed to carry out the array twice in the event of a technical problem with the execution of the array and to carry out mRNA candidate validation by qRT-PCR. A minimum of three biological replicates is recommended as described above. Analysis of microarray data should follow four main steps:

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(1) quality control to identify microarray artifacts from hybridization and correct for chip batch effects; (2) data preprocessing to eliminate outliers; (3) identification of differentially expressed genes (DEG); and (4) extraction of biological knowledge from DEG. It is recommended that, given the expense of microarray experiments, appropriate expertise in statistical and computational analysis also be available either through collaborative or fee-forservice arrangements. The most basic approach in step 3 uses basic statistical analysis to identify genes that showed the greatest quantitative changes in their expression [43, 44–46]. In this context this type of approach can be used to identify both known and novel genes that show the greatest changes in expression over the time course of healing. Other more sophisticated statistical approaches have been carried out that cluster genes based on common temporal profiles of expression and that examine gene functions in each cluster across the time course of fracture healing [6, 47, 48].

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Conclusion In this brief introductory chapter, we have reviewed the biological events of fracture repair. We have also laid out a number of the general methodological approaches including a summary of various radiographic and molecular techniques that are used to assess bone repair, using fracture healing as the example. General guidance is presented on integrating multiple technical approaches to best assess skeletal repair that may be also be applicable for other areas of skeletal biology research.

Acknowledgments This work is supported by NIH grants AR056637 and AR062642. References 1. Bolander ME (1992) Regulation of fracture repair by growth factors. Proc Soc Exp Biol Med 200(2):165–170 2. Einhorn TA (1998) The cell and molecular biology of fracture healing. Clin Orthop Relat Res 355(Suppl):S7–S21 3. Ferguson C et al (1999) Does adult fracture repair recapitulate embryonic skeletal formation? Mech Dev 87:57–66 4. Gerstenfeld LC et al (2003) Fracture healing as a post-natal developmental process: molecular, spatial, and temporal aspects of its regulation. J Cell Biochem 88(5):873–884

5. Vortkamp A et al (1998) Recapitulation of signals regulating embryonic bone formation during postnatal growth and in fracture repair. Mech Dev 71:65–76 6. Bais M et al (2009) Transcriptional analysis of fracture healing and the induction of embryonic stem cell-related genes. PLoS One 4(5): e5393 7. Phillips AM (2005) Overview of the fracture healing cascade. Injury 36S:55–57 8. Buckwalter JA, Einhorn TA, Marsh JL (2001) Bone and joint healing. In: Bucholz RW, Heckman JD (eds) Rockwood and Green’s fractures

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in adults. Lippincott, Williams, and Wilkins, Philadelphia, pp 245–271 9. Gerstenfeld LC et al (2003) Impaired fracture healing in the absence of TNF-alpha signaling: the role of TNF-alpha in endochondral cartilage resorption. J Bone Miner Res 18 (9):1584–1592 10. Gardner TN et al (2000) The influence of mechanical stimulus on the pattern of tissue differentiation in a long bone fracture—an FEM study. J Biomech 33:415–425 11. Axelrad TW, Einhorn TA (2011) Use of clinical assessment tools in the evaluation of fracture healing. Injury 42(3):301–305 12. Bhandari M et al (2002) A lack of consensus in the assessment of fracture healing among orthopaedic surgeons. J Orthop Trauma 16 (8):562–566 13. Goldhahn J et al (2008) Clinical evaluation of medicinal products for acceleration of fracture healing in patients with osteoporosis. Bone 43:343–347 14. Whelan DB et al (2010) Development of the radiographic union score for tibial fractures for the assessment of tibial fracture healing after intramedullary fixation. J Trauma 2 (68):629–632 15. Litrenta J et al (2015) Determination of radiographic healing: an assessment of consistency using RUST and Modified RUST in metadiaphyseal fractures. J Orthop Trauma 29 (11):516–520 16. Tawonsawatruk T, Hamilton DF, Simpson AH (2014) Validation of the use of radiographic fracture-healing scores in a small animal model. J Orthop Res 32(9):1117–1119 17. Cooke ME et al (2017) Correlation between RUST assessments of fracture healing to structural and biomechanical properties. J Orthop Res 36(3):945–953 18. Fiset S et al (2018) Experimental validation of the radiographic union score for tibial fractures (RUST) using micro-computed tomography scanning and biomechanical testing in an in-vivo rat model. J Bone Joint Surg Am 100 (21):1871–1878 19. Bouxsein ML et al (2010) Guidelines for assessment of bone microstructure in rodents using micro-computed tomography. J Bone Miner Res 25(7):1468–1486 20. Morgan EF et al (2009) Micro-computed tomography assessment of fracture healing: relationships among callus structure, composition, and mechanical function. Bone 44:335–344 21. Hayward LN et al (2012) MRT letter: contrast-enhanced computed tomographic

imaging of soft callus formation in fracture healing. Microsc Res Tech 75(1):7–14 22. Duvall CL et al (2004) Quantitative microcomputed tomography analysis of collateral vessel development after ischemic injury. Am J Physiol Heart Circ Physiol 287:H302–H310 23. Duvall CL et al (2007) Impaired angiogenesis, early callus formation, and late stage remodeling in fracture healing of osteopontin-deficient mice. J Bone Miner Res 22:286–297 24. Morgan EF et al (2012) Vascular development during distraction osteogenesis proceeds by sequential intramuscular arteriogenesis followed by intraosteal angiogenesis. Bone 51:535–545 25. Li W et al (2006) High-resolution quantitative computed tomography demostrating selective enhancement of medium-size collaterals by placental growth factor-1 in the mouse ischemic hindlimb. Circulation 113:2445–2453 26. Fei J et al (2010) Imaging and quantitative assessment of long bone and vasculature. Microsc Res Tech 293:215–224 27. Schneider PK et al (2009) Simultaneous 3D visualization and quantification of murine bone and bone vasculature using microcomputed tomography and vascular replica. Microsc Res Tech 72:690–701 28. Sider KL, Song J, Davies JE (2010) A new bone vascular perfusion compound for the simultaneous analysis of bone and vasculature. Microsc Res Tech 73:665–672 29. Foux A, Black RC, Uhthoff HK (1990) Quantitative measures for fracture healing: an in-vitro biomechanical study. J Biomech Eng 112:401–406 30. Tsiridis E et al (2007) Effects of OP-1 and PTH in a new experimental model for the study of metaphyseal bone healing. J Orthop Res 25:1193–1203 31. Ulrich-Vinther M, Andreassen TT (2005) Osteoprotegerin treatment impairs remodeling and apparent material properties of callus tissue without influencing structural fracture strength. Calcif Tissue Int 76:280–286 32. Leong PL, Morgan EF (2008) Measurement of fracture callus material properties via nanoindentation. Acta Biomater 4 (5):1569–1575 33. Manjubala I (2009) Spatial and temporal variations of mechanical properties and mineral content of the external callus during bone healing. Bone 45:185–192 34. Parfitt AM et al (1987) Bone histomorphometry: standardization of nomenclature, symbols and units. J Bone Miner Res 2:595–610

Overview of Skeletal Repair 35. Gerstenfeld LC et al (2005) Perspective: the application of histomorphometric methods to the study of bone repair. J Bone Miner Res 20:1715–1722 36. Gerstenfeld LC et al (2006) Three dimensional reconstruction of fracture callus morphogenesis demonstrates asymmetry in callus development. J Histochem Cytochem 54 (11):1215–1228 37. Hacker SA et al (1997) A methodology for the quantitative assessment of articular cartilage histomorphometry. Osteoarthr Cartil 5:343–355 38. O’Driscoll SW et al (1999) Method for automated cartilage histomorphometry. Tissue Eng 5:13–23 39. Kon T et al (2001) Expression of osteoprotegerin, receptor activator of NF-kappaB ligand (osteoprotegerin ligand) and related proinflammatory cytokines during fracture healing. J Bone Miner Res 16(6):1004–1014 40. Tiyapatanaputi P et al (2004) A novel murine segmental femoral graft model. J Orthop Res 22:1254–1260 41. Jepsen KJ et al (2008) Genetic variation in the patterns of skeletal progenitor cell differentiation and progression during endochondral bone formation affects the rate of fracture healing. J Bone Miner Res 23(8):1204–1216

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42. Kakar S et al (2007) Enhanced chondrogenesis and Wnt-signaling in parathyroid hormone treated fractures. J Bone Miner Res 22 (12):1903–1912 43. Salisbury Palomares KT et al (2010) Transcriptional profiling and biochemical analysis of mechanically induced cartilaginous tissues in a rat model. Arthritis Rheum 62(4):1108–1118 44. Hadjiargyrou M et al (2002) Transcriptional profiling of bone regeneration. Insight into the molecular complexity of wound repair. J Biol Chem 277(33):30177–30182 45. Wang K et al (2006) Analysis of fracture healing by large-scale transcriptional profile identified temporal relationships between metalloproteinase and ADAMTS mRNA expression. Matrix Biol 25(5):271–281 46. Rundle CH (2006) Microarray analysis of gene expression during the inflammation and endochondral bone formation stages of rat femur fracture repair. Bone 38(4):521–529 47. Wise JK et al (2010) Temporal gene expression profiling during rat femoral marrow ablationinduced intramembranous bone regeneration. PLoS One 5(10):pii: e12987 48. Grimes R et al (2011) The transcriptome of fracture healing defines mechanisms of coordination of skeletal and vascular development during endochondral bone formation. J Bone Miner Res 26(11):2597–2609

Chapter 3 Advantages and Limitations of Cre Mouse Lines Used in Skeletal Research Florent Elefteriou and Greig Couasnay Abstract The Cre-LoxP technology permits gene ablation in specific cell lineages, at chosen differentiation stages of this lineage and in an inducible manner. It has allowed tremendous advances in our understanding of skeleton biology and related pathophysiological mechanisms, through the generation of loss/gain of function or cell tracing experiments based on the creation of an expanding toolbox of transgenic mice expressing the Cre recombinase in skeletal stem cells, chondrocytes, osteoblasts, or osteoclasts. In this chapter, we provide an overview of the different Cre-LoxP systems and Cre mouse lines used in the bone field, we discuss their advantages, limitations, and we outline best practices to interpret results obtained from the use of Cre mice. Key words Cre-recombinase, Floxed gene, Specificity, Transgenic mice, Mouse model, Off target, Osteoblasts, Osteocytes, Chondrocytes, Osteoclasts

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Generalities About the Cre-LoxP Technology

1.1 Global Versus Conditional Gene Targeting

Global (whole body) gene inactivation is a powerful approach to investigate gene function in vivo and has led to major findings in the field of bone biology and many others [1, 2]. It involves the use of mouse embryonic stem cells and gene targeting by homologous recombination, leading to gene inactivation or insertion of point mutations in every cell of the mouse model created. This approach is particularly suited for investigating gene function at early developmental stages, when expression of the targeted gene first arises. In that embryonic context, the phenotype likely stems from direct consequences of gene inactivation on cell function, tissue structure or homeostasis. In contrast, care has to be taken when analyzing the consequences of global gene inactivation during later stages of development in adults or during aging. In this context, observed phenotypes may reflect indirect mechanisms caused by early developmental events that are secondary to gene inactivation or compensatory pathways. Another significant limitation of global gene

Matthew J. Hilton (ed.), Skeletal Development and Repair: Methods and Protocols, Methods in Molecular Biology, vol. 2230, https://doi.org/10.1007/978-1-0716-1028-2_3, © Springer Science+Business Media, LLC, part of Springer Nature 2021

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ablation is the potential embryonic lethality of mutant mice, which precludes postnatal analyses. Despite these limitations, global gene deletion or mutation strategies are particularly useful to generate preclinical models of Mendelian genetic diseases affecting the skeleton, for which the natural history of the disease can be characterized in great detail and compared to clinical data. Examples of such models include mice with constitutive activation of Fgfr3, leading to the various forms of dwarfism [3] or mice deficient for Alpl leading to hypophosphatasia and lack of skeletal mineralization [4]. Conditional gene ablation addresses the major limitations of global gene inactivation and over the last 20 years, has been invaluable for our understanding of bone and cartilage biology. The generation of conditional mouse models requires two mouse lines: one with LoxP sites flanking the chosen DNA sequence to ablate, and a second one in which the expression of a constitutive or inducible form of the Cre recombinase is driven by a construct containing a chosen promoter often including enhancer elements [5, 6]. The Cre recombinase is an enzyme derived from the P1 bacteriophage that can recognize the 34 bp DNA sequences of LoxP sites introduced into a gene of interest by homologous recombination. It first creates a DNA loop between two LoxP sites and then excises the DNA fragment in between. When two LoxP sites flank a critical exon in the same direction, action of the Cre-recombinase leads to gene inactivation in those cells where the Cre-recombinase is expressed. In contrast, when two LoxP sites are positioned in an inverted direction, the flanked DNA-fragment will instead be inverted [7]. The yeast Flippase/FRT recognition target system, which recognizes FRT sites rather than LoxP sites, is otherwise mechanistically identical to the Cre/Lox system [8]. The combination of both recombination systems can be used to induce sequential deletion in the same cell lineage, and it is commonly used to delete antibiotic-resistance selection cassettes in ES cells during the generation of conditional knockout (KO) mice [9]. The Cre recombinase can be used to delete an entire gene or part of it depending on the position of the LoxP sites. It can also be used to turn on the expression of a gene, transgene, or reporter gene whose expression is silenced in absence of Cre expression by a floxed stop signal (usually a floxed polyadenylation signal sequence or a codon stop cassette, which is removed upon Cre-mediated recombination) placed upstream of the transgene. This is commonly used for tracing and cell fate lineage studies where reporter genes can be expressed in cells where the Cre-recombinase has been active (see below). Breeding between “Cre” and “floxed” mouse lines allows investigators to inactivate a gene or to activate transgenes in the cell lineage where the Cre recombinase is expressed. However, the system has a number of limitations to be considered during

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experimental design and analyses of results, which will be discussed in Subheading 2 [10, 11]. 1.2 Inducible Gene Deletion

Although cell-specific control of Cre-recombinase expression can in many cases bypass the embryonic lethality of mutant mice generated via global gene deletion, it can also lead to embryonic lethality due to an essential role of the targeted gene for developing tissues. In addition, embryonic gene inactivation in a specific cell lineage, especially if induced early during the differentiation of this lineage, can result in significant phenotypes affecting multiple tissues and biological processes. Indeed, the engineered gene modification is irreversible and will be transmitted to all viable daughter cells that may contribute to the structure and function of tissues other than the one originally targeted. One might also want to study gene function at various stages of development, or during aging, or at specific stages of certain pathological conditions, such as bone healing in adults. In these situations, the timing of Cre recombinase activity must be controlled. To this end, the Cre recombinase has been fused to receptors that are responsive to either tetracycline/ doxycycline [12–14], tamoxifen [2, 6, 15] or type I interferon [16], enabling temporal control of Cre recombinase activity. Thus far, the tamoxifen and the Tet/Doxycycline systems have mainly been used in the bone field.

1.2.1 Tamoxifen-Based Models

Currently, the majority of available skeletal Cre transgenic or Cre knockin mice are based on modified Cre recombinases that are responsive to tamoxifen or 4-hydroxy-tamoxifen (OHT) treatment. In these lines, the Cre recombinase is fused with the human ligand binding domain of the estrogen receptor (hER). This receptor is normally sequestered in the cytoplasm but translocates to the nucleus upon ligand binding. However, the sensitivity of this chimeric Cre-hER protein to endogenous estrogen led to spontaneous Cre-mediated recombination upon use in physiological systems. To solve this issue, several Cre-hER fusion proteins have been mutated with the goal of reducing sensitivity to endogenous estrogen and increasing sensitivity to synthetic estrogen analogs such as tamoxifen. The Chambon laboratory developed the first Cre-hER fusion protein in the late 1990s and obtained proof of concept evidence that Cre-hER fusion proteins were able to specifically recombine DNA between LoxP sites and only after OHT treatment [2, 17]. To further decrease the sensitivity of the Cre-hER fused protein to endogenous estrogen, a mutation at glycine 521 (G521R) of the human ER ligand-binding domain was introduced, creating a “Cre-hERT” fusion protein that is insensitive to endogenous estrogen but is responsive to tamoxifen. Using synthetic estrogen molecules like tamoxifen and OHT improved the applicability of the Cre-ER system. However, these drugs can have significant effects on bone cells (see Subheading

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2.6), which can potentially affect the interpretation of results. To circumvent this issue, additional mutations were introduced in the Cre-hERT transgene (G400V, M543A, and M544A) to generate the so called “Cre-hERT2” fusion protein, which has a high affinity for synthetic estrogen ligands but none for endogenous ones [18]. Most of the transgenic mouse models available to date are built with a Cre recombinase transgene that is fused to the human ligand binding domain of the ER gene, but the so-called Cre-ERTM transgenic mice have the mutated ligand binding domain (G525R) of the murine estrogen receptor [19, 20]. 1.2.2 TetOff/On Based Models

The TetOff/On system, created in 1992 by Gossen and Bujard, was first designed to allow transient tunable expression of a target gene [21]. This system is based on the activity of an inducible transcriptional activator/repressor (tTA for the TetOff system and rtTA for the TetOn system), whose expression is regulated by a tissue-specific promoter. In the TetOff system, expressed tTA proteins bind to cognate operator tetO sequences and activate transcription of a selected gene (Cre recombinase for instance) from a minimal human CMV promoter. In this system, the addition of tetracycline/doxycycline chelates the tTA proteins and prevents the transactivation of the gene of interest, leading to an “OFF” state. The TetOn system is identical, with the exception of four amino acid exchanges that converts the tTA transcriptional activator into a repressor (rtTA). The latter can only bind to the tetO sequences and activates transcription of the target gene in the presence of tetracycline/doxycycline (“ON” state). Since the creation of the first transgenic mouse relying on the TetOff/On system in 1994 [22], over 100 tetracycline-responsive strains have been generated. Similar to the Cre-ER system, the TetOff/On system can be used for cell-type specific conditional gene induction or deletion, using specific promoters/enhancers to express the tTA/rtTA proteins in selected cells [23]. The Osx:: GFP-tTA-Cre [24] and the Col2a1-tTA-Cre mice [25] are two examples of lines based on this strategy to induce floxed gene recombination in bone and cartilage tissues, respectively.

2 Important Considerations for Interpretation of Results Derived from the Use of Cre Mice The Cre-LoxP technology has allowed for significant progress in the study of gene function and in our understanding of developmental and degenerative mechanisms, however it is not without limitations. Awareness and careful consideration of these limitations are necessary to use this approach and to rigorously analyze results derived from it.

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2.1 Insertion of LoxP Sites

Since the optimal size of the DNA fragment between two LoxP sites is 2 kb or shorter and since the average size of a murine gene is about 8 kb [26], the entire gene can usually not be deleted. Instead, the first coding exon(s) or functionally important parts of the gene (e.g., regions encoding catalytic or essential protein–protein interaction domains) are usually targeted. This approach, however, has the risk of generating truncated proteins that may have a dominantnegative effect and impact other proteins and processes in a way that is difficult to predict [27]. If the biology of the protein is well known, an alternative approach is to introduce point mutations that can ablate enzymatic activity or interactions with other proteins, which can be best accomplished by CRISPR/Cas9 technology. Of note, LoxP sites are recommended to be placed about 150–200 bp away from splice acceptor/donor sites [28].

2.2 Impact of Cre Recombinase Insertion into the Genome

In most cases, the construct containing the selected promoter and Cre transgene integrates randomly in the genome. Hence, there is a risk of the Cre transgene landing within the coding sequence of a gene, thereby disrupting its open reading frame. The promoter-Cre recombinase construct can also integrate in an intronic or regulatory region and alter expression of the gene(s) under the control of this region. In both cases, the insertion of the promoter-Cre recombinase construct can cause a phenotype that is not directly related to inactivation of the targeted floxed gene. It is therefore recommended (a) to compare CreTg/+ mice to Cre+/+ (WT) littermates to ensure there is no phenotype induced by insertion of the Cre transgene into an important genomic regions involved in the phenotype/mechanism under study and (b) to avoid generating Cre-transgenic mice with two copies of the Cre transgene, which can lead to a homozygous state of the genomic region where the construct inserted and thus to higher chance for a nonspecific, floxed gene-independent phenotype. In some Cre lines, only the Cre recombinase sequence is inserted through homologous recombination downstream of a selected endogenous promoter in the 50 /30 UTR; or is even nested within a particular gene in order to obtain a pattern of Cre recombinase expression that mimics the gene of interest. The targeted nature of this strategy eliminates the issue of potential disruption of genes different from the floxed one under study by random promoter-Cre transgene genomic insertion. However, it is possible in that case that integration of the Cre recombinase construct disrupts the expression of the “host” gene. This can be a problem for cases where haploinsufficiency of the gene driven by the selected promoter gives rise to a phenotype, as observed in the Acan-Cre ERT2 mice [29]. Hence, it is in general preferable to keep mice heterozygous for the Cre transgene to avoid any phenotype unrelated to inactivation of the floxed allele.

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Do not generate CreTg/Tg mice

Although the majority of Cre mouse lines develop normally, several studies have highlighted the effect of the Cre recombinase on genomic DNA stability. Indeed, Cre “toxicity” in neuronal progenitor-specific mouse lines and in cardiac-specific Cre lines was found to lead to cardiac [30–33] and brain developmental defects [34]. Cre activation was also shown to lead to reduced proliferation and increased apoptosis in hematopoietic cells, cardiomyocytes, insulin-producing β-cells, and fibroblasts [34–36]. One study showed reduced proliferation, numerous chromosomal aberrations and an increased number of sister chromatid exchanges in the presence of Cre recombinase and that this Cre toxicity correlated with the level of Cre activity [37]. Because limiting the intensity and duration of Cre expression can attenuate Cre toxicity in mammalian cells [38], constructs in which the Cre recombinase excises itself once a threshold expression required for floxed gene excision is reached can be an effective way of reducing Cre toxicity. In the bone field, the Osx-GFP::Cre line reportedly displays reduced stature at an early age [39, 40]. Such “toxic” effects of Cre activation can stem from illegitimate chromosomal rearrangements, from the presence of a passenger gene in the used BAC construct [41] or from micronuclei formations and other forms of DNA alterations that may depend on the presence of pseudo LoxP or cryptic sites [42, 43]. In fact, the Cre recombinase requires as few as 8–10 matches in its 13 bp binding site for efficient recombination [44, 45]. Early studies have shown that the Cre recombinase is capable of catalyzing recombination between cryptic LoxP sites naturally present in E. coli, yeast [46] and the mammalian genome [42]. A recent bioinformatics evaluation estimates that such sites are present in the mouse genome at a frequency of 1 per 1.2 megabases [43]. Therefore, upon use of Cre-expressing mice, efforts must be made to detect and report any phenotype caused by Cre recombinase activity in the absence of engineered LoxP sites. Cre toxicity: Compare [WT versus CreTg/+] 2.3 Irreversibility of Cre-mediated Gene Recombination

It is important to realize that the genetic modification subsequent to floxed gene recombination is irreversible and conserved in all viable daughter cells of the initial cell that underwent Cre-mediated recombination. For instance, osteocytes may have a floxed allele recombined by the Prx1-Cre transgene, although Prx1 is not expressed in osteocytes. This is because recombination occurred in Prx1-Cre + progenitor cells which then differentiate into

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osteoblasts and osteocytes. For this reason, the choice of a promoter that drives Cre recombinase expression in progenitor cells will usually generate a much broader pool of cells that have undergone gene recombination and a less specific tissue expression pattern when compared to a line where the Cre recombinase is expressed in more differentiated cells. This must be kept in mind when interpreting phenotypes generated by gene ablation at an early developmental stage. 2.4 Importance of Genotype Verification

The advent of the Cre/LoxP system has led to the expansion of Cre mouse lines available to the research community, which are often shared within and between laboratories. Although convenient and commonly used, genotyping these Cre lines with a generic set of primers that merely amplifies the Cre transgene does not distinguish between various Cre lines. This practice poses the significant risk of mistakenly swapping these Cre lines, as laboratories often host and handle several lines at a time and may utilize multiple lines per project. It is therefore highly recommended to design and validate specific sets of primers that can discriminate between Cre mouse lines, as recently reported by Couasnay et al. [47]. Bringing a reporter gene into a Cre/LoxP colony can also permits confirmation of the site of Cre recombination activity (see Subheading 3). To prevent accidental swap of Cre lines, use genotyping primers able to discriminate between Cre lines

2.5 Efficiency of Recombination

The efficiency of recombination of floxed alleles upon Cre activity is rarely 100% and thus typically gives rise to mosaic tissues containing recombined, partially recombined and nonrecombined cells. It is dependent on a number of factors, including the level of Cre recombinase expression [48], its activity upon induction (dependent of the concentration and regimen of tamoxifen or doxycycline) as well as accessibility to the LoxP sites, which can vary based on nuclear architecture or transcriptional activity. In the bone field, there are several cases of Cre lines in which Cre expression, activity or induction are suboptimal. Those include the Prx1-CreERT and the Col2-CreERT lines that show reduced activity at postnatal stages, or the mouse 2.3 kb Col1a1-Cre mice in which Cre activity was lost upon time and multiple generations in the original colony. It is therefore critical to check for DNA recombination and gene/ protein knock down for each conditional mutant mouse model generated. This can be done at several levels, including PCR on genomic DNA to visualize the presence of a recombined allele and by RT- qPCR and Western blot analyses to quantify the relative reduction in RNA and protein expression in mutant versus WT mice.

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It should be noted that the result of recombination efficiency analysis can be confounded by the cellular complexity of the tissues where the Cre recombinase is active. Bone is a typical example for this, with its different cellular lineages in marrow, growth plate, or cortical bone. Albeit more labor and time intensive, separation of the various bone compartments or sorting of specific cellular lineages by fluorescent-activated cell sorting (FACS) can improve accuracy for the quantification of recombination efficiency. Checking for recombination efficiency in an osteoblast or osteocytespecific Cre driver line for instance can benefit from performing measurements on flushed bones that mainly contain bone surface, adherent mature osteoblasts, and embedded osteocytes. Lastly, cases of Cre silencing, possibly via methylation or epigenetic changes, have been reported [49, 50], further reinforcing the need to assess allele deletion and reduction in expression of the encoded protein, in addition to detecting the presence of the Cre transgene. Cre recombinase efficiency: Measure gDNA recombination + mRNA and protein knockdown

2.6 Impact of Tamoxifen and Doxycycline on Bone Cells

Tamoxifen, OHT or tet/doxycycline are products that can impact skeletal physiology, especially when given at high dose. They can also affect other tissues that can in turn impact bone cells. For example, toxicity of tamoxifen and its derivatives are well-known for a wide range of tissues and biological pathways [51], including stomach [52], liver [53, 54], retinal pigment epithelial cells [55], and bone marrow stromal cells [56]. Despite its toxicity, tamoxifen remains the most common form of induction of the Cre recombinase because the majority of Cre lines are generated with the ERT2 moiety. To alleviate the deleterious effects of tamoxifen, its concentration and the frequency of its administration by injection or gavage can be titrated in a way to achieve maximal recombination at the lowest dose/regimen. Importantly, age, gender as well as the type of tissue that is targeted can require different amount and frequency of tamoxifen administration for optimal recombination efficiency. A review of the literature reveals a wide variability in the concentration of tamoxifen that is injected (5–300 μg/g of body weight) and the regimen of administration (from one injection up to daily injections for several weeks) [57–60], in line with the need of a tailored treatment for each Cre mouse line. One convenient solution to optimize the protocol of tamoxifen administration is to introduce a reporter allele such as the mTmG reporter in the background of a Flox/Cre mouse line to visualize occurrence and site of Cre activity. This transgenic mouse line expresses a membrane-localized fluorescent “tomato” reporter globally

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(driven by the CAG promoter), and Cre-mediated excision of this floxed Tomato transgene in cells where the Cre recombinase is expressed/activated switches expression of Tomato to a membrane-localized EGFP reporter. However, as discussed in the next paragraph, this approach is convenient to detect site of Cre activity, but may not provide an accurate quantification of recombination efficiency for the floxed allele that is under study. In addition to the mode of injection and dose/regimen protocol, the form of tamoxifen used can contribute to diverse outcomes. As reported previously by Kiermayer et al., tamoxifen citrate seems as effective as tamoxifen or its derivative, OHT as well as potentially safer. However, the oral delivery proposed in this approach can be affected by diet refusal by the animals, resulting in possible insufficient recombination, weight loss and starvation-related phenotypes [61]. Similar to tamoxifen, doxycycline has its own disadvantages. It is a matrix metalloproteinase inhibitor that can itself influence bone/cartilage homeostasis [62, 63]. Therefore, regardless of the model and inducer that is used, it is always a good idea to account for these and other (un)known effects of the Cre inducer by including in the analyses inducer-treated, Cre-negative mice that share the same genetic background. Choosing one inducible system over another largely depends on the goal and context of the study. As discussed above, the potential impacts of tamoxifen or doxycycline on the measured outcomes must be considered. The inducible nature of ON systems (ER and Tet-based) allows for transient activation of Cre recombinase activity necessary for pulse-type experiments in the context of tracing and for gene inactivation studies. The ON system also offers the advantage of only activating the Cre recombinase for short periods of time, thus limiting the nonspecific activity of the Cre recombinase on cryptic sites in the host genome. Because of the transient nature of Cre expression induced by the ON system and depending on the fate of the cell targeted (expansion, contribution to other lineages or cell death), this system may or may not be the most useful. The TetOff system on the other hand may be advantageous in cases where a gene needs to be deleted at an adult stage and for an extended period of time without further use of Cre inducers. In all cases, proper controls, usually Cre-negative (+/+); Flox/Flox mice  inducer, must be included to account for the toxicity of Cre inducers. Toxicity of the Cre inducer: Compare iCre+/+, GeneX f/f mice treated with Cre inducer or vehicle

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2.7 Off Target Activity of the Cre Recombinase

Although promoter-driven Cre mouse lines are commonly referred to as tissue- or cell-specific, Cre expression or activity can often, if not always, be detected in unexpected sites. Unfortunately, this so-called leaky expression is not always reported in the papers that describe or use these mice. Cre lines with “off target” Cre activity can be grouped into two main categories. One includes the cases of Cre transgene expression at early differentiation and/or early developmental stages that cause observed gene recombination down the lineage. This is not a true off target effect, as in this case the inactivation of the floxed allele in the targeted progenitor is transmitted to all viable daughter cells, whose number may vastly increase through the various stages of development. These cells can also contribute to different sublineages and to the formation of multiple tissues. An example of this would be promoters used for targeting mesenchymal stem cells (Prx1-Cre for instance) that recombine floxed alleles in mesenchymal stem cells, leading to gene inactivation in osteoblasts, osteocytes, chondrocytes, tenocytes, or adipocytes. The second category of Cre lines with off target Cre activity includes the ones with unexpected activity in tissues that are unrelated to bone, such as for instance the brain or heart. This Cre “off target” activity can be due to our incomplete knowledge of the expression pattern of the native gene in different organs or tissues, or to insertion of the transgene in the proximity of enhancers, suppressors or other regulatory elements that induce expression of the Cre recombinase in unexpected tissues, or to failure in incorporating all regulatory elements in the promoter sequence that was cloned in front of the Cre recombinase gene. It is noteworthy that many “bone-specific” Cre lines induce recombination in the central nervous system. Those include Wnt1-Cre (although this one is expected based on the early neural crest cell specificity of this promoter) [19, 64], Sox9-CreERT [65], Col2-Cre [66, 67], Osx-Cre [68, 69], Rat 2.3 kb Col1-Cre and Rat 3.6 kb Col1-Cre [70], 10 kb Dmp1-Cre [71], and LysM-Cre [72]. Along the same line, one must be aware of possible “leakiness” of the CreERT2 recombinase in absence of induction [73]. Although less common, germline “leakage” of Cre activity can be observed and problematic since it will lead to transmission of the deleted gene to subsequent generations. Such cases of germline leakage were reported in Dermo1-Cre males [74], 2.4kbPrx1-Cre females [75], Rat 2.3kbCol1-Cre females [76], and in the Ctsk-Cre line [77]. In these cases, the breeding strategy must be adapted by appropriately selecting the gender carrying the Cre transgene. The high sensitivity of fluorescent reporters such as the Ai9 tdTomato reporter compared to the originally used chromogenic ß-galactosidase activity has facilitated the detection and precise characterization of unexpected expression patterns of many Cre transgenes. However, one should keep several points in mind

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when using these reporters: First, “leaky” Cre expression in a tissue other than the one targeted is not always consequential phenotypically, particularly if the gene to be deleted is not expressed in the tissues where off target Cre activity is detected. Second, the recombination pattern obtained with a particular Lox-Stop-Lox reporter line does not necessarily mimic that of the floxed gene of interest [78]. This is because the Lox-Stop-Lox reporter allele and the floxed allele under study are different and their recombination can be locus-dependent. Cell type and developmental stage-specific variation in chromatin structure at different loci, state of DNA methylation and transcriptional activity can indeed affect DNA accessibility. Of note is that reporters used to assess Cre activity or in cell tracing studies can be expressed endogenously. For instance, osteoclasts express the ß-galactosidase [79, 80] and collagen is autofluorescent [81]. There are several strategies to increase the specificity of Cre expression when generating a new Cre-mouse line. One is to use larger constructs, such as bacterial artificial chromosomes (BACs), to generate BAC transgenic mice that harbor a maximal number of regulatory sequences that control Cre recombinase expression. Another is to insert the promoter-Cre unit construct in an active but neutral genomic site to reduce the variation of expression that is due to the site of transgene insertion. “Knocking in” the Cre transgene within the endogenous locus of a selected promoter is an alternative solution, although as noted earlier this option can lead to deleterious hemizygous effects. The latter issue can be avoided by using an Internal Ribosome Entry Site (IRES)-Cre cassette that would leave the “hijacked” gene unaffected. When trying to optimize the use of existing Cre lines with off target activity, using an inducible Cre system to ablate a gene of interest at postnatal stages can aid in restricting expression of the Cre transgene to a more selective pool of cells. For instance, the 10 kb Dmp1-CreERT2 line allows a more specific recombination in osteocytes than the 10 kb Dmp1-Cre line, which displays Cre activity in osteocytes but also skeletal muscles and bone marrow. Another strategy, although more onerous, is to analyze multiple cKO models in parallel, in which the same floxed allele is deleted using various Cre lines that share expression/activity in the tissue/ lineage of interest, but have differential off target patterns. In that case, the phenotype can be attributed to the common site of Cre expression only. A complementary approach to assign the origin of an in vivo phenotype to a selective population of cells consists of inducing deletion of a floxed gene ex vivo in this selected population of cells. This can be achieved by purifying the cells of interest from the “floxed” mouse line, and subsequently infecting these cells with viral particles expressing the Cre recombinase. This approach has been used extensively for bone marrow stromal cells and chondrocytes [82, 83].Of note is that cell infection with virus

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particles may alter cell behavior and viability, and hence requires appropriate controls (GFP-expressing viruses for instance) and recovery time post infection. Alternatively, the same ex vivo manipulation can be performed on cells extracted from mice carrying an inducible Cre recombinase, by adding the Cre-inducer ex vivo in the cell culture medium. In this case, OHT, the active form of the prodrug tamoxifen is required, because tamoxifen is activated via a hepatic hydroxylation step which will not occur ex vivo. Unexpected sites of Cre activity: Assess genomic recombination of the floxed allele in multiple tissues, or use of a reporter mouse

2.8 Novel Options to Improve Specificity of the Cre/Lox System

Overall, the Cre-LoxP system remains the most widely used strategy to investigate the role of selected genes in physiological and pathophysiological processes, but as discussed above, it is associated with a significant number of drawbacks. There is thus room for improvement, and work toward this goal is ongoing. Apart from increasing the selective sensitivity for tamoxifen, Cre-variants that have different affinity for distinct LoxP sites have been created by a directed evolution mutation approach, thus opening the possibility for the creation of Cre lines with higher Cre recombinase specificity and lower potential recognition of cryptic LoxP sites present in the mammalian genome [38, 84]. As mentioned above, self-excising Cre constructs have also been created to reduce Cre toxicity [85]. A novel advancement, the spilt Cre strategy also shows potential in improving specificity of Cre activity. This technology expresses two parts of the Cre recombinase under the control of two different promoters. By itself, one half of Cre recombinase is unable to mediate recombination. Only when the two fragments are expressed in the same cell and can interact with each other, is recombination between LoxP sites induced. Hence, Cre activity in this approach is restricted to cells/tissues where both promoters are active [86–88]. For instance, a Split-Cre system based on the use of the Dmp1 and Sost promoters would restrict Cre activity and gene recombination in osteocytes by eliminating Cre recombinase expression in skeletal muscles and in the hematopoietic lineage, as observed in the Dmp1-Cre and Sost-Cre transgenic mouse models, respectively. A disadvantage of this approach is that it will require at least three alleles together in the same mouse and more controls than usual. An extensive list of Cre mice targeting skeletal tissues can be found on the MGI website (http://www.informatics.jax.org/ home/recombinase). Table 1 summarizes the main Cre lines used in the bone field and includes the original report describing each line following a cross to reporter mice.

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Table 1 List of the most commonly used Cre mice in the bone field and associated reference of the original work describing each line Lineage

Promoter

References

“Skeletal stem cell” Cre lines

Dermo1-Cre 2.4b Prx1-Cre 2.4 kb Prx1-CreERT LepR-Cre Mx1-Cre aSma-CreERT2 Sox9-Cre Sox9-CreERT2 Gremlin1-CreERT Axin2-CreERT2

[91] [75] [92] [93–95] [96] [97] [98] [65] [99] [100] [101]

“Chondrocyte” Cre lines

Col2-Cre Col2-Cre Col2-Cre Col2-Cre hCol2-Cre Col2-CreERT Col2-CreERT2 Col2-CreERTM Col2rtTA-Cre AcanCre-ERT2 Col10a1-Cre Col10a1-Cre Col10a1-Cre Col10a1-CreERT GDF5-Cre Prg4-GFPCreERT2

[102] [66] [103] [67] [104] [105] [60] [106] [25] [58] [107] [108] [109] [110] [111] [112]

“Osteoblast” Cre lines

Runx2-iCre Osx1-GFP::Cre Osx-CreERT2 2.3 k Col1a1-Cre Rat 2.3 kb Col1a1-Cre Rat 3.6 kb Col1a1-Cre 3.2 kb Col1a1-CreERTM hOC-Cre hOC-CreERT2

[113] [24] [59] [114] [76] [76] [59] [115] [116]

“Osteocyte” Cre lines

8 kb Dmp1-Cre 10 kb Dmp1-Cre 10 kb Dmp1-CreERT2 Sost-Cre

[117, 118] [119] [57] [120]

“Osteoclast” Cre lines

Ctsk-Cre Ctsk-CreERT2 TRAP-Cre LysM-Cre hCD11b-Cre RANK-Cre Mb1-Cre

[121] [122] [121] [123] [124] [125] [126]

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Use of the Cre-LoxP Technology for Lineage Tracing Because activation of a reporter gene by Cre-mediated recombination indelibly marks Cre-expressing cells and their descendants, Cre-based strategies are increasingly used for in vivo cell lineage tracing analyses. The development of tissue clearing methods adapted to bone tissues has also promoted the use of Cre-based tracing tools [89]. In this approach, a gene encoding a chromogenic (LacZ, ALP) or fluorescent (GFP, Tomato, etc.) moiety is activated upon Cre-based excision of an upstream engineered LoxStop-Lox sequence. The choice of the promoter used to drive Cre expression thus leads to activation of the reporter transgene in the targeted cell population and its descendants, and the use of inducible cre-recombinases and inductions at well-chosen stages allows lineage tracing. With a judicious selection of the Cre-activated fluorescent constructs in combination with additional transgenes that label specific cell lineages, this approach can be used postmortem or in vivo to track the position, density or migration of cells in tissues, and their transition to differentiated stages [90].

4

Conclusions The Cre-LoxP system remains one of the most powerful tools to investigate gene function in skeletal biology research. Considering both advantages and limitations of this technology, it has become better recognized that the use and comparison of multiple Cre lines to target ablation of a floxed gene has multiple benefits. It is required for being able to dissociate the impact of targeted versus unexpected Cre activity on the phenotype(s) observed and thus for robust interpretations of results and proper understanding of gene function in time and space. The Cre-LoxP system is continuously evolving and it is likely that specificity and toxicity limitations associated with current Cre lines will be circumvented by the creation of new Cre recombinases or Flox sequences.

Acknowledgments This work was supported by funding from the National Institute of Health (RO1AG055394, R21AR072483) and the Department of Defense (GRANT12693412).

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References 1. Lewandoski M (2001) Conditional control of gene expression in the mouse. Nat Rev Genet 2:743–755. https://doi.org/10.1038/ 35093537 2. Metzger D, Clifford J, Chiba H, Chambon P (1995) Conditional site-specific recombination in mammalian cells using a liganddependent chimeric Cre recombinase. Proc Natl Acad Sci U S A 92:6991–6995 3. Wang Y, Spatz MK, Kannan K, Hayk H, Avivi A, Gorivodsky M, Pines M, Yayon A, Lonai P, Givol D (1999) A mouse model for achondroplasia produced by targeting fibroblast growth factor receptor 3. Proc Natl Acad Sci U S A 96:4455–4460 4. Fedde KN, Blair L, Silverstein J, Coburn SP, Ryan LM, Weinstein RS, Waymire K, Narisawa S, Milla´n JL, MacGregor GR, Whyte MP (1999) Alkaline phosphatase knock-out mice recapitulate the metabolic and skeletal defects of infantile hypophosphatasia. J Bone Miner Res 14:2015–2026. https://doi.org/10.1359/jbmr.1999.14.12. 2015 5. International Mouse Knockout Consortium, Collins FS, Rossant J, Wurst W (2007) A mouse for all reasons. Cell 128:9–13. https://doi.org/10.1016/j.cell.2006.12. 018 6. Feil R, Brocard J, Mascrez B, LeMeur M, Metzger D, Chambon P (1996) Ligandactivated site-specific recombination in mice. Proc Natl Acad Sci U S A 93:10887–10890 7. Sauer B, Henderson N (1988) Site-specific DNA recombination in mammalian cells by the Cre recombinase of bacteriophage P1. Proc Natl Acad Sci U S A 85:5166–5170 8. Andrews BJ, Proteau GA, Beatty LG, Sadowski PD (1985) The FLP recombinase of the 2 micron circle DNA of yeast: interaction with its target sequences. Cell 40:795–803 9. Scho¨nhuber N, Seidler B, Schuck K, Veltkamp C, Schachtler C, Zukowska M, Eser S, Feyerabend TB, Paul MC, Eser P, Klein S, Lowy AM, Banerjee R, Yang F, Lee C-L, Moding EJ, Kirsch DG, Scheideler A, Alessi DR, Varela I, Bradley A, Kind A, Schnieke AE, Rodewald H-R, Rad R, Schmid RM, Schneider G, Saur D (2014) A nextgeneration dual-recombinase system for time- and host-specific targeting of pancreatic cancer. Nat Med 20:1340–1347. https://doi. org/10.1038/nm.3646

10. Birling M-C, Gofflot F, Warot X (2009) Sitespecific recombinases for manipulation of the mouse genome. Methods Mol Biol 561:245–263. https://doi.org/10.1007/ 978-1-60327-019-9_16 11. Nagy A (2000) Cre recombinase: the universal reagent for genome tailoring. Genesis 26:99–109 12. St-Onge L, Furth PA, Gruss P (1996) Temporal control of the Cre recombinase in transgenic mice by a tetracycline responsive promoter. Nucleic Acids Res 24:3875–3877 13. Baron U, Bujard H (2000) Tet repressorbased system for regulated gene expression in eukaryotic cells: principles and advances. Methods Enzymol 327:401–421 14. Bockamp E, Maringer M, Spangenberg C, Fees S, Fraser S, Eshkind L, Oesch F, Zabel B (2002) Of mice and models: improved animal models for biomedical research. Physiol Genomics 11:115–132. https://doi.org/10. 1152/physiolgenomics.00067.2002 15. Brocard J, Feil R, Chambon P, Metzger D (1998) A chimeric Cre recombinase inducible by synthetic, but not by natural ligands of the glucocorticoid receptor. Nucleic Acids Res 26:4086–4090 16. Ku¨hn R, Schwenk F, Aguet M, Rajewsky K (1995) Inducible gene targeting in mice. Science 269:1427–1429 17. Metzger D, Chambon P (2001) Site- and time-specific gene targeting in the mouse. Methods 24:71–80. https://doi.org/10. 1006/meth.2001.1159 18. Feil R, Wagner J, Metzger D, Chambon P (1997) Regulation of Cre recombinase activity by mutated estrogen receptor ligandbinding domains. Biochem Biophys Res Commun 237:752–757. https://doi.org/ 10.1006/bbrc.1997.7124 19. Danielian PS, Muccino D, Rowitch DH, Michael SK, McMahon AP (1998) Modification of gene activity in mouse embryos in utero by a tamoxifen-inducible form of Cre recombinase. Curr Biol 8:1323–1326 20. Littlewood TD, Hancock DC, Danielian PS, Parker MG, Evan GI (1995) A modified oestrogen receptor ligand-binding domain as an improved switch for the regulation of heterologous proteins. Nucleic Acids Res 23:1686–1690 21. Gossen M, Bujard H (1992) Tight control of gene expression in mammalian cells by tetracycline-responsive promoters. Proc Natl Acad Sci U S A 89:5547–5551

54

Florent Elefteriou and Greig Couasnay

22. Furth PA, St Onge L, Bo¨ger H, Gruss P, Gossen M, Kistner A, Bujard H, Hennighausen L (1994) Temporal control of gene expression in transgenic mice by a tetracycline-responsive promoter. Proc Natl Acad Sci U S A 91:9302–9306 23. Cox BC, Dearman JA, Brancheck J, Zindy F, Roussel MF, Zuo J (2014) Generation of Atoh1-rtTA transgenic mice: a tool for inducible gene expression in hair cells of the inner ear. Sci Rep 4:6885. https://doi.org/10. 1038/srep06885 24. Rodda SJ, McMahon AP (2006) Distinct roles for Hedgehog and canonical Wnt signaling in specification, differentiation and maintenance of osteoblast progenitors. Development 133:3231–3244. https://doi. org/10.1242/dev.02480 25. Grover J, Roughley PJ (2006) Generation of a transgenic mouse in which Cre recombinase is expressed under control of the type II collagen promoter and doxycycline administration. Matrix Biol 25:158–165. https://doi. org/10.1016/j.matbio.2005.11.003 26. Jareborg N, Birney E, Durbin R (1999) Comparative analysis of noncoding regions of 77 orthologous mouse and human gene pairs. Genome Res 9:815–824 27. Yang SH, Bergo MO, Farber E, Qiao X, Fong LG, Young SG (2009) Caution! Analyze transcripts from conditional knockout alleles. Transgenic Res 18:483–489. https://doi. org/10.1007/s11248-008-9237-9 28. protocol4_cko_design.pdf 29. Rashid H, Chen H, Hassan Q, Javed A (2017) Dwarfism in homozygous Agc1CreERT mice is associated with decreased expression of aggrecan. Genesis 55(10):10.1002/ dvg.23070. https://doi.org/10.1002/dvg. 23070 30. Buerger A, Rozhitskaya O, Sherwood MC, Dorfman AL, Bisping E, Abel ED, Pu WT, Izumo S, Jay PY (2006) Dilated cardiomyopathy resulting from high-level myocardial expression of Cre-recombinase. J Card Fail 12:392–398. https://doi.org/10.1016/j. cardfail.2006.03.002 31. Hall ME, Smith G, Hall JE, Stec DE (2011) Systolic dysfunction in cardiac-specific ligandinducible MerCreMer transgenic mice. Am J Physiol Heart Circ Physiol 301:H253–H260. https://doi.org/10.1152/ajpheart.00786. 2010 32. Koitabashi N, Danner T, Zaiman AL, Pinto YM, Rowell J, Mankowski J, Zhang D, Nakamura T, Takimoto E, Kass DA (2011) Pivotal role of cardiomyocyte TGF-β signaling

in the murine pathological response to sustained pressure overload. J Clin Invest 121:2301–2312. https://doi.org/10.1172/ JCI44824 33. Bersell K, Choudhury S, Mollova M, Polizzotti BD, Ganapathy B, Walsh S, Wadugu B, Arab S, Ku¨hn B (2013) Moderate and high amounts of tamoxifen in αMHC-MerCreMer mice induce a DNA damage response, leading to heart failure and death. Dis Model Mech 6:1459–1469. https://doi.org/10.1242/ dmm.010447 34. Forni PE, Scuoppo C, Imayoshi I, Taulli R, Dastru` W, Sala V, Betz UAK, Muzzi P, Martinuzzi D, Vercelli AE, Kageyama R, Ponzetto C (2006) High levels of Cre expression in neuronal progenitors cause defects in brain development leading to microencephaly and hydrocephaly. J Neurosci 26:9593–9602. https://doi.org/10.1523/ JNEUROSCI. 2815-06.2006 35. Takebayashi H, Usui N, Ono K, Ikenaka K (2008) Tamoxifen modulates apoptosis in multiple modes of action in CreER mice. Genesis 46:775–781. https://doi.org/10. 1002/dvg.20461 36. Naiche LA, Papaioannou VE (2007) Cre activity causes widespread apoptosis and lethal anemia during embryonic development. Genesis 45:768–775. https://doi.org/10.1002/ dvg.20353 37. Loonstra A, Vooijs M, Beverloo HB, Allak BA, van Drunen E, Kanaar R, Berns A, Jonkers J (2001) Growth inhibition and DNA damage induced by Cre recombinase in mammalian cells. Proc Natl Acad Sci U S A 98:9209–9214. https://doi.org/10.1073/ pnas.161269798 38. M€aho¨nen AJ, Airenne KJ, Lind MM, Lesch HP, Yl€a-Herttuala S (2004) Optimized selfexcising Cre-expression cassette for mammalian cells. Biochem Biophys Res Commun 320:366–371. https://doi.org/10.1016/j. bbrc.2004.05.175 39. Huang W, Olsen BR (2015) Skeletal defects in Osterix-Cre transgenic mice. Transgenic Res 24:167–172. https://doi.org/10.1007/ s11248-014-9828-6 40. Wang L, Mishina Y, Liu F (2015) Osterix-Cre transgene causes craniofacial bone development defect. Calcif Tissue Int 96:129–137. https://doi.org/10.1007/s00223-0149945-5 41. Thomas C, Pellicciari R, Pruzanski M, Auwerx J, Schoonjans K (2008) Targeting bile-acid signalling for metabolic diseases. Nat Rev Drug Discov 7:678–693. https:// doi.org/10.1038/nrd2619

Overview for the Use of Skeleton-Related Cre Mouse Lines 42. Thyagarajan B, Guimara˜es MJ, Groth AC, Calos MP (2000) Mammalian genomes contain active recombinase recognition sites. Gene 244:47–54 43. Semprini S, Troup TJ, Kotelevtseva N, King K, Davis JRE, Mullins LJ, Chapman KE, Dunbar DR, Mullins JJ (2007) Cryptic loxP sites in mammalian genomes: genomewide distribution and relevance for the efficiency of BAC/PAC recombineering techniques. Nucleic Acids Res 35:1402–1410. https://doi.org/10.1093/nar/gkl1108 44. Abremski K, Hoess R (1985) Phage P1 Cre-loxP site-specific recombination. Effects of DNA supercoiling on catenation and knotting of recombinant products. J Mol Biol 184:211–220 45. Abremski K, Frommer B, Wierzbicki A, Hoess RH (1988) Properties of a mutant Cre protein that alters the topological linkage of recombination products. J Mol Biol 202:59–66 46. Sauer B (1992) Identification of cryptic lox sites in the yeast genome by selection for Cre-mediated chromosome translocations that confer multiple drug resistance. J Mol Biol 223:911–928 47. Couasnay G, Frey C, Elefteriou F (2019) Promoter Cre-specific genotyping assays for authentication of Cre-driver mouse lines. JBMR Plus 3(4):e10128. https://doi.org/ 10.1002/jbm4.10128 48. Araki K, Imaizumi T, Okuyama K, Oike Y, Yamamura K (1997) Efficiency of recombination by Cre transient expression in embryonic stem cells: comparison of various promoters. J Biochem 122:977–982 49. Long MA, Rossi FMV (2009) Silencing inhibits Cre-mediated recombination of the Z/AP and Z/EG reporters in adult cells. PLoS One 4:e5435. https://doi.org/10. 1371/journal.pone.0005435 50. Schulz TJ, Glaubitz M, Kuhlow D, Thierbach R, Birringer M, Steinberg P, Pfeiffer AFH, Ristow M (2007) Variable expression of Cre recombinase transgenes precludes reliable prediction of tissue-specific gene disruption by tail-biopsy genotyping. PLoS One 2:e1013. https://doi.org/10.1371/journal. pone.0001013 51. Birge SJ, McEwen BS, Wise PM (2001) Effects of estrogen deficiency on brain function. Implications for the treatment of postmenopausal women. Postgrad Med Spec No:11–16 52. Huh WJ, Khurana SS, Geahlen JH, Kohli K, Waller RA, Mills JC (2012) Tamoxifen

55

induces rapid, reversible atrophy, and metaplasia in mouse stomach. Gastroenterology 142:21.e7–24.e7. https://doi.org/10. 1053/j.gastro.2011.09.050 53. Phillips DH, Carmichael PL, Hewer A, Cole KJ, Hardcastle IR, Poon GK, Keogh A, Strain AJ (1996) Activation of tamoxifen and its metabolite alpha-hydroxytamoxifen to DNA-binding products: comparisons between human, rat and mouse hepatocytes. Carcinogenesis 17:89–94 54. Stearns V, Gelmann EP (1998) Does tamoxifen cause cancer in humans? J Clin Oncol 16:779–792. https://doi.org/10.1200/ JCO.1998.16.2.779 55. Kim LA, Amarnani D, Gnanaguru G, Tseng WA, Vavvas DG, D’Amore PA (2014) Tamoxifen toxicity in cultured retinal pigment epithelial cells is mediated by concurrent regulated cell death mechanisms. Invest Ophthalmol Vis Sci 55:4747–4758. https:// doi.org/10.1167/iovs.13-13662 56. McHaffie SL, Hastie ND, Chau Y-Y (2016) Effects of CreERT2, 4-OH tamoxifen, and gender on CFU-F assays. PLoS One 11: e0148105. https://doi.org/10.1371/jour nal.pone.0148105 57. Powell WF, Barry KJ, Tulum I, Kobayashi T, Harris SE, Bringhurst FR, Pajevic PD (2011) Targeted ablation of the PTH/PTHrP receptor in osteocytes impairs bone structure and homeostatic calcemic responses. J Endocrinol 209:21–32. https://doi.org/10.1530/JOE10-0308 58. Henry SP, Jang C-W, Deng JM, Zhang Z, Behringer RR, de Crombrugghe B (2009) Generation of aggrecan-CreERT2 knockin mice for inducible Cre activity in adult cartilage. Genesis 47:805–814. https://doi.org/ 10.1002/dvg.20564 59. Maes C, Kobayashi T, Selig MK, Torrekens S, Roth SI, Mackem S, Carmeliet G, Kronenberg HM (2010) Osteoblast precursors, but not mature osteoblasts, move into developing and fractured bones along with invading blood vessels. Dev Cell 19:329–344. https://doi.org/10.1016/j.devcel.2010.07. 010 60. Chen M, Lichtler AC, Sheu T-J, Xie C, Zhang X, O’Keefe RJ, Chen D (2007) Generation of a transgenic mouse model with chondrocyte-specific and tamoxifen-inducible expression of Cre recombinase. Genesis 45:44–50. https://doi.org/10.1002/dvg. 20261 61. Kiermayer C, Conrad M, Schneider M, Schmidt J, Brielmeier M (2007) Optimization of spatiotemporal gene inactivation in mouse

56

Florent Elefteriou and Greig Couasnay

heart by oral application of tamoxifen citrate. Genesis 45:11–16. https://doi.org/10. 1002/dvg.20244 62. Fowlkes JL, Nyman JS, Bunn RC, Cockrell GE, Wahl EC, Rettiganti MR, Lumpkin CK, Thrailkill KM (2015) Effects of long-term doxycycline on bone quality and strength in diabetic male DBA/2J mice. Bone Rep 1:16–19. https://doi.org/10.1016/j.bonr. 2014.10.001 63. do Nascimento Gomes K, APNN A, PGP D, de GSB V (2017) Doxycycline induces bone repair and changes in Wnt signalling. Int J Oral Sci 9:158–166. https://doi.org/10. 1038/ijos.2017.28 64. Lewis AE, Vasudevan HN, O’Neill AK, Soriano P, Bush JO (2013) The widely used Wnt1-Cre transgene causes developmental phenotypes by ectopic activation of Wnt signaling. Dev Biol 379:229–234. https://doi. org/10.1016/j.ydbio.2013.04.026 65. Soeda T, Deng JM, de Crombrugghe B, Behringer RR, Nakamura T, Akiyama H (2010) Sox9-expressing precursors are the cellular origin of the cruciate ligament of the knee joint and the limb tendons. Genesis 48:635–644. https://doi.org/10.1002/dvg. 20667 66. Sakai K, Hiripi L, Glumoff V, Brandau O, Eerola R, Vuorio E, Bo¨sze Z, F€assler R, Aszo´di A (2001) Stage-and tissue-specific expression of a Col2a1-Cre fusion gene in transgenic mice. Matrix Biol 19:761–767 67. Long F, Zhang XM, Karp S, Yang Y, McMahon AP (2001) Genetic manipulation of hedgehog signaling in the endochondral skeleton reveals a direct role in the regulation of chondrocyte proliferation. Development 128:5099–5108 68. Chen J, Shi Y, Regan J, Karuppaiah K, Ornitz DM, Long F (2014) Osx-Cre targets multiple cell types besides osteoblast lineage in postnatal mice. PLoS One 9:e85161. https://doi. org/10.1371/journal.pone.0085161 69. Park J-S, Baek W-Y, Kim YH, Kim J-E (2011) In vivo expression of Osterix in mature granule cells of adult mouse olfactory bulb. Biochem Biophys Res Commun 407:842–847. https://doi.org/10.1016/j.bbrc.2011.03. 129 70. Scheller EL, Leinninger GM, Hankenson KD, Myers MG, Krebsbach PH (2011) Ectopic expression of Col2.3 and Col3.6 promoters in the brain and association with leptin signaling. Cells Tissue Org 194:268–273. https:// doi.org/10.1159/000324745

71. Lim J, Burclaff J, He G, Mills JC, Long F (2017) Unintended targeting of Dmp1-Cre reveals a critical role for Bmpr1a signaling in the gastrointestinal mesenchyme of adult mice. Bone Res 5:16049. https://doi.org/ 10.1038/boneres.2016.49 72. Orthgiess J, Gericke M, Immig K, Schulz A, Hirrlinger J, Bechmann I, Eilers J (2016) Neurons exhibit Lyz2 promoter activity in vivo: implications for using LysM-Cre mice in myeloid cell research. Eur J Immunol 46:1529–1532. https://doi.org/10.1002/ eji.201546108 73. Seime T, Kolind M, Mikulec K, Summers MA, Cantrill L, Little DG, Schindeler A (2015) Inducible cell labeling and lineage tracking during fracture repair. Develop Growth Differ 57:10–23. https://doi.org/10.1111/dgd. 12184 74. He Y, Sun X, Wang L, Mishina Y, Guan J-L, Liu F (2017) Male germline recombination of a conditional allele by the widely used Dermo1-cre (Twist2-cre) transgene Genesis 55(9):10.1002/dvg.23048. https://doi. org/10.1002/dvg.23048 75. Logan M, Martin JF, Nagy A, Lobe C, Olson EN, Tabin CJ (2002) Expression of Cre recombinase in the developing mouse limb bud driven by a Prxl enhancer. Genesis 33:77–80. https://doi.org/10.1002/gene. 10092 76. Liu F, Woitge HW, Braut A, Kronenberg MS, Lichtler AC, Mina M, Kream BE (2004) Expression and activity of osteoblast-targeted Cre recombinase transgenes in murine skeletal tissues. Int J Dev Biol 48:645–653. https:// doi.org/10.1387/ijdb.041816fl 77. Winkeler CL, Kladney RD, Maggi LB, Weber JD (2012) Cathepsin K-Cre causes unexpected germline deletion of genes in mice. PLoS One 7:e42005. https://doi.org/10. 1371/journal.pone.0042005 78. Liu J, Willet SG, Bankaitis ED, Xu Y, Wright CVE, Gu G (2013) Non-parallel recombination limits Cre-LoxP-based reporters as precise indicators of conditional genetic manipulation. Genesis 51:436–442. https:// doi.org/10.1002/dvg.22384 79. Kopp H-G, Hooper AT, Shmelkov SV, Rafii S (2007) Beta-galactosidase staining on bone marrow. The osteoclast pitfall. Histol Histopathol 22:971–976. https://doi.org/10. 14670/HH-22.971 80. Odgren PR, MacKay CA, Mason-Savas A, Yang M, Mailhot G, Birnbaum MJ (2006) False-positive beta-galactosidase staining in osteoclasts by endogenous enzyme: studies in neonatal and month-old wild-type mice.

Overview for the Use of Skeleton-Related Cre Mouse Lines Connect Tissue Res 47:229–234. https:// doi.org/10.1080/03008200600860086 81. Prentice AI (1967) Autofluorescence of bone tissues. J Clin Pathol 20:717–719 82. Tahaei SE, Couasnay G, Ma Y, Paria N, Gu J, Lemoine BF, Wang X, Rios JJ, Elefteriou F (2018) The reduced osteogenic potential of Nf1-deficient osteoprogenitors is EGFRindependent. Bone 106:103–111. https:// doi.org/10.1016/j.bone.2017.10.012 83. de la Croix NJ, Makowski AJ, Uppuganti S, Vignaux G, Ono K, Perrien DS, Joubert S, Baglio SR, Granchi D, Stevenson DA, Rios JJ, Nyman JS, Elefteriou F (2015) Corrigendum: asfotase-α improves bone growth, mineralization and strength in mouse models of neurofibromatosis type-1. Nat Med 21:414. https://doi.org/10.1038/nm0415-414c 84. Eroshenko N, Church GM (2013) Mutants of Cre recombinase with improved accuracy. Nat Commun 4:2509. https://doi.org/10. 1038/ncomms3509 85. Silver DP, Livingston DM (2001) Selfexcising retroviral vectors encoding the Cre recombinase overcome Cre-mediated cellular toxicity. Mol Cell 8:233–243 86. Hirrlinger J, Requardt RP, Winkler U, Wilhelm F, Schulze C, Hirrlinger PG (2009) Split-CreERT2: temporal control of DNA recombination mediated by split-Cre protein fragment complementation. PLoS One 4: e8354. https://doi.org/10.1371/journal. pone.0008354 87. Casanova E, Lemberger T, Fehsenfeld S, Mantamadiotis T, Schu¨tz G (2003) Alpha complementation in the Cre recombinase enzyme. Genesis 37:25–29. https://doi.org/ 10.1002/gene.10227 88. Seidi A, Mie M, Kobatake E (2007) Novel recombination system using Cre recombinase alpha complementation. Biotechnol Lett 29:1315–1322. https://doi.org/10.1007/ s10529-007-9406-6 89. Tainaka K, Kuno A, Kubota SI, Murakami T, Ueda HR (2016) Chemical principles in tissue clearing and staining protocols for wholebody cell profiling. Annu Rev Cell Dev Biol 32:713–741. https://doi.org/10.1146/ annurev-cellbio-111315-125001 90. Park D, Spencer JA, Lin CP, Scadden DT (2014) Sequential in vivo imaging of osteogenic stem/progenitor cells during fracture repair. J Vis Exp. https://doi.org/10.3791/ 51289 91. Yu K, Xu J, Liu Z, Sosic D, Shao J, Olson EN, Towler DA, Ornitz DM (2003) Conditional inactivation of FGF receptor 2 reveals an

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essential role for FGF signaling in the regulation of osteoblast function and bone growth. Development 130:3063–3074 92. Kawanami A, Matsushita T, Chan YY, Murakami S (2009) Mice expressing GFP and CreER in osteochondro progenitor cells in the periosteum. Biochem Biophys Res Commun 386:477–482. https://doi.org/10. 1016/j.bbrc.2009.06.059 93. DeFalco J, Tomishima M, Liu H, Zhao C, Cai X, Marth JD, Enquist L, Friedman JM (2001) Virus-assisted mapping of neural inputs to a feeding center in the hypothalamus. Science 291:2608–2613. https://doi. org/10.1126/science.1056602 94. Zhou BO, Yu H, Yue R, Zhao Z, Rios JJ, Naveiras O, Morrison SJ (2017) Bone marrow adipocytes promote the regeneration of stem cells and haematopoiesis by secreting SCF. Nat Cell Biol 19:891–903. https://doi. org/10.1038/ncb3570 95. Zhou BO, Yue R, Murphy MM, Peyer JG, Morrison SJ (2014) Leptin-receptor-expressing mesenchymal stromal cells represent the main source of bone formed by adult bone marrow. Cell Stem Cell 15:154–168. https://doi.org/10.1016/j.stem.2014.06. 008 96. Park D, Spencer JA, Koh BI, Kobayashi T, Fujisaki J, Clemens TL, Lin CP, Kronenberg HM, Scadden DT (2012) Endogenous bone marrow MSCs are dynamic, fate-restricted participants in bone maintenance and regeneration. Cell Stem Cell 10:259–272. https:// doi.org/10.1016/j.stem.2012.02.003 97. Grcevic D, Pejda S, Matthews BG, Repic D, Wang L, Li H, Kronenberg MS, Jiang X, Maye P, Adams DJ, Rowe DW, Aguila HL, Kalajzic I (2012) In vivo fate mapping identifies mesenchymal progenitor cells. Stem Cells 30:187–196. https://doi.org/10.1002/ stem.780 98. Akiyama H, Kim J-E, Nakashima K, Balmes G, Iwai N, Deng JM, Zhang Z, Martin JF, Behringer RR, Nakamura T, de Crombrugghe B (2005) Osteo-chondroprogenitor cells are derived from Sox9 expressing precursors. Proc Natl Acad Sci U S A 102:14665–14670. https://doi.org/10. 1073/pnas.0504750102 99. Worthley DL, Churchill M, Compton JT, Tailor Y, Rao M, Si Y, Levin D, Schwartz MG, Uygur A, Hayakawa Y, Gross S, Renz BW, Setlik W, Martinez AN, Chen X, Nizami S, Lee HG, Kang HP, Caldwell J-M, Asfaha S, Westphalen CB, Graham T, Jin G, Nagar K, Wang H, Kheirbek MA, Kolhe A, Carpenter J, Glaire M, Nair A, Renders S,

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Manieri N, Muthupalani S, Fox JG, Reichert M, Giraud AS, Schwabe RF, Pradere J-P, Walton K, Prakash A, Gumucio D, Rustgi AK, Stappenbeck TS, Friedman RA, Gershon MD, Sims P, Grikscheit T, Lee FY, Karsenty G, Mukherjee S, Wang TC (2015) Gremlin 1 identifies a skeletal stem cell with bone, cartilage, and reticular stromal potential. Cell 160:269–284. https://doi.org/10. 1016/j.cell.2014.11.042 100. Tan SH, Senarath-Yapa K, Chung MT, Longaker MT, Wu JY, Nusse R (2014) Wnts produced by Osterix-expressing osteolineage cells regulate their proliferation and differentiation. Proc Natl Acad Sci U S A 111: E5262–E5271. https://doi.org/10.1073/ pnas.1420463111 101. van Amerongen R, Bowman AN, Nusse R (2012) Developmental stage and time dictate the fate of Wnt/β-catenin-responsive stem cells in the mammary gland. Cell Stem Cell 11:387–400. https://doi.org/10.1016/j. stem.2012.05.023 102. Ovchinnikov DA, Deng JM, Ogunrinu G, Behringer RR (2000) Col2a1-directed expression of Cre recombinase in differentiating chondrocytes in transgenic mice. Genesis 26:145–146 103. Terpstra L, Prud’homme J, Arabian A, Takeda S, Karsenty G, Dedhar S, St-Arnaud R (2003) Reduced chondrocyte proliferation and chondrodysplasia in mice lacking the integrin-linked kinase in chondrocytes. J Cell Biol 162:139–148. https://doi.org/10. 1083/jcb.200302066 104. Haigh JJ, Gerber HP, Ferrara N, Wagner EF (2000) Conditional inactivation of VEGF-A in areas of collagen2a1 expression results in embryonic lethality in the heterozygous state. Development 127:1445–1453 105. Nakamura E, Nguyen M-T, Mackem S (2006) Kinetics of tamoxifen-regulated Cre activity in mice using a cartilage-specific CreER(T) to assay temporal activity windows along the proximodistal limb skeleton. Dev Dyn 235:2603–2612. https://doi.org/10. 1002/dvdy.20892 106. Hilton MJ, Tu X, Long F (2007) Tamoxifeninducible gene deletion reveals a distinct cell type associated with trabecular bone, and direct regulation of PTHrP expression and chondrocyte morphology by Ihh in growth region cartilage. Dev Biol 308:93–105. https://doi.org/10.1016/j.ydbio.2007.05. 011 107. Yang G, Cui F, Hou N, Cheng X, Zhang J, Wang Y, Jiang N, Gao X, Yang X (2005) Transgenic mice that express Cre recombinase

in hypertrophic chondrocytes. Genesis 42:33–36. https://doi.org/10.1002/gene. 20120 108. Kim Y, Murao H, Yamamoto K, Deng JM, Behringer RR, Nakamura T, Akiyama H (2011) Generation of transgenic mice for conditional overexpression of Sox9. J Bone Miner Metab 29:123–129. https://doi.org/ 10.1007/s00774-010-0206-z 109. Gebhard S, Hattori T, Bauer E, Schlund B, Bo¨sl MR, de Crombrugghe B, von der Mark K (2008) Specific expression of Cre recombinase in hypertrophic cartilage under the control of a BAC-Col10a1 promoter. Matrix Biol 27:693–699. https://doi.org/10.1016/j. matbio.2008.07.001 110. Yang L, Tsang KY, Tang HC, Chan D, Cheah KSE (2014) Hypertrophic chondrocytes can become osteoblasts and osteocytes in endochondral bone formation. Proc Natl Acad Sci U S A 111:12097–12102. https://doi.org/ 10.1073/pnas.1302703111 111. Rountree RB, Schoor M, Chen H, Marks ME, Harley V, Mishina Y, Kingsley DM (2004) BMP receptor signaling is required for postnatal maintenance of articular cartilage. PLoS Biol 2:e355. https://doi.org/10. 1371/journal.pbio.0020355 112. Kozhemyakina E, Zhang M, Ionescu A, Ayturk UM, Ono N, Kobayashi A, Kronenberg H, Warman ML, Lassar AB (2015) Identification of a Prg4-expressing articular cartilage progenitor cell population in mice. Arthritis Rheumatol 67:1261–1273. https://doi.org/10.1002/art.39030 113. Rauch A, Seitz S, Baschant U, Schilling AF, Illing A, Stride B, Kirilov M, Mandic V, Takacz A, Schmidt-Ullrich R, Ostermay S, Schinke T, Spanbroek R, Zaiss MM, Angel PE, Lerner UH, David J-P, Reichardt HM, Amling M, Schu¨tz G, Tuckermann JP (2010) Glucocorticoids suppress bone formation by attenuating osteoblast differentiation via the monomeric glucocorticoid receptor. Cell Metab 11:517–531. https://doi.org/10. 1016/j.cmet.2010.05.005 114. Dacquin R, Starbuck M, Schinke T, Karsenty G (2002) Mouse alpha1(I)-collagen promoter is the best known promoter to drive efficient Cre recombinase expression in osteoblast. Dev Dyn 224:245–251. https://doi. org/10.1002/dvdy.10100 115. Zhang M, Xuan S, Bouxsein ML, von Stechow D, Akeno N, Faugere MC, Malluche H, Zhao G, Rosen CJ, Efstratiadis A, Clemens TL (2002) Osteoblast-specific knockout of the insulinlike growth factor (IGF) receptor gene reveals

Overview for the Use of Skeleton-Related Cre Mouse Lines an essential role of IGF signaling in bone matrix mineralization. J Biol Chem 277:44005–44012. https://doi.org/10. 1074/jbc.M208265200 116. Yoshikawa Y, Kode A, Xu L, Mosialou I, Silva BC, Ferron M, Clemens TL, Economides AN, Kousteni S (2011) Genetic evidence points to an osteocalcin-independent influence of osteoblasts on energy metabolism. J Bone Miner Res 26:2012–2025. https://doi. org/10.1002/jbmr.417 117. Bivi N, Condon KW, Allen MR, Farlow N, Passeri G, Brun LR, Rhee Y, Bellido T, Plotkin LI (2012) Cell autonomous requirement of connexin 43 for osteocyte survival: consequences for endocortical resorption and periosteal bone formation. J Bone Miner Res 27:374–389. https://doi.org/10.1002/ jbmr.548 118. Kalajzic I, Braut A, Guo D, Jiang X, Kronenberg MS, Mina M, Harris MA, Harris SE, Rowe DW (2004) Dentin matrix protein 1 expression during osteoblastic differentiation, generation of an osteocyte GFP-transgene. Bone 35:74–82. https:// doi.org/10.1016/j.bone.2004.03.006 119. Lu Y, Xie Y, Zhang S, Dusevich V, Bonewald LF, Feng JQ (2007) DMP1-targeted Cre expression in odontoblasts and osteocytes. J Dent Res 86:320–325. https://doi.org/10. 1177/154405910708600404 120. Xiong J, Piemontese M, Onal M, Campbell J, Goellner JJ, Dusevich V, Bonewald L, Manolagas SC, O’Brien CA (2015) Osteocytes, not osteoblasts or lining cells, are the main source of the RANKL required for osteoclast formation in remodeling bone. PLoS One 10: e0138189. https://doi.org/10.1371/jour nal.pone.0138189

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121. Chiu WSM, McManus JF, Notini AJ, Cassady AI, Zajac JD, Davey RA (2004) Transgenic mice that express Cre recombinase in osteoclasts. Genesis 39:178–185. https://doi.org/ 10.1002/gene.20041 122. Sanchez-Fernandez MA, Sbacchi S, CorreaTapia M, Naumann R, Klemm J, Chambon P, Al-Robaiy S, Blessing M, Hoflack B (2012) Transgenic mice for a tamoxifen-induced, conditional expression of the Cre recombinase in osteoclasts. PLoS One 7:e37592. https://doi.org/10.1371/journal.pone. 0037592 123. Clausen BE, Burkhardt C, Reith W, Renkawitz R, Fo¨rster I (1999) Conditional gene targeting in macrophages and granulocytes using LysMcre mice. Transgenic Res 8:265–277 124. Ferron M, Vacher J (2005) Targeted expression of Cre recombinase in macrophages and osteoclasts in transgenic mice. Genesis 41:138–145. https://doi.org/10.1002/ gene.20108 125. Maeda K, Kobayashi Y, Udagawa N, Uehara S, Ishihara A, Mizoguchi T, Kikuchi Y, Takada I, Kato S, Kani S, Nishita M, Marumo K, Martin TJ, Minami Y, Takahashi N (2012) Wnt5a-Ror2 signaling between osteoblast-lineage cells and osteoclast precursors enhances osteoclastogenesis. Nat Med 18:405–412. https://doi. org/10.1038/nm.2653 126. Hobeika E, Thiemann S, Storch B, Jumaa H, Nielsen PJ, Pelanda R, Reth M (2006) Testing gene function early in the B cell lineage in mb1-cre mice. Proc Natl Acad Sci U S A 103:13789–13794. https://doi.org/10. 1073/pnas.0605944103

Part II Skeletal Repair, Parabiosis, Transplantations, and Organ Cultures

Chapter 4 Generation of Closed Transverse Fractures in Small Animals Anthony De Giacomo, Elise F. Morgan, and Louis C. Gerstenfeld Abstract The most common procedure that has been developed for use in rats and mice to model fracture healing is described. The nature of the regenerative processes that may be assessed and the types of research questions that may be addressed with this model are briefly outlined. The detailed surgical protocol to generate closed simple transverse fractures is presented and general considerations when setting up an experiment using this model are described. Key words Fracture healing, Surgical model, Rodent

1

Introduction

1.1 General Information on the Closed Model of Fracture Healing

Models of fracture healing generally are developed to assess repair after fracture of appendicular bones and have mainly focused on the long bones of the hind limbs. These models primarily heal through an endochondral bone formation process and with the development of an external callus, although the extent of callus formation is dependent on the type of fixation and the degree to which the fixation method stabilizes the fractured bone and is greatly influenced by mechanical signals that the healing callus experiences [1–4]. The most common model of bone repair used in rats and mice is produced by externally applied blunt trauma to generate a closed, simple transverse fracture. The most widespread application of this model was first described in Bonnarens and Einhorn [5], for use in rats and has been subsequently adapted in various forms for use in mouse by numerous investigators [6–9]. The fracture is generated via three-point bending to a long bone (usually the femur or tibia). Stabilization of the fracture is achieved by inserting an intramedullary pin prior to generating the fracture. The use of this model is the closest in anatomical site, etiology and fixation method to the most common fractures seen clinically since these fractures tend to be closed injuries that are produced by a traumatic event such as falls and other accidents. The model is well suited for high-throughput

Matthew J. Hilton (ed.), Skeletal Development and Repair: Methods and Protocols, Methods in Molecular Biology, vol. 2230, https://doi.org/10.1007/978-1-0716-1028-2_4, © Springer Science+Business Media, LLC, part of Springer Nature 2021

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screening, owing to the simplicity, speed (~15 min per animal), and reproducibility of the procedure [9]. The model can also be used to assess basic molecular processes that affect endochondral bone formation, and can be extrapolated to both embryological development [10–14] and post-natal epiphyseal growth of long bones [13, 15] (See these reviews for discussion of the comparison between developmental and fracture endochondral bone formation [16–18]). 1.2 Applications and Limitations of the Closed Model of Fracture Healing

2 2.1

It has been widely applied to assess the safety and efficacy of systemic pharmaceuticals that might affect fracture healing [19– 22]. Due to its closed nature, it has a lesser degree of reproducibility for the local delivery of biological therapeutics and pharmaceuticals than an open procedure. This is due to the fact that delivery of the therapeutics is via percutaneous injection at the fracture site [23], and its actual anatomical delivery in the callus can only be approximated by palpitation. Similarly, placement of the site of the fracture is more subjective than in an open osteotomy procedure since control over placement is achieved only by visible inspection of the positioning of the leg and by palpitation of the bone through skin and muscle before fracture. Fractures generated in this model can also have some degree of comminution [9]. Figure 1 shows a series of radiographs of fractures in the murine tibia and femur (Fig. 1a), and compares these optimal fractures to cases that would be excluded from a study, due to the fractures being displaced, poorly positioned, or comminuted (Fig. 1b).

Materials Animals

1. For rat studies, Sprague Dawley rats 350- to 450 g in weight are typically used with no more than a 50 g variation in group weights. 2. For mice, ages between 10 and 18 weeks can be used although within a group of mice that is used for a study individual mice should be within 2 weeks of each other (see Note 1).

2.2 Instruments and Reagents

The instruments and materials that are needed for carrying out the surgical procedure in either rats or mice is shown in Fig. 2a. 1. Electric hair trimmer. 2. Scalpel with (no. 15) disposable blades. 3. Forceps (Dumont Vessel Cannulation Forceps Inox .5 mm). 4. Scissors (Castroviejo Micro Dissecting scissors). 5. Hemastatic clamp (Halstead Mosquito Forceps 500 Straight, 1.3 mm tip).

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Fig. 1 Radiographic examples of closed simple transverse fracture model. (a) Examples of the pin placement and fracture of mouse femur (upper) and mouse tibia (lower) panels. Images were generated using dental X-ray device. (b) Three examples of fractures that would be excluded from study. The exclusion criteria are denoted in the figure with the arrow indicating the position of the fracture on the radiograph. Images were generated using a Faxitron® device

6. Wire cutters. 7. 1 ml syringe with (27 gauge) needle. 8. 100% isopropyl alcohol wipes. 9. Surgical gauze. 10. Povidone-iodine solution. 11. Warming pad or slide warmer. 12. Small animal X-ray imaging device. 13. Small animal fracture device. 14. Isoflurane or ketamine–xylazine. 15. Buprenorphine. 16. Dremel tool or small electric drill (for rat fractures). 17. ~0.9 mm stainless steel K-wire with a threaded tip (fixation device for rats).

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Fig. 2 Materials for carrying out closed fracture procedure. (a) The instruments and materials that are needed for carrying out the surgical procedure in either rats or mice. (b) Fracture device for the generation of closed transverse fractures by controlled blunt trauma and three-point bending. (1) Device as generated from the

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18. Stainless steel 23- to 27-gauge spinal needle stylet (fixation device for mice). 19. 5-0 sutures. Simple schematic drawings for making a fracture device for generating controlled blunt trauma are provided in Bonnarens and Einhorn [5]. The size of the device can be scaled appropriately for rats or mice. A more recent modification of this device that provides for more accurate positioning of the animal and better control for release of the weight that drives the blunt striker to generate the fractures was reported by Marturano et al. [9], and is currently in use in our laboratory (Fig. 2b). Figure 2c shows a display of the three types of fixation pins that we have used.

3

Methods The surgical steps for generating a closed fracture are shown in Fig. 3. The general protocol is as follows:

3.1 Preparation of an Animal Protocol

1. For all animal studies, a protocol approved by an Institutional Care and Use Committee should be generated to define the scientific rationale and goals for the study, number of animals needed, operative procedure, operative anesthesia, and postoperative analgesia and care. Detailed information on animal welfare, selection of methods for anesthesia, monitoring animals while anesthetized, sterile surgical techniques, and postoperative monitoring are available at https://www. aalaslearninglibrary.org/.

3.2 Preparation of the Surgical Site

1. Wipe down the surgical site with sterile 100% isopropyl alcohol wipes and then remove the hair from the surgical area with a small animal shaver. 2. Wipe down the site with surgical gauze that has been dipped in a solution of povidone–iodine. 3. Perform the surgery on a warming pad under sterile conditions.

3.3 Induction of Anesthesia

An isoflurane anesthesia machine may be used or a mixture of ketamine and xylazine may also be used.

ä Fig. 2 (continued) schematic drawings courtesy of Dr. Kristen Billiar, and as described in Marturano et al. [9]. (2) Drop weight and electromagnet striker release assembly. (3) Calibration scale to adjust distance of drop. (4) Blunt striking blade and anvil for positioning of femur and generation of three-point bending. (c) Three types of fixation pins used to stabilize closed fractures in rats and mice

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Fig. 3 The surgical steps of the closed fracture procedure. (1) Manual palpitation and positioning to localize line for incision over the central patellar groove. (2) Exposure of the center of the groove on the femoral and tibia condyle for pin insertion. (3) Lateral subluxation of the patella and extensor mechanism. (4) Creation of the entry hole for pin insertion. (5) Pin insertion. (6) Position of the femur for fracture. (7) Positioning of the mouse for postoperative X-ray and immediate postoperative X-ray assessment showing a successful mid-diaphyseal fracture

1. For isoflurane induction, the animal is induced in a closed chamber with a 4% isoflurane/oxygen mixture. Once induced the animal is maintained on a 2% isoflurane/oxygen mixture. 2. For ketamine and xylazine induction, the following dosage is used for mice (80–200 mg/kg ketamine and 7–20 mg/kg xylazine) and the dosage for rats (80–100 mg/kg ketamine and 5–10 mg/kg xylazine). 3. Prior to incision, the animals are also given a dose of .01 mg/ kg buprenorphine (Buprenex) to ensure that there will be immediate postoperative pain management. 3.4 Insertion of the Fixation Pin (See Note 2)

1. Pin insertion is carried out prior to fracture by making an anterior longitudinal midline incision centered over the knee joint. This incision is followed by identifying the extensor mechanism, which consists of the quadriceps, patellar tendon, patella, and patellar ligament. Careful attention is made to not disrupt this mechanism in order to allow immediate ambulation of either the rat or mouse following surgery. 2. A subsequent incision is made just medial to the patella and extensor mechanism, which is followed by elevating and displacing the quadriceps and extensor mechanism in a lateral fashion. After subluxating the patella and extensor mechanism

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laterally, the distal end of the femur as well as the proximal end of the tibia will be exposed. From this exposure, an entry hole is created in the center of the groove on either the femoral or tibia condyle for pin insertion (see Note 3). 3. The pin is inserted down the length of the medullary canal in either a retrograde manner for the femur or an antegrade manner for the tibia. The depth of insertion can be manually felt since the insertion into the canal encounters minimal resistance until meeting the corticals bone of the greater trochanter of the proximal femur or the distal epiphysis of the tibia (see Note 4). 4. The tip of the pin is then buried under the surface of the condyle. The length of the pin may be further trimmed using wire cutters if it is too long (see Note 5). 5. The incision is then closed with 5-0 absorbable gut suture. 3.5 Generation of the Fracture

1. Following the surgical procedure, the fracture is generated by dropping a weight onto the operated extremity using the fracture device described in Fig. 2. The weight is set at a defined initial height that will generate a large enough bending moment upon impact to fracture the bone. The combination of weight and initial height should be empirically determined for the specific strain, age, and sex of the animals.

3.6 Intraoperative Assessment of Quality of the Fracture

1. Immediately after fracture and before the animal revives from anesthesia, an X-ray should be taken (such as with a mobile dental X-ray unit) to check that placement of the intramedullary pin is adequate and that the fracture is mid-diaphyseal without comminution (see Fig. 1).

3.7 Postoperative Management

1. Animals should be monitored until awake and should be observed for their ability to freely ambulate over a 48-h period. 2. Analgesia is maintained with buprenorphine for 48 h at 12-h increments. Animals should be able to regain free mobility in 48 h, if not they should be euthanized (see Note 6). 3. Animals should be sacrificed for analysis of fracture repair at predetermined time points following fracture (see Note 7).

4

Notes 1. In general, unless an experiment is specifically designed to examine fracture healing in the context of juvenile development or aging, skeletally mature animals at the end of their juvenile growth period are used. In a prior study in which we examined the effects of Denosumab, male mice ranging in age

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from 11 to 18 weeks were used. For this study, we saw no differences in callus structure, composition, or mechanical properties associated for mice of varying ages within a test group. It should be noted however that in aged mice [24, 25] and rats [26, 27] that there are definable differences in the rate of healing that are affected by changes in the observable molecular mechanism that effect healing. Finally it is important to note that there is considerable sexual dimorphism in bone structure and strength [28, 29] such that only one sex should be used in a given study or comparisons between sexes should be planned as part of the experimental design. Recent National Institutes of Health Guidelines have proscribed that experimental comparisons should be made between sexes to account for she sexual dimorphic variations. Thus unless cost prohibitive and for justifiable scientific reasons most studies should be carried out in both sexes. https://orwh.od.nih. gov/sites/orwh/files/docs/NOT-OD-15-102_Guidance.pdf 2. The exact size of the pin should be empirically determined for each experiment by considering the ratio between pin diameter and intramedullary diameter. This ratio, along with the purchase of the pin in the proximal or distal metaphysis, and the stiffness of the pin material determine the rigidity of the fixation, which greatly affects the extent of the external callus formation. We have found that using a fully threaded K-wire in the rat model produces a very rigid construct which will change the extent to which endochondral bone formation takes place due to the much greater stabilization of the fixation. Pins of different materials have been used experimentally to model the effects of varying amounts of micromotion on bone healing [30]. In a more extreme case, not including a pin for fixation has also been used to increase the induction of periosteal endochondral bone formation. However, in the absence of any stabilization, the model can only be used for qualitative study of healing [11] because of the large degree of variability in the timing and quantities of new bone formation. 3. For rats, the hole is mechanically drilled. Rechargeable carpenter’s drills, a Dremel Tool® or dental handpiece with a drill attachment are all suitable. For mice, the hole can be generated by using the beveled end of a 23-gauge syringe needle with manual rotation. 4. For rat surgeries, pins are precut to an approximate length of the femur plus about 5 mm by sizing the length though palpitation of the bone through the muscle and visible inspection of

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the leg. For the mouse, pins are cut at the time of surgery when they are inserted. For C57B6 mice, the stylet of a 25-gauge spinal needle is used as the intramedullary pin. The pin is buried by twisting it either manually (mouse) or mechanically driving the pin into the underlying bone by affixing the pin to a drill (rat). 5. Occasional pin retraction is observed in cases in which the pin has not been fully buried and is tightly in place in the bone. In such cases the pin will be seen protruding through the skin at the knee. Such cases should be immediately euthanized since the fracture fixation will not be stable. 6. Use of nonsteroidal anti-inflammatory drugs (NSAIDs) should not be used as postoperative analgesics since they have been shown to inhibit bone healing after surgery [19, 31]. 7. Studies focusing on developmental processes related to endochondral bone formation may be purposely restricted to the early and intermediate periods of endochondral bone formation. On the other hand, if a study is directed at examining coupled remodeling then later periods will need to be examined. In healthy rats, the periods of endochondral formation through cartilage resorption can last until 28 days, while the period of coupled remodeling initiates around day 21 and lasts up until 12 weeks. In mice, the period of endochondral bone formation through resorption is 21 days, with the period of coupled remodeling initiating around day 14 days and lasting up until 8 weeks. Since specific experimental conditions can greatly alter the time-course of healing, pilot studies using several animals per group should be carried out for each new set of experimental conditions. For these studies, series of X-rays over a defined time-period can help determine the timeframe that should be experimentally examined. In studies examining therapeutics, end points should be chosen to appropriately assess regain of mechanical function and if a study is assessing therapeutic efficacy in the context of promoting healing, multiple time-points are needed to determine the rate of regain of mechanical strength. Because of the time-evolving nature of fracture healing, it is optimal to examine multiple time-points to capture times when key biological processes are taking place and to relate these processes to the regain in the functional properties of the callus.

Acknowledgments This work was supported by NIH grants AR056637 and AR062642.

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References 1. Salisbury-Palomares KT et al (2009) Mechanical stimulation alters tissue differentiation and molecular expression during bone healing. J Orthop Res 27:1123–1132 2. Miclau T et al (2007) Effects of delayed stabilization on fracture healing. J Orthop Res 25 (12):1552–1558 3. Lu C et al (2011) Mechanical stability affects angiogenesis during early fracture healing. J Orthop Trauma 25(8):494–499 4. Yu YY et al (2012) Creating rigidly stabilized fractures for assessing intramembranous ossification, distraction osteogenesis, or healing of critical sized defects. J Vis Exp 11:62 5. Bonnarens F, Einhorn T (1984) Production of a standard closed fracture in laboratory animal bone. J Ortho Res 2(1):97–101 6. Hiltunen A, Vuorio E, Aro H (1993) A standardized experimental fracture in the mouse tibia. J Ortho Res 11(2):305–312 7. Kon T et al (2001) Expression of osteoprotegerin, receptor activator of NF-kappaB ligand (osteoprotegerin ligand) and related proinflammatory cytokines during fracture healing. J Bone Miner Res 16(6):1004–1014 8. Gerstenfeld LC et al (2006) Three dimensional reconstruction of fracture callus morphogenesis demonstrates asymmetry in callus development. J Histochem Cytochem 54 (11):1215–1228 9. Marturano JE et al (2008) An improved murine femur fracture device for bone healing studies. J Biomech 41(6):1222–1228 10. Zhang X et al (2002) Cyclooxygenase-2regulates mesenchymal cell differentiation into the osteoblast lineage and is critically involved in bone repair. J Clin Invest 109 (11):1405–1415 11. Colnot C et al (2003) Altered fracture repair in the absence of MMP9. Development 130 (17):4123–4133 12. Tsuji K et al (2006) BMP2 activity, although dispensable for bone formation, is required for the initiation of fracture healing. Nat Genet 38 (12):1424–1429 13. Jepsen KJ et al (2008) Genetic variation in the patterns of skeletal progenitor cell differentiation and progression during endochondral bone formation affects the rate of fracture healing. J Bone Miner Res 23(8):1204–1216 14. Grimes R et al (2011) The transcriptome of fracture healing defines mechanisms of coordination of skeletal and vascular development during endochondral bone formation. J Bone Miner Res 26(11):2597–2609

15. Wigner NA et al (2010) Acute phosphate restriction leads to impaired fracture healing and resistance to BMP-2. J Bone Miner Res 25(4):724–733 16. Vortkamp A et al (1998) Recapitulation of signals regulating embryonic bone formation during postnatalgrowth and in fracture repair. Mech Dev 71:65–76 17. Ferguson C et al (1999) Does adult fracture repair recapitulate embryonic skeletal formation? Mech Dev 87:57–66 18. Gerstenfeld LC et al (2003) Fracture healing as a post-natal developmental process: molecular, spatial, and temporal aspects of its regulation. J Cell Biochem 88:873–884 19. Simon AM, Manigrasso MB, O’Connor JP (2002) Cyclo-oxygenase 2 function is essential for bone fracture healing. J Bone Miner Res 17 (6):963–976 20. Alkhiary YM et al (2005) Enhancement of experimental fracture-healing by systemic administration of recombinant human parathyroid hormone (PTH 1-34). J Bone Joint Surg Am 87(4):731–741 21. Kakar S et al (2007) Enhanced chondrogenesis and Wnt-signaling in parathyroid hormone treated fractures. J Bone Miner Res 22 (12):1903–1912 22. Gerstenfeld LC et al (2008) Comparison of bisphosphonate alendronate versus the RANKL inhibitor denosumab on murine fracture healing. J Bone Miner Res 24(2):196–208 23. Einhorn TA (2003) A single percutaneous injection of recombinant human bone morphogenetic protein-2 accelerates fracture repair. J Bone Joint Surg Am 85-A (8):1425–1423 24. Lu C et al (2005) Cellular basis for age-related changes in fracture repair. J Orthop Res 23 (6):1300–1307 25. Lu C et al (2008) Effect of age on vascularization during fracture repair. J Orthop Res 26 (10):1384–1389 26. Meyer J et al (2001) Age and ovariectomy impair both the normalization of mechanical properties and the accretion of mineral by the fracture callus in rats. J Ortho Res 19:428–435 27. Meyer RA Jr et al (2003) Gene expression in older rats with delayed union of femoral fractures. J Bone Joint Surg Am 85-A:1243–1254 28. Halloran BP et al (2002) Changes in bone structure and mass with advancing age in the male C57BL/6J mouse. J Bone Miner Res 17 (6):1044–1050

Mouse Model of Skeletal Repair 29. Glatt V et al (2007) Age-related changes in trabecular architecture differ in female and male C57BL/6J mice. J Bone Miner Res 22 (8):1197–1207 30. Willie B et al (2009) Mechanical characterization of external fixator stiffness for a rat femoral fracture model. J Orthop Res 27(5):687–693

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31. Gerstenfeld LC et al (2007) Selective and nonselective cyclooxygenase-2 inhibitors and experimental fracture-healing: reversibility of effects after short-term treatment. J Bone Joint Surg Am 89(1):114–125

Chapter 5 A Mouse Femoral Ostectomy Model to Assess Bone Graft Substitutes Ryan P. Trombetta, Emma K. Knapp, and Hani A. Awad Abstract The shortcomings of autografts and allografts in bone defect healing have prompted researchers to develop suitable alternatives. Numerous biomaterials have been developed as bone graft substitutes each with their own advantages and disadvantages. However, in order to test if these biomaterials provide an adequate replacement of the clinical standard, a clinically representative animal model is needed to test their efficacy. In this chapter, we describe a mouse model that establishes a critical sized defect in the mid-diaphysis of the femur to evaluate the performance of bone graft substitutes. This is achieved by performing a femoral ostectomy and stabilization utilizing a femoral plate and titanium screws. The resulting defect enables the bone regenerative potential of bone graft substitutes to be investigated. Lastly, we provide instruction on assessing the torsional strength of the healed femurs to quantitatively evaluate the degree of healing as a primary outcome measure. Key words Bone graft substitute, Critical sized defect, Mouse model, Scaffold, Healing, Bone, Biomechanics, Torsional testing

1

Introduction Bone grafting is a common and necessary procedure for healing bone defects that result from trauma, disease, cancer, or congenital anomalies. Due to the high demand for bone grafts, bone is the second most transplanted tissue [1]. In the United States, over half a million patients receive bone defect repairs with an estimated cost exceeding 2.5 billion US dollars [2]. The current gold standard for bone grafting procedures are autografts harvested from the patient’s own tissue typically from the iliac crest. Although autografts provide the osteoinduction, osteoconduction, and osteogenesis needed for bone regeneration, only a limited amount of tissue can be harvested from the patient, which results in donor site morbidity and severe blood loss [3, 4]. Because of these caveats, allografts are the clinical standard for large bone defects. However, prior to being implanted allografts are decellularized to abate the

Matthew J. Hilton (ed.), Skeletal Development and Repair: Methods and Protocols, Methods in Molecular Biology, vol. 2230, https://doi.org/10.1007/978-1-0716-1028-2_5, © Springer Science+Business Media, LLC, part of Springer Nature 2021

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host’s immune rejection of the graft and gamma irradiated to remove viruses, bacteria, and fungi [5, 6]. This renders the allograft a nonviable scaffold incapable of remodeling the damage accumulated during daily activities as living bone normally does. Thus, allogenic failure rates reported as high as 37% are attributed to the rigorous decellularization and sterilization process of allografts, which result in loss of osteogenic properties that limits the bone healing potential. Due to the various shortcomings of allogenic and autogenic bone transplantations, researchers have thoroughly investigated alternative biomaterials that could be used as bone graft substitutes. Various biomaterials have been utilized as bone graft substitutes. The primary objective of all these substitutes is to enable osseous healing with complete bone union restoring the biomechanical strength and function. The most commonly investigated bone graft alternatives are synthetic bone grafts. These consist of calcium salt derivatives (i.e., calcium sulfate [7] and calcium phosphate [8]), bioactive glass [9], synthetic (and natural) polymers [10], and composite grafts [11]. These, materials have been produced in multiple physical forms such as powders [12], pastes [13], cements [14], pellets [15], spacers [16], putties [17], and coatings [18]. These biomaterial formulations also share the key characteristic of being bioresorbable. The design of these scaffolds generally aims to have resorption rates comparable to new bone formation rates [19]. However, the major limiting factor is that synthetic bone grafts are solely osteoconductive, lacking osteoinductive and osteogenic properties. Hence the host tissue is required to initiate and orchestrate the bone healing and regeneration process. To circumvent this limitation, orthobiologics are added to the synthetic scaffolds in order to incorporate osteoinductive components to initiate and enhance bone regeneration [20]. These include growth factors such as bone morphogenetic proteins [21], vascular endothelial growth factors [22], insulin-like growth factors [23], fibroblast growth factors [24], and plateletderived growth factors [25]. Demineralized bone matrix (DBM) [26] and platelet-rich plasma (PRP) [27] can also be added to these biomaterials, which contain a multitude of undefined growth factors. Bone graft substitutes can also be seeded with bone forming cells, such as osteoblasts [28], mesenchymal stem cells [29], and osteoprogenitor cells [30] in order to enhance bone formation. An emerging technique currently being investigated is incorporation of engineered periosteum around bone graft substitutes in order to augment osteogenesis [31]. Stem cell sheet technology is utilized to create the thin outermost portion of bone consisting of MSCs and osteoprogenitor cells, which has been shown to be critical for bone healing [32]. Despite the various grafting materials and combinations of orthobiologics, there is minimum level 1 evidence supporting the use of bone graft substitutes over conventional

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grafting standards [20]. Therefore, the field of tissue engineering must continue to push the scientific development and evaluation of bone graft substitutes that promise to replace the clinical standards of allografts in preclinical models. Equally as important as the grafting material itself are the preclinical models used to investigate and assess the regeneration of bone in these critical defects. Clinically relevant animal models are essential for the preclinical evaluation of promising bone graft substitutes. Bone healing models encompass various modalities ranging from fracture to large critical sized defect reconstruction. Numerous species have been implemented, and each has their advantages and disadvantages. This variety of models provides researchers with numerous options for investigating the efficacy of their bone graft substitutes. Ultimately the researcher must choose a model that will provide the appropriate answers for their underlying research questions. With regard to bone graft substitutes, a critical sized defect (does not heal spontaneously) is ideal for testing the regenerative properties of bone grafts and their biomaterial substitutes. Critical sized defects are associated with high rates of complications and poor functional outcomes in the clinic [33]. In animal models, critical sized defects are achieved by one of three ways, (1) cylindrical metaphyseal defects (tibial plateau, femoral plug), (2) segmental mid-diaphysis defects, or (3) circular calvaria defects. Metaphyseal and calvaria defects provide inexpensive and easy surgical approaches due to the lack of required stabilization hardware. Metaphyseal and calvaria defects are not fully load bearing, while segmental mid-diaphysis defects are load bearing. Studies have shown that controlled loading positively affect bone healing [34]. Segmental mid-diaphysis defects are commonly associated with trauma and infection scenarios and require stabilization hardware that replicates clinical practice. The degree of load bearing is determined by both the choice of the stabilization hardware and biomaterial scaffold used as a bone graft substitute. These models are often performed in larger animals, such as canines or sheep, which can be expensive. However, due to advances in micromachining, small fracture-fixation hardware has been developed for use in rodents, including mice. Mice are easily handled, housed, and offer a cost-effective animal model allowing for high sample sizes. Furthermore, the mouse genome is easily manipulated enabling an abundance of precise disease models for investigation. However, due to their small size and limited bone volume researchers face challenges to rigor and reproducibility when performing invasive surgical procedures that are associated with segmental bone defects. This concern is eliminated with mouse-specific stabilization hardware, specialized surgical tools, and an established and repeatable surgical approach that we present herein to enable reproducible reconstruction of a critical-size femoral defect in a mouse.

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In this chapter, we describe the methods of establishing a critical-size segmental defect by performing an ostectomy in a murine femur stabilized by using a polyether ether ketone (PEEK) femoral plate and titanium screws. This model enables the study of bone graft substitutes made from various biomaterials to induce and orchestrate bone healing and regeneration in the defect as determined by torsional mechanical testing as the definitive standard outcome of healing in bone.

2 2.1

Material Presurgical Prep

1. Hair clipper. 2. Povidone prep solution and 70% isopropyl alcohol. 3. Ophthalmic ointment. 4. Surgical pad. 5. General surgical disposables consisting of face masks, sterile drapes, sterile gloves, and sterile gauze.

2.2 Anesthetic and Analgesic Drugs

1. Anesthesia drugs: 130 mg/kg ketamine hydrochloride and 12 mg/kg xylazine. Drugs are combined and administered as a single intraperitoneal (IP) injection. The following regimen will produce a surgical level of anesthesia lasting 45–60 min and sedation 2–3 h: combine 1.0 ml of 130 mg/ml ketamine hydrochloride with 1.0 ml of 12 mg/ml xylazine and 8.0 ml 1
PBS. The combined drugs are to be administered at 0.1 ml/ 10 g body weight via an IP injection using a 1 ml syringe with 25G 5/8 in. needle. 2. Analgesic drugs: 0.3 mg/ml buprenorphine hydrochloride (Buprenex®). 0.1 mg/kg Buprenex® is administered subcutaneously preoperatively and every 12 h up to 3 days postoperatively (see Note 1). Subcutaneous injections are given using a 1 ml syringe with a 25–30 G 5/8 in. needle.

2.3 Femoral Ostectomy

1. RISystems MouseFix plate 6-hole, polyether ether ketone (PEEK) (see Note 2). 2. RISystems MouseFix screw, L ¼ 2.00 mm. 3. RISystems MouseFix saw guide (2–3 mm) (see Note 3). 4. RISystems drill bit 0.30 mm. 5. RISystems Hand Drill. 6. Saw guide mount (Fig. 1a, b, see Note 4). 7. Dremel. 8. Forceps. 9. Surgical scissors.

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Fig. 1 Photograph of the saw guide mount

10. Curved forceps. 11. Hemostatic forceps. 12. Gigli wire saw 0.22 mm. 13. 5-0 monofilament sutures. 14. Sterile drapes. 15. No. 10 scalpel. 16. LX-60 X-ray cabinet (Faxitron Bioptics© or equivalent). 2.4 Sample Prep for Biomechanical Torsion Testing

1. Harvested femurs, cleaned of all soft tissue.

2.5 Set-up Prep for Biomechanical Torsion Testing

1. Square aluminum tubing (outer dimension of 1/400 , cut to 3/400 lengths).

2. Scalpel. 3. Forceps.

2. Plumber’s putty. 3. Plastic alignment jig—with 6 mm gauge length opening. 4. Alignment release bars. 5. Aluminum compacting bar.

2.6 Alignment Fixation for Biomechanical Torsion Testing

1. 5 ml syringes. 2. Autopolymerizable bone cement mix (acrylic powder and liquid). 3. Wooden tongue depressors. 4. Graduated cylinder.

2.7 Sample Rehydration for Biomechanical Torsion Testing

1. 1
phosphate buffer solution. 2. Petri dish.

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2.8 Biomechanical Torsion Testing

1. EnduraTec Test Bench™, with 200N.mm torque cell (Bose Corp. Minnetonka, MN). 2. WinTest7 software.

3

Methods Please note that all personnel performing this protocol should be familiar with their institutional animal welfare policies as well as the anatomy of the mouse’s hind limb. Adherence to guidelines such as the ARRIVE (Animal Research: Reporting In Vivo Experiments) Guidelines Checklist is highly recommended.

3.1 Surgical Preparation of Mouse Hind Limb

1. Anesthetize the mouse as describe in Subheading 2.2, item 1. This will provide a surgical plane of anesthesia that will last for approximately 45–60 min, which is enough time for an experienced rodent surgeon to complete this procedure. The average time for this procedure is 25–30 min. 2. Administer pain management drugs preoperatively as per Subheading 2.2, item 2, with a subcutaneous injection of buprenorphine. 3. Place the mouse on the surgical pad. Using the clipper, shave the hair on the posterior surface of the hind limb spanning from the distal end of the tibia to halfway up the spine and just beyond the medial plane of the spine. Shave any remaining hair on the anterior surface of the hind limb (Fig. 2a). 4. Using a gauze pad or Q-tip applicator, prep the posterior surface on the hind leg with alternating scrubs of povidone– iodine and 70% isopropanol. Repeat this procedure three times. 5. Prepare a sterile drape and organize sterile instruments. 6. Position the mouse in the prone position (Fig. 2a).

3.2 Femoral Ostectomy

1. Using sterile gloves and aseptic technique, create a longitudinal incision in line with the femur spanning from the hip joint to the knee using surgical scissors. To initiate the cut, pinch excess skin at the hip with teethed forceps to make initial cut. Use minimum number of cuts to establish surgical wound. 2. Perform a blunt dissection to separate the vastus lateralis and biceps femoris muscles to expose the full length of the femur while preserving the sciatic nerve. To perform the blunt dissection, stretch exposed tissue with forceps downward to look for a gap between the lateralis and biceps femoris muscles (Fig. 2b). Insert closed surgical scissors and open in order to separate the two muscles (Fig. 2c). Repeat this action multiple times if necessary to expose the femur.

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Fig. 2 Procedure of the femoral ostectomy

Fig. 3 Planar X-ray image post-surgery of an implanted calcium phosphate bone graft substitute in a 3 mm defect

3. Uncover and skeletonize the femur using forceps. Then use curved forceps to perform a circular preparation of the femur at the transition from the middle third to the distal third of the femur (Fig. 2d). 4. Use the forceps with teeth to remove any soft tissue attached to the femur by lightly grasping the soft tissue and pulling away

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from the center of the bone. This must be performed very carefully as to not damage the periosteum (outermost vascular connective tissue of the bone), which plays a critical role in bone healing. Further expose the length of the femur by using the surgical scissors to cut any attached soft tissue at the proximal and distal ends of the femur. 5. Place the six-hole PEEK plate to the anterolateral surface of the femur (see Note 5). Temporarily suture the PEEK plate to femur via the middle notch (Fig. 2e). As a visual reference to align the plate at the center of the femur, the second most proximal hole is adjacent to the third trochanter. 6. Ensure that the plate is centered and use the hemostatic forceps in the nondominant hand to grip the plate at the middle and rotate away from the mouse (toward the surgeon). 7. Use the drill bit inserted into the Dremel to drill the most distal hole (Fig. 2f). During the act of drilling, the surgeon should feel two gives corresponding to the drill bit penetrating through each cortex of the femur. When drilling it is important to make sure the drill is perpendicular to the bone in order to ensure stability and avoid fracture. 8. Use the hand drill to insert a Titanium screw perpendicular to the bone. Tighten slowly until the screw head is flush with the plate (this is the part that sits right on top of the threads for the plate). When finished, shear off the pin by rotating away from the center of the plate in a clockwise manner (Fig. 2g). 9. Repeat this process for the most proximal and then the second most proximal screws. 10. Insert the final screw into the second most distal hole. Do not shear off the pin, leave intact for the saw guide (Fig. 2h). 11. Remove the sutures with either surgical scissors or a No. 10 scalpel. 12. Install the saw guide by aligning with PEEK plate and insert the intact screw pin into the distal most screw hole of the saw guide. Grasp the intact screw pin with the forceps and push the saw guide down until it is flush with the plate (see Note 6). 13. Clamp the saw guide in place with the hemostatic forceps (Fig. 2i). Use only the first locking point to prevent bone fracture. 14. Move the mouse by gripping the curved forceps and saw guide to the saw guide mount and lock into place (Fig. 2j). 15. Use approximately 8 in. of the Gigli saw wire to perform the osteotomy. Run the Gigli saw wire underneath the bone and align to saw guide’s slots using two forceps. Using a narrow grip, slowly initiate the cut with short strokes. Gradually widen

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your grip and perform longer strokes until the bone is completely cut (see Note 7). 16. Unlatch the clamp and remove the mouse. Unclamp the hemostats and remove the saw guide. Check to ensure the bone was cleanly cut at both ends through both cortices. Remove the bone with forceps (Fig. 2k). 17. Shear off the intact screw pin by reinserting hand drill and grasping the exposed plate with the hemostatic forceps. 18. Suture proximal and distal ends of the femur using a 5-0 nylon monofilament suture in order to prevent screw pull-out (see Note 8). 19. Place gauze pad underneath the mouse and spray the defect area with sterile 1
PBS from a syringe in order to irrigate the surgical wound and remove any bone debris. 20. Insert any grafting or scaffolding material into the defect using forceps. Secure the graft of scaffold in a cerclage fashion using a 6-0 nylon braided suture and cut carefully with surgical scissors (Fig. 2l). 21. Remove the curved forceps. 22. Suture the muscle with two to four 5-0 sutures ensuring that surgical wound is closed. 23. Close the skin with 5-0 sutures. 24. Perform a radiographic evaluation of the mouse’s hind limb to assess the placement and success of the surgical hardware and bone graft substitute (Fig. 3). Acquire planar X-ray images using a LX-60 X-ray cabinet or similar device. Scans are performed with an energy of 55 kV, intensity of 145 μA, and 300 ms integration time (see Note 9). 25. Place mouse on a heated pad for anesthesia recovery. 3.3 Sample Prep for Biomechanical Torsion Testing

1. At the end of the study (see Note 10), euthanize the mice and harvest the femur ensuring the proximal and distal ends are intact. 2. Remove all soft tissue and surgical hardware from the harvested femur. If necessary, some soft tissue can be kept around the implanted graft/callus to ensure that it remains stable (see Note 11).

3.4 Set-up for Biomechanical Torsion Testing

1. For each femur, two square aluminum tubes, one plastic alignment jig and one alignment release bar will be used. Label each tube (Fig. 4a). 2. Fill one tube completely full of plumber’s putty. Fill the other tube half way with plumber’s putty. 3. Insert the release bar into the alignment jig (Fig. 4b).

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Fig. 4 Schematic depicting the alignment jig used to line up the harvested femur in the aluminum tubing with putty and bone cement

4. Insert the proximal end of the bone into the tubing that is filled with putty. Using forceps, orient the bone, so the mid-diaphysis is centered and in line with the tubing (Fig. 4c). 5. Place the tube with the bone into the plastic alignment jig, so the edge of the tube with the bone protruding is flush with the inner edge of the alignment jig. 3.5 Alignment Fixation for Biomechanical Torsion Testing

1. Mix a small batch of autopolymerizable bone cement (see Note 12). 2. Pour the bone cement into the tube that is half filled with plumber’s putty. Make sure that bone cement is filled to the top of the tube. 3. Insert the bottom tube, so that the distal end of the bone is submerged in wet cement. Slide the tube up, so the edge of the tube is flush with the inner edge of the alignment jig. Set the jig upright, to allow the bone cement to harden (Fig. 4d). This will create an exposed bone gauge length of 6 mm. If any excess bone cement gets on the exposed gauge length, gently scrap off the bone cement before it hardens, being careful to keep the bone aligned.

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4. Allow the bone cement to harden for 15–20 min. 5. Using the alignment release bar, remove the specimen and tubes from the alignment jig. The distal end of the bone should be fixed within the tube at this point. 6. Remove the proximal end of the bone from the tube and use the compacting bar to remove half of the putty, leaving the tube half full. 7. Reinsert the alignment release bar into the alignment jig, and insert the specimen, so the tube with the distal end cemented is now flush with the inner edge of the alignment jig. 8. Repeat steps 1–5 to cement the proximal end of the bone in the tube. 9. Once the sample has both the proximal and distal end firmly fixed in bone cement, place the sample in a petri dish with 1
phosphate buffer solution for at least 2 h, to rehydrate the sample. 3.6 Biomechanical Torsion Testing

1. Turn on the EnduraTec Test Bench™ and open the WinTest7 software. 2. The samples are inserted into the clamps, and four screws are used to secure the clamps. Custom fabricated clamps firmly grip and anchor the square tubes (Fig. 5a–c). 3. Initialize the program, with rotation and torque being recorded. Continue the test until the femur is completely fractured. 4. Test the samples at a rotation speed of 1 /s and a data collection rate of 20 samples/s. 5. The WinTest7 software produces a .txt file for each sample tested. This .txt file is then imported into Excel to determine the torsional rigidity, maximum torque and rotation at maximum torque of each sample.

4

Notes 1. As a substitution for 12 h doses of 0.1 mg/kg buprenorphine, extended release buprenorphine can be adminstered once preopertatively at a dose of 3.25 mg/kg subcutaneously. 2. A PEEK plate is radiolucent and will not be visible on radiographic imaging, such as X-rays and micro-computed tomography (μCT). This was chosen for instances where μCT is performed as an added outcome measure to quantify bone regeneration because metal plates will produce metallic artifiacts and noise making quantification difficult.

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Fig. 5 Custom machined clamps fabricated to clasp the square tubes containing the potted femur

3. Various saw guide sizes can be purchased or fabricated. For a mouse critical-sized femoral defect, a defect size of 2 mm is require. 4. The saw guide mount was created in-house using a low-profile hold-down toggle clamp seated at the end of a cantilever arm attached to a weighted base. 5. Installing the first screw is crucial because at this moment the alignment of the plate is determined. Pay extra attention to aligning the surface prior to implantation. Adjust the oritentation of the long axis so that the plate is sitting flush on the anterolateral surface of the femur and is centered. 6. Rotating the bone can help align the saw guide, if the plate is sitting perfectly in the middle of the bone the saw guide will fit

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flushly with the plate. Otherwise, attempt to adjust orientation of the saw guide so that is fits over the plate and bone. 7. It is important to ensure the bone is cut all the way through for both slots. The surgeon should be able to feel the difference in friction between cutting the bone verse cutting plate indicating the bone is completely cut. If bone was not completely cut and the ostectomy was incomplete, the bone cannot be removed. If this occurs install the saw guide and continue cutting with the gigli saw until the osteoctomy is complete. On the contrary, cutting too much can damage the plate and this must be avoided to ensure stabilization of the femur. 8. This step is not required but is performed as a safety precaution. 9. When assessing quality of surgery, examine the position of the defect and screws to ensure the defect is centered in the femur. Confirm that the bone graft subsitute is sucessfully implanted in the defect. Also examine if any bone fragments or fractures resulted from the installation of the titanium screws and ostectomy. 10. Study duration is determined by the time needed to heal the defect. Larger defects will require a longer healing time. A minimum of 12 weeks is suggested as the time required to heal a 2 mm critical sized femoral defect. 11. When potting, the bone cement will only adhere to exposed bone, and excess soft tissue can cause an air pocket to form around the ends of the bone when curing, comprising the structural integrity of the fixation. 12. The brand used was Bosworth Fastray. Approximately 10 g of powder and 3 ml of liquid were used to make the bone cement.

Acknowledgments This work was supported by the NIAMS/NIH grants P30AR069655 and P50AR072000, and the AOTrauma Clinical Priority Program. References 1. Giannoudis PV, Dinopoulos H, Tsiridis E (2005) Bone substitutes: an update. Injury 36 (Suppl 3):S20–S27. https://doi.org/10. 1016/j.injury.2005.07.029 2. Amini AR, Laurencin CT, Nukavarapu SP (2012) Bone tissue engineering: recent advances and challenges. Crit Rev Biomed Eng 40(5):363–408

3. Laurencin CT, Khan Y (2009) Bone graft substitute materials. Expert Rev Med Devices 3 (1):49–57 4. Tan MH, Mankin HJ (1997) Blood transfusion and bone allografts. Effect on infection and outcome. Clin Orthop Relat Res 340:207–214. https://doi.org/10.1097/ 00003086-199707000-00027

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5. Nguyen H, Morgan DA, Forwood MR (2007) Sterilization of allograft bone: effects of gamma irradiation on allograft biology and biomechanics. Cell Tissue Bank 8(2):93–105. https://doi.org/10.1007/s10561-006-90201 6. Brigman BE, Hornicek FJ, Gebhardt MC, Mankin HJ (2004) Allografts about the knee in young patients with high-grade sarcoma. Clin Orthop Relat Res 421:232–239. https:// doi.org/10.1097/01.blo.0000127132. 12576.05 7. Beuerlein MJ, McKee MD (2010) Calcium sulfates: what is the evidence? J Orthop Trauma 24(Suppl 1):S46–S51. https://doi.org/10. 1097/BOT.0b013e3181cec48e 8. Bohner M, Galea L, Doebelin N (2012) Calcium phosphate bone graft substitutes: failures and hopes. J Eur Ceram Soc 32 (11):2663–2671. https://doi.org/10.1016/j. jeurceramsoc.2012.02.028 9. Fu Q, Saiz E, Rahaman MN, Tomsia AP (2011) Bioactive glass scaffolds for bone tissue engineering: state of the art and future perspectives. Mater Sci Eng C Mater Biol Appl 31 (7):1245–1256. https://doi.org/10.1016/j. msec.2011.04.022 10. Sabir MI, Xu X, Li LI (2009) A review on biodegradable polymeric materials for bone tissue engineering applications. J Material Sci 44 (21):5713–5724. https://doi.org/10.1007/ s10853-009-3770-7 11. Murugan R, Ramakrishna S (2005) Development of nanocomposites for bone grafting. Compos Sci Technol 65(15):2385–2406. https://doi.org/10.1016/j.compscitech. 2005.07.022 12. Whiteman D, Gropper PT, Wirtz P, Monk P (1993) Demineralized bone powder. Clinical applications for bone defects of the hand. J Hand Surg 18(4):487–490 13. Johal HS, Buckley RE, Le IL, Leighton RK (2009) A prospective randomized controlled trial of a bioresorbable calcium phosphate paste (alpha-BSM) in treatment of displaced intra-articular calcaneal fractures. J Trauma 67 (4):875–882. https://doi.org/10.1097/TA. 0b013e3181ae2d50 14. Zhang J, Liu W, Schnitzler V, Tancret F, Bouler J-M (2014) Calcium phosphate cements for bone substitution: chemistry, handling and mechanical properties. Acta Biomater 10 (3):1035–1049. https://doi.org/10.1016/j. actbio.2013.11.001 15. Urban RM, Turner TM, Hall DJ, Infanger SI, Cheema N, Lim TH, Moseley J, Carroll M, Roark M (2004) Effects of altered crystalline

structure and increased initial compressive strength of calcium sulfate bone graft substitute pellets on new bone formation. Orthopedics 27(1 Suppl):s113–s118 16. Joosten U, Joist A, Gosheger G, Liljenqvist U, Brandt B, von Eiff C (2005) Effectiveness of hydroxyapatite-vancomycin bone cement in the treatment of Staphylococcus aureus induced chronic osteomyelitis. Biomaterials 26(25):5251–5258. https://doi.org/10. 1016/j.biomaterials.2005.01.001 17. Bohner M (2010) Design of ceramic-based cements and putties for bone graft substitution. Eur Cell Mater 20:1–12 18. Daculsi G (1998) Biphasic calcium phosphate concept applied to artificial bone, implant coating and injectable bone substitute. Biomaterials 19(16):1473–1478 19. Petite H, Viateau V, Bensaid W, Meunier A, de Pollak C, Bourguignon M, Oudina K, Sedel L, Guillemin G (2000) Tissue-engineered bone regeneration. Nat Biotechnol 18(9):959–963. https://doi.org/10.1038/79449 20. Peterson JR, Chen F, Nwankwo E, Dekker TJ, Adams SB (2019) The use of bone grafts, bone graft substitutes, and orthobiologics for osseous healing in foot and ankle. Surgery 4 (3):2473011419849019. https://doi.org/10. 1177/2473011419849019 21. Carragee EJ, Hurwitz EL, Weiner BK (2011) A critical review of recombinant human bone morphogenetic protein-2 trials in spinal surgery: emerging safety concerns and lessons learned. Spine J 11(6):471–491. https://doi. org/10.1016/j.spinee.2011.04.023 22. Street J, Bao M, deGuzman L, Bunting S, Peale FV Jr, Ferrara N, Steinmetz H, Hoeffel J, Cleland JL, Daugherty A, van Bruggen N, Redmond HP, Carano RAD, Filvaroff EH (2002) Vascular endothelial growth factor stimulates bone repair by promoting angiogenesis and bone turnover. Proc Natl Acad Sci U S A 99 (15):9656–9661. https://doi.org/10.1073/ pnas.152324099 23. Meinel L, Zoidis E, Zapf J, Hassa P, Hottiger MO, Auer JA, Schneider R, Gander B, Luginbuehl V, Bettschart-Wolfisberger R, Illi OE, Merkle HP, Rechenberg BV (2003) Localized insulin-like growth factor I delivery to enhance new bone formation. Bone 33 (4):660–672. https://doi.org/10.1016/ S8756-3282(03)00207-2 24. Kawaguchi H, Nakamura K, Tabata Y, Ikada Y, Aoyama I, Anzai J, Nakamura T, Hiyama Y, Tamura M (2001) Acceleration of fracture healing in nonhuman primates by fibroblast growth factor-2. J Clin Endocrinol Metab 86

Mouse Femoral Ostectomy Model to Asses Bone Graft Substitutes (2):875–880. https://doi.org/10.1210/jcem. 86.2.7199 25. Marx RE, Carlson ER, Eichstaedt RM, Schimmele SR, Strauss JE, Georgeff KR (1998) Platelet-rich plasma: growth factor enhancement for bone grafts. Oral Surg Oral Med Oral Pathol Oral Radiol Endod 85 (6):638–646 26. Wildemann B, Kadow-Romacker A, Haas NP, Schmidmaier G (2007) Quantification of various growth factors in different demineralized bone matrix preparations. J Biomed Mater Res A 81(2):437–442. https://doi.org/10.1002/ jbm.a.31085 27. Freymiller EG (2004) Platelet-rich plasma: evidence to support its use. J Oral Maxillofac Surg 62(8):1046. Author reply 1047-1048 28. Ishaug SL, Crane GM, Miller MJ, Yasko AW, Yaszemski MJ, Mikos AG (1997) Bone formation by three-dimensional stromal osteoblast culture in biodegradable polymer scaffolds. J Biomed Mater Res 36(1):17–28. https://doi. org/10.1002/(sici)1097-4636(199707) 36:13.0.Co;2-o 29. Wei X, Liu B, Liu G, Yang F, Cao F, Dou X, Yu W, Wang B, Zheng G, Cheng L, Ma Z, Zhang Y, Yang J, Wang Z, Li J, Cui D, Wang W, Xie H, Li L, Zhang F, Lineaweaver WC, Zhao D (2019) Mesenchymal stem cellloaded porous tantalum integrated with biomimetic 3D collagen-based scaffold to repair large osteochondral defects in goats. Stem

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Cell Res Ther 10(1):72. https://doi.org/10. 1186/s13287-019-1176-2 30. Schliephake H, Knebel JW, Aufderheide M, Tauscher M (2001) Use of cultivated osteoprogenitor cells to increase bone formation in segmental mandibular defects: an experimental pilot study in sheep. Int J Oral Maxillofac Surg 30(6):531–537. https://doi.org/10. 1054/ijom.2001.0164 31. Long T, Zhu Z, Awad HA, Schwarz EM, Hilton MJ, Dong Y (2014) The effect of mesenchymal stem cell sheets on structural allograft healing of critical sized femoral defects in mice. Biomaterials 35(9):2752–2759. https://doi. org/10.1016/j.biomaterials.2013.12.039 32. Zhang X, Xie C, Lin AS, Ito H, Awad H, Lieberman JR, Rubery PT, Schwarz EM, O’Keefe RJ, Guldberg RE (2005) Periosteal progenitor cell fate in segmental cortical bone graft transplantations: implications for functional tissue engineering. J Bone Miner Res 20 (12):2124–2137. https://doi.org/10.1359/ JBMR.050806 33. Schemitsch EH (2017) Size matters: defining critical in bone defect size! J Orthop Trauma 31:S20–S22. https://doi.org/10.1097/bot. 0000000000000978 34. Claes LE, Heigele CA, Neidlinger-Wilke C, Kaspar D, Seidl W, Margevicius KJ, Augat P (1998) Effects of mechanical factors on the fracture healing process. Clin Orthop Relat Res (355 Suppl):S132–S147. https://doi. org/10.1097/00003086-199810001-00015

Chapter 6 Surgical Induction of Posttraumatic Osteoarthritis in the Mouse Robert D. Maynard, David A. Villani, William G. Schroeder, Douglas J. Adams, and Michael J. Zuscik Abstract Given the prevalence and the scope of the personal and societal burden of osteoarthritis (OA), investigators continue to be deeply interested in understanding the pathogenic basis of disease and developing novel disease modifying OA therapies. Because joint trauma/injury is considered a leading predisposing factor in the development of OA, and since posttraumatic OA is one of the most common forms of OA in general, large animal and rodent models of knee injury that accurately recapitulate the OA disease process have become increasingly widespread over the past decade. To enable study in the context of defined genetic backgrounds, investigative teams have developed standardized protocols for injuring the mouse knee that aim to induce a reproducible degenerative process both in terms of severity and temporal pacing of disease progression. The destabilization of the medial meniscus (DMM) is one of the most commonly employed surgical procedure in rodents that reproducibly models posttraumatic OA and allows for the study of disease progression from initiation to end-stage disease. The description provided here sets the stage for both inexperienced and established investigators to employ the DMM procedure, or other similar surgical destabilization methods, to initiate the development of posttraumatic OA in the mouse. Successful application of this method provides a preclinical platform to study the mechanisms driving the pathogenesis of posttraumatic OA and for testing therapeutic strategies to treat it. Key words Osteoarthritis, Knee, Trauma, Destabilization of medial meniscus (DMM), Surgery, Meniscus, Patellar tendon, Articular cartilage

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Introduction Osteoarthritis (OA) is the most common form of arthritis, with forecasts indicating that 25.9% of the adult US population (>18 years old), or nearly 78.4 million people, will have physician-diagnosed disease by 2040 [1]. OA is a degenerative joint disease characterized by dysfunction of articular chondrocytes, articular cartilage degradation involving fibrillation and clefting of the matrix, osteophyte formation, and subchondral sclerosis (reviewed in [2]). Despite the major health crisis that is rooted in the increasing population of afflicted individuals, only recently have

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there been advances in understanding the seminal molecular, cellular, and tissue events that drive these hallmarks of joint disease. Recent advances have been mainly driven by work in animal models of disease, which range from injury-induced models in large animals and rodents to genetic models in mice. While the pathogenic basis of OA is complex and includes genetic factors [3–5] epigenetic factors [6], obesity and metabolic dysfunction [7–9], and aging [10], joint trauma is widely accepted to be a central pathogenic mechanism of disease [11], with an estimated sixfold increase in the risk of developing radiographic OA of the knee within 21 years postinjury/surgery [12]. Based on this, study of the OA disease process has involved the use of animal models of joint trauma because of their clinical and translational relevance. The most popular posttrauma models of OA involve disruption of structures of the knee joint (menisci and ligaments), with early studies involving large animal models including rabbit [13], dog [14], and sheep [15]. Study of posttraumatic OA (PTOA) in rodent models led to the subsequent development of connective tissue injuries to the knee joint of the rat [16], setting the stage for pursuit of murine models which provide the advantage of studying the progression of trauma-induced OA on defined genetic backgrounds. For example, joint degeneration initiated via surgical detachment of ligaments connecting the menisci to the tibial plateau can be accelerated based on which additional structures are disrupted. Kamekura et al. summarize the injury– disease relationship in a number of mouse strains in a widely cited publication [17]. Several different types of knee joint injury have been employed in the mouse to initiate OA-like cartilage degeneration and associated joint changes, with the severity of the injury (i.e. the number of disrupted structures) dictating the pace of disease progression [17]. Destabilization of the medial meniscus (DMM), in which the medial meniscotibial ligament (MMTL) is transected to induce mechanical stress at the articulating surfaces in the knee joint, is a widely used surgical procedure to induce PTOA in rodents [18]. Detachment of the MMTL leads to the reproducible emergence of OA over a 2–6 month period [19], with mid-stage degeneration at 3 months (Fig. 1), allowing for the study of the progression of PTOA on defined genetic backgrounds. The DMM model was initially employed to evaluate the ability of Adamts4 and/or Adamts5 gene deletion in mice to protect against the development of cartilage degeneration. Based on these studies, it was established that deletion of Adamts5 [20] or both Adamts4 and 5 [21] significantly slows/prevents the progression of articular cartilage degeneration following injury. Using this DMM injury model, deletion of Mmp13 renders mice resistant to cartilage erosion posttrauma [22]. It has also been shown that haploinsufficiency of Runx2 protects against the development of OA-like

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Fig. 1 DMM induces progressive degeneration of articular cartilage. Compared to sham-operated joints at 3 weeks post-surgery (3 week, Sham), joints from mice administered DMM showed initial stages of cartilage degeneration that further progressed to mid-stage disease by 12 weeks, including loss of proteoglycan staining (red stain) and reduced thickness of the superficial zone cartilage. The tidemark is represented by a dashed yellow line. Evidence of fibrillation and clefting is denoted with black arrows. Presented mid-sagital sections of the medial compartment stained with Safranin O/Fast green histology are representative of groups of 5 or more mice for each of the time points. F femur, M meniscus, T tibia

degeneration in mice administered injuries to the meniscus and medial collateral ligament [23]. Since these early studies employed the DMM injury, the use of this model has become mainstream, with >290 published studies utilizing this model to study OA pathogenesis and to develop therapeutic paradigms. This literature was identified based on a PubMed search for “destabilization of the medial meniscus” or “DMM” and “osteoarthritis.” Other mouse models of PTOA include anterior cruciate ligament transaction (ACLT) with or without partial or complete meniscectomy, meniscal-ligamentous injury (MLI), and noninvasive models using applied external forces to the knee joint. The ACLT model was introduced in dogs [24] and has since been utilized in mouse models of PTOA [17, 18]. Transection of the ACL results in joint instability and increased mechanical load to the posterior compartment of the knee joint, resulting in the onset of moderate to severe OA. One study reported that ACLT led to cartilage destruction and damage to the subchondral bone as deep as the growth plate in about one third of mice by 8 weeks post-surgery [18] and resulted in a more severe phenotype than the DMM surgery. The meniscal ligamentous injury (MLI) model involves transection of the medial collateral ligament (MCL) in addition to partial meniscectomy. This leads to a more rapid progression of OA (terminal disease/eburnation within 4 months) [25], thus shortening the experimental time frame required to follow the development of pathology or the efficacy of a therapeutic intervention. However, this accelerated disease progression comes with an increased risk of variability between mice, and the surgical procedure itself can be more challenging. Additionally,

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inflammation from increased tissue damage due to surgery and the observation of osteophyte formation is more common with this procedure and can obfuscate the assessment of cartilage damage alone. A tibial compression model is currently in use as a noninvasive, nonsurgical method to induce posttraumatic OA that simulates human ACL rupture [26–28]. In this model, an axial compressive load is applied to the mouse knee to the point of overload leading to rupture of the ACL and degeneration of articular cartilage by 8 weeks postinjury [26, 27]. It is reported that this model also leads to inflammation at the injury site, synovitis, and a substantial loss of trabecular bone at the femoral and tibial epiphyses initially followed by partial recovery of bone volume (BV/TV) [26–28]. Because objective evaluation of the literature suggests that DMM is the most widely employed model for PTOA in rodents, in this chapter we fully describe the DMM procedure; the general surgical approach described here applies to any combination of meniscus injury and ligament disruption in the mouse, with the main surgical differences specifically involving accessing the targeted structure in a manner that protects against injury to other adjacent tissues.

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Materials An array of the key supplies and instrumentation required to perform the DMM procedure in the mouse is shown in Fig. 2. Based on the anticipated number of experimental mice used in a given surgical session, the investigative team should be sure an adequate supply of consumables have been gathered. Also, obtain an adequate number of fresh cages with fresh litter, food, and water to provide a clean environment for the mice in the immediate postoperative period and when the mice are returned to their housing rooms in the vivarium. 1. C57BL/6 J male mice (or similar mice with a defined genetic mutation), 12+ weeks of age (see Notes 1 and 2). 2. Isoflurane solution and the appropriate, certified delivery equipment (and murine nose cone) and scavenging equipment (see Note 3). 3. Light source with fiber optic cable to allow directing and positioning of the light beam. 4. Dissecting microscope (see Note 4). 5. Small animal hair clippers. 6. Student Vannas Spring microdissection scissors. 7. Dumont #5 fine forceps.

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Fig. 2 Supplies and instruments. For the most efficient performance of a series of DMM procedures on a cohort of mice, the depicted supplies and equipment are suggested and should be collected prior to initiating animal work. The presurgical punch list includes: (a) antiseptic scrub (e.g., povidone–iodine), (b) light source with fiber optic cable (or ring light) to allow directing and positioning of the light beam, (c) dissecting microscope (Leica shown), (d) isoflurane anesthesia (and the equipment necessary for the delivery and scavenging of waste gas), (e) sterile gauze, (f) 5-0 nylon suture, (g) sterile absorbent surgical pads, (h) sterile drapes, (i) hair clippers, (j) 25-gauge needles, (k) 1 ml syringes, (l) Student Vannas Spring microdissection scissors, (m) Dumont #5 fine forceps, (n) general small surgical scissors, (o) general small hemostat, (p) #11 scalpel, and (q) #10 scalpel

8. General small surgical scissors. 9. General small hemostat. 10. Sterile absorbent surgical pads. 11. Sterile gauze. 12. Sterile drapes. 13. #10 and #11 scalpels. 14. 25-gauge needles. 15. 1 ml syringes. 16. 5-0 nylon suturing kits. 17. Antiseptic scrub (e.g. povidone–iodine).

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Methods As with any animal surgical procedure, appropriate institutional approval of the experimental and operative protocol will be required, and a clean surgical suite/area (either in the vivarium or laboratory) needs to be established. The procedure is best carried out with two individuals, one performing the surgery and an assistant to manage anesthesia, to pass instruments as necessary, to assist with wound closure, to apply analgesia (typically subcutaneously administered buprenorphine delivered perioperatively), and to monitor the mice as they recover during the postanesthesia period (see Note 5). To minimize the potential of infection, the surgeon and the assistant should wear surgical garb, a hair net, surgical mask, and sterile gloves. When multiple procedures are to be performed serially, regular changing into fresh sterile gloves and mask is recommended. Regarding the DMM procedure specifically, following the surgery, mice are maintained for periods up 6 months, with experimental time points dictating euthanasia via induction of carbon dioxide narcosis followed by cervical dislocation. Typical time points for tissue harvest following DMM include 3, 4, 8, 12, 16, 20, and 24 weeks with 12 weeks representing the disease midpoint (loss of 50% of tibial plateau cartilage) (Fig. 1) [25]. After sterilizing the location where the surgery will be performed using an antibacterial/antiviral wash solution, gather all of the supplies and equipment required and bring the animals (in their housing units) to a holding area adjacent to where the procedure will be performed. Once everything is gathered and the surgical team is garbed-up, the following steps are executed: 1. Anesthesia/Analgesia: Administer the anesthesia of choice and ensure that a surgical plane is achieved (see Note 3). Following confirmation of plane via absence of the plantar/toe-pinch reflex, immediately deliver the first dose of buprenorphine (0.05 mg/kg, subcutaneous injection) for pain management. Note that this dose of buprenorphine is delivered every 12 h for the first 72 h postsurgery. If available, slow-release buprenorphine can be administered as a single dose that provides pain relief for the duration of surgical recovery. 2. Surgical Site Preparation: Using veterinary hair clippers, remove all hair from both knees (assuming either bilateral administration of DMM, or use of one knee for the DMM and the contralateral knee as a sham control, see Note 6). Using povidone–iodine, scrub the sheared surgical site, and apply a surgical drape to window off the surgical site(s). 3. Incision and Displacement of Patellar Tendon: Using the #10 or #11 scalpel blade, create a 5–10 mm longitudinal incision along the medial side of the knee to expose the knee joint and

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Fig. 3 Identifying joint structures following the skin incision. (a) Following application of a longitudinal skin incision medial to the knee joint (not shown, skin removed for illustration purposes), the patellar tendon, patella, and joint capsule are visible. (b) Based on visualization of the patellar tendon and the patella, the location of other key structures can be approximated. (c) A second longitudinal incision is performed medial to the patellar tendon to open the joint space and expose the medial meniscus and medial meniscotibial ligament. F femur, T tibia, LM lateral meniscus, MM medial meniscus, ACL anterior cruciate ligament, MMTL medial meniscotibial ligament

underlying tissues (Fig. 3a, b). Make a second longitudinal incision through the knee joint capsule directly beneath the initial incision (Fig. 3c). Be careful not to damage the articular surfaces or ligaments (see Note 7 for control of bleeding). Using forceps, remove the fat pad and displace any soft tissue obstructing the view of the medial meniscus. While holding the distal hind limb, apply gentle pressure laterally on the patellar tendon using jeweler’s forceps to perform a lateral dislocation of the patella and expose more of the joint space. Alternatively, loop a 5-0 suture around the patellar tendon and gently pull it away from the surgical site to expose the medial meniscus. Once open, identify the medial meniscus (MM) and locate the medial meniscotibial ligament (MMTL) attached to the tibial plateau (Fig. 4a, b). Apply sterile saline using a dropper or pipette to the joint space to prevent dehydration of the articular cartilage and surrounding tissues.

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Fig. 4 Transection of the MMTL. (a, b) Visualization of the MMTL and medial meniscus can be made following an incision medial to the patellar tendon and subsequent lateral dislocation of the patella (c) View of the intact MMTL before transection. Black arrow denotes the MMTL prior to transection (d) Representative image of a transected MMTL in the knee joint space. F femur, T tibia, MCL medial collateral ligament, MMTL medial meniscotibial ligament

4. Transect the MMTL: Using an inverted #11 scalpel blade, carefully place the tip of the scalpel underneath the MMTL and lift to transect the MMTL (Fig. 4c, d). If preferred, spring microdissection scissors can be used to cut the MMTL. Once the MMTL is severed, the medial meniscus will be destabilized and move more freely within the joint space (Fig. 4d). Gently reposition the patella and patellar tendon using forceps. If a suture was used to displace the patellar tendon, unloop the suture to allow the patellar tendon to be repositioned. 5. Closure: In the mouse, because it is only a few cell layers thick, it is difficult to suture the capsular membrane (i.e. the synovium) following disruption of the meniscus. If desired, close

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the joint capsule with 6-0 absorbable sutures. Alternatively, simply close the skin incision with 5-0 nylon sutures, applied in an interrupted pattern. Apply approximately 1 suture for every 1–2 mm of incision length. Following closure, articulate the joint several times to verify the integrity of the sutures. Apply additional sutures as necessary to maintain closure. 6. Postoperative Recovery: If isoflurane is the anesthetic used, stop the flow of gas and remove the face cone to permit access to ambient atmosphere (see Note 8 if isoflurane anesthesia is not used). Place the mouse in a clean “post-op” cage to recover, where within 3–5 min it should begin emerging from the effects of the isoflurane. Once it is fully able to ambulate (within 10 min), place the mouse in a fresh housing cage or with other mice that have just recovered from the procedure. Return the housing cage to the vivarium housing room as soon as possible. As mentioned, administer analgesia (buprenorphine, 0.05 mg/kg) every 12 h for 3 days or, if available, administer a single dose of slow-release buprenorphine. 7. Harvest & Analyze Tissue: At the predetermined experimental time points, perform euthanasia, remove the hind limbs, and dissect the joint from the mid-femur to the mid-tibia, removing as much soft tissue as possible without disrupting the joint capsule. For cartilage RNA isolation and analysis, disarticulate the joint, remove the tibial plateau and femoral condyle articular cartilage, and perform nucleic acid isolation and analysis as we have previously described [29]. For tissue analysis, follow the OARSI-recommended scoring and histomorphometry following fixation, decalcification and sectioning of the joints as recommended by the OARSI task force [30], and as we have previously described [25, 29]. Histology sections prepared this way will also be useful for molecular analyses involving in situ hybridization and immunohistochemistry. Many of the tissue preparation, fixation and staining methods are described elsewhere in this book.

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Notes 1. The DMM procedure described in this chapter has been routinely used to initiate posttraumatic OA in C57BL/6 mice. Other inbred and hybrid strains have been used in various posttrauma models of OA, with strain-specific differences in the pacing of the disease process noted previously [17]. Thus, forethought related to strain difference is required when designing the experimental approach. An additional key factor that dictates choice of strain relates to the genetic background carrying a specific genetic alteration that is required for addressing hypotheses of interest. To minimize indirect effects of

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skeletal growth and to maximize the size of the joint (enabling better visualization of structures and accurate application of the injury), mice that are used for the DMM procedure are not younger than 12 weeks of age. 2. It is common for male mice to be used preferentially when studying posttraumatic OA. The main rationale for this is that male mice display a more rapid pace of degeneration postDMM relative to female counterparts. Females can also be used, however, with the caveat that the degenerative process takes about twice as long. Animals are usually group-housed (5 per cage) following the surgery, but close attention should be paid to the interaction between the mice in the immediate postsurgery period due to potential fighting and surgical wound opening that can occur when males are grouped. It is best to allow littermate males to acclimatize to each other for 1–2 weeks prior to surgery to identify possible conflicts. This grouping should be maintained post-DMM to minimize fighting. Finally, mice should be housed using a 12-h light–dark cycling to maintain diurnal rhythm and avoid potential effects that an altered rhythm might have on cartilage homeostasis [31]. 3. Deep anesthesia is critical to prevent spurious tissue damage. The safest and most reliable method to achieve adequate anesthetic restraint as quickly as possible involves the use of isoflurane gas. A murine face cone will be required as well as an anesthesia machine that allows for titering of the gas–air mixture to achieve (5% isoflurane) and maintain (2% isoflurane) a surgical plane. Also, the necessary gas scavenging equipment will be required to prevent occupational exposure of the surgical team. If isoflurane delivery and scavenging equipment is not available, an alternative would be to use a mix of ketamine and xylazine that is administered via intraperitoneal injection in a dose that is carefully titered for the body weight of the mouse (88 mg/kg ketamine, 8 mg/kg xylazine). The key caveats to the use of a ketamine–xylazine mix is the extended period required to achieve the surgical plane, the variability in the amount of anesthetic necessary based on a body weight measurement, and the extended recovery period following completion of the surgery, which can be up to an hour depending on dose administered. 4. To maximize visualization of structures within the joint, a dissecting microscope is recommended. Alternatively, surgical loops can be used, although this requires that the surgeon have a pair made to meet their visual specifications and fit. 5. DMM (and other similar types of mouse knee trauma surgery to induce OA) is a microsurgical procedure that requires practice to master. It is recommended that the surgical team

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perform the procedure on a cohort of mice that will be followed histologically to assess reproducibility of the OA disease process. Even slight damage to the joint surface or other nontargeted structures in the joint will lead to more rapid progression of disease and thus add to the size of experimental error when performing quantitative analyses (histomorphometry in particular). In our experience, from a stereotactic perspective, DMM surgery is easiest to administer to the mouse’s right hind limb when the surgeon is right-handed. Accordingly, lefthanded surgeons have a more straightforward approach to the left hind limb. 6. It is common to perform a knee-destabilizing surgical procedure on one limb, using the contralateral limb as a sham-operated control. The specific procedural approach for administering sham surgery is identical through step 3 of the surgical protocol described in this chapter, followed by steps 5 and 6. It is important to perform the procedure through to the point when capsulotomy is performed to control for effects that synovial injury might have on otherwise healthy joint cartilage. It is widely felt that an un-operated joint, or application of a skin incision only does not provide an appropriate negative control. 7. An experienced surgeon should be able to complete the surgical procedure without significant bleeding. If necessary, however, astriction can be used to arrest hemorrhaging. Failing this, application of one drop of a 1:1000 solution epinephrine (in 0.9% NaCl) can be employed. 8. If the ketamine–xylazine mix was used as the anesthetic, place the recovering mouse in a clean “post-op” cage that is resting on a 37  C warming pad to prevent hypothermia. Once fully ambulatory (30–60 min), transfer to a housing cage, return to the vivarium, and provide buprenorphine analgesia as described.

Acknowledgments We would like to thank Sarah Mack, from the Histology, Biochemistry and Molecular Imaging Core in the Center for Musculoskeletal Research at the University of Rochester Medical Center, for outstanding support in the generation of histological images of murine knee joint tissue. Support for establishment of the DMM protocol at the Center for Musculoskeletal Research at the University of Rochester and the Center for Orthopedic Research at the University of Colorado was provided by NIH NCATS TR000042 (M.J.Z.), NIH NIAMS P50 AR054041-5471 (M.J.Z.), Core services supported by NIH NIAMS P30 AR061307 and P30 AR069665, and DOD W81XWH1910807 (M.J.Z.).

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References 1. Hootman JM, Helmick CG (2006) Projections of US prevalence of arthritis and associated activity limitations. Arthritis Rheum 54 (1):226–229 2. Buckwalter JA, Mankin HJ, Grodzinsky AJ (2005) Articular cartilage and osteoarthritis. Instr Course Lect 54:465–480 3. Loughlin J, Dowling B, Chapman K, Marcelline L, Mustafa Z, Southam L, Ferreira A, Ciesielski C, Carson DA, Corr M (2004) Functional variants within the secreted frizzled-related protein 3 gene are associated with hip osteoarthritis in females. Proc Natl Acad Sci USA 101(26):9757–9762 4. Reynard LN, Loughlin J (2013) Insights from human genetic studies into the pathways involved in osteoarthritis. Nat Rev Rheumatol. https://doi.org/10.1038/nrrheum.2013.121 5. Reynard LN, Loughlin J (2013) The genetics and functional analysis of primary osteoarthritis susceptibility. Expert Rev Mol Med 15:e2. https://doi.org/10.1017/erm.2013.4 6. Goldring MB, Marcu KB (2012) Epigenomic and microRNA-mediated regulation in cartilage development, homeostasis, and osteoarthritis. Trends Mol Med 18(2):109–118. https://doi.org/10.1016/j.molmed.2011.11. 005 7. Mooney RA, Sampson ER, Lerea J, Rosier RN, Zuscik MJ (2011) High-fat diet accelerates progression of osteoarthritis after meniscal/ ligamentous injury. Arthritis Res Ther 13(6): R198. https://doi.org/10.1186/ar3529 8. Griffin TM, Huebner JL, Kraus VB, Yan Z, Guilak F (2012) Induction of osteoarthritis and metabolic inflammation by a very high-fat diet in mice: effects of short-term exercise. Arthritis Rheum 64(2):443–453. https://doi. org/10.1002/art.33332 9. Louer CR, Furman BD, Huebner JL, Kraus VB, Olson SA, Guilak F (2012) Diet-induced obesity significantly increases the severity of posttraumatic arthritis in mice. Arthritis Rheum 64(10):3220–3230. https://doi.org/ 10.1002/art.34533 10. Vo N, Niedernhofer LJ, Nasto LA, Jacobs L, Robbins PD, Kang J, Evans CH (2013) An overview of underlying causes and animal models for the study of age-related degenerative disorders of the spine and synovial joints. J Orthop Res 31(6):831–837. https://doi.org/ 10.1002/jor.22204 11. Englund M (2010) The role of biomechanics in the initiation and progression of OA of the

knee. Best Pract Res Clin Rheumatol 24 (1):39–46 12. Roos H, Lauren M, Adalberth T, Roos EM, Jonsson K, Lohmander LS (1998) Knee osteoarthritis after meniscectomy: prevalence of radiographic changes after twenty-one years, compared with matched controls. Arthritis Rheum 41(4):687–693 13. Suzuki Y, Takeuchi N, Sagehashi Y, Yamaguchi T, Itoh H, Iwata H (1998) Effects of hyaluronic acid on meniscal injury in rabbits. Arch Orthop Trauma Surg 117(6–7):303–306 14. Goto H, Shuler FD, Niyibizi C, Fu FH, Robbins PD, Evans CH (2000) Gene therapy for meniscal injury: enhanced synthesis of proteoglycan and collagen by meniscal cells transduced with a TGFbeta(1)gene. Osteoarthr Cartilage 8(4):266–271 15. Murphy JM, Fink DJ, Hunziker EB, Barry FP (2003) Stem cell therapy in a caprine model of osteoarthritis. Arthritis Rheum 48 (12):3464–3474. https://doi.org/10.1002/ art.11365 16. Janusz MJ, Bendele AM, Brown KK, Taiwo YO, Hsieh L, Heitmeyer SA (2002) Induction of osteoarthritis in the rat by surgical tear of the meniscus: inhibition of joint damage by a matrix metalloproteinase inhibitor. Osteoarthr Cartilage 10(10):785–791 17. Kamekura S, Hoshi K, Shimoaka T, Chung U, Chikuda H, Yamada T, Uchida M, Ogata N, Seichi A, Nakamura K, Kawaguchi H (2005) Osteoarthritis development in novel experimental mouse models induced by knee joint instability. Osteoarthr Cartilage 13 (7):632–641 18. Glasson SS, Blanchet TJ, Morris EA (2007) The surgical destabilization of the medial meniscus (DMM) model of osteoarthritis in the 129/SvEv mouse. Osteoarthr Cartilage 15(9):1061–1069 19. Clements KM, Price JS, Chambers MG, Visco DM, Poole AR, Mason RM (2003) Gene deletion of either interleukin-1beta, interleukin1beta-converting enzyme, inducible nitric oxide synthase, or stromelysin 1 accelerates the development of knee osteoarthritis in mice after surgical transection of the medial collateral ligament and partial medial meniscectomy. Arthritis Rheum 48(12):3452–3463 20. Glasson SS, Askew R, Sheppard B, Carito B, Blanchet T, Ma HL, Flannery CR, Peluso D, Kanki K, Yang Z, Majumdar MK, Morris EA (2005) Deletion of active ADAMTS5 prevents

Surgical Induction of Osteoarthritis in Mice cartilage degradation in a murine model of osteoarthritis. Nature 434(7033):644–648 21. Majumdar MK, Askew R, Schelling S, Stedman N, Blanchet T, Hopkins B, Morris EA, Glasson SS (2007) Double-knockout of ADAMTS-4 and ADAMTS-5 in mice results in physiologically normal animals and prevents the progression of osteoarthritis. Arthritis Rheum 56(11):3670–3674 22. Little CB, Barai A, Burkhardt D, Smith SM, Fosang AJ, Werb Z, Shah M, Thompson EW (2009) Matrix metalloproteinase 13-deficient mice are resistant to osteoarthritic cartilage erosion but not chondrocyte hypertrophy or osteophyte development. Arthritis Rheum 60 (12):3723–3733. https://doi.org/10.1002/ art.25002 23. Kamekura S, Kawasaki Y, Hoshi K, Shimoaka T, Chikuda H, Maruyama Z, Komori T, Sato S, Takeda S, Karsenty G, Nakamura K, Chung UI, Kawaguchi H (2006) Contribution of runt-related transcription factor 2 to the pathogenesis of osteoarthritis in mice after induction of knee joint instability. Arthritis Rheum 54(8):2462–2470 24. Pond MJ, Nuki G (1973) Experimentallyinduced osteoarthritis in the dog. Ann Rheum Dis 32(4):387–388. https://doi.org/10. 1136/ard.32.4.387 25. Sampson ER, Beck CA, Ketz J, Canary KL, Hilton MJ, Awad H, Schwarz EM, Chen D, O’Keefe RJ, Rosier RN, Zuscik MJ (2011) Establishment of an index with increased sensitivity for assessing murine arthritis. J Orthop Res. https://doi.org/10.1002/jor.21368 26. Hsia AW, Anderson MJ, Heffner MA, Lagmay EP, Zavodovskaya R, Christiansen BA (2017) Osteophyte formation after ACL rupture in

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mice is associated with joint restabilization and loss of range of motion. J Orthop Res 35 (3):466–473. https://doi.org/10.1002/jor. 23252 27. Hsia AW, Tarke FD, Shelton TJ, Tjandra PM, Christiansen BA (2018) Comparison of knee injury threshold during tibial compression based on limb orientation in mice. J Biomech 74:220–224. https://doi.org/10.1016/j. jbiomech.2018.04.014 28. Rai MF, Duan X, Quirk JD, Holguin N, Schmidt EJ, Chinzei N, Silva MJ, Sandell LJ (2017) Post-traumatic osteoarthritis in mice following mechanical injury to the Synovial joint. Sci Rep 7:45223. https://doi.org/10. 1038/srep45223 29. Sampson ER, Hilton MJ, Tian Y, Chen D, Schwarz EM, Mooney RA, Bukata SV, O’Keefe RJ, Awad H, Puzas JE, Rosier RN, Zuscik MJ (2011) Teriparatide as a chondroregenerative therapy for injury-induced osteoarthritis. Sci Transl Med 3(101):101ra193. https://doi.org/10.1126/scitranslmed. 3002214 30. Glasson SS, Chambers MG, Van Den Berg WB, Little CB (2010) The OARSI histopathology initiative—recommendations for histological assessments of osteoarthritis in the mouse. Osteoarthr Cartil 18(Suppl 3):S17–S23. https://doi.org/10.1016/j.joca.2010.05.025 31. Gossan N, Zeef L, Hensman J, Hughes A, Bateman JF, Rowley L, Little CB, Piggins HD, Rattray M, Boot-Handford RP, Meng QJ (2013) The circadian clock in chondrocytes regulates genes controlling key aspects of cartilage homeostasis. Arthritis Rheum. https:// doi.org/10.1002/art.38035

Chapter 7 Parabiosis: Assessing the Effects of Circulating Cells and Factors on the Skeleton Benjamin Alman and Gurpreet Baht Abstract The circulatory system carries within it numerous types of cells, proteins, and other factors that are able to influence the local biology of tissues. Within this chapter, we present a protocol for parabiosis, a surgical model which results in shared circulation between two mice. Such chimeras have recently been used to probe the impact of age-associated changes in the circulation on skeletal, muscular, and neural biology. In conjunction with transgenic mouse models, parabiosis can be used as a tool to investigate the effects of specific factors on local tissues. Here we discuss our adaptation of this surgical procedure including technique details, pitfalls, and suggestions for optimization. Key words Circulating factors, Circulating cells, Parabiosis, Anastomosis, Rejuvenation

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Introduction In recent years, cells, proteins, and small molecules within the circulation have garnered attention for their ability to influence bone biology during development, homeostasis, and repair. Thus, efforts are now being made to better understand the role of these cells and factors on the skeleton. Most strikingly, the resurgence of parabiosis models in concert with the development of transgenic mouse models has led to the burgeoning study of circulating cells and factors, as they affect age and disease, in the context of tissue regeneration and homeostasis [1–4]. Parabiosis is a surgical procedure that results in the anastomosis of two organisms. In this way, the circulatory system of two mice can be connected to result in the sharing of circulating cells, proteins, and other factors. This classical method has helped to identify the existence and function of leptin [5, 6], the origin of the osteoclast [7, 8], and recently we and others have used parabiosis to identify changes in circulatory factors with age and to determine how these changes manifest an “aged bone” phenotype [9–12]. It should be noted that while this model is powerful in addressing

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the importance of circulation in bone biology, it is difficult to separate the variable of mechanotransduction. For example, the forced locomotion of an aged parabiont by a young parabiont could lead to altered bone and skeletal metrics, independent of the effect of circulating factors. To control for this anomaly follow-up models (such as bone marrow transplantation and/or transgenic mouse models) are often required. Here we outline our adaptation of the originally reported procedure and discuss considerations when using such a protocol [13].

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Materials All work reported within this protocol has been approved by the Duke University Institutional Animal Care and Use Committee (IACUC). Prior to surgical setup, all surfaces are cleaned with 70% ethanol and all instruments are washed and autoclaved. 1. Two laboratory mice. See Notes 1–3. 2. Sutures (4-0). See Note 4. 3. Ophthalmic liquid gel. 4. Saline solution. 5. Buprenorphine-SR (sustained release). 6. Anesthetic setup including two vaporizers, each with independent adjustable O2 flow meter and their own nose cones. See Note 5. 7. Oxygen tank. 8. Isoflurane. 9. 70% ethanol. 10. Povidone–iodine 10%. 11. 1.0 mL syringes. 12. 25-G needles. 13. Heating pad. 14. Scalpel. 15. Scissors. 16. Forceps. 17. Needle driver/needle holder. 18. Trimmer/shaver suitable for rodent fur.

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Methods

3.1 Preparation for Surgery

1. Select the pair of mice to be anastomosed. House each pair in a separate cage for a minimum of 2 weeks, in order for the mice to acclimate themselves with each other. A workflow of the parabiosis pairing is diagramed in Fig. 1. 2. At the time of surgery, weigh each mouse and ensure the individual weights of the two mice are within 10% of each other. 3. Prepare the surgical work area: include an area designated for shaving, an operative area, and an area to house sterile instruments. 4. Using 2% isoflurane and 1 L/min O2 flow, place the first mouse under sedation using an induction chamber. 5. Once the mouse is under sedation, place it in the shaving area and use a nose cone to administer 2% isoflurane. 6. Shave the full flank of the mouse; remove any loose fur with an ethanol wipe. 7. Administer 500 μL of warmed saline subcutaneously. See Note 6. 8. Move the mouse to the operative area and use a nose cone to administer 2% isoflurane. 9. Repeat steps 4 through 8 for the second mouse. 10. Clean the surgical site of both mice with three rounds of 70% ethanol followed by povidone-iodine. 11. Apply ophthalmic liquid gel to the eyes. See Note 7.

Fig. 1 Workflow of parabiosis-based analysis. Mice to be anastomosed together are commonly weaned into the same cage, with no other cage mates. If this is not possible, the mice are paired into the same cage for a minimum of 2 weeks. After parabiosis surgery the mice are allowed to heal fully for 4 weeks. During this time blood sharing is established and can be measured using flow cytometry. Upon confirmation of blood sharing, skeletal assessment can be performed

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Fig. 2 Regions of interest in anastomosis surgery. (a) A longitudinal incision is made along the flank of the mice. (b) The forelimbs of each parabiont are attached with sutures proximal to the elbow. (c) The hindlimbs of each parabiont are attached with sutures proximal to the knee

3.2 Anastomosis Surgery

1. With a scalpel, make a longitudinal incision (superficial, skindeep) from just proximal to the knee, continuous to the posterior shoulder (Fig. 2a). Perform this for each mouse. 2. Using absorbable suture material, introduce two interrupted sutures to attach the lateral aspects of the lateral head of the triceps brachii of each mouse (Fig. 2b). See Note 8. 3. Using absorbable suture material, attach the lateral aspect of the adjacent latissimus dorsi from the two mice, bringing together the two rib cages. 4. Tie a simple suture knot but do not trim the suture material. 5. Place seven to nine continuous sutures to bring together the adjacent body walls of the mice. See Note 9. 6. Using absorbable suture material, introduce two interrupted sutures to attach the lateral aspects of the vastus lateralis of the quadriceps femoris of each mouse, resulting in attaching the legs together (Fig. 2c). 7. Using nonabsorbable suture material, close the incision site using interrupted sutures, bringing the skin of one mouse together with the skin of the other mouse. See Note 10. 8. Administer 500 μL of warmed saline, subcutaneously on the outer flank, to each mouse. 9. Allow the pair to wake from anesthetic on a heating pad. 10. Monitor the behavior upon waking. See Notes 11 and 12.

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Fig. 3 Example of blood analysis to determine blood sharing. The most common method to determine blood sharing in parabiosis pairs is measurement of a circulating label. In the example depicted above, GFP+ cells within the circulation are measured using flow cytometry. Solitary wild-type (WT) and GFP-positive mice are used as comparative controls. Within the pairings, both WT and GFP mice are assessed for the presence of GFP+ cells in circulation

11. Mice should be allowed to heal from parabiosis surgery for 3–4 weeks. See Notes 13 and 14. 12. Blood sharing can be determined using flow cytometry. See Notes 15 and 16. Figure 3. 3.3

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Final Thoughts

This protocol results in the anastomosis of two mice. In this way, the importance of circulating factors on the skeleton can be identified. We have recently investigated how circulating factors change with age and how these molecules affect fracture healing [1, 2, 9]. Others have used similar experimental approaches to identify age-associated circulatory effect in muscle regeneration [14, 15], cardiac tissue biology [16], and cognition [17]. After establishing the parabiosis pair, skeletal phenotyping (as discussed in later chapters) can be used to determine the role of circulating cells and factors on the skeleton.

Notes 1. As this protocol results in a shared circulatory system, graft versus host disease (referred to as “parabiosis disease” in the context of laboratory anastomosis) is a possible complication [18, 19]. To minimize this occurrence, it is suggested to use an inbred mouse strain (such as C57/BL6). Additionally, backcrossing the mice for a number of generations (>20) has also been noted to decrease such complications.

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2. For anastomosis, female mice are suggested as they are more docile. Certainly this is a limitation as sex-related changes in bone phenotype may be overlooked; however, some studies have reported the successful use of male mice provided they are littermates and have been housed together from weaning [20, 21]. 3. As the mice will be physically joined for a period of time, differences in the relative sizes of the mice can present itself as a physical stressor reducing mobility and accessibility to food. To minimize this complication, when possible, mice of similar sizes (ideally within 10% body weight) should be used. 4. Sutures for internal use should be long-lasting absorbable material such as poly(p-dioxanone). Sutures for external use should be of nonabsorbable material such as polypropylene. 5. Two separate vaporizers are suggested to control the depth of anesthesia of each mouse individually during the procedure. 6. During the parabiosis surgery, the mice are placed on a heating pad to help maintain body temperature. Due to the large surgical site, a significant area of the internal tissue will be exposed to the ambient air. Collectively, the heated environment and the prolonged exposure of internal tissue surfaces to air cause increased dehydration of the mice. We have found that administering saline pre- and postoperatively is helpful in maintaining hydration. 7. Due to the duration of the procedure and the frequent need to adjust the mice and the nose cones, ophthalmic liquid gel may need to be reapplied during the procedure. 8. After this step the mice should be in a natural supine position, adjacent to one another, attached at the proximal region of the forelimbs, without any stress being placed on the attachment due to pressure from the head or the shoulders. 9. The continuous sutures should be taut so that there are no gaps within the adjoining fascia. 10. Interrupted suturing is suggested for closing in this surgical procedure as it leads to less complications; however, the use of dermal staples would be faster and has been proven effective in parabiosis [22]. 11. Upon waking, the mice should be able to ambulate and have full use of their attached limbs. The mice should not appear to be in any discomfort. 12. Due to the buprenorphine-SR and the extended time under anesthetic, the mice may take 5–15 min to fully awaken. 13. During recovery from parabiosis surgery, health monitoring of the pair should be performed daily for the duration of the

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experiment—until pair sacrifice and tissue harvest. The most common complications from parabiosis surgery are dehydration, lethargy due to dehydration, excessive weight loss, and lethargy due to wasting. These symptoms can be treated with subcutaneous injections of saline and ad libitum feeding of hydrating food gel. 14. A less frequent complication of the surgery is lack of mobility of the attached forelimb or hindlimb. If care is not taken during surgery and tissues not indicated in this protocol are attached, it is possible for the mice to lose full range of motion. This will present as a lame limb, limping, or preference of one limb over the other. In severe situations, blood vessels may be impinged which can lead to injury and even loss of the limb. To avoid these complications, this procedure should be extensively practiced and planned using sacrificed mice prior to attempting survival surgery. 15. In parabiosis studies, blood sharing is conventionally assessed using flow cytometry. To measure and monitor blood sharing over time, whole blood from pairs in which labeled mice have been surgically attached to nonlabeled mice (eg. eGFP mouse to C57B/L6 mouse) can be collected at longitudinal time points. The appearance of labeled cells within the nonlabeled mouse is assessed and related to the number of labeled cells within the labeled mouse. Ideally, the number of labeled cells within the two mice would be equal, indicating equal blood sharing. 16. Attachment within the parabiosis surgery can be divided into four areas: (1) triceps brachii, (2) quadriceps femoris, (3) skin closing, and (4) the body wall. These areas can be classified as serving one of two primary functions: (1) Physical attachment of the mice—triceps brachii, quadriceps femoris, and skin closing. These suture points serve to anchor the mice together and to ensure they are not injured secondary to movement. (2) Induction of blood sharing–body wall. Suturing of the body wall causes slight injury within both mice and healing of the two injured sights in combination with the close proximity of the injured sites leads to angiogenesis and healing of the tissue of one mouse into the adjacent mouse. Through this process, blood sharing is made possible. Indeed, some studies have used interfaces such as polymer disks to prevent touching of the body walls after surgical attachment and this action prevented blood sharing [16].

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References 1. Baht GS, Silkstone D, Vi L, Nadesan P, Amani Y, Whetstone H, Wei Q, Alman BA (2015) Exposure to a youthful circulaton rejuvenates bone repair through modulation of beta-catenin. Nat Commun 6:7131. https:// doi.org/10.1038/ncomms8131 2. Xiong C, Zhang Z, Baht GS, Terrando N (2018) A mouse model of orthopedic surgery to study postoperative cognitive dysfunction and tissue regeneration. J Vis Exp 132:56701. https://doi.org/10.3791/56701 3. Conboy IM, Conboy MJ, Wagers AJ, Girma ER, Weissman IL, Rando TA (2005) Rejuvenation of aged progenitor cells by exposure to a young systemic environment. Nature 433 (7027):760–764. https://doi.org/10.1038/ nature03260 4. Brack AS, Conboy MJ, Roy S, Lee M, Kuo CJ, Keller C, Rando TA (2007) Increased Wnt signaling during aging alters muscle stem cell fate and increases fibrosis. Science 317 (5839):807–810. https://doi.org/10.1126/ science.1144090 5. Coleman DL (1973) Effects of parabiosis of obese with diabetes and normal mice. Diabetologia 9(4):294–298. https://doi.org/10. 1007/bf01221857 6. Coleman DL (1978) Obese and diabetes: two mutant genes causing diabetes-obesity syndromes in mice. Diabetologia 14(3):141–148. https://doi.org/10.1007/bf00429772 7. Walker DG (1975) Bone resorption restored in osteopetrotic mice by transplants of normal bone marrow and spleen cells. Science 190 (4216):784–785. https://doi.org/10.1126/ science.1105786 8. Walker DG (1975) Spleen cells transmit osteopetrosis in mice. Science 190(4216):785–787. https://doi.org/10.1126/science.1198094 9. Vi L, Baht GS, Soderblom EJ, Whetstone H, Wei Q, Furman B, Puviindran V, Nadesan P, Foster M, Poon R, White JP, Yahara Y, Ng A, Barrientos T, Grynpas M, Mosely MA, Alman BA (2018) Macrophage cells secrete factors including LRP1 that orchestrate the rejuvenation of bone repair in mice. Nat Commun 9 (1):5191. https://doi.org/10.1038/s41467018-07666-0 10. Boban I, Jacquin C, Prior K, Barisic-DujmovicT, Maye P, Clark SH, Aguila HL (2006) The 3.6 kb DNA fragment from the rat Col1a1 gene promoter drives the expression of genes in both osteoblast and osteoclast lineage cells. Bone 39(6):1302–1312. https://doi.org/10. 1016/j.bone.2006.06.025

11. Boban I, Barisic-Dujmovic T, Clark SH (2010) Parabiosis model does not show presence of circulating osteoprogenitor cells. Genesis 48 (3):171–182. https://doi.org/10.1002/dvg. 20602 12. Huang R, Zong X, Nadesan P, Huebner JL, Kraus VB, White JP, White PJ, Baht GS (2019) Lowering circulating apolipoprotein E levels improves aged bone fracture healing. JCI Insight 4(18). https://doi.org/10.1172/jci. insight.129144 13. Bunster E, Meyer R (1933) An improved method of parabiosis. Anat Rec 57:339–343 14. Sinha M, Jang YC, Oh J, Khong D, Wu EY, Manohar R, Miller C, Regalado SG, Loffredo FS, Pancoast JR, Hirshman MF, Lebowitz J, Shadrach JL, Cerletti M, Kim MJ, Serwold T, Goodyear LJ, Rosner B, Lee RT, Wagers AJ (2014) Restoring systemic GDF11 levels reverses age-related dysfunction in mouse skeletal muscle. Science 344(6184):649–652. https://doi.org/10.1126/science.1251152 15. Conboy IM, Rando TA (2012) Heterochronic parabiosis for the study of the effects of aging on stem cells and their niches. Cell Cycle 11 (12):2260–2267. https://doi.org/10.4161/ cc.20437 16. Loffredo FS, Steinhauser ML, Jay SM, Gannon J, Pancoast JR, Yalamanchi P, Sinha M, Dall’Osso C, Khong D, Shadrach JL, Miller CM, Singer BS, Stewart A, Psychogios N, Gerszten RE, Hartigan AJ, Kim MJ, Serwold T, Wagers AJ, Lee RT (2013) Growth differentiation factor 11 is a circulating factor that reverses age-related cardiac hypertrophy. Cell 153(4):828–839. https://doi.org/10.1016/j.cell.2013.04.015 17. Villeda SA, Plambeck KE, Middeldorp J, Castellano JM, Mosher KI, Luo J, Smith LK, Bieri G, Lin K, Berdnik D, Wabl R, Udeochu J, Wheatley EG, Zou B, Simmons DA, Xie XS, Longo FM, Wyss-Coray T (2014) Young blood reverses age-related impairments in cognitive function and synaptic plasticity in mice. Nat Med 20(6):659–663. https://doi.org/10.1038/nm.3569 18. Conboy MJ, Conboy IM, Rando TA (2013) Heterochronic parabiosis: historical perspective and methodological considerations for studies of aging and longevity. Aging Cell 12 (3):525–530. https://doi.org/10.1111/acel. 12065 19. Finerty JC, Panos TC (1951) Parabiosis intoxication. Proc Soc Exp Biol Med 76 (4):833–835

The Effect of Circulation on the Skeleton 20. Kumagai K, Vasanji A, Drazba JA, Butler RS, Muschler GF (2008) Circulating cells with osteogenic potential are physiologically mobilized into the fracture healing site in the parabiotic mice model. J Orthop Res 26 (2):165–175. https://doi.org/10.1002/jor. 20477 21. Brazelton TR, Blau HM (2005) Optimizing techniques for tracking transplanted stem cells

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in vivo. Stem Cells 23(9):1251–1265. https:// doi.org/10.1634/stemcells.2005-0149 22. Ajami B, Bennett JL, Krieger C, Tetzlaff W, Rossi FM (2007) Local self-renewal can sustain CNS microglia maintenance and function throughout adult life. Nat Neurosci 10 (12):1538–1543. https://doi.org/10.1038/ nn2014

Chapter 8 Murine Limb Bud Organ Cultures for Studying Musculoskeletal Development Martin Arostegui and T. Michael Underhill Abstract The biological signals that coordinate the three-dimensional outgrowth and patterning of the vertebrate limb bud have been well delineated. These include a number of vital embryonic signaling pathways, including the fibroblast growth factor, WNT, transforming growth factor, and hedgehog. Collectively these signals converge on multiple progenitor populations to drive the formation of a variety of tissues that make up the limb musculoskeletal system, such as muscle, tendon, cartilage, stroma, and bone. The basic mechanisms regulating the commitment and differentiation of diverse limb progenitor populations has been successfully modeled in vitro using high density primary limb mesenchymal or micromass cultures. However, this approach is limited in its ability to more faithfully recapitulate the assembly of progenitors into organized tissues that span the entire musculoskeletal system. Other biological systems have benefitted from the development and availability of three-dimensional organoid cultures which have transformed our understanding of tissue development, homeostasis and regeneration. Such a system does not exist that effectively models the complexity of limb development. However, limb bud organ cultures while still necessitating the use of collected embryonic tissue have proved to be a powerful model system to elucidate the molecular underpinning of musculoskeletal development. In this methods article, the derivation and use of limb bud organ cultures from murine limb buds will be described, along with strategies to manipulate signaling pathways, examine gene expression and for longitudinal lineage tracking. Key words Limb bud, Skeletal development, Organ culture, In situ hybridization

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Introduction The development of the vertebrate limb is a carefully choreographed program that relies on the precise coordination of diverse cellular signals, cell-cell interactions, and mechanical forces. Mesodermal derivatives from various sources respond to distinct signals present in the limb bud microenvironment and generate muscle, tendon, and cartilage tissues within the limb musculoskeletal system. At early stages of development, the limb bud consists of a seemingly morphologically homogeneous population of mesenchymal cells encased in an overlying simple ectoderm [1]. These mesenchymal cells are derived from lateral plate mesoderm (LPM),

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which is the subdivision of mesoderm that surrounds the trunk of the embryo. The LPM will give rise to the appendicular cartilaginous skeletal elements and associated connective tissues such as tendon and ligament [2–4]. In contrast, limb musculature is formed by muscle progenitors that originate from the ventrolateral dermomyotome of somites and migrate into the nascent limb buds at specific stages of development [5–7]. Once within the limb, myoblasts undergo a period of rapid proliferation followed by aggregation into dorsal and ventral muscle masses [8]. Afterward, myoblasts partition into subgroups representing future muscle bodies and begin to form myotubes effectively generating mature muscle groups [9]. For limbs to create motion, muscles must transmit their generated forces onto the skeletal framework via tendons. Tendons are specialized connective tissues formed primarily by type I collagen fibrils organized in parallel to the tendon axis [10]. The identification of Scleraxis, a basic helix–loop–helix transcription factor, as a marker of connective tissues that mediate the attachment of muscle to bone [11], has been instrumental in uncovering the cellular and molecular mechanisms underlying embryonic tenogenesis. Through analysis of Scx expression, Brent and colleagues were able to identify the syndetome, a fourth somitic compartment of Scx-expressing axial tendon progenitors [12]. Unlike axial tendons, limb tendons are not derived from the syndetome [13]. Instead, limb tendon progenitors are derived from the lateral plate mesoderm of the forming limb bud and differentiate into mature tendons upon interactions with cartilage and muscle precursors [14]. Tendon development is closely associated with that of musculature, and the examination of muscleless limbs has shown that tendon progenitors initiate their differentiation but further formation of mature tendons requires the presence of muscle [11, 15]. Tendons transmit muscle-generated forces onto the skeletal structure resulting in motion and/or stability. The vertebrate limb skeleton arises through endochondral ossification of a pre-existing cartilaginous template. The first noticeable step leading to the formation of this template is the condensation of prechondrogenic mesenchymal cells [16]. These cells differentiate into cartilage-forming cells, chondrocytes, which produce a cartilaginous template that is eventually remodeled into bone. Critical to the chondrogenic program is the transcription factor SOX9, which belongs to a family of transcription factors characterized by a highmobility group DNA binding domain [17]. Analysis of conditional null mutants of Sox9 has demonstrated its requirement in the formation of precartilaginous condensations and in subsequent chondrocyte differentiation [18]. Other Sox genes are expressed in the developing skeleton, in particular, L-Sox5 and Sox6, and together are necessary for cartilage formation, but appear to act downstream of Sox9 [19, 20]. Together, these factors coordinate

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the expression of the chondrocytic phenotype and the expression of a matrix rich in type II collagen (Col2a1) and aggrecan (Acan) [21– 23]. As chondrocytes mature, they exit the cell cycle, increase in size, and become hypertrophic [24, 25]. Hypertrophic chondrocytes are characterized in part by high levels of alkaline phosphatase (ALP) activity and by expression of type X collagen (Col10a1). During subsequent ossification, it is believed that most hypertrophic chondrocytes undergo apoptosis. However, an increasing body of evidence suggests that some chondrocytes may have specific functions associated with bone formation, including recruitment of bone cells [26] and transdifferentiation into osteoblasts [27]. Indeed, it has been shown that, during development, hypertrophic chondrocytes re-enter the cell cycle and differentiate into osteoblasts, osteocytes, and bone marrow stromal cells thus contributing to the formation of trabecular bone, the endosteum, and the hematopoietic niche [28, 29]. Historically, musculoskeletal development has been studied with the aid of animal models that are easily amenable to experimental manipulations during gestation. Examples include birds, amphibians, and reptiles [30]. The advent of in vitro cultures of embryonic tissues allowed for the interrogation of musculoskeletal development in species more closely related to humans such as mice and rats. These in vitro techniques consisted of culturing whole or dissociated embryonic tissues under conditions that permit for the differentiation of precursor cells into more mature cell types such as chondrocytes, myocytes, and tenocytes. Techniques in this category include primary limb bud mesenchyme cultures [31], calvarial and mandibular organ cultures [32, 33], and embryonic limb bud cultures [30]. In contrast to culturing dissociated tissues, limb bud cultures are not only a good model for studying cell differentiation and dynamics but also provide valuable information on organ morphogenesis in three dimensions (Fig. 1). Ex vivo culture systems are robust, easy to manipulate, and more ethically acceptable than in vivo systems [34]. Although culturing limb buds ex vivo leads to some skeletal abnormalities, anlagen of the major bone elements appear in a clearly recognizable fashion (Fig. 2a). In the context of musculoskeletal malformations or diseases, limb bud cultures serve as a robust experimental model to explore the molecular basis of such deficiencies. In this manner, the system is highly amenable to genetic and chemical manipulation such that gene and/or function and fate can be efficiently interrogated. The use of appropriate experimental perturbation systems in conjunction with limb cultures routinely give rise to 90% of abnormalities seen in vivo [35]. In this chapter, we seek to introduce an updated version of the limb bud organ culture method that leverages the large number of genetically engineered mouse models. In combination with modern-day methods and technologies, limb bud cultures remain

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Fig. 1 Overview of the limb bud organ culture system and associated applications. Organ cultures are typically initiated from E10–E11 embryos. Limbs are carefully dissected away from the body wall and placed into a dedicated culture apparatus. Limbs are typically cultured for various time frames from 1 to 6 days. Shown are examples of limb buds that have been cultured and processed for X-gal staining (LacZ detection; E11.5 limb bud cultured for 6 days), lineage tracing in a Cre-dependent tdTomato reporter background (TAM was delivered to the pregnant dam at E10; E11.5 limb bud cultured for 4 days), detection of MyHC (anti-MyHC skeletal, Fast Green) and SOX9 (red) using whole mount immunofluorescence of 3 day cultured E11.5 limb bud, and WISH detection of Aldh1a2 in limb buds that were cultured for 24 h (collected at E11.75) following implantation of Affi-Blue beads containing BMP4 [38]

a relevant, highly informative, and practical model for studying musculoskeletal development and diseases. Here, we show how to combine limb bud cultures with genetic lineage tracing, whole mount in situ hybridization, tissue clearing, whole mount immunofluorescence staining, and confocal microscopy to achieve a powerful characterization of musculoskeletal development and any associated perturbations.

2

Materials

2.1 Preparation of Limb Culture Incubation Chambers

1. Stainless steel fine mesh cut into 2.5
2.5 cm squares (see Note 1). 2. Center-well polystyrene organ culture dish 60
15 mm, polystyrene, tissue culture treated by vacuum gas plasma (Falcon).

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Fig. 2 Representative examples of limb bud organ cultures processed for different purposes. (a) Alcian Blue stained images that show the developing cartilaginous elements with the limb bud organ cultures over time. Similar to normal limb development, differentiation and elaboration of the chondrogenic elements occurs in a proximal to distal direction. The chondrogenic digit arrays can be easily visualized after long culture periods (b) Glass and metal imaging apparatus for whole imaging of BABB-cleared samples. (c) Limb bud organ culture setup with stainless steel screen and membrane in an organ culture dish. (d) Gross images of cultured limb buds at the day of collection and 1–2 days postculture. Note the cartilaginous elements begin to become visible by day 1 and these can be visualized as opaque regions within the limb bud. (e) Images of 3D renditions of 3 day cultured, whole mount stained, BABB cleared limb buds. The orientation is noted as distal, dist; proximal, prox; anterior, ant; and posterior, post

3. Scalpel with No. 10 stainless steel surgical blade. 4. No. 5 Dissection forceps (Fine Science Tools) or similar. 5. Cell culture insert, 6 well format, 1.0 μm pore size PET tracketched membrane (Falcon). 6. BGJb media (Gibco, Life Technologies). 7. Penicillin/Streptomycin (Gibco, Life Technologies), L-Glutamine (Gibco, Life Technologies). 8. Humidified tissue culture incubator set at 37  C and 5% CO2. 9. Laminar flow hood.

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2.2 Collection of Embryonic Limb Buds

1. Set up timed matings with mouse lines of interest (see Note 2). 2. A humane method for euthanizing mice according to the animal care committee of the host institution (see Note 3). 3. Stereo dissection microscope with low power magnification. 4. Dissection scissors, #5 dissection forceps (Fine Science Tools), microdissection scissors (World Precision Instruments), and embryo spoon (Fine Science Tools). 5. Cut or wide-bore P1000 pipette tips. 6. 10 cm sterile bacterial petri dishes. 7. Sterile 1
phosphate buffered saline (PBS): 137 mM NaCl, 2.7 mM KCl, 8 mM Na2HPO4, and 2 mM KH2PO4. Sterilize through autoclaving before use and dispense in a laminar flow tissue culture hood (store at 4  C).

2.3 Whole Mount LacZ Staining

1. 1
PBS. 2. LacZ Fixative solution: 100 mM MgCl2, 0.2% glutaraldehyde, 5 mM ethylene glycol tetraacetic acid (EGTA) in 1
PBS. 3. 4% paraformaldehyde (PFA) (see Note 4) in 1
PBS. 4. 20 mM MgCl2 (10
stock) prepared in ddH2O (store at room temperature). 5. 0.1% sodium deoxycholate, 0.2% NP-40 (10
detergent solution) prepared in ddH2O (store at room temperature). 6. 100 mM potassium ferricyanide (20
stock) prepared in ddH2O (store at 4  C). 7. 100 mM potassium ferrocyanide (20
stock) prepared in ddH2O (store at 4  C). 8. 40 mg/ml 5-bromo-4-chloro-3-indolyl-β-d-galactopyranoside (40
X-gal stock) is prepared in dimethylformamide and aliquoted in appropriate “experiment-size” aliquots and stored at 20  C. 9. LacZ staining solution: 2 mM MgCl2, 0.01% sodium deoxycholate, 0.02% NP-40, 5 mM potassium ferricyanide, 5 mM potassium ferrocyanide prepared from stock solutions above. Add X-gal (or other β-galactosidase substrates) to a final 1
concentration or 1 mg/ml. Use immediately and protect from light (see Note 5). 10. 37  C hybridization oven (no rocking necessary). 11. 12-well cell culture cluster plate, flat bottom with lid (Costar or similar).

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2.4 Whole Mount Immunofluorescence Staining, Tissue Clearing, and Imaging

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1. 2% PFA made up fresh in 1
PBS. Prepare from 4% stock described in Subheading 2.3, item 3. 2. 2.0 ml microcentrifuge tubes. 3. Dent’s fix: mix dimethyl sulfoxide (DMSO) and methanol in a 1:4 ratio. 4. Dent’s bleach: mix hydrogen peroxide and Dent’s fix in a 1:2 ratio, prepare fresh. 5. Block solution: 2.5% bovine serum albumin (BSA) (Fraction V, fatty acid free, protease free; Sigma-Aldrich), 2.5% serum from the secondary antibody species in 1
PBS. 6. Primary and secondary antibodies of choice (see Note 6). 7. Methanol. 8. Benzyl alcohol–benzyl benzoate (BABB) clearing solution: 1:2 mixture of benzyl alcohol and benzyl benzoate. Can be stored for long-term use. Prepare in a glass container with cap, as BABB will dissolve polystyrene and polypropylene tubes. 9. 20 ml borosilicate glass scintillation vials. 10. Glass pipettes 10 or 25 ml for dispensing large volumes of organic solvents. 11. Custom-modified slide to contain sample immersed in BABB clearing solution. Alternatively, glass or metal imaging slides/ plates can be used such as the 35 mm stainless steel dish from Aireka Scientific Corporation (Fig. 2b) (see Note 7). 12. Inverted confocal microscope with high laser power capable of deep imaging (see Note 8).

2.5 Whole Mount Alcian Blue Staining

1. Parafilm “M” laboratory film (Fisher Scientific). 2. Formalin (3.7% formaldehyde in water). 3. 0.2 M HCl. 4. Alcian Blue 8GX powder. 5. Alcian Blue stock: 0.5% Alcian Blue in 95% ethanol, filter to remove nondissolved particles. 6. Alcian Blue working solution: mix 0.2 M HCl and 0.5% Alcian Blue in a 4:1 ratio.

2.6 Lineage Tracing in Whole Mount Limb Bud Cultures

1. E11 to E13 embryos collected from a Cre or CreERT2 mouse line along with a Cre-dependent reporter (see Note 9). 2. 2% paraformaldehyde (PFA), make fresh from 4% PFA stock in 1
PBS as in Subheading 2.3, item 3. 3. CUBIC clearing solution: 25% Urea, 25% N,N,N0 ,N0 -tetrakis (2-hydroxy-propyl) ethylenediamine, 15% Triton X-100 in ddH2O.

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4. ScaleA2 clearing solution: 4 M urea, 10% glycerol, 0.1% Triton X-100 in ddH2O. 5. 35 mm imaging dish (MatTek Corporation or similar), No. 1.5 cover glass, 14 mm microwell. 2.7 Whole Mount Double ISH

1. For riboprobe generation, a plasmid containing the insert of interest (300–1000 bp) along with Sp6, T3, or T7 transcriptional elements. 2. RNase-free double distilled water. Diethylpyrocarbonate (DEPC, Sigma-Aldrich) is added to double distilled water at 0.1%, shake vigorously and autoclave. 3. Transcription buffer 10 (Sigma-Aldrich), 1 mM ATP, GTP, and CTP (each); 0.65 mM UTP, 0.35 mM Digoxigenin-11-UTP (Sigma-Aldrich), or Fluorescein-12-UTP (for double whole mount in situ hybridization, WISH); 20–50 units of RNase inhibitor (Sigma-Aldrich); and 20 units of RNA polymerase (New England Biolabs of Sigma-Aldrich). 4. DNA ladder, lambda DNA Hind III digest (New England Biolabs). 5. Riboprobe precipitation, 0.2 M EDTA, 4 M LiCl, and ethanol. 6. Riboprobe quantification, DNA gel apparatus, agarose and Tris-acetate-EDTA buffer (Life Technologies), DNA ladder (lambda DNA-HindIII digest, New England Biolabs) and SYBR green (Life Technologies). 7. PBS, pH 7.4, sterile and RNase free. Prepared in diethyl pyrocarbonate-treated double distilled water. 8. Hybridization oven or heating block with rocking capabilities. 9. 4% PFA in PBS from Subheading 2.3, item 3. 10. PBS-T: 0.1% Triton X-100 detergent in PBS. 11. Methanol. 12. 10 mg/ml Proteinase K (Sigma-Aldrich): Make up in RNasefree water, aliquot and store frozen at 20  C. Thaw a new aliquot each time to assure consistent activity (see Note 10). 13. 0.2% glutaraldehyde (Sigma-Aldrich). 14. Standard saline citrate (SSC) buffer: 20
stock is 3 M NaCl and 0.3 M sodium citrate. Prepare in RNase-free water and autoclave. 15. Hybridization buffer: 50% deionized formamide, 5
SSC, 2% blocking reagent (powder is added directly to the hybridization buffer mix), 0.1% Triton X-100, 0.5% CHAPs (Sigma), 5 mM EDTA, 100 μg/ml tRNA (from a 10 mg/ml stock solution in RNase-free water, Sigma-Aldrich— molecular biology grade tRNA from E. coli MRE600), and 100 μg/ml heparin (Sigma-Aldrich, Grade 1-A: from porcine intestinal mucosa). Make up remaining volume with RNase-free water.

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16. Digoxygenin and/or fluorescein-labeled RNA probes (see Note 11). 17. Posthybridization wash Solution 1: 50% formamide, 5
SSC, 0.1% Triton X-100 and 0.5% CHAPs (Sigma-Aldrich). 18. Blocking reagent (Sigma-Aldrich): 10
stock is prepared in Maleic acid buffer (100 mM Maleic acid, 250 mM NaCl, pH 7.5 adjust with NaOH). For dissolution, heat to at least 60o C for 1 h with agitation, do not boil. Aliquot and store at 20  C. 19. TBST: (0.14 M NaCl, 2.5 mM KCl, 25 mM Tris–HCl 7.5, and 0.1% Tween 20). 20. NTMT: (100 mM NaCl, 100 mM Tris–HCl pH 9.5, 50 mM MgCl2, and 0.1% Tween 20, freshly prepared). 21. Alkaline phosphatase–conjugated anti-digoxigenin and antifluorescein antibodies, Fab fragments (Sigma-Aldrich). For single WISH use anti-digoxigenin and for double WISH the second probe is detected using anti-fluorescein antibody. 22. Staining solutions to produce purple, light blue or red stains: Purple: 4.5 μl/ml NBT (75 mg/ml in 70% dimethylformamide), BCIP 3.5 μl/ml (50 mg/ml in dimethylformamide) in NTMT. Light blue: BCIP 3.5 μl/ml (50 mg/ml in dimethylformamide) in NTMT. Red: 7.5 μl/ml of INT/BCIP stock solution (33 mg/ml INT and 33 mg/ml BCIP in DMSO; Sigma-Aldrich) in NTMT (see Note 12). Prepare all staining solutions fresh just before use. 2.8 Bead Implantation

1. Affi-Gel blue gel beads (Bio-Rad). 2. PBS. 3. 10 cm petri dish. 4. Laminar flow hood. 5. Recombinant protein(s) of interest. 6. Microcentrifuge tubes. 7. Graefe knives (Fine Sciences Tools). 8. Dissection microscope as described above.

3

Methods

3.1 Preparation of Limb Culture Incubation Chambers

1. Optimal culture of the embryonic limb bud with retention of “normal” morphological development, requires that the limb buds are cultured at the air-medium interface (see Note 13). To achieve this, create scaffolds by cutting stainless steel grid

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sections to generate 2.5
2.5 cm squares. Create as many squares as necessary for the desired number of simultaneous cultures (see Note 14). Autoclave and ethanol-sterilize the stainless-steel grids before use. 2. To create an opening for exposure of the limb buds to culture media, generate a hole within the center of each square using a standard (6 mm) hole-punch. 3. The following steps are performed in a Class II biosafety cabinet. 4. Cut the PET membrane off a 1.0 μm-pore size transwell filter using a sterile scalpel by creating an incision around the whole membrane perimeter and pushing it free into a sterile 10 cm dish. Once loose, split the membrane into four equal sections by performing two orthogonal slices through its center. These slices must be done in a single firm and rapid motion to avoid crumpling or folding of the membrane (see Note 15). 5. Place the sterilized stainless steel mesh squares into the center well of 60
15 mm polystyrene organ culture dishes in such a way that the edges maintain the square mesh above the innermost well (Fig. 2c). Place a quarter of the PET membrane cut in the last step on top of the circular hole previously made in the center of the steel mesh. 6. Using a P1000 pipette, dispense BGJb media containing 100 mM Penicillin/Streptomycin and 100 mM L-glutamine into the center well until it encounters the PET membrane, about 1.5 mL is required (see Note 16). 7. Fill the outer trough of the culture dish with sterile dH2O, replace lid and equilibrate at 37  C prior to use. 3.2 Collection of Embryonic Limb Buds

1. Set up timed-pregnancy matings in order to collect E11 to E13 embryos (see Note 17). Breeding pairs are introduced together at the end of the day before the dark cycle and the morning after the copulatory plug is considered embryonic day 0.5 (E0.5). 2. On the day of the desired embryonic time point, sacrifice the pregnant mouse according to the animal care committee of the host institution. Disinfect the abdominal surface by spraying with 70% ethanol and make a small superficial incision in the lower abdomen. Separate the skin from underlying connective tissue using the blunt end of dissection scissors and cut subcutaneously until the lower trunk is accessible. Pull the liberated skin rostrally to reveal the peritoneal membrane. Cut the membrane using dissection scissors to expose the peritoneal cavity. Carefully excise both uterine horns by making a horizontal cut at the cervix and removing excess adipose and connective

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tissues surrounding the uterus. Place the uterus into a 10 cm petri dish containing cold sterile 1
PBS. 3. Under a dissection microscope, carefully dissect out each embryo and remove all extraembryonic membranes. Exchange the cold sterile 1
PBS dish with a fresh aliquot as needed. Confirm the embryos are at the appropriate developmental stage and, using an embryo spoon, transfer them into a new 10 cm dish containing cold sterile 1
PBS (see Note 18). 4. Remove the forelimb and/or hindlimb buds from each embryo by holding the body with a pair of #5 forceps and using another pair to dissect the limb bud off at the body wall (microdissection scissors can also be used for increased precision). Transfer limbs into a new 10 cm dish containing cold/sterile 1
PBS or 6-well dishes if the embryos must be kept separate (i.e., different genotypes). 5. The following steps are performed in a Class II biosafety cabinet. 6. Using a pipettor with a modified 1 mL pipette tip lacking the first ~0.5 cm of the tip (alternatively could use a wide-bore pipette tip), transfer each limb bud into a limb culture incubation chamber previously prepared and equilibrated to 37  C. If needed, two to three limb buds can be cocultured in one dish if spaced appropriately. Avoid contact between limbs on the same plate as they will fuse together after prolonged time in culture. 7. Place the plates containing limb buds into a well-humidified tissue culture incubator (37  C, 5% CO2) and incubate for 1–6 days. Change media every other day by tilting the plate at a 45 angle and carefully aspirating off media through spaces in the stainless steel mesh using a Pasteur pipette attached to a vacuum source. Gently dispense fresh media into the bottom well using this method. Skeletal elements will become evident 2 days after culture (Fig. 2d), the autopod, stylopod, and zeugopod cartilages are clearly distinguishable by day 4 (see Note 19). 3.3 Whole Mount LacZ Staining

1. Set up timed pregnancies between mice that will yield offspring containing an integrated LacZ reporter gene within a locus of interest or a Cre/FLP-dependent LacZ reporter. Embryos of E11 (can be earlier) to E13 (can be slightly later, but problems with stain penetration) can be used. As above, the morning after the copulatory plug is considered E0.5 days postcoitus. 2. Follow steps 2–7 of Subheading 3.2 to obtain the desired number of limb bud cultures for experimentation. 3. On the desired day postincubation, transfer the PET membranes containing cultured limb buds into a 10 cm dish containing cold 1
PBS. Carefully remove the limbs from the

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membrane using #5 forceps. To do this, bend the membrane so that the limbs lose adherence and gently push the limbs off the membrane and into the 1
PBS dish using another pair of #5 forceps. 4. Using a wide-bore P1000 pipette tip, transfer the limbs into a 12-well plate containing ~1 mL of LacZ fix. Incubate for 60 min at 4  C. 5. Wash limbs 3
10 min with cold 1
PBS. 6. Incubate limbs in LacZ staining solution for 3 h to overnight (see Note 20). 7. To remove excess stain, wash 3
5 min in room temperature 1
PBS. 8. For imaging and storage transfer and maintain in 70% ethanol. 3.4 Whole Mount Immunofluorescence Staining, Tissue Clearing and Imaging

1. Follow steps 1–7 of Subheading 3.2. Retrieve cultured limb buds from incubator at desired time point (see Note 21). 2. Using a 1 ml wide-bore pipette tip, transfer limbs into a 12-well plate. Wash 3
5 min with 1
PBS and fix in 2% PFA for 2–3 days. Tightly cover the topside of the culture plate using Parafilm to reduce loss of solutions due to evaporation. For this purpose, we take a rectangular piece of Parafilm large enough to cover the plate wells and this is covered with the lid so that the Parafilm is sandwiched between the lid and the top of the wells. 3. Wash 3
10 min in 1
PBS. 4. Bleach limbs in 1 ml Dent’s bleach overnight at 4  C. 5. Briefly wash 5
in 100% methanol. 6. Fix in Dent’s fix for 2–3 days (see Note 22). 7. Wash 3
1
PBS for 1 h each, on a rocking platform at room temperature. 8. Block using blocking solution for 1 h at room temperature. 9. Add primary antibodies (in blocking solution) for 1–2 days on a rocking platform at room temperature. 10. Wash 5
1
PBS for 1 h each. Use large volumes (~5 ml) for these washes. 11. Add secondary antibodies (in blocking solution) for 1–2 days on a rocking platform at room temperature. Cover the plate with aluminum foil to protect limbs from light. 12. Rinse 3
with 1
PBS. Followed by 3
1
PBS washes for 1 h each. 13. Replace half of 1
PBS with methanol, mix and let stand for 5 min. 14. Wash with methanol 3
20 min.

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15. Incubate in a 1:1 mixture of methanol and BABB for 10 min. 16. Clear in BABB overnight. 17. Transfer limbs into a receptacle suitable for confocal imaging and resistant to organic solvents (see Note 23). 18. Image using an inverted confocal microscope. Acquire multiple z-stacks to generate 3-D renditions (Fig. 2e) (see Note 24). 3.5 Whole Mount Alcian Blue Staining

1. Follow steps 1–7 of Subheading 3.2. Retrieve cultured limb buds from incubator at desired time point. 2. Rinse 3
with 1
PBS. 3. Transfer to a 12-well plate and fix in formalin for 30 min. 4. Wash 3
10 min with 1
PBS. 5. Wash 1
10 min with 0.2 M HCl. 6. Add stain: 4:1 ratio of 0.2 M HCl and 0.5% Alcian Blue, 1 ml per well is sufficient. 7. Cover the topside of the plate with Parafilm to avoid evaporation and use the plate lid to aid sealing. 8. Stain overnight on a rocking platform at room temperature. 9. To remove excess stain, wash with 70% ethanol. Store longterm in 70% ethanol.

3.6 Lineage Tracing in Whole Mount Limb Bud Cultures

1. To perform lineage tracing, timed pregnancies are arranged between mice containing a Cre or a CreERT2 fusion protein knockin allele under the control of the promoter of a gene of interest and a separate transgenic allele coding for Cre-dependent expression of a reporter protein (i.e., a fluorescent protein). 2. Administer 200–400 mg/kg tamoxifen dissolved in sunflower oil to the pregnant dam via oral gavage at appropriate embryonic time point to allow for maximum recombination efficiency (see Note 25). 3. At E11.0–13.0, sacrifice the pregnant mouse and collect the embryos according to Subheading 3.2. 4. At desired day postcollection, retrieve the cultured limbs and wash 3
10 min with 1
PBS. 5. Carefully transfer limbs to a 12-well plate and fix with 2% PFA for 2–3 days. Tightly cover the topside of the culture plate using Parafilm to reduce loss of solutions due to evaporation. 6. Wash 3
5 min with 1
PBS. 7. Add enough CUBIC clearing reagent into every well to completely submerge the limb buds (see Note 26). Incubate overnight at room temperature on a rocking platform. Cover plate with aluminum foil to avoid photobleaching.

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8. Rinse and store in ScaleA2 clearing solution (see Note 27). 9. To image, transfer limbs into a glass-bottom 35 mm tissue culture dish. Add just enough ScaleA2 clearing solution to completely submerge the limbs. Adding too much solution may cause the limbs to move while imaging (see Note 28). 10. Image cleared limbs in ScaleA2 using an inverted confocal microscope (see Note 29). 3.7 Whole Mount Single and Double In Situ Hybridization

1. This whole mount in situ protocol for detection of one or two transcripts has been adapted from Hargrave et al. [36]. This protocol can also be used with slight modification to detect transcripts in embryos. 2. Riboprobes are generated from ~10 μg linearized DNA templates that contain the gene of interest along with flanking SP6, T3, or T7 transcriptional elements. The template is generated from plasmid DNA using appropriate endonuclease digestion to produce linearized DNAs that will yield sense (control) and anti-sense riboprobes (see Note 30). 3. To generate riboprobes. For a 20 μl reaction add 1 μg of linearized DNA; 2 μl transcription buffer 10
(Roche); 1 mM ATP, GTP, and CTP (each); 0.65 mM UTP; 0.35 mM Digoxigenin-11-UTP (Roche); 20–50 units of RNase inhibitor; and 20 units of the appropriate RNA polymerase (Sp6, T3, or T7) and incubate for 2 h at 37  C. 4. At the end of the reaction, remove 1 μl for quantitation and skip to step 5. To quantify the probe, run the sample (in loading buffer) on a 1% agarose TAE gel along with a DNA ladder (lambda DNA Hind III digest). The amount of riboprobe can be estimated by the intensity of the riboprobe band in comparison to a known quantity of loaded DNA. For these purposes we typically use a HindIII lambda DNA digestion where the riboprobe band can be matched to a band of approximately equal intensity in the ladder. 5. To the completed reaction, add 2 μl 0.2 M EDTA, 2.5 μl LiCl and 75 μl EtOH, precipitate at 20  C for 2 h. 6. Pellet and resuspend pellet in 22.5 μl H2O at 37  C for 30 min, then add 2.5 μl LiCl and 75 μl EtOH and precipitate at 20  C. 7. Pellet and resuspend in 200 μl hybridization buffer. The riboprobe can be stored at 20  C. 8. Prepare timed pregnancies and collect embryo limb buds as described in Subheading 3.2. 9. At desired day postcollection, retrieve the cultured limbs and wash 3
5 min with 1
PBS.

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10. All subsequent steps up until the addition of the antibody should be carried out in DEPC-treated solutions or solutions treated with DEPC. Use RNase-free plasticware. 11. Transfer to a 12-well plate and fix with 4% PFA rocking for 1 h at room temperature to overnight at 4  C. Tightly cover the topside of the culture plate using Parafilm to reduce loss of solutions due to evaporation. 12. Wash in PBS-T, 2 times for 5 min each. 13. Dehydrate through 25%, 50%, 75% and twice 100% methanol in PBS-T, 5 min. For each step. Limb buds can be stored in methanol 100% at 20  C up to several months. 14. Rehydrate through 75%, 50%, 25% methanol in PBS-T and twice in PBS-T, 5 min. Each step. 15. Wash 2–3
in PBS-T for 5 min. 16. Treat with 10 μg/ml proteinase K in PBS-T (at room temperature). As described above, the duration of the proteinase K treatment needs to be optimized and is dependent upon on the limb bud stage and the batch of proteinase K. 17. Wash 2
5 min with PBS-T. 18. Fix in freshly prepared 0.2% glutaraldehyde/4% PFA in PBS-T for 20 min. 19. Wash in PBS-T, 2
5 min. 20. Prehybridize at least 1 h at 65  C with hybridization buffer in a hybridization oven or heating block with rocking or rotating capabilities. 21. Replace with hybridization buffer containing 1 μg/ml digoxygenin- and/or fluorescein-labeled RNA probes. Incubate at 65  C overnight. 22. Following hybridization, a series of posthybridization washes are carried out with varying amounts of Solution 1 and 2
SSC. All hybridizations are carried out at 65  C for 5 min. Each. 23. First wash, 100% Solution 1; second wash 75% Solution 1: 25% 2
SSC; third wash, 50% Solution 1: 50% 2
SSC; and fourth wash, 25% Solution 1: 75% 2
SSC. 24. Wash with 2
SSC, 0.1% CHAPs buffer for 2
30 min at 65  C. 25. Wash with 0.2
SSC, 0.1% CHAPs buffer for 2
30 min at 65  C. 26. Wash 2
10 min with TBST. 27. Incubate embryos in 1% blocking reagent (Sigma-Aldrich) in TBST (or PBS-T) for at least 1 h.

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28. Replace with 1% blocking reagent in TBST (or PBS-T) containing anti-digoxigenin or anti-fluorescein antibody conjugated with alkaline phosphatase (1:1000). Incubate overnight at 4  C. 29. Wash 3
in TBST for 10 min at room temperature. 30. Wash 5
40 min in TBST at rt., longer washes will improve signal-to-noise. 31. Wash 2
20 min in NTMT at rt. 32. To start the color reaction by adding the desired staining solution (purple, light blue or red) to culture wells and incubate in the dark at room temperature (see Notes 31 and 32) for varying periods of time. Monitor staining intensity by checking periodically under low light conditions. Staining can sometimes require an overnight incubation. 33. Wash 3
for 5 min with PBS-T. 34. Postfix the stained limb buds in 4% PFA in 1
PBS overnight at 4  C. This step will also inactivate the alkaline phosphatase enzyme. 35. Wash 4
for 5 min with PBS-T. 36. For detection of the fluorescein-containing riboprobe repeat blocking and washing steps, starting from step 21. 3.8 Bead Implantation

1. Wash Affi-Gel blue gel beads 5
with 1
PBS and store at 4  C until required. 2. Prepare timed pregnancies and collect embryo limb buds as described in Subheading 3.2. 3. In a laminar flow hood, mix an aliquot of gel beads and create several 20 μl spots into a petri dish. Once dry, rehydrate the beads with control and stock solutions of protein(s) of interest. 4. Transfer the beads into microcentrifuge tubes on ice until immediately prior to implantation into the limb (see Note 33). 5. Etch a crosshatch pattern into the center of each dish using a sterile blade, this will aid with adherence of the limb to the bottom of the dish. 6. Place limbs over the etched surface in the appropriate petri dishes for each sample and dispense beads adjacent to the limb on the etched surface. 7. Using a dissection microscope and a pair of Graefe knives, make a small incision into the region of interest in the limb. Adjusting the light source such that the light is semiparallel to the stage will aid visualization of the areas in the limb paddle (at later stages > E12) that are undergoing thickening prior to giving rise to the digits.

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8. Using the Graefe knives, guide the beads into the incision by gently applying pressure to ensure the entry of the beads into the mesenchyme. 9. Transfer beads into limb culture apparatus described in the previous section. 10. Culture limbs at 37  C with 5% CO2 and change medium daily. 11. Once finished the limbs can be processed for visualization to examine the impact of bead implantation on skeletogenesis by examining Alcian Blue staining, cell fate (lineage tracing) or gene expression (WISH or section WISH).

4

Notes 1. The source of material for the scaffolds must be a rustproof and autoclavable wire mesh. We routinely generate scaffolds from stainless steel coffee strainers; however other sources such as kitchen strainers are also acceptable. Using wire cutters, cut strips of ~2.5 cm thickness through the whole length of the wire mesh. Next, cut across each strip at ~2.5 cm intervals to generate 2.5
2.5 cm squares. Store extra cut and uncut materials for future use. 2. The morning of the postcoitum plus is considered embryonic day (E) 0.5. For mouse embryonic limb bud cultures, collect the embryos within E11.0 to E13.0. 3. The method of euthanasia will be dependent on Institutional animal care committee policies and/or guidelines. For these purposes, it is ideal to collect embryos from the uteri as soon as possible after euthanization. 4. For preparation of stock 4% PFA, dissolve electron microscopy grade PFA in PBS on a stirring hot plate at ~60  C and gently stir until the solution becomes clear. Allow the solution to come to room temperature, then aliquot and store at 20  C. As PFA is an irritant and suspected carcinogen, any work involving PFA should be carried out in a fume hood. 5. The LacZ staining solution should always be prepared fresh immediately before use. To avoid the formation of salt crystals, the solution containing all ingredients except X-gal should be warmed to 37  C prior to the addition of X-gal (or other β-galactosidase substrates) and subsequent staining [37]. 6. When deciding on secondary antibodies, it is best to pair the brightest fluorophore to the protein/epitope of lowest abundance and the weaker fluorophores to more abundant targets. This will allow for optimal fluorescence visualization of the proteins of interest after staining.

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7. For visualization of BABB-immersed samples with a confocal microscope, the optical surface should ideally utilize a #1.5 cover glass. 8. Long-wavelength lasers are ideal for imaging whole mount cleared samples as biological specimens do not absorb light as efficiently at longer wavelengths and the low-energy light can penetrate deeper into the sample. Therefore, fluorescent dyes or proteins that absorb and emit light at longer wavelengths (i.e., Alexa 647, mCherry) are ideal for this purpose. 9. Cre and CreERT2 genetic recombination systems are useful tools for generating conditional mutations in mice. For the purposes of lineage tracing, the Cre/LoxP recombination system is simple and effective as the only materials needed are a Cre-transgenic mouse and a floxed-reporter mouse. If temporal control of recombination is required, this can be achieved through the use of a line that contains a Cre fused to a portion of the estrogen receptor (CreERT or newer optimized version CreERT2) encompassing the ligand binding domain. In the absence of tamoxifen, CreER is retained in the cytoplasm and following exposure to 4-hydroxytamoxifen the CreER translocates to the nucleus where it can induce recombination at LoxP sites. The CreERT2 mouse lines have the advantages of spatial and temporal control over reporter gene expression but require an extra amount of expertise with tamoxifen dosing and methods of administration. 10. Determine the optimum working concentration of a new stock of proteinase K by performing parallel experiments using different concentrations for a constant digestion time. If the concentration is too low, the signals from the hybridized probes will be reduced, and if it is too high, the limb buds are visually degraded. 11. For single whole mount in situ hybridization, digoxigeninlabeled riboprobes are preferred and the inclusion of a fluorescein-labeled riboprobe enables detection of an additional transcript. 12. Combinations of the stains have been used for detection in double whole mount in situ hybridization experiments. In whole-limb, the combination of the light blue and purple enables robust detection of two distinct transcripts [38]. Optimization of staining duration may need to be carried out to aid contrast between the two different stains. 13. Limb culture incubation chambers are prepared as described in Barak et al. [39], with some modifications. 14. Each incubation chamber will require one metal mesh scaffold square and can fit two to three forelimbs depending on developmental time point. We don’t recommend placing more than

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three forelimbs in one incubation chamber as the limbs will grow and adhere to each other if placed in close proximity. Instead, create enough metal mesh squares to culture several incubation chambers simultaneously. 15. The slices must be performed in a single rapid motion to avoid crumpling or folding of the membrane. Alternatively, a sterile pair of scissors can also be used to create the slices. It is crucial to avoid crumpling or generating creases while cutting as this will weaken the structural stability of the membrane. 16. To dispense media into the inner-most well without disturbing the wire mesh, tilt the culture plate at a ~35 angle and direct the pipette tip away from the PET membrane. Dispense media slowly through one of the openings created by the space between the wire mesh and the walls of the inner well. Sufficient media has been added once the solution encounters the PET membrane. If the membrane protrudes above the level of the metal mesh, carefully remove media until it is flush with the mesh. 17. In mice, embryonic limb cultures typically utilize limbs from E11 to E13 embryos. We have found that collecting E11.5 forelimbs and culturing them for 2–3 days is ideal for the study of musculoskeletogenesis [38]. However, suitable time points will vary depending on the research question at hand. We have found E11.5 forelimbs to be ideal for the study of development of limb precartilaginous and subsequent cartilaginous elements from the humerus to the distal digit rays. 18. Depending on the nature of the experiment (i.e., different genotypes), a small amount of embryo or extraembryonic membranes can be collected for genotyping. Under these conditions, the limb buds should be processed in separate vessels or wells. 19. Cartilage condensations can be observed as soon as 24 h after culture (Day 1). By day 2, the cartilage anlagen of the humerus, radius, and ulna can begin to be distinguished. The digits become apparent at day 3 and the limb continues to grow past days 4 and 5. Myotubes and tendon primordia can also be observed by the end of day 2. 20. Shorter incubation times are appropriate for E11.5 or Day 1 cultured limb buds, longer incubation times may be necessary for >Day 3 cultured limb buds. Transcription of the LacZ transgene will vary between different genetic constructs; as such, it may also be necessary to adjust incubation times accordingly. 21. This protocol is adapted from the whole mount immunofluorescence protocol available for download at the Kardon lab (University of Utah) website (REF: Lewis and Kardon

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(October 2013). Whole mount antibody staining. Retrieved from http://www.kardonlab.org/protocols. 22. A longer fixation time may be necessary for older limbs. Likewise, for younger/smaller limbs, incubation in Dent’s fixation could be carried out overnight. 23. BABB reacts with plastic surfaces and creates a white precipitate that will impede proper imaging and reduce the quality of the sample. Therefore, imaging must be performed in a metal or glass vessel that has a #1.5 coverslip glass as the optical surface. Several options are available. In our lab, we have two stainlesssteel Aireka Cells 35 mm imaging dishes (Part No. SC15032) that allow us to whole mount image relatively large samples in BABB. These dishes require two 18 mm round coverslips to seal the top and bottom which are not included in the kit. 24. A Nikon A1R HD25 confocal microscope allows us to routinely image day 4–5 whole limb buds at ~250 μm depth. Two of the main limitations in achieving deep imaging are light scattering and light absorption by the tissue. This protocol should allow the limb buds to become mostly transparent and therefore reduce light scattering. To address the issue of light absorption, longer wavelength lasers are recommended as they can achieve greater penetration into the tissue and biological samples do not absorb light as efficiently within a 640–800 nm range. A two-photon microscope may be ideal if available. 25. Tamoxifen dose and route of administration will be variable depending on the CreERT2/LoxP genetic constructs used. We provide a recommended treatment course for a typical CreERT2/LoxP construct. 26. CUBIC (Clear, Unobstructed Brain Imaging Cocktails) [40] clearing solutions are a set of two water-based clearing reagents (reagent-1 and reagent-2) that maintain fluorescent proteins in their native conformation thereby allowing fluorescent signal to be observed without the need of any indirect methods. We have found that the use of CUBIC reagent-1 is sufficient to clear embryonic forelimb buds, making reagent-2 unnecessary. CUBIC clearing as described in this chapter works well for cultured forelimbs; however, other aqueous clearing techniques such as Scale [41], SeeDB [42], and ClearT2 [43] may achieve similar results. 27. Samples can be stored in ScaleA2 solution at 4  C for several weeks. However, it is recommended to image the sample as soon as possible since tissue integrity will be compromised if stored in ScaleA2 for a prolonged period of time. 28. A whole cultured limb can usually be imaged at 4
and/or 10
magnification depending on the field of view capabilities of the microscope. However, if it becomes necessary to acquire

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a large image consisting of several frames, we recommend decreasing the microscope X-Y stage speed to minimal settings. This will help avoid movement of the limb caused by abrupt changes in stage X-Y position. 29. Final imaging results may vary depending on the abundance and brightness of the fluorescent protein being analyzed. Fluorescent signal can be enhanced by following the “whole mount immunofluorescence staining, tissue clearing and imaging” section of this chapter and staining with the appropriate antibody (anti-GFP, anti-RFP, etc.) to increase fluorescent protein signal. 30. Double in situ hybridization is performed by simultaneous hybridization of two different riboprobes (digoxigenin and fluorescein). In both cases, the digoxigenin and fluorescein are detected using antibody Fab fragments conjugated to alkaline phosphatase. To optimize double transcript detection, we usually pair the lower abundance transcript with the digoxigenin-riboprobe and the higher abundance transcript with fluorescein. 31. To increase detection of weak signals color development can be out carried out in 10% polyvinyl alcohol (PVA) [44]. For these purposes, 10% PVA (10 ml solution) is made up as follows: 1 g PVA (Sigma-Aldrich, 18,000–23,000 avg. mol. weight), 200 μl 5 M NaCl, 1 ml 1 M Tris–HCl pH 9.5, and H2O to 9.5 ml. Heat solution to 80o C, mix occasionally by vortexing and continue heating until PVA is fully dissolved (usually takes a couple of hours). Once dissolved cool to room temperature and add 500 μl 1 M MgCl2 and 1 μl Tween-20. 32. There is another AP substrate staining solution from Boehringer (Roche) called BM Purple. Do not use Tween-20 in the buffer if using this stain. 33. To minimize the chance of improper bead implantation, once beads are introduced into the limb bud, that limb bud is moved to a new well.

Acknowledgements This work was supported by the following grants: Canadian Institutes of Health (CIHR) PJT-149026 (T.M.U.) and PJT-148816 (T.M.U.). M.A. was supported by a UBC graduate scholarship.

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References 1. Benazet JD, Zeller R (2009) Vertebrate limb development: moving from classical morphogen gradients to an integrated 4-dimensional patterning system. Cold Spring Harb Perspect Biol 1:a001339 2. Hurle JM, Ros MA, Ganan Y et al (1990) Experimental analysis of the role of ECM in the patterning of the distal tendons of the developing limb bud. Cell Differ Dev 30:97–108 3. Tabin CJ (1991) Retinoids, homeoboxes, and growth factors: toward molecular models for limb development. Cell 66:199–217 4. Tickle C, Eichele G (1994) Vertebrate limb development. Annu Rev Cell Biol 10:121–152 5. Chevallier A, Kieny M, Mauger A (1977) Limb-somite relationship: origin of the limb musculature. J Embryol Exp Morphol 41:245–258 6. Christ B, Jacob HJ, Jacob M (1977) Experimental analysis of the origin of the wing musculature in avian embryos. Anat Embryol (Berl) 150:171–186 7. Lee KK, Sze LY (1993) Role of the brachial somites in the development of the appendicular musculature in rat embryos. Dev Dyn 198:86–96 8. Kardon G, Harfe BD, Tabin CJ (2003) A Tcf4positive mesodermal population provides a prepattern for vertebrate limb muscle patterning. Dev Cell 5:937–944 9. Kardon G (1998) Muscle and tendon morphogenesis in the avian hind limb. Development 125:4019–4032 10. Gaut L, Duprez D (2016) Tendon development and diseases. Wiley Interdiscip Rev Dev Biol 5:5–23 11. Schweitzer R, Chyung JH, Murtaugh LC et al (2001) Analysis of the tendon cell fate using Scleraxis, a specific marker for tendons and ligaments. Development 128:3855–3866 12. Brent AE, Schweitzer R, Tabin CJ (2003) A somitic compartment of tendon progenitors. Cell 113:235–248 13. Kieny M, Chevallier A (1979) Autonomy of tendon development in the embryonic chick wing. J Embryol Exp Morphol 49:153–165 14. Tozer S, Duprez D (2005) Tendon and ligament: development, repair and disease. Birth Defects Res C Embryo Today 75:226–236 15. Chen JW, Galloway JL (2014) The development of zebrafish tendon and ligament progenitors. Development 141:2035–2045

16. Hall BK, Miyake T (2000) All for one and one for all: condensations and the initiation of skeletal development. BioEssays 22:138–147 17. Bi W, Deng JM, Zhang Z et al (1999) Sox9 is required for cartilage formation. Nat Genet 22:85–89 18. Akiyama H, Chaboissier MC, Martin JF et al (2002) The transcription factor Sox9 has essential roles in successive steps of the chondrocyte differentiation pathway and is required for expression of Sox5 and Sox6. Genes Dev 16:2813–2828 19. Lefebvre V, Behringer RR, de Crombrugghe B (2001) L-Sox5, Sox6 and Sox9 control essential steps of the chondrocyte differentiation pathway. Osteoarthr Cartilage 9(Suppl A): S69–S75 20. Smits P, Li P, Mandel J et al (2001) The transcription factors L-Sox5 and Sox6 are essential for cartilage formation. Dev Cell 1:277–290 21. Bell DM, Leung KK, Wheatley SC et al (1997) SOX9 directly regulates the type-II collagen gene. Nat Genet 16:174–178 22. Bridgewater LC, Walker MD, Miller GC et al (2003) Adjacent DNA sequences modulate Sox9 transcriptional activation at paired Sox sites in three chondrocyte-specific enhancer elements. Nucleic Acids Res 31:1541–1553 23. Han Y, Lefebvre V (2008) L-Sox5 and Sox6 drive expression of the aggrecan gene in cartilage by securing binding of Sox9 to a far-upstream enhancer. Mol Cell Biol 28:4999–5013 24. Kronenberg HM (2006) PTHrP and skeletal development. Ann N Y Acad Sci 1068:1–13 25. Lefebvre V, Smits P (2005) Transcriptional control of chondrocyte fate and differentiation. Birth Defects Res C Embryo Today 75:200–212 26. Gerstenfeld LC, Shapiro FD (1996) Expression of bone-specific genes by hypertrophic chondrocytes: implication of the complex functions of the hypertrophic chondrocyte during endochondral bone development. J Cell Biochem 62:1–9 27. Aghajanian P, Mohan S (2018) The art of building bone: emerging role of chondrocyteto-osteoblast transdifferentiation in endochondral ossification. Bone Res 6:19 28. Mizuhashi K, Ono W, Matsushita Y et al (2018) Resting zone of the growth plate houses a unique class of skeletal stem cells. Nature 563:254–258 29. Yang L, Tsang KY, Tang HC et al (2014) Hypertrophic chondrocytes can become

Limb Bud Organ Culture osteoblasts and osteocytes in endochondral bone formation. Proc Natl Acad Sci U S A 111:12097–12102 30. Friedman L (1987) Teratological research using in vitro systems. II. Rodent limb bud culture system. Environ Health Perspect 72:211–219 31. Underhill TM, Dranse HJ, Hoffman LM (2014) Analysis of chondrogenesis using micromass cultures of limb mesenchyme. Methods Mol Biol 1130:251–265 32. Curtin P, Youm H, Salih E (2012) Threedimensional cancer-bone metastasis model using ex-vivo co-cultures of live calvarial bones and cancer cells. Biomaterials 33:1065–1078 33. Sloan AJ, Taylor SY, Smith EL (2012) Organotypic mandibular cultures for the study of inflammatory bone pathology. In: Davies J (ed) Replacing animal models. Wiley, New York, pp 159–166 34. Smith EL, Kanczler JM, Oreffo RO (2013) A new take on an old story: chick limb organ culture for skeletal niche development and regenerative medicine evaluation. Eur Cell Mater 26:91–106. discussion 106 35. Neubert D, Barrach H-J (1977) Techniques applicable to study morphogenetic differentiation of limb buds in organ culture. In: Neubert D, Merker H-J, Kwasigroch TE (eds) Methods in prenatal toxicology. Georg Thieme Publishers, Stuttgart, pp 241–251 36. Hargrave M, Bowles J, Koopman P (2006) In situ hybridization of whole-mount embryos. Methods Mol Biol 326:103–113

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37. Scott RW, Underhill TM (2016) Methods and strategies for lineage tracing of mesenchymal progenitor cells. Methods Mol Biol 1416:171–203 38. Hoffman LM, Garcha K, Karamboulas K et al (2006) BMP action in skeletogenesis involves attenuation of retinoid signaling. J Cell Biol 174:101–113 39. Barak H, Boyle SC (2011) Organ culture and immunostaining of mouse embryonic kidneys. Cold Spring Harb Protoc 2011:pdb.prot5558 40. Gomez-Gaviro MV, Balaban E, Bocancea D et al (2017) Optimized CUBIC protocol for three-dimensional imaging of chicken embryos at single-cell resolution. Development 144:2092–2097 41. Hama H, Kurokawa H, Kawano H et al (2011) Scale: a chemical approach for fluorescence imaging and reconstruction of transparent mouse brain. Nat Neurosci 14:1481–1488 42. Ke MT, Fujimoto S, Imai T (2013) SeeDB: a simple and morphology-preserving optical clearing agent for neuronal circuit reconstruction. Nat Neurosci 16:1154–1161 43. Kuwajima T, Sitko AA, Bhansali P et al (2013) ClearT: a detergent- and solvent-free clearing method for neuronal and non-neuronal tissue. Development 140:1364–1368 44. Barth J, Ivarie R (1994) Polyvinyl alcohol enhances detection of low abundance transcripts in early stage quail embryos in a nonradioactive whole mount in situ hybridization technique. BioTechniques 17(324):326–327

Chapter 9 Murine Limb Explant Cultures to Assess Cartilage Development Manuela Wuelling and Andrea Vortkamp Abstract To investigate chondrocyte biology in an organized structure, limb explant cultures have been established that allow for the cultivation of the entire cartilaginous skeletal elements. In these organ cultures, the arrangement of chondrocytes in the cartilage elements and their interaction with the surrounding perichondrium and joint tissue are maintained. Chondrocyte proliferation and differentiation can thus be studied under nearly in vivo conditions. Growth factors and other soluble agents can be administered to the explants and their effect on limb morphogenesis, gene expression and cell–matrix interactions can be studied. Cotreatment with distinct growth factors and their inhibitors as well as the use of transgenic mice will allow one to decipher the epistatic relationship between different signaling systems and other regulators of chondrocyte differentiation. Here we describe the protocol to culture cartilage explants ex vivo and discuss the advantages and disadvantages of the culture system. Key words Organ culture, Cartilage explant culture, Ex vivo analysis, Chondrocyte differentiation, Bone, Chondrocyte

1

Introduction The complex physiological mechanisms regulating endochondral ossification and growth plate formation are extremely difficult to study in vivo. Different approaches are thus required, each having specific advantages and disadvantages. For most investigations living mice would be preferable, and the use of transgenic mouse lines has given considerable insight into the mechanisms regulating chondrocyte proliferation and differentiation. However, such in vivo investigations are cost intensive and time consuming. Furthermore, the role of an individual gene at the relatively late stages of chondrocyte differentiation is often masked by its role at earlier developmental stages. For a more direct investigation, various cell culture systems have been established in which chondrocytes can be treated with different growth factors and other soluble agents, or can be transfected with overexpression constructs and—more

Matthew J. Hilton (ed.), Skeletal Development and Repair: Methods and Protocols, Methods in Molecular Biology, vol. 2230, https://doi.org/10.1007/978-1-0716-1028-2_9, © Springer Science+Business Media, LLC, part of Springer Nature 2021

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recently—with siRNA or shRNA expressing vectors. In these systems, chondrocyte differentiation can be manipulated at different time points allowing the investigation of a certain signaling system in the isolated cells at a specific step of differentiation. Although many important questions can be addressed a major disadvantage of cell culture systems is the lack of interaction between different cell types and tissues in an organized structure that takes place in the developing skeletal anlagen in vivo. To investigate the reaction of chondrocytes, maintained in an organized structure, to soluble factors, organ culture systems have been established that permit the cultivation and analysis of the embryonic cartilaginous skeletal anlagen with the surrounding perichondrium [1]. In these ex vivo cultures, the integrity of the tissue and the arrangement of cells in typical regions of round (low proliferating), columnar (high proliferating) and hypertrophic (nonproliferating) chondrocytes as well as their interaction with the flanking perichondrium are maintained. Cells can thus communicate similarly as they do in vivo, at least for a restricted period of time. Alterations in the cartilaginous structures under different culture conditions can then be investigated during the culture period. Initial experiments with chick cartilage organ cultures were already reported in 1926 [2] and chicken limb or femur organ culture is still an attractive model system [3]. The first mammalian cartilage explant cultures were established from rat and rabbit limb tissue in 1970 [4, 5]. Since then, the conditions of cartilage explant cultures were modified in many ways. Today, the commonly used culture media are based on the so-called BGJ medium described by Biggers et al. in 1961. One of the most important advances of this medium is the replacement of serum by chemically defined nutrition and salt components [6, 7]. As serum contains many growth factors in undefined concentrations, only the use of a serum free medium allows the reproducible investigation of growth factors under defined conditions. Growth factors and other agents can be added to the serum free medium as single components or in combination with other factors or inhibitors to investigate their specific role during chondrocytes differentiation as well as their interaction and epistatic relationship with other signaling systems [8–10]. The differentiation of the skeletal elements can be assessed on a morphological level by measuring width and length over time in culture (Fig. 1). Furthermore, the differentiation process can be monitored on a cellular level by analyzing changes in the respective chondrocyte populations after sectioning. These can be defined based on their morphology after staining or, in more sophisticated approaches, by analyzing the expression of characteristic chondrocyte markers either by mRNA in situ hybridization or by immunodetection of the respective proteins [11–14] (see other chapters for details).

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Fig. 1 The elongation of the skeletal elements can be measured during culture and the effect of growth factors and their respective inhibitors, as shown for BMP2 and Noggin treatment, can be directly evaluated in the cultures

Today, the most widely used cartilage explant cultures are either forelimbs isolated from mouse embryos between embryonic day 13.5 (E13.5) and E18.5 or mouse hindlimb metatarsals isolated between E16.5 and postnatal day 3 (P3). Similar to metatarsal cultures, explant cultures of other single skeletal elements, like tibia or femur, have successfully been established [15]. To choose the right object for the experiment, one has to consider that cultures of single skeletal elements and whole limb explants offer distinct advantages. While metatarsal cultures are less affected by distortion during the time in culture, whole limb cartilage explants permit the simultaneous assessment of differently advanced skeletal elements as the individual cartilage anlagen develop from proximal to distal. Additionally, the joint regions are maintained in these cultures and effects of the investigated factors on the differentiation of the joint structure can be studied in parallel. On the other hand, cultures of the complete forelimb skeleton are more difficult to handle, as the longer cartilage structures have the tendency to bend in culture. In both systems the cartilage explants can successfully be cultured for several days [9, 13], theoretically for up to at least 2 weeks [16, 17]. One has, however, to keep in mind that vascularization and ossification, which are important steps during normal development, will not be maintained under culture conditions. Therefore, the analysis of the cultures is restricted to the proliferation and differentiation of chondrocytes until late hypertrophy [13], whereas cartilage matrix degradation in terminal hypertrophic cells and ossification cannot be monitored. As a consequence of this limitation, the zone of hypertrophic chondrocytes will

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expand over time leading to the elongation, but also thinning of the hypertrophic zone [9, 18], which in its center will ultimately (after long culture periods) consist of dead cells. Interestingly, a recent study described the formation of calcified zones in limb cultures resembling the mineralized cartilage matrix after supplementation of the cultures with ascorbic acid, ß-glycerol phosphate, and calcium [3]. These agents have been reported to induce mineralization of mesenchymal progenitor cells in vitro [19] and seem to be sufficient to induce mineralization in organ cultures. The calcified areas were accompanied by expression of the osteogenic marker Collagen Type 1 in chondrocytes [20]. Recent studies provide evidence that chondrocytes contain a pool of skeletal stem cells or can transdifferentiate to form osteoblasts [21, 22]. The induction of mineralization in limb cultures might provide a model system to track this differentiation process ex vivo. However, even under adequate culture conditions, the length of the cultures is limited and the developmental progression will slow down with culture age. This retardation can be visualized by labeling proliferating cells with bromodeoxyuridine (BrdU) for a defined period before ending the culture. The number of labeled, proliferating cells will decrease gradually with the age of the culture [8, 9, 18]. Nevertheless, in spite of these limitations, limb cartilage explant cultures provide an excellent method to study the orderly progression of chondrocyte subpopulations for a limited time under nearly in vivo conditions. As a rule of thumb 4 days in culture are optimal to observe differences in total size and the relative contribution of distinct chondrocyte populations, whereas 2 days are ideal to investigate treatment dependent changes in proliferation and differentiation on a molecular level, for example, by mRNA in situ hybridization or immunodetection of differentiation markers. For mineralization, at least 5 days of culture are necessary and the culture period can be extended up to 10 days. Using these conditions, limb organ cultures treated with agonist and antagonist of different signaling systems have given important insights into the epistatic interactions of signaling systems during embryonic chondrocyte differentiation, especially if explants of transgenic animals were included in the investigations ([23] and others). Table 1 gives an overview of the used growth factors or inhibitors tested in limb and metatarsal culture studies.

2 2.1

Materials Mice

1. Timed pregnancies are set up at 1:1 ratio between females and males in a 12 h light-dark cycle. Female mice are examined for evidence of successful copulation determined by vaginal plugs each morning between 8:00 and 9:00 AM. The day of finding

10

1–100

[27]

[9]

[27]

7 days

1

[9]

[8]

[28]

[28]

6 days

10

48–96 h

10 days

50

48–96 h [8]

[28, 29]

8 days

0.1–1

48–96 h

[29]

10 days

100

[8]

48–96 h

[30]

3 days

50

[8]

48 h

[30]

3 days

50

[24]

96 h

[25]

48 h

The concentration used and duration of the treatment is given. If not stated specifically, the concentration is applicable for developmental stages from E14.5 to E16.5 [8, 9, 31]

[26]

References

5

7 days

100–500

Treatment time 4 days

Concentration

Metatarsal culture

[8]

48–96 h

BMP (ng/ml) Cyclopamine (μM) SAG (μM) IGF (ng/ml) LY 294002 (μM) CNP (μM) IGF (ng/ml) Insulin (nM) Rapamycin (μM)

7

HS (μg/ ml)

Treatment

3  10

TGFß (ng/ml)

[9]

10

Su5402 (μg/ PTHrP ml) (M)

References

50

Fgf2 E16. 5 (ng/ml)

12–48 h

250

Fgf2 E14.5 (ng/ml)

12–48 h

100

Noggin (ng/ml)

Treatment time

500

BMP2 (ng/ml)

10

Shh (μg/ Cyclopamine ml) (μM)

Concentration 5

Limb culture

Treatment

Table 1 Overview of growth factors and their respective inhibitors tested in limb explant cultures

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the plug is designated E0.5 of embryonic development. Pregnant mice are sacrificed by cervical dislocation at the desired stages (see Notes 1 and 2). 2.2

Supplies

1. Two fine forceps such as Dumont #55 (Fine Science Tools), cleaned and disinfected to prevent contamination of the cultures. 2. 10 cm petri dishes. 3. Two-chambered “in vitro fertilization dishes” (Falcon) or, alternatively, “trans-well” cell culture plates of other providers (see Note 3). 4. Triangular or rectangular metal grids, cell culture grade, autoclaved (see Note 4). 5. 0.8 μm nitrocellulose filter (Millipore), cut into suitable pieces.

2.3 Reagents and Media

All solutions should be prepared in sterile deionized water and sterile filtered through a 0.2 μm filter prior to use. All cell culture plastic ware should be single-use. All chemicals and reagents should be obtained as cell culture grade. 1. 1 phosphate buffered saline (1PBS), pH 7.4 (Invitrogen): 1.06 mM KH2PO4, 155 mM NaCl, 2.97 mM Na2HPO4 + 7H2O. 2. Antibiotic/Antimycotic Stock Solution (Invitrogen): 100 Stock Solution containing 10,000 units/ml penicillin G, 10 mg/ml streptomycin sulfate, and 25 μg/ml amphotericin B. 3. 1PBS with Antibiotic/Antimycotic: Add 1 ml of the 100 Antibiotic/Antimycotic stock solution to 100 ml 1PBS. 4. 100 ml Biggers Medium (ICN Biomedicals see Note 5), 0.1% bovine serum albumin (BSA), 700 μl 200 mM L-glutamine stock solution, and 1 antibiotic/antimycotic stock solution (Invitrogen, see Note 6). For metatarsal cultures, DMEM-F12 medium has been described supplemented with 0.2% bovine serum albumin, 5 μg/ml L-ascorbic acid phosphate, 1 mM β-glycerophosphate, 0.05 mg/ml gentamicin, and 1.25 μg/ ml fungizone [32]. 5. For the induction of mineralization of cartilage matrix, 10% FCS, 10 mM CaCl2, 10 mM ß-glycerol phosphate, and 50 μg/ ml ascorbic acid can be supplemented [20].

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3 3.1

145

Methods Limb Preparation

The subsequent steps are conducted under a dissection microscope at room temperature in sterile 1PBS. 1. Mouse embryos are dissected from the uterus of time-pregnant females in sterile 1PBS in petri dishes. 2. Embryos are transferred to fresh 1PBS in clean, sterile petri dishes (see Note 7). 3. Limbs are removed from the torso next to the shoulder blade (see Note 8). 4. Using two fine forceps, the skin and the surrounding soft tissue is carefully removed from the skeletal elements (see Notes 9– 14). 5. For whole limb cultures, care should be taken to leave the joints intact. For single skeletal elements, excess connective tissue should be removed (see Note 15). 6. If limb cultures will be treated with growth factors or inhibitors, the prepared limbs should be collected in pairs to allow pair-wise comparison of the treated and untreated cartilage elements. 7. Each limb explant should be numbered and measured prior to culture to follow the individual growth.

3.2

Explant Culture

Assemble the in vitro culture dish (Falcon) as shown in Fig. 2: 1. Triangular metal grids are centered in the middle well of the in vitro fertilization dish [33].

Fig. 2 Experimental setup of the limb culture system showing the assembly of the limb on the nylon membrane on top of the metal grid, which is inserted in the inner well of an “in vitro fertilization” dish. The limb is cultured at the interface between air and media

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2. A 0.8 μm nitrocellulose filter is positioned in the center of the metal grid. 3. The outer rim of the culture dish is filled with 5 ml sterile 1 PBS with Antibiotic/Antimycotic to maintain humidity in the culture dish. 4. The inner well is filled with 1 ml limb culture medium (Biggers Medium). 5. Using forceps, the skeletal elements are carefully transferred onto the nylon filter in the culture dish preferably with the palm facing up and incubated at the liquid–air interface (see Note 16). 6. Cultures are grown in a humid, 37  C incubator under an atmosphere of 5% CO2 (see Note 17). 7. The culture media needs to be replaced every day; the 1 PBS in the outer rim needs not be replaced (see Note 18). 8. Limbs can successfully be cultured for at least 5 days (see Note 19).

4

Notes 1. The chosen embryonic stage is defined by the scientific question and can range from E13.5 to P3. Note that the result of the experiment is dependent on the similarity of the developmental stage of the embryos used. The initial size and cellular compositions of the skeletal elements greatly varies between litters and stages. Therefore, the age and the stage of the developing limbs should be carefully determined and compared prior to culture. 2. For the analysis of growth factors and other soluble agents, the left and right limb of an embryo should be compared as treated and nontreated to reduce the biological variability of the experiment. 3. As an alternative to the described “in vitro fertilization” dishes, “transwell” cell culture plates can be used for cultivation. In that case, the nylon membrane must be placed on the transwell inset. It is important to fill the well with sufficient culture medium to allow culturing at the liquid–air interface. 4. The metal grids can be cleaned and reused after sterilization. 5. Biggers Medium is also sold as BGJb medium by other companies. Both media can be obtained supplemented with Lglutamine. Alternatively, DMEM/Ham12 medium with supplementation of 10% fetal calf serum (FCS) has been described for cultivation up to 10 days [20]. The addition of FCS has several

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disadvantages. FCS contains undefined amounts of several growth factors which can affect culture conditions and are highly variable between suppliers and lot numbers. If defined concentrations of growth factors or hormones should be tested in the cultures, addition of FCS might mask the intended effects. 6. Alternatively, each solution can be added to the final concentration of 100 U/ml penicillin, 100 μg/ml streptomycin, and 250 ng/ml Fungizone. 7. All embryos should be dissected out of the uterus and transferred to a new petri dish without blood or dissected tissues. 8. The shoulder blade can stay attached during preparation to facilitate handling of the skeletal elements, but it should be removed prior to culture to prevent bending. 9. The joints, particularly the elbow joints, should stay intact, to support stability of the skeletal elements. 10. The tendons at the wrist should be dissected to prevent bending at wrist level. 11. The skin surrounding the digits should be removed by opening the skin at the palm and carefully shifting it to the outside from proximal to distal. 12. The perichondrium should stay intact; avoid over dissection of the supporting muscle and tendons to prevent damage to the perichondrium. 13. Residual mesenchymal tissue surrounding the skeletal elements, the elbow joints and the metacarpals will eventually degrade during culture. 14. Keeping the limbs at room-temperature during dissection is preferable over cooling them on ice. 15. For the preparation of metatarsal cultures, an instructional video is available online [34]. 16. Make sure the limbs are completely placed on the nitrocellulose filter, as they may stick to the metal grid. 17. During culture, the residual mesenchymal cells tend to stick to the supporting nylon filter. To prevent distortion of the skeletal elements during growth, the limbs should carefully be lifted from the filter membrane once a day. Especially if regions of chondrocyte subpopulations shall be measured, the straight growth of the limbs is essential to allow sectioning along the longitudinal axis of the skeletal elements. Non parallel sections will disturb precise measurements of expression domains. 18. To follow elongation of the cartilage elements, the length of the limbs or the respective bone should be measured on a daily basis.

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19. If desired, growth factors or inhibitors can be administered to the culture medium [8, 9, 12, 18, 31]. To avoid nonphysiological effects the minimal effective concentration should be determined before the experiment. Growth factors and inhibitors need to be replaced daily. Table 1 shows a list of growth factors tested and the concentrations used.

Acknowledgments This work was supported by grants from the BMBF (01EC1408E) and the DFG (FOR2407; Vo620/14-1). References 1. Friedman L (1987) Teratological research using in vitro systems. II. Rodent limb bud culture system. Environ Health Perspect 72:211–219 2. Strangeways TSP, Fell GB (1926) Experimental studies on the differnetiation od embryonic tissues growing in vivo and in vitro. I the development of the undifferentiated limb-bud (a) when subcutaneously grafted into the pos-embryonic chick and (b) when cultivated in vitro. Proc R Soc Lond B Biol Sci B99:340–366 3. Smith EL, Kanczler JM, Oreffo RO (2013) A new take on an old story: chick limb organ culture for skeletal niche development and regenerative medicine evaluation. Eur Cell Mater 26:91–106; discussion 106 4. Shepard TH, Bass GL (1970) Organ culture of limb buds from riboflavin-deficient and normal rat embryos in normal and riboflavin-deficient media. Teratology 3(2):163–167 5. Shepard TH, Bass GL (1971) Organ-culture studies of achondroplastic rabbit cartilage: evidence for a metabolic defect in glucose utilization. J Embryol Exp Morphol 25(3):347–363 6. Biggers JD, Gwatkin RB, Heyner S (1961) Growth of embryonic avian and mammalian tibiae on a relatively simple chemically defined medium. Exp Cell Res 25:41–58 7. Biggers JD, Heyner S (1961) Studies on the amino acid requirements of cartilaginous long bone rudiments in vitro. J Exp Zool 147:95–111 8. Minina E et al (2002) Interaction of FGF, Ihh/Pthlh, and BMP signaling integrates chondrocyte proliferation and hypertrophic differentiation. Dev Cell 3(3):439–449 9. Minina E et al (2001) BMP and Ihh/PTHrP signaling interact to coordinate chondrocyte

proliferation and differentiation. Development 128(22):4523–4534 10. Sahni M et al (1999) FGF signaling inhibits chondrocyte proliferation and regulates bone development through the STAT-1 pathway. Genes Dev 13(11):1361–1366 11. Andrade AC et al (2011) Methods to study cartilage and bone development. Endocr Dev 21:52–66 12. Hung IH et al (2007) FGF9 regulates early hypertrophic chondrocyte differentiation and skeletal vascularization in the developing stylopod. Dev Biol 307(2):300–313 13. Mak KK et al (2008) Indian hedgehog signals independently of PTHrP to promote chondrocyte hypertrophy. Development 135 (11):1947–1956 14. Serra R, Karaplis A, Sohn P (1999) Parathyroid hormone-related peptide (PTHrP)-dependent and -independent effects of transforming growth factor beta (TGF-beta) on endochondral bone formation. J Cell Biol 145 (4):783–794 15. Guo J et al (2006) PTH/PTHrP receptor delays chondrocyte hypertrophy via both Runx2-dependent and -independent pathways. Dev Biol 292(1):116–128 16. Klement BJ, Spooner BS (1992) Endochondral bone formation in embryonic mouse pre-metatarsals. Trans Kans Acad Sci 95 (1-2):39–44 17. Klement BJ, Spooner BS (1993) Embryonic mouse pre-metatarsal development in organ culture. J Exp Zool 265(3):285–294 18. Mau E et al (2007) PTHrP regulates growth plate chondrocyte differentiation and proliferation in a Gli3 dependent manner utilizing hedgehog ligand dependent and independent mechanisms. Dev Biol 305(1):28–39

Murine Limb Explant Cultures 19. Fiorentini E et al (2011) Effects of osteogenic differentiation inducers on in vitro expanded adult mesenchymal stromal cells. Int J Artif Organs 34(10):998–1011 20. Masuda E et al (2015) A newly established culture method highlights regulatory roles of retinoic acid on morphogenesis and calcification of mammalian limb cartilage. BioTechniques 58(6):318–324 21. Wuelling M, Vortkamp A (2019) A newly discovered stem cell that keeps bones growing. Nature 567(7747):178–179 22. Zhou X et al (2014) Chondrocytes transdifferentiate into osteoblasts in endochondral bone during development, postnatal growth and fracture healing in mice. PLoS Genet 10(12): e1004820 23. Vortkamp A (2000) The Indian hedgehog-PTHrP system in bone development. Ernst Schering Res Found Workshop 29:191–209 24. Bouche M et al (1995) Rapid activation and down-regulation of protein kinase C alpha in 12-O-Tetradecanoylphorbol-13-acetateinduced differentiation of human rhabdomyosarcoma cells. Cell Growth Differ 6 (7):845–852 25. Koziel L et al (2004) Ext1-dependent heparan sulfate regulates the range of Ihh signaling during endochondral ossification. Dev Cell 6 (6):801–813 26. Oichi T et al (2019) Adamts17 is involved in skeletogenesis through modulation of BMP-Smad1/5/8 pathway. Cell Mol Life Sci 76(23):4795–4809

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27. Hojo H et al (2013) Hedgehog-Gli activators direct osteo-chondrogenic function of bone morphogenetic protein toward osteogenesis in the perichondrium. J Biol Chem 288 (14):9924–9932 28. Ulici V et al (2010) Regulation of gene expression by PI3K in mouse growth plate chondrocytes. PLoS One 5(1):e8866 29. Mushtaq T et al (2004) Insulin-like growth factor-I augments chondrocyte hypertrophy and reverses glucocorticoid-mediated growth retardation in fetal mice metatarsal cultures. Endocrinology 145(5):2478–2486 30. Phornphutkul C et al (2008) mTOR signaling contributes to chondrocyte differentiation. Dev Dyn 237(3):702–712 31. Koziel L et al (2005) Gli3 acts as a repressor downstream of Ihh in regulating two distinct steps of chondrocyte differentiation. Development 132(23):5249–5260 32. Dobie R et al (2015) Increased linear bone growth by GH in the absence of SOCS2 is independent of IGF-1. J Cell Physiol 230 (11):2796–2806 33. Trowell OA (1954) A modified technique for organ culture in vitro. Exp Cell Res 6 (1):246–248 34. Houston DA et al (2016) Culture of murine embryonic metatarsals: a physiological model of endochondral ossification. J Vis Exp 118:54978

Chapter 10 Renal Capsule Transplantation to Assay Angiogenesis in Skeletal Development and Repair Anais Julien, Simon Perrin, Rana Abou-Khalil, and Ce´line Colnot Abstract Renal capsule transplantation is a very helpful method to grow embryonic tissues or tumors in a vascular environment, allowing for long-term engraftment and biological analyses. This chapter describes the surgical procedure for the transplantation of embryonic skeletal elements in the renal capsule of adult mice and points out the manipulations that can be applied for assaying the role of angiogenesis during bone development and repair. Key words Skeletal development, Skeletal repair, Angiogenesis, Renal capsule transplantation

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Introduction Bone is a highly vascularized tissue with tight connections between blood vessels, bone marrow, and bone cells to maintain skeletal integrity. Angiogenesis plays a pivotal role in skeletal development and particularly during endochondral ossification as an angiogenic switch is required for the replacement of cartilage by bone marrow and bone [1–3]. Numerous tools and methodologies have been used to study the impact of angiogenesis on osteogenesis both in vitro and in vivo [4–8]. Although in vitro angiogenesis assays have provided direct evidence for bidirectional interactions between osteoblasts and endothelial cells, which are crucial for osteogenesis, other cell types, circulating factors, and extracellular matrix proteins are involved in bone vascularization. Therefore, in vivo angiogenic assays are also essential to study the role of supporting cells (smooth muscle cells, pericytes, and fibroblastic cells) and other factors in the tissue environment. Moreover, in vitro assays do not allow the development of the hematopoietic compartment of bone, which is required for establishing the stromal compartment of bone and providing osteoclasts that are together necessary for bone formation and remodeling. Since the

Matthew J. Hilton (ed.), Skeletal Development and Repair: Methods and Protocols, Methods in Molecular Biology, vol. 2230, https://doi.org/10.1007/978-1-0716-1028-2_10, © Springer Science+Business Media, LLC, part of Springer Nature 2021

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kidney is one of the most vascularized organs in the body, the renal capsule constitutes a permissive environment to grow cells, tumors, or embryonic tissues [9–13]. The renal capsule of adult mice has been used as a host environment to dissect the role of angiogenesis in skeletal development [13, 14]. With the growing number of genetically modified mouse models, this approach can help distinguish the effects of specific gene mutations in skeletal tissues versus blood vessels and their impact on angiogenesis and subsequent bone development [15, 16]. Any skeletal element from the developing embryo can potentially be collected prior to its vascularization in vivo and transplanted in the adult host renal capsule. More recently, this model was used to grow human embryonic skeletal elements [17]. Vascularization of the grafts occurs within 3 days, and the renal capsule environment can support normal bone development and growth, including establishment of the bone marrow, cortical bone, and surrounding periosteum (Fig. 1). Finally, skeletal stem/progenitors within bone marrow and periosteum can be isolated from long bones grown in the renal capsule and mobilized to repair bone after skeletal injury thus extending the use of this system to study bone repair mechanisms (Fig. 2) [18].

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Materials

2.1 Anesthetics and Analgesic

1. Anesthetics: Prepare the solution of ketamine–medetomidine by mixing 1 volume of ketamine with 1 volume of medetomidine. 2. Anesthetic reversal solution: Atipamezole comes as a ready to use reagent. 3. Analgesics: Prepare the solution of buprenorphine in NaCl 0.09%.

2.2 Isolation of E14– E14.5 Mouse Femora

1. Pregnant female mice with embryos at E14–E14.5 (see Note 1). 2. Surgical instruments (Fine forceps Dumont #5 and #55, scissors, Fine Science Tools). 3. Ice-cold phosphate buffer saline solution (1 PBS). 4. 70% ethanol. 5. 24-well plate. 6. Petri dish 100 mm diameter. 7. Binocular microscope.

2.3 Renal Capsule Transplantation

1. 24-well plate. 2. Insulin micro-fine syringe (30G). 3. Wound clipper.

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Fig. 1 Steps of the surgical procedure and development of skeletal elements in the renal capsule. (a) Anesthetized host mouse prior to transplantation; note the

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Fig. 2 Steps of the surgical procedure and repair after cortical defect or fracture in long bones grown under the renal capsule. (a–d) Steps of cortical defect repair and (e–h) steps of fracture repair. (a, e) Exteriorized kidney 6 weeks posttransplantation of an E14.5 femoral skeletal element. (b, f) Cortical defect (b) and bone fracture (f) at the time of procedure (white arrows point to the injury site). (c, g) Healing cortical defect (c) and bone fracture (g) 2 weeks after the procedure (white arrows indicate the repair site). (d, h) Longitudinal sections of cortical defect (d) and fracture (h) stained with Safranin-O showing a fully ossified callus (orange dotted line) composed of newly formed bone (black arrows) ä Fig. 1 (continued) position of the skin incision. (b) Exteriorized kidney posttransplantation of one E14.5 femoral skeletal element (denoted by a white dotted line). The white arrow indicates the incision in the renal capsule. (c) Femoral skeletal elements at days 0 (d0), 5 (d5), 7 (d7), and 60 (d60) posttransplantation. By day 60, the skeletal element is fully ossified and has grown to reach almost the size of a two-month-old mouse femur (approximately 1 cm in length). (d) PECAM immunostaining reveals blood vessels (black arrows) on longitudinal sections of femoral skeletal elements at d0 (E14.0), d5, and d7 posttransplantation. At the time of transplantation (d0), the perichondrium (pc) is vascularized but not the cartilage (c). The cartilage becomes vascularized by day 5 to form the primary ossification center. The bone marrow (bm) and periosteum (po) are well developed by day 7; the epiphyseal cartilage (c) is not yet invaded by blood vessels to form the secondary ossification center

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4. Male mice (8–12 weeks old) (see Note 1). 5. Pregnant mouse female (E14–E14.5). 6. Binocular microscope. 7. Betadine soap and Betadine solution. 8. Cotton swab. 9. 1 PBS. 10. Plastic Pasteur pipette. 11. “L”-shape glass rod (Home made): Using a fire, separate the narrow end of a glass Pasteur pipette (4–5 cm in length); make a thin “L”-shaped glass rod with a rounded closed end of approximately 1 mm in diameter. 12. Surgical instruments (Fine forceps, Fine Vanna Scissors, tweezers, hemostatic forceps, scissors) (Fine Science Tools) (see Note 2). 13. 4-0 absorbable sutures. 14. Clips and wound clipper. 2.4 Bone Fracture and Cortical Defect in the Renal Capsule

1. Wound clipper. 2. Binocular microscope. 3. Betadine soap and Betadine solution. 4. Surgical instruments (Fine forceps, Fine Vanna Scissors, tweezers, scissors) (Fine Science Tools) (see Note 2). 5. Drill with 0.8 mm drill bit. 6. 4-0 absorbable sutures. 7. Clips and wound clipper.

2.5 Analysis of Vascularization and Angiogenesis

1. Glass jar. 2. 4% paraformaldehyde (PFA) fixative solution. 3. 0.5 M ethylenediaminetetraacetic acid (EDTA) pH 7.4. 4. 70% ethanol. 5. 95% ethanol. 6. 100% ethanol. 7. Xylene. 8. Paraffin. 9. Superfrost microscope slides. 10. Histoclear. 11. 1 PBS. 12. Rotary microtome. 13. Deionized water.

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14. Hydrophobic pen. 15. Hydrogen peroxidase (H2O2). 16. Methanol. 17. Ficin solution. 18. Glycine. 19. Ovalbumin. 20. Nonfat powdered milk. 21. Normal goat serum. 22. Rat anti-PECAM primary antibody (BD Biosciences). 23. Goat biotinylated (BD Biosciences). 24. Horseradish peroxidase (BD Biosciences).

anti-Rat

secondary

(HRP)-conjugated

antibody Streptavidin

25. Diaminobenzidine (DAB): Prepare working solution according to supplier manual (Life Technologies). 26. Fast Green. 27. Permount. 28. Cover slides.

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3.1 Isolation of Mouse Embryonic Femora

Prepare the cartilage grafts by finely dissecting the skeletal elements of E14-E14.5 mouse embryos using 2 pairs of fine forceps (see Note 3). 1. Sacrifice pregnant mouse by cervical dislocation under anesthesia (IP injection of ketamine–medetomidine: 50 mg of ketamine and 0.5 mg of medetomidine per kg of body weight) and position the mouse in a supine posture. 2. Soak the abdomen with 70% ethanol, and make a small incision at the midline. Continue with a V-shaped incision through the skin and pull the skin toward the head to expose the abdomen. 3. Cut the peritoneum to expose the abdominal cavity. 4. Locate the 2 uterine horns, the uterus and oviduct in the dorsal region of the abdomen cavity. 5. Explant the uterus by cutting the mesometrium and the surrounding fat tissue. Place the uterus in ice-cold 1 PBS (see Note 4). 6. Discard the pregnant mouse and proceed for embryo dissection.

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7. Separate each embryo by cutting between implantation sites along the uterine explant. 8. Make a small incision through the decidua tissue surrounding each embryo and with a pair of fine forceps, tear decidua apart and the embryo can be shelled out. 9. Once embryo is removed, Reichert’s membrane may still be attached as well as the ectoplacental cone (see Note 5). 10. Place the embryo in a clean petri dish with clean ice-cold 1 PBS and proceed to carefully dissecting the embryo under the binocular microscope. 11. Using fine forceps, carefully separate the upper and bottom parts of the embryo body by cutting through the abdomen. Discard the upper body. 12. Carefully peel the skin to visualize the femora. 13. Use the surrounding soft tissue to hold the hindlimb with the forceps and separate the hindlimb from the hip. 14. Using a pair of fine forceps, pinch the soft tissue on both sides of the femora and the soft tissue on both sides of the tibia. Pull to separate the femora and the tibia. Discard tibia. 15. Take off the surrounding soft tissue. Keep some to be used to grasp the femoral cartilage (see Note 6). 16. Place the femoral cartilage grafts in ice-cold 1 PBS or DMEM medium in a 24-well plate on ice for no longer than 2 h for optimal development after transplantation. 3.2 Renal Capsule Transplantation

1. Weigh male (8–12 week old) mice and induce general anesthesia with an IP injection of ketamine–medetomidine (50 mg of ketamine and 0.5 mg of Medetomidine per kg of body weight). 2. Perform a subcutaneous injection of analgesics solution (0.1 mg buprenorphine in NaCl 0.09% per kg of body weight) (see Note 7). 3. With the mouse under anesthesia, shave the left flank with the electric clipper. 4. Position the mouse on its side with the left shaved flank facing up under the binocular microscope (Fig. 1). 5. Swab the shaved area center-out with Betadine soap followed by Betadine solution. 6. Locate the left kidney and make a small longitudinal incision of approximately 1–1.5 cm through the skin and the body wall (Fig. 1) (see Note 8). 7. Expose the kidney outside the body by pulling with forceps the fat located at the distal pole of the kidney and simultaneously applying a slight pressure to both sides of the incision with the

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forefinger and thumb to pop the kidney out of the abdominal cavity. The exteriorized kidney will rest on the body wall. Keep the kidney moist by applying a PBS solution with a Pasteur pipette (see Note 9). 8. Prepare the graft site by making a small 2 mm hole in the renal capsule at the base of the kidney using small Vanna scissors (Fig. 1, arrow) (see Note 10). 9. Insert the “L”-shape glass rod into the hole and carefully slide it in between the capsule and the kidney parenchyma to make a small pouch for the graft (see Note 11). 10. Transfer the graft to the surface of the kidney using a pair of fine forceps (see Notes 12–14). 11. Insert the graft into the pouch by gently lifting the capsule with one pair of fine forceps and by placing the graft under the capsule with another pair of forceps. Once the graft is entirely covered with the capsule, guide it with the forceps to position it in the mid-axial part of the kidney (Fig. 1, white dotted line). 12. Reposition the kidney into the body cavity and close the body wall layer with 2 stiches using a 4-0 silk absorbable suture. 13. Align both sides of the skin incision together and close the skin with 2 or 3 clips using a wound clipper. 14. If needed, clean the skin of the mouse using a Betadine solution swab. 15. Inject the anesthetic reversal solution (0.1 mg atipamezole per kg body weight) via IP injection and place the mouse on a heating blanket set at approximately 37  C for recovery. Monitor the mice closely until fully awake. Let the mice ambulate freely to access food and water. 16. Monitor mice daily and remove skin staples after 2 weeks (see Note 15). 3.3 Bone Fracture and Cortical Defect in the Renal Capsule

1. Six to eight weeks after transplantation of E14–E14.5 femora in the renal capsule, weight and anesthetize the host mice as described in Subheading 3.2. 2. Shave the left flank of the mice and clean the shaved area with Betadine soap then Betadine solution. 3. At the level of the left kidney, make an incision (approximately 1–1.5 cm) longitudinally within the skin and the body wall. 4. Expose the kidney with the bone transplant to make it accessible as described in Subheading 3.2 (see Note 16) (Fig. 2a, e). 5. To induce a cortical defect, drill a hole of 0.8 mm in diameter into one cortex in the diaphysis (Fig. 2b). To induce a bone fracture, cut the bone in the mid diaphysis with scissors (see Note 17) (Fig. 2f).

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6. Reposition the kidney into the body cavity and suture the body wall using a 4-0 silk absorbable suture. 7. Close the skin incision with 2 or 3 clips using a wound clipper. 8. If needed, clean the skin of the mouse using a Betadine solution swab. 9. Inject the anesthetic reversal solution (0.1 mg atipamezole per kg body weight) via IP injection and place the mouse on a heating blanket set at approximately 37  C for recovery. Monitor the mice closely until fully awake. Let the mice ambulate freely to access food and water. 10. Monitor mice daily and remove skin staples after 2 weeks (see Note 15). 3.4 Analysis of Vascularization and Angiogenesis

Blood vessels are visualized with anti-PECAM (CD31) immunostaining (see Notes 18 and 19). 1. Harvest renal capsule transplanted femora and fix the tissue with 4% PFA fixative solution for 24 h (see Notes 20 and 21). 2. Decalcify samples in 0.5 M EDTA for 24 h—7 days on a rocking platform shaker at 4  C. Change EDTA solution every day (see Notes 22 and 23). 3. Dehydrate skeletal tissues by immersing tissue in graded ethanol series followed by xylene three times for 20 min each at room temperature (see Note 24). 4. Embed the tissue in paraffin at 58  C. 5. Cut 5–7 μm thick tissue sections using a rotary microtome. Float the sections in a 56  C water bath and mount the sections onto microscope slides. 6. Dry the slides at room temperature for 1 h and proceed with anti-PECAM immunostaining (see Notes 25 and 26). 7. Rehydrate sections by immersing the slides in Histoclear two times for 5 min each. 8. Immerse the slides in 100% ethanol two times for 5 min each. 9. Immerse the slides in 95% ethanol for 5 min. 10. Immerse the slides in 70% ethanol for 5 min. 11. Immerse the slides with deionized H2O for 5 min. 12. Rehydrate the slides with 1 PBS for 5 min using a glass jar with lid. 13. Surround the tissue with a hydrophobic barrier using a barrier pen. 14. Block endogenous peroxidase activity with fresh 0.3% of hydrogen peroxide (H2O2) diluted in methanol for 45 min at room temperature (see Notes 27 and 28).

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15. Proceed for enzymatic antigen retrieval step by incubating sections with ready to use Ficin solution for 5 min at room temperature (see Notes 29). 16. Wash slides three times for 5 min each in 1 PBS. 17. Block non-specific staining by incubating sections with 0.1 M glycine solution diluted in 1 PBS for 60 s at room temperature (see Note 30). 18. Wash slides three times for 5 min each in 1 PBS. 19. Block nonspecific staining by incubating sections with 5% nonfat powdered milk solution diluted in 1 PBS for 10 min at room temperature. 20. Wash slides three times for 5 min each in 1 PBS. 21. Block nonspecific staining by incubating sections with 0.1% ovalbumin solution diluted in 1 PBS for 10 min at room temperature. 22. Wash slides three times for 5 min each in 1 PBS. 23. Block nonspecific staining by incubating sections with 5% normal goat serum diluted in 1 PBS for 30 min at room temperature (see Note 31). 24. Apply rat anti-PECAM primary antibodies solution at 1:50 diluted in serum blocking solution (5% normal goat serum diluted in 1 PBS) and incubate overnight at 4  C (see Notes 32 and 33). 25. Rinse one time with 1 PBS to drain the excess of primary antibodies. 26. Wash slides three times for 5 min each in 1 PBS. 27. Block non-specific staining by incubating sections with 5% normal goat serum diluted in 1 PBS for 30 min at room temperature (see Note 31). 28. Apply goat biotinylated anti- rat secondary antibodies solution at 1:200 diluted in serum blocking solution (5% normal goat serum diluted in 1 PBS) and incubate for 1 h at room temperature. 29. Rinse the slides once with 1 PBS to remove excess secondary antibody. 30. Wash the slides three times for 5 min each in 1 PBS. 31. Apply HRP-conjugated streptavidin solution at 1:100 diluted in serum blocking solution (5% normal goat serum diluted in 1 PBS) and incubate for 45 min at room temperature. 32. Rinse the slides once with 1 PBS to remove excess HRP-streptavidin solution. 33. Wash the slides three times for 10 min each in 1 PBS.

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34. Apply DAB substrate working solution and develop for 30 s to 1 min (see Notes 34–36). 35. Rinse the slides for 2 min with ddH2O. 36. Wash the slides three times for 5 min each in ddH2O. 37. Proceed to counterstaining by incubating sections with 0.01% Fast Green solution diluted in deionized H2O for 15 s at room temperature. 38. Dehydrate the sections by immersing tissue in 70% ethanol for 3 min at room temperature. 39. Immerse the slides in 95% ethanol for 3 min at room temperature. 40. Immerse the slides in 100% ethanol for 3 min at room temperature. 41. Immerse the slides in Histoclear solution for 5 min at room temperature. 42. Apply a drop of Permount and coverslip. 43. Let slides dry at room temperature.

4

Notes 1. Use donor and host mice from the same genetic background to avoid graft rejection. Host mice should be preferably male as remodeling of the graft is accelerated in female hosts. 2. All surgical instruments and reagents must be sterile to avoid any risks of infection. 3. For the transplantation of stylopods and zeugopods, E14– E14.5 embryonic stage is the ideal time point as hypertrophic cartilage is well differentiated and will efficiently attract host blood vessels, but endogenous blood vessels have not invaded the cartilage yet and will not for another 24 h. For other skeletal elements that are less advanced in their development, such as autopods, later embryonic stages may be more appropriate. 4. Avoid excessive compression on the uterine explant, which could deform and compromise the embryonic tissues. 5. Embryo can be handled by grasping the attached Reichert’s membrane as well as the ectoplacental cone using fine forceps. 6. Use the soft tissue surrounding the stylopod (cartilage femoral graft) to handle the embryonic tissue. At E14–E14.5 embryonic stage, the stylopod is very soft. Excessive compression may deform and compromise the normal development under the renal capsule.

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7. At least one additional injection of analgesic is performed the day following the surgery. Please refer to your institutional guidelines concerning animal care and welfare. All of our procedures received approval from the Paris Descartes University Ethical committee. 8. The kidney is retroperitoneal. It is not necessary to cut into the peritoneum. The cut through the body wall should start just above the hip level and should be long enough (1–1.5 cm) for the kidney to be “popped out” but not longer to avoid the risk of it falling back into the body cavity during the procedure (Fig. 1). Avoid cutting major vessels and nerves. 9. It is important to keep the capsule moist during the entire process; otherwise it will be easily torn. 10. The size of the incision in the capsule is determined by the size of the graft, but it should not exceed 4 mm as it may cause a loss of the graft (Fig. 1). 11. The L-shape glass should be manipulated under the capsule tangential to the surface of the kidney to avoid tearing the capsule. Great care should be taken while creating the pouch to not damage the kidney parenchyma which if damaged will bleed. 12. The skeletal elements should be transplanted with intact perichondrium to allow optimal vascularization and development. Some remaining soft tissues can be kept around the graft, as it will not interfere with bone development and growth. 13. Numerous treatments and manipulations can be applied to the graft prior to transplantation or at the time of transplantation (for example incubation in a solution of blocking antibody, or placing beads soaked in a protein solution adjacent to the graft under the kidney capsule) [16]. 14. Several grafts can be transplanted in one kidney capsule depending on the length of the study (3 grafts for up to 1 week of development in the renal capsule, 2 grafts for up to 2 weeks, one graft for longer time points). Bilateral grafting is not recommended. 15. Potential adverse effects include infection, parenchyma bleeding and graft rejection. Although these effects occur very rarely, mice should be monitored daily following the transplantation. 16. Avoid pulling on bone transplant to expose the kidney. 17. These procedures are performed through the outer layer of the kidney. Make a small incision and maintain one extremity of the bone with forceps to create fracture or cortical defect. This will not affect the attachment of the bone to the kidney surface.

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18. Femurs or other long bones grown under the renal capsule repair after a cortical defect or a fracture (Fig. 2c, d, g–h, respectively) following a similar regeneration process as observed in adult bone [18]. 19. PECAM immunostaining can be realized on uninjured transplanted embryonic femur (Fig. 1d) or after a cortical defect or fracture (data not shown) following the same protocol. 20. The volume of fixative solution should be 50 times greater than the size of the immersed tissue to ensure a proper fixation of the tissue. 21. Avoid fixing the tissue for more than 24 h since tissue antigens may either be masked or destroyed. 22. Decalcification using chelator reagents such as EDTA works by capturing the calcium ions from the bone. EDTA acts slowly but is compatible with many immunostaining protocols. 23. The time of decalcification varies from 24 h to 7 days depending on the mineral density of the sample determined by the size of the skeletal element and the time point of harvest. 24. Paraffin is immiscible with water. Tissue must be dehydrated before adding paraffin wax. 25. Slides with paraffin-embedded sections can be stored either at room temperature or at 2–8  C for several years in slide storage boxes. However, PECAM immunostaining should be performed within a week after sectioning for optimum results. 26. This PECAM immunostaining protocol can also be performed on cryo-embedded tissues (starting the procedure at step 12). 27. Some cells or tissues contain endogenous peroxidase. Using HRP conjugated antibody may result in high, non-specific background staining. Incubation with Peroxide (H2O2) suppresses endogenous peroxidase activity and therefore reduces background staining. 28. Hydrogen peroxide should be stored in the refrigerator and protected from sunlight in order to slow its thermal decomposition. Always use fresh H2O2 working solution. 29. The Ficin enzymatic antigen retrieval method serves as a proteolytic digestion to expose the antigenic sites that are covered when the tissue is fixed, making antibody-antigen binding easier during the staining procedure. 30. The non-specific staining blocking step is most often performed just prior to incubating the sample with the primary and secondary antibodies. Non-specific staining blocking solution reduces the background signal produced by non-specific interaction of primary and secondary antibodies with proteins in the tissue section.

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31. Serum is required in the blocking solution to block immunoglobulin Fc receptors present on cells in the section. The serum should be of the same species as the secondary antibody. 32. Overnight incubation at 4  C with primary antibodies allows proper and optimal specific binding of antibodies to tissue targets and reduces nonspecific background staining. 33. A negative control is critical for an accurate interpretation of the immunostaining results. A negative control could be using the incubation buffer with no primary antibody to identify non-specific staining of the secondary reagents. Additional controls can be employed to support the specificity of staining generated by the primary antibody. These include absorption controls, isotype-matched controls (for monoclonal primary antibodies), and tissue-type controls. 34. Upon reaction with HRP, DAB substrate will produce a brown colored deposit. Signal development should be monitored under microscope. 35. DAB is extremely carcinogenic. Necessary precautions should be taken (wear gloves and use only glass containers). 36. DAB is photosensitive: Keep the DAB working solution away from light and always use freshly prepared DAB working solution.

Acknowledgments This work was supported by INSERM ATIP-AVENIR, Osteosynthesis and Trauma Care Foundation, ANR-18-CE14-0033 and NIH-NIAMS R01AR072707 grants to CC. References 1. Vu TH, Shipley JM, Bergers G, Berger JE, Helms JA, Hanahan D, Shapiro SD, Senior RM, Werb Z (1998) MMP-9/gelatinase B is a key regulator of growth plate angiogenesis and apoptosis of hypertrophic chondrocytes. Cell 93(3):411–422 2. Gerber HP, Hillan KJ, Ryan AM, Kowalski J, Keller GA, Rangell L, Wright BD, Radtke F, Aguet M, Ferrara N (1999) VEGF is required for growth and survival in neonatal mice. Development 126(6):1149–1159 3. Gerber HP, Vu TH, Ryan AM, Kowalski J, Werb Z, Ferrara N (1999) VEGF couples hypertrophic cartilage remodeling, ossification and angiogenesis during endochondral bone formation. Nat Med 5(6):623–628

4. Gerber HP, Ferrara N (2000) Angiogenesis and bone growth. Trends Cardiovasc Med 10 (5):223–228 5. Zelzer E, Mamluk R, Ferrara N, Johnson RS, Schipani E, Olsen BR (2004) VEGFA is necessary for chondrocyte survival during bone development. Development 131 (9):2161–2171 6. Maes C, Kobayashi T, Selig MK, Torrekens S, Roth SI, Mackem S, Carmeliet G, Kronenberg HM (2010) Osteoblast precursors, but not mature osteoblasts, move into developing and fractured bones along with invading blood vessels. Dev Cell 19(2):329–344 7. Grellier M, Ferreira-Tojais N, Bourget C, Bareille R, Guillemot F, Amedee J (2009) Role of vascular endothelial growth factor in

Renal Capsule Transplantation the communication between human osteoprogenitors and endothelial cells. J Cell Biochem 106(3):390–398 8. Schipani E, Maes C, Carmeliet G, Semenza GL (2009) Regulation of osteogenesisangiogenesis coupling by HIFs and VEGF. J Bone Miner Res 24(8):1347–1353 9. Vu TH, Alemayehu Y, Werb Z (2003) New insights into saccular development and vascular formation in lung allografts under the renal capsule. Mech Dev 120(3):305–313 10. Wiesen JF, Young P, Werb Z, Cunha GR (1999) Signaling through the stromal epidermal growth factor receptor is necessary for mammary ductal development. Development 126(2):335–344 11. Wang Y, Revelo MP, Sudilovsky D, Cao M, Chen WG, Goetz L, Xue H, Sadar M, Shappell SB, Cunha GR, Hayward SW (2005) Development and characterization of efficient xenograft models for benign and malignant human prostate tissue. Prostate 64(2):149–159 12. Szot GL, Koudria P, Bluestone JA (2007) Transplantation of pancreatic islets into the kidney capsule of diabetic mice. J Vis Exp 9:404 13. Chan CK, Seo EY, Chen JY, Lo D, McArdle A, Sinha R, Tevlin R, Seita J, Vincent-Tompkins J, Wearda T, Lu WJ, Senarath-Yapa K, Chung

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MT, Marecic O, Tran M, Yan KS, Upton R, Walmsley GG, Lee AS, Sahoo D, Kuo CJ, Weissman IL, Longaker MT (2015) Identification and specification of the mouse skeletal stem cell. Cell 160(1-2):285–298 14. Colnot C, Lu C, Hu D, Helms JA (2004) Distinguishing the contributions of the perichondrium, cartilage, and vascular endothelium to skeletal development. Dev Biol 269 (1):55–69 15. Colnot C (2005) Cellular and molecular interactions regulating skeletogenesis. J Cell Biochem 95(4):688–697 16. Colnot C, de la Fuente L, Huang S, Hu D, Lu C, St-Jacques B, Helms JA (2005) Indian hedgehog synchronizes skeletal angiogenesis and perichondrial maturation with cartilage development. Development 132 (5):1057–1067 17. Chan CKF, Gulati GS, Sinha R, Tompkins JV, Lopez M et al (2018) Identification of the human skeletal stem cell. Cell 175(1):43–56. e21 18. Duchamp de Lageneste O, Julien A, AbouKhalil R, Frangi G, Carvalho C, Cagnard N, Cordier C, Conway SJ, Colnot C (2018) Periosteum contains skeletal stem cells with high bone regenerative potential controlled by Periostin. Nat Commun 9(1):773

Part III Structural, Histological, and Molecular Analyses on Skeletal Tissues and Tissue Sections

Chapter 11 MicroCT for Scanning and Analysis of Mouse Bones Yung Kim, Michael D. Brodt, Simon Y. Tang, and Matthew J. Silva Abstract The purpose of this Chapter is to present a detailed description of methods for performing bone MicroComputed Tomography (microCT) scanning and analysis. MicroCT is an x-ray imaging method capable of visualizing bone at the micro-structural scale, that is, 1-100 μm resolution. MicroCT is the gold-standard method for assessment of 3D bone morphology in studies of small animals. As applied to the small bones of mice or rats, microCT can efficiently and accurately assess bone structure (e.g., cortical bone area [Ct.Ar]) and micro-structure (e.g., trabecular bone volume fraction [Tb.BV/TV]). The particular application described herein is for post mortem mouse femur specimens. The material presented should be generally applicable to many commercially available laboratory microCT systems, although some details are specific to the system used in our lab (Scanco mCT 40; SCANCO Medical AG, Bruttisellen, Switzerland). Key words microCT, Bone imaging, Bone morphology, Mouse femur

1

Introduction The purpose of this chapter is to present a detailed description of methods for performing bone micro-computed tomography (microCT) scanning and analysis. The particular application described below is for postmortem mouse femur specimens. First, we present a concise review of basic concepts and terminology. The material presented should be generally applicable to many commercially available laboratory microCT systems, although some details are specific to the system used in our lab (Scanco μCT 40; SCANCO Medical AG, Bruttisellen, Switzerland). For a comprehensive overview of microCT principles and guidelines for assessing and reporting bone microstructure, the reader is referred to Bouxsein et al. [1]. MicroCT is an X-ray imaging method capable of visualizing bone at the microstructural scale, that is, 1–100 μm resolution. It is similar to clinical CT, but achieves higher resolution by combining a smaller field-of-view, micro-focus X-ray source, and higher resolution detector. MicroCT is the gold-standard method for

Matthew J. Hilton (ed.), Skeletal Development and Repair: Methods and Protocols, Methods in Molecular Biology, vol. 2230, https://doi.org/10.1007/978-1-0716-1028-2_11, © Springer Science+Business Media, LLC, part of Springer Nature 2021

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Fig. 1 Attenuation of X-ray energy through bone depends on thickness (t) and mineral density. The thicker and denser the object, the greater the attenuation (loss)

assessment of 3D bone morphology in studies of small animals. As applied to the small bones of mice or rats, microCT can efficiently and accurately assess bone structure (e.g., cortical bone area [Ct. Ar]) and microstructure (e.g., trabecular bone volume fraction [Tb.BV/TV]) [1–3]. In brief, when an X-ray source is focused on an object, the intensity of the X-rays is attenuated based on the thickness and atomic number density of the constituent material (Fig. 1). An X-ray detector array opposite the source measures the X-ray energy, and essentially converts the X-ray shadow of the object into a projection/image. Clinical CT (and in vivo microCT) works by acquiring multiple projections as the source and detector rotate around the object (e.g., patient, mouse), and reconstructing these projections into a 2D tomogram or CT slice (Fig. 2). By contrast, specimen microCT scanners rotate the object while the source and detector remain stationary. In either case, a set of 2D slices is stacked to form a 3D image, which is represented as an X-Y-Z array of voxels. A voxel (a.k.a., volume element) is the 3D version of a pixel; for Scanco microCT systems, voxels are “isotropic,” that is, of equal dimension in X, Y and Z directions. By convention, the Z-direction corresponds to the longitudinal axis, and the X–Y plane is the transverse plane of the CT slice. Because bone mineral is relatively dense, it attenuates X-ray energy much more than marrow or soft tissue, and thus CT provides a clear contrast between bone and adjacent nonmineralized tissue (Fig. 3). Likewise, bone regions of lower density have less X-ray attenuation than regions of higher density, allowing for discrimination of variations in bone mineral density. The relative X-ray attenuation is expressed as a linear attenuation coefficient [1/cm], which may be converted to Hounsfield units (HU), or simply scaled per mille, that is, ranging from
1000 to +1000. The latter values are typically used to represent the data for display as a grayscale image, and for thresholding. For bone microCT, standard practice is that the linear attenuation is converted to mineral density based on a hydroxyapatite (HA) calibration phantom, as bone mineral is similar to hydroxyapatite. Thus, the units of bone mineral density (BMD) from microCT are [mg HA/cm3]. In summary,

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Fig. 2 At each position, the detector acquires a projection (P) of the object (e.g., forearm). The CT reconstruction algorithm essentially averages the intensities of these multiple projections at each spot on the detector array, and generates a 2D tomogram (slice) which is like a density map. Note that for clinical CT and in vivo microCT, the source and detector rotate around the subject, as shown above. By contrast, for specimen microCT, multiple projections are acquired by rotating the object while the source and detector are fixed

Fig. 3 MicroCT slice through sample tube containing a mouse lower leg. This grayscale image illustrates the differing attenuations of mineralized bone (tibia and fibula) compared to surrounding nonmineralized soft tissue and air (background). (Brightness of this image was increased to more easily visualize the soft tissue.) By convention the higher attenuating material is shown as brighter (whiter), although this can be inverted or even reassigned to color. A rubber band was placed with the sample in the tube as a position landmark

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Fig. 4 Schematic illustrating source–object–detector geometry. The field-of-view size is the diameter (“D”) of the sample holder (tube). The nominal voxel size is the field-of-view divided by the matrix size of the detector array. Thus, using the smallest sample holder that will contain your samples is recommended to give the best resolution, that is, smallest voxel size. Similarly, selecting a larger matrix size will improve resolution, although at the cost of larger file sizes and slower reconstruction times

microCT attenuation values may be expressed per mille or as BMD [mg HA/cm3]. There are many parameters that affect how a microCT scan is acquired and the quality of the resulting image. A critical parameter is the nominal voxel size, which is a measure of spatial resolution. The smaller the voxel size, the better the ability to accurately render microstructural features of bone such as trabecular thickness. Assuming isotropic (cubic) voxels, a single dimension is sufficient to describe voxel size. Estimated voxel size is computed as field-ofview [mm] divided by the matrix size [voxel number] used for image reconstruction (Fig. 4). The field-of-view is determined by geometry, namely, the focal distances from the X-ray source to the object, and the object to detector distance. In practice, for Scanco specimen microCT systems the user selects from specimen holders of varying diameters, and the field-of-view equates to the holder diameter. The matrix size is scanner dependent, and can range from 512  512 to 4096  4096 or higher, depending on the resolution of the X-ray detector. In general, to achieve the highest possible resolution (i.e., smallest voxel size) keep your specimen size as small as possible, that is, use the smallest holder possible. Higher resolution can also be achieved by using a larger matrix size. A common matrix size is 1024  1024, corresponding to “standard” resolution on the Scanco μCT 40 system. For example, if you select a field-of-view of 16.4 mm (i.e, 16.4 mm specimen holder), and a matrix size of 1024  1024, the nominal voxel size is 16.4/ 1024 ¼ 0.016 mm, or 16 μm. For the same field-of-view, a matrix size of 2048  2048 (‘high’ resolution) gives a voxel size of 8 μm.

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Table 1 Nominal voxel size for different combinations of sample holder tube size and resolution for Scanco μCT 40 Sample holder (tube) diameter 12 mm 16 mm 20 mma 25 mm

30 mm 35 mm

Low (1024  1024)

12 μm 16 μm 20 μm

25 μm

30 μm 35 μm

Medium (1024  1024)

12 μm 16 μm 20 μm

25 μm

30 μm 35 μm

Nominal voxel size Resolution (matrix size)

Higha (2048  2048) 6 μm

8 μm

10 μma

12.5 μm 15 μm 17.5 μm

a

Denotes values used for protocol described in Subheading 3

Additional tube size and resolution combinations for Scanco systems are shown below (Table 1). A recent report illustrated the importance of voxel size for quantification of trabecular bone microstructure in mice [4]. Based on these results, a voxel size of 6–10 μm is recommended for accurate determination of key trabecular parameters, including bone volume fraction (BV/TV), number (Tb.N), thickness (Tb. Th), and separation (Tb.Sp). A voxel size of 15–20 μm is acceptable, but a voxel size larger than this is not recommended. These results make sense when considering that the thickness of a mouse trabecula is approximately 40 μm, and as a rule the ratio of feature size to voxel size should be at least 3 or 4:1. Most commercially available scanners are capable of 10 μm voxel size, typically as a “high resolution” option. The trade-offs for choosing the highest possible resolution are increased scan time, increased reconstruction time, and increased data file size. (As a rule, if voxel size is decreased by half, the file size increases fourfold.) Over the past decade as computing power has increased (especially with the use of graphical processing units (GPUs)), and disk storage costs have declined, the use of higher-resolution scanning has become more practical and common. A number of parameter settings are selected by the user in the Control File. Two of these are essential to report in methods sections of papers [1], and are described briefly here. X-ray energy is a critical factor influencing attenuation. This is most commonly specified as the X-ray tube potential (peak voltage [kVp]); a higher kVp produces greater X-ray energy and results in less attenuation through mineral phantoms [5]. It is generally recommended that thicker, denser samples should be scanned at higher voltages (e.g., 70 kVp), yet, on the other hand, the contrast between marrow and bone may be better at lower voltage settings [1]. Another key parameter is integration time [ms], which is analogous to exposure time on a camera. A longer integration time allows more photons to

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Table 2 Key parameters for microCT scanning Parameter

Description

Units

Voxel size

Measure of nominal spatial resolution; smaller the value the better the resolution

μm

X-ray tube potential (peak voltage)

Measure of X-ray energy; higher voltage results in less attenuation through denser/thicker specimens

kVp

X-ray intensity (current)

Electrical current through X-ray tube; higher current produces more photons and better signal-to-noise

μA

Integration time

Duration of time for photons to hit detector for each projection; longer ms times give better signal-to-noise but increases scan time

Frame averaging (average data)

Number of repeated measurements at each projection; higher number gives better signal-to-noise but increases scan time

n

(Modified from Table 1 of Bouxsein et al. [1])

reach the detector and generally improves the image quality, or signal-to-noise ratio (SNR) [1]. The trade-off with longer integration times is longer time for scan acquisition and higher radiation exposure of the sample. (For postmortem bone samples, radiation exposure is not a practical concern.) The other settings that influence SNR are tube current [μA] and frame averaging [number]; although not essential to report, a complete description of scan settings would include these settings as well (Table 2).

2

Materials 1. Scanco μCT 40 (Scanco Medical, Switzerland). 2. Calibration Phantom (Scanco Medical). 3. Sample holder (Scanco Medical). 4. Dissection equipment (scissors, forceps, etc.). 5. 4% paraformaldehyde. 6. 70% ethanol. 7. Phosphate buffered saline. 8. Parafilm®. 9. 2 ml snap lock Eppendorf tube. 10. 15 ml conical tube. 11. 6 cm petri dish (60  15 mm). 12. 2% agarose: Add 100 ml distilled water to 2 g agarose powder in a glass bottle. Stir to suspend agarose. Cover the bottle with a cap (do not close tightly) and heat in a microwave until all agarose is dissolved. Allow it to cool to approximately 38  C before using (see Note 1).

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Methods The methods below are for microCT scanning and analysis of intact (whole) mouse femurs, using a specimen microCT scanner (see Note 2). An overview of these steps is shown in Fig. 5. MicroCT scanning is nondestructive, so it may be followed by other assays, typically either (1) histology (paraffin or plastic embedding) or (2) mechanical testing.

3.1 Sample Preparation, Fixation, and Embedding

1. Dissection: Within 10–15 min (see Note 3) after euthanasia, use scalpel and scissors to remove femurs from mice. Take care to keep the entire femur intact; the femoral head and distal (knee) epiphysis can easily separate from the rest of the bone, especially in younger mice. 2. If doing histology after microCT, remove enough muscle to facilitate good infiltration of solutions, but leave the bone covered by a layer of muscle and leave the periosteum intact (Fig. 6a) (see Note 4). 3. If doing mechanical testing after microCT, remove as much soft tissue as possible, especially along the diaphysis, which should be “down to the bone”. It is acceptable to leave small amounts of tissue at the two ends (Fig. 6b) (see Note 4).

Fig. 5 MicroCT workflow

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Fig. 6 Dissected mouse femurs (a) with and (b) without muscle

4. If doing histology after microCT, fix the bones according to your lab’s histology protocol (e.g., 4% paraformaldehyde for 48 h), transfer to 70% Ethanol, then store at 4 until ready to scan (see Note 5). 5. If doing mechanical testing after microCT or if no other assays are planned, wrap the bones in gauze or paper towels that have been soaked with 1 PBS. Wrap in plastic wrap, then place in individual labeled tubes, for example, 2 ml snap lock Eppendorf or 15 ml conical tube. Freeze at
20  C until ready to scan. Bones can be stored for up to 1 year. When ready to scan, thaw samples at room temperature for 30–60 min, keeping them soaked with 1 PBS. 6. Choose the 20 mm diameter sample holder (see Note 6) (Fig. 7). 7. To minimize sample movement (and avoid motion artifact) during scanning, the specimen is embedded in medium before placement in the sample holder. We use 2% agarose, as indicated above in Subheading 2, item 12. This also will keep the sample hydrated during scanning, and provide a water/softtissue like background for the CT images. The method here describes staging for 10 bones to be placed in a 20 mm tube and scanned in one session. Bones are arranged in two stacks of five per stack. Use an asymmetric arrangement to facilitate identifying the individual bones (Fig. 8). It is essential to record in your lab notebook which bone is in which position. You can work with variations of this method to scan one or more femur per session (see Note 7).

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Fig. 7 Scanco PEI (polyetherimide) sample tube. Outer diameter is 20 mm; inner diameter is 18.5 mm. Stainless steel pin aligns tube in the scanner

8. Add a small amount of warm (approx. 38  C), liquid agarose to cover the bottom of a Petri dish to make a base layer. Let gel cool until firm. 9. Place a rubber band that is about the length of one of the bones into the dish for a fiducial marker to help locate which bone is where in the scan (Fig. 9a). 10. Place two bones next to the rubber band (the first layer). Try to align the mid-point of bones. Space between bones is approximately 2.5 mm. The rubber band and bones should fit inside a width of approximately 18 mm. Add liquid agarose to cover bones. 11. Pour a layer of agarose over the first layer to make a barrier between the first and the second layer, and allow to cool. 12. Place three bones on the barrier (approx. 18 mm total width) and add agarose to cover (Fig. 9b). 13. After agarose is solidified, use a razor to cut a block containing the samples to the approximate dimensions of: 18  18  18 mm. Place the block into the 20 mm sample tube so that the long axis of the femurs aligns with the long axis of the tube (Z direction). By convention we position the knee (distal) end toward the bottom of the tube and the femoral head (proximal) toward the top.

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Fig. 8 Sketch illustrates asymmetric positioning for five femurs in a single stack, allowing for multiple bones to be scanned simultaneously in a single measurement. Additional stacks can be placed in the sample holder and scanned as additional measurements in batch mode. Further, if your system is equipped with a sample changer, you can load up to 10 holders for batch mode scanning

Fig. 9 Agarose Embedding. (a) First layer with rubber band marker and two femurs. (b) Second layer with three femurs. (c) Top view looking down on femoral heads. (d) Side View

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14. (Optional) Follow Subheading 3.1, steps 8–13 to make another stack of five bones and add it into the tube. 15. Fill the tube with liquid agarose and let it harden. 16. Cover the tube with Parafilm®. Be sure that the film is tightly adhered to the tube, and trim any excess (see Note 8). 3.2 Labeling Samples, Defining Region of Interest (ROI), and Generating Measurements

1. Place tube(s) in the scanner (Fig. 10). If the system has a carousel, place the first sample holder in the carousel and note which position (e.g., position 1). Add additional sample holders as necessary and record their positions. Prior to scanning any samples, the system needs to have a sample number that is system generated. Each sample number requires a name. 2. Click on the “Create/Change Sample Data” (Fig. 11, #1). 3. Enter a descriptive sample name in the “Name” field (Fig. 12) (see Note 9). 4. Click “Save.” Record the sample number. 5. Begin measurements by clicking on the “Perform Measurements” (Fig. 11, #2). 6. Click on the “Yes” for starting a measurement (Fig. 13a). 7. Enter Sample Number (Fig. 13b)—enter the sample number that was created in Subheading 3.2, step 4. 8. Check carousel positions (Fig. 14). The program shows the valid (occupied) carousel positions (marked in green). Select one highlighted carousel position. 9. In the Measurement window, create a new or select an existing control file.

Fig. 10 Placing the sample holder in the scanner. Here the holder is in position 1 of the sample changer (carousel). This changer can hold up to 10 sample tubes

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Fig. 11 Scanco software main menu

Fig. 12 Create new sample number with descriptive name (see Note 9)

10. If creating a new control file, set the scanning parameters in Subheading 3.2, steps 11–17) (Fig. 15). If using a saved control file, check/edit the parameters. 11. Holder type (Fig. 15, #1): choose the sample holder that matches the size of the holder in this carousel position. For our example, select “’20 mm  H 75 mm.” 12. Energy/intensity (Fig. 15, #2): choose 70 kVp, 114 μA, 8 W from one of the preset combinations (see Note 10). 13. Resolution (Fig. 15, #3): select “High” resolution (see Note 11). 14. FOV/Diameter [mm]: leave setting at ‘20.5’ to match the sample holder size (see Note 12). 15. Voxelsize (μm) (Fig. 15, #4): based on sample holder size and resolution, the system sets the Voxelsize automatically (Table 1). For this example it is 10 μm.

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Fig. 13 (a) Start a new measurement. (b) Enter sample number created in previous step

Fig. 14 Carousel Position. Example shows holders in positions 5 and 7 (green)

16. Integration time (ms) (Fig. 15, #5): select 300 ms (see Note 13). 17. Average data (Fig. 15, #6): set to 1 (see Note 14). 18. After creating or selecting the Control file, click on the “Scout View” in the main measurement window (Fig. 16a) to bring up the Scout View window (Fig. 16b). This window is used to define the initial ‘Scout scan’ which is a low-resolution 2D projection image of the specimen holder. 19. Adjust the Startposition and Endposition sliders to 0 and 80 mm, respectively, to acquire a scout scan of the entire sample holder. Click on the “Scout View” button in the upper right of the window (Fig. 16b) to acquire the scout scan, which may take a few minutes. 20. We recommend doing the cortical and cancellous bone regions as two separate measurements (Fig. 17). Define the cortical region first, then add a new scan for the cancellous region (see Note 15).

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Fig. 15 Controlfile settings

Fig. 16 (a) Measurement main window; selecting Scout View. (b) Scout-View window. (c) Scout-view of a bone biopsy with reference line (green) selections (Scanco Medical)

21. After the Scout View scan is complete and displayed on the screen, click on “Reference-Line” (Fig. 16b). The Reference lines are used to determine the longitudinal (z-axis) extent of the scan region (see Note 16). Use the mouse to set reference lines indicating the start and end positions for the scan (Fig. 16c). 22. Click on the left mouse button to fix the reference lines once ROI is defined. 23. Click on the “Add Scan” (Fig. 16b).

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Fig. 17 Scan and Analysis Regions for Cortical and Cancellous/Trabecular regions

24. To add Cancellous region, repeat Subheading 3.2, steps 21– 23. Similarly, if you wish to scan another stack of bones in the same sample holder, repeat these same steps. 25. When done adding scans, click on the “OK” (Fig. 16b). 26. If you wish to scan bones in another sample holder, go to Subheading 3.2, step 8 and select a new carousel position. Then repeat Subheading 3.2, steps 9–25. 27. When all the scans are added for all sample holders, click on the “Task List” (Fig. 16a) to display each scan that was added and then click on the “Submit Batch Scans” button. The Measurement window will close. 28. Enter “que” in command prompt window to see the list of active measurements. Once a measurement is complete, its entry will no longer be on the list. The system will start to acquire your raw images, followed by image reconstruction resulting in two files per measurement (see Note 17). When reconstruction is complete, you can proceed to Subheading 3.3 for Image Analysis and Evaluation. 3.3 Image Analysis and Evaluation

1. Start the evaluation program (Fig. 11, #3). Enter or select the sample number and choose the measurement from the list. Measurement numbers are assigned by the system during reconstruction. The images are loaded and you can check them in the Evaluation window (Fig. 18). 2. If there is more than one bone in the scan, go to the “Zoom” menu and select magnification to zoom in on the bone to be analyzed (Fig. 19).

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Fig. 18 Overview of Evaluation Program (Scanco Medical)

Fig. 19 Example CT slice showing mid-diaphyseal cross-sections of five femurs, with rubber band fiducial marker. Orientation of bones relative to rubber band is as shown in Fig. 8. Dashed box illustrates zoom selection of femur A for analysis

3. To contour the bones (Fig. 20), delineate the region of interest by manual or automatic methods. For cortical bone perform Subheading 3.3, steps 4–8, and for trabecular bone perform Subheading 3.3, steps 9–16. 4. Draw manual contours on the first and last slices of the cortical analysis region (Figs. 17 and 21) by first zooming in on the region of interest and clicking “Draw Contour” (Fig. 21, #1). 5. Draw contour around the outside of the cortical bone (counter-clockwise) for first and last slices (Fig. 21, #2).

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Fig. 20 Contouring menu

Fig. 21 Manual Contouring (Cortical Bone)

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Fig. 22 Automatic Contouring (Cortical Bone)

6. Select the first/starting slice and click on the “C. . .” (Fig. 22, #1). 7. Choose “Forwards” in the Selection column (Fig. 22, #2). 8. Click on the “Iterate forwards” (Fig. 22, #3). Once the automatic contours have been drawn, scroll through the slices to check for accuracy. Then go to Subheading 3.3, step 17. 9. For trabecular bone contouring (see Note 18), begin manual contour starting at the first slice. 10. Zoom in on the region of interest and click on “Draw Contour” (Fig. 21, #1). 11. Draw contour around medullary area (just inside of cortical bone, counterclockwise). 12. Repeat steps 10 and 11 for every tenth slice of the trabecular analysis region (Fig. 17). 13. We will now use the Morph automatic contour function. 14. Select the first slice and click on the “C. . .”. 15. Choose “all” in the Selection column (Fig. 23, #1). 16. Click on “Morph” (Fig. 23, #2). Once the automatic contours have been drawn, scroll through the slices to check for accuracy. Proceed to Subheading 3.3, step 17. 17. Evaluation of the data requires segmentation (see Note 19), which involves applying filters to the image data (see Note 20) and selecting a threshold (see Note 21). Begin this process by selecting 3D Evaluation “T. . .” (Fig. 18, bottom left) in the Evaluation window.

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Fig. 23 Automatic Contouring Window (Trabecular Bone)

Fig. 24 3D-Evaluation Window

18. In the 3D Evaluation window click on “Select” (Fig. 24, #1) and choose the proper Task (Script). Choose the default cortical or trabecular analysis script created by Scanco during your system set-up. (On our system, these are named “Bone midshaft more than e.g. 50 slices Evaluation” for cortical bone analysis, and “Bone Trab. Morphometry” for trabecular bone analysis.)

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Fig. 25 Choosing the threshold (adapted from Bouxsein et al. [1]). (a) Grayscale image of femur metaphysis. (b) A good threshold value results in a segmented image that closely matches the grayscale. (c) Too high a threshold results in erosion of bone and underestimating bone volume. (d) Too low a threshold results in excessive bone

19. Adjust the sliders to select values for “Gauss Sigma” and “Gauss Support” (Fig. 24, #2). We recommend starting with the combination of Sigma ¼ 0, Support ¼ 0 (see Note 20). 20. Adjust the sliders to select a “Lower Threshold” and “Upper Threshold” values (Fig. 24, #3). For cortical bone, a lower threshold value of approximately 300 (per 1000) may be appropriate. For trabecular bone, a lower threshold of approximately 220 (per 1000) may be appropriate. The values need to be confirmed by user inspection. Typically, the lower threshold value is chosen iteratively by comparing grayscale vs. segmented images and adjusting the threshold value until the two images match as closely as possible (Fig. 25, see Note 21). To assist with selecting a lower threshold, you may wish to check the histogram for the grayscale image (see Note 22). Generally, you will not adjust the Upper Threshold value; it should be set to maximum value (1000). 21. Click on the “Start Evaluation” (Fig. 24, #4) and “Yes” to save contours. You must wait until evaluation is completed before collecting data or doing any additional analyses (e.g., a second bone in the same measurement). If you attempt concurrent evaluations, the results will be meaningless. Novice users should collect the data (Subheading 3.3, step 22) before doing additional analyses. 22. To acquire quantitative data (see Note 23), click on “Applications” in Session Manager window (Fig. 26). 23. Click on the “DECterm” to open a terminal window and enter “uct_list ‘measurement number’ ” in the terminal window. Results of the bone analysis will be displayed for the current bone and current contours (Fig. 27). You may wish to record the results at this time. Interpretation of the many outcomes generated by the analysis is addressed below (see Notes 24–35).

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Fig. 26 Session Manager Window

24. To observe the image data as a 3D display, first start 3D Display (Fig. 11, #4) and enter “Sample number” and select “measurement number.” The retrieved images should be contoured and evaluated. 25. Next, click on “Start” in 3D display window (Fig. 29). 26. The image can now be manipulated via Object Rotation, Elevation of the light source, or Cropping (see Note 36). 27. To analyze another bone, close the Evaluation window. Open a new Evaluation window and delete previous contour (s) (Fig. 20, #8). Go to Subheading 3.3, step 1 to repeat the steps for a new bone. 3.4 Third-Party Image Analysis Software

1. Users can export their images in DICOM (Digital Imaging and Communications in Medicine) file format and then use third party software for 3D image visualization (see Note 37).

3.5 Quality Control (QC)

1. System managers should perform weekly density calibration check (‘QC1’) using the Scanco provided phantom. This will confirm accuracy of mineral density values (Fig. 30) (see Note 38). 2. System managers should perform a monthly check of scanner alignment (geometry), also called “QC2.” This routine uses three fine aluminum wires in the phantom to check alignment.

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Fig. 27 Structural and densitometric indices of bone from (a) Cortical bone analysis, and (b) Trabecular bone analysis. For trabecular analysis, the ‘VOX’ parameters are the most straightforward, as they are based simply on counting voxels; the ‘DT’ parameters are based on distance transformation analysis which is the only method recommended for these analyses. (Scanco Medical)

4

Notes 1. Do not use boiling or hot agarose. It causes damage to the sample and will negatively affect later histology or mechanical testing. 2. While the principles described here will be instructive for other systems, many of the details are specific to SCANCO microCT systems, and in particular to the system used in our lab (SCANCO μCT 40). At the time of this writing, we were running version 6.4 of the Scanco Tomography program, and version 6.6 of the Scanco Evaluation program.

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3. For samples that will go to histology after microCT, minimizing the time from animal death to sample collection is important. The precise time window depends on the downstream staining protocols; the user should consult with histology personnel to determine what is recommended. For example, if immunostaining is planned a short time to dissection (1 mL volume), place (uncapped) under vacuum (17 Hg; longer is better) for at least 2 h. 4. Remove the vials from the vacuum and reorient specimen, if necessary, for sectioning. Then, fasten the lid tightly and place in refrigerator overnight (or over weekend). 5. Remove the specimen from the refrigerator. Check the orientation and solution volume. Top with embedding media as necessary to keep specimen covered, always fastening lid tightly afterward. Allow media to polymerize on countertop at room temperature (see Note 15).

3.3.2 Embedding in Glass: Embedding Media II, 0.8% Perkadox

1. Fill the 6  50 mm glass culture tube (see Fig. 2) with embedding media (0.8% catalyst, 1Ct between technical replicates may indicate a problem with RNA quality. 25. Typically, we perform RT-qPCR with at least two technical replicates per sample and biological replicates ranging from 4 to 10 within an experiment. The number of biological replicates is determined by considering the effect size and performing power analysis. 26. Decalcified bone paraffin sections should be on charged slides to ensure slides adhere during the antigen retrieval step. 27. All incubations should be done in humid chambers. We recommend using the commercially available histochemistry staining trays with black lids (Simport stain tray™). These trays are designed with a reservoir to hold water beneath the slides. 28. Primary antibody concentration should be optimized for various samples. 29. DAB working solution must be made fresh every time, right before adding it onto the sections. 30. The diazonium salt solution must be made fresh and not incubated longer than 5 min. 31. The Fast Green solution can reused for staining at least five times before its final discard. 32. Bring the fixative solution to RT before use and discard if evaporation is noted. 33. It is possible that the sections might stain positive for TRAP within 30 min of exposure. Thus, we recommend monitoring the stain development within the 1 h. At times incubations longer than an hour might be required. We have conducted TRAP staining for 2 h for better development of stain color. 34. Osteocyte TRAP positive cells are visible on sagittal sections of long bones only. For axial sections of long bones, we recommend performing IHC using Cathepsin K antibody.

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Acknowledgments The authors gratefully acknowledge funding from NIH R01DE019284, P30AR061312, R21AR070403; NSF 1636331 and Center for Disruptive Musculoskeletal Innovation, DOD OR170044, and the Read Research Foundation (TA), NIA-1F31AG063402-01A1 (CS), and NIH P01AG039355 and R21AR054449 (SD). We acknowledge use of the UMKC Confocal Microscopy Core supported by NIH grants S10RR027668 and S10OD021665. References 1. Bonewald LF (2011) The amazing osteocyte. J Bone Miner Res 26(2):229–238 2. Dallas SL, Prideaux M, Bonewald LF (2013) The osteocyte: an endocrine cell ... and more. Endocr Rev 34(5):658–690 3. You LD, Weinbaum S, Cowin SC, Schaffler MB (2004) Ultrastructure of the osteocyte process and its pericellular matrix. Anat Rec A Discov Mol Cell Evol Biol 278(2):505–513 4. Buenzli PR, Sims NA (2015) Quantifying the osteocyte network in the human skeleton. Bone 75:144–150 5. Schaffler MB, Henderson SC, Wang Y, Wang L, Weinbaum S, Majeska RJ, Han Y (2005) In situ measurement of solute transport in the bone lacunar-canalicular system. Proc Natl Acad Sci 102(33):11911–11916 6. Klein-Nulend J, Bakker AD, Bacabac RG, Vatsa A, Weinbaum S (2013) Mechanosensation and transduction in osteocytes. Bone 54 (2):182–190 7. Scheiner S, The´oval A, Pivonka P, Smith DW, Bonewald LF (2014) Investigation of nutrient transport mechanisms in the lacunae-canaliculi system. IOP Conf Ser Mater Sci Eng 10(1):1–8 8. Fritton SP, Weinbaum S (2009) Fluid and solute transport in bone: flow-induced Mechanotransduction. Annu Rev Fluid Mech 41:347–374 9. Wang L (2018) Solute transport in the bone lacunar-canalicular system (LCS). Curr Osteoporos Rep 16(1):32–41 10. van Hove RP, Nolte PA, Vatsa A, Semeins CM, Salmon PL, Smit TH, Klein-Nulend J (2009) Osteocyte morphology in human tibiae of different bone pathologies with different bone mineral density — is there a role for mechanosensing? Bone 45(2):321–329 11. Tsourdi E, J€ahn K, Rauner M, Busse B, Bonewald LF (2018) Physiological and pathological

osteocytic osteolysis. J Musculoskelet Neuronal Interact 18(3):292–303 12. Qing H, Bonewald LF (2009) Osteocyte remodeling of the perilacunar and pericanalicular matrix. Int J Oral Sci 1(2):59–65 13. Wysolmerski JJ (2012) Osteocytic osteolysis: time for a second look? Bonekey Rep 1:229 14. Belanger LF (1969) Osteocytic osteolysis. Calcif Tissue Res 4(1):1–12 15. Jande SS, Be´langer LF (1970) Electron microscopy of osteocytes and the pericellular matrix in rat trabecular bone. Calcif Tissue Res 6 (1):280–289 16. Bonewald LF (2007) Osteocytes as dynamic multifunctional cells. Ann N Y Acad Sci 1116:281–290 17. McGee-Lawrence ME, Carey HV, Donahue SW (2008) Mammalian hibernation as a model of disuse osteoporosis: the effects of physical inactivity on bone metabolism, structure, and strength. Am J Physiol Integr Comp Physiol 295(6):R1999–R2014 18. Qing H, Ardeshirpour L, Divieti Pajevic P, Dusevich V, J€ahn K, Kato S, Wysolmerski J, Bonewald LF (2012) Demonstration of osteocytic perilacunar/canalicular remodeling in mice during lactation. J Bone Miner Res 27 (5):1018–1029 19. Teti A, Zallone A (2009) Do osteocytes contribute to bone mineral homeostasis? Osteocytic osteolysis revisited. Bone 44(1):11–16 20. Yee CS, Schurman CA, White CR, Alliston T (2019) Investigating osteocytic perilacunar/ canalicular remodeling. Curr Osteoporos Rep 17(4):157–168 21. Inoue K, Mikuni-Takagaki Y, Oikawa K, Itoh T, Inada M, Noguchi T, Park JS, Onodera T, Krane SM, Noda M, Itohara S (2006) A crucial role for matrix metalloproteinase 2 in osteocytic canalicular formation and

322

Neha S. Dole et al.

bone metabolism. J Biol Chem 281 (44):33814–33824 22. Tang SY, Herber R-P, Ho SP, Alliston T (2012) Matrix metalloproteinase-13 is required for osteocytic perilacunar remodeling and maintains bone fracture resistance. J Bone Miner Res 27(9):1936–1950 23. J€ahn K, Kelkar S, Zhao H, Xie Y, Tiede-Lewis LAM, Dusevich V, Dallas SL, Bonewald LF (2017) Osteocytes acidify their microenvironment in response to PTHrP in vitro and in lactating mice in vivo. J Bone Miner Res 32 (8):1761–1772 24. Lotinun S, Ishihara Y, Nagano K, Kiviranta R, Carpentier VT, Neff L, Parkman V, Ide N, Hu D, Dann P, Brooks D, Bouxsein ML, Wysolmerski J, Gori F, Baron R (2019) Cathepsin K–deficient osteocytes prevent lactation-induced bone loss and parathyroid hormone suppression. J Clin Invest 129 (8):3058–3071 25. Yamada S, Billinghurst RC, Chrysovergis K, Inoue S, Poole AR, Holmbeck K, Pidoux I, Birkedal-Hansen H, Bianco P, Wu W (2004) The metalloproteinase MT1-MMP is required for normal development and maintenance of osteocyte processes in bone. J Cell Sci 118 (1):147–156 26. Tokarz D, Martins JS, Petit ET, Lin CP, Demay MB, Liu ES (2018) Hormonal regulation of osteocyte perilacunar and canalicular remodeling in the Hyp mouse model of X-linked hypophosphatemia. J Bone Miner Res 33 (3):499–509 27. Kaya S, Basta-Pljakic J, Seref-Ferlengez Z, Majeska RJ, Cardoso L, Bromage T, Zhang Q, Flach CR, Mendelsohn R, Yakar S, Fritton SP, Schaffler MB (2017) Lactation induced changes in the volume of osteocyte lacunar-canalicular space Alter mechanical properties in cortical bone tissue. J Bone Miner Res 32(4):688–697 28. Dole NS, Mazur CM, Acevedo C, Lopez JP, Monteiro DA, Fowler TW, Gludovatz B, Walsh F, Regan JN, Messina S, Evans DS, Lang TF, Zhang B, Ritchie RO, Mohammad KS, Alliston T (2017) Osteocyte-intrinsic TGF-β signaling regulates bone quality through perilacunar/canalicular remodeling. Cell Rep 21(9):2585–2596 29. Tang SY, Herber RP, Ho SP, Alliston T (2012) Matrix metalloproteinase-13 is required for osteocytic perilacunar remodeling and maintains bone fracture resistance. J Bone Miner Res 27(9):1936–1950 30. Amling M, Pu¨schel K, Rolvien T, Yorgan T, Jeschke A, Busse B, Krause M, Demay MB, Schinke T (2017) Vitamin D regulates

osteocyte survival and perilacunar remodeling in human and murine bone. Bone 103:78–87 31. Kogawa M, Wijenayaka AR, Ormsby RT, Thomas GP, Anderson PH, Bonewald LF, Findlay DM, Atkins GJ (2013) Sclerostin regulates release of bone mineral by osteocytes by induction of carbonic anhydrase 2. J Bone Miner Res 28(12):2436–2448 32. Dole NS, Mazur CM, Acevedo C, Lopez JP, Monteiro DA, Fowler TW, Gludovatz B, Walsh F, Regan JN, Messina S, Evans DS, Lang TF, Zhang B, Ritchie RO, Mohammad KS, Alliston T (2017) Osteocyte-intrinsic TGF-β signaling regulates bone quality through perilacunar/canalicular remodeling. Cell Rep 21(9):2585–2596 33. Heller-Steinberg M (1951) Ground substance, bone salts, and cellular activity in bone formation and destruction. Am J Anat 89 (3):347–379 34. Chappard D, Basle´ MF, Legrand E, Audran M (2011) New laboratory tools in the assessment of bone quality. Osteoporos Int 22 (8):2225–2240 35. Tazawa K, Hoshi K, Kawamoto S, Tanaka M, Ejiri S, Ozawa H (2004) Osteocytic osteolysis observed in rats to which parathyroid hormone was continuously administered. J Bone Miner Metab 22(6):524–529 36. Baylink DJ, Wergedal JE (1971) Bone formation by osteocytes. Am J Phys 221(3):669–678 37. Webster DJ, Schneider P, Dallas SL, Mu¨ller R (2013) Studying osteocytes within their environment. Bone 54(2):285–295 38. Fowler TW, Acevedo C, Mazur CM, HallGlenn F, Fields AJ, Bale HA, Ritchie RO, Lotz JC, Vail TP, Alliston T (2017) Glucocorticoid suppression of osteocyte perilacunar remodeling is associated with subchondral bone degeneration in osteonecrosis. Sci Rep 7:44618 39. Tiede-Lewis LM, Xie Y, Hulbert MA, Campos R, Dallas MR, Dusevich V, Bonewald LF, Dallas SL (2017) Degeneration of the osteocyte network in the C57BL/6 mouse model of aging. Aging 9(10):2190–2208 40. Kamel-ElSayed SA, Tiede-Lewis LM, Lu Y, Veno PA, Dallas SL (2015) Novel approaches for two and three dimensional multiplexed imaging of osteocytes. Bone 76:129–140 41. Schneider P, Meier M, Wepf R, Mu¨ller R (2010) Towards quantitative 3D imaging of the osteocyte lacuno-canalicular network. Bone 47(5):848–858 42. Dong P, Pacureanu A, Zuluaga MA, Olivier C, Grimal Q, Peyrin F (2014) Quantification of the 3d morphology of the bone cell network

Assessment of Osteocytes from synchrotron micro-CT images. Image Anal Stereol 33(2):157 43. Schneider P, Meier M, Wepf R, Mu¨ller R (2011) Serial FIB/SEM imaging for quantitative 3D assessment of the osteocyte lacunocanalicular network. Bone 49(2):304–311 44. Sano H, Kikuta J, Furuya M, Kondo N, Endo N, Ishii M (2015) Intravital bone imaging by two-photon excitation microscopy to identify osteocytic osteolysis in vivo. Bone 74:134–139 45. Fowler TW, Acevedo C, Mazur CM, HallGlenn F, Fields AJ, Bale HA, Ritchie RO, Lotz JC, Vail TP, Alliston T (2017) Glucocorticoid suppression of osteocyte perilacunar remodeling is associated with subchondral bone degeneration in osteonecrosis. Sci Rep 7:44618 46. Mazur CM, Woo JJ, Yee CS, Fields AJ, Acevedo C, Bailey KN, Kaya S, Fowler TW, Lotz JC, Dang A, Kuo AC, Vail TP, Alliston

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T (2019) Osteocyte dysfunction promotes osteoarthritis through MMP13-dependent suppression of subchondral bone homeostasis. Bone Res 7:34 47. Gaudin-Audrain C, Gallois Y, PascarettiGrizon F, Hubert L, Massin P, Basle´ M-F, Chappard D (2008) Osteopontin is histochemically detected by the AgNOR acid-silver staining. Histol Histopathol 23(4):469–478 48. Skinner RA (2003) Decalcification of bone tissue BT. In: An YH, Martin KL (eds) Handbook of histology methods for bone and cartilage. Humana Press, Totowa, NJ, pp 167–184 49. Monleau M, Bonnel S, Gostan T, Blanchard D, Courgnaud V, Lecellier C-H (2014) Comparison of different extraction techniques to profile microRNAs from human sera and peripheral blood mononuclear cells. BMC Genomics 15 (1):395

Chapter 18 Cell Lineage Tracing: Colocalization of Cell Lineage Markers with a Fluorescent Reporter Yan Jing, Patricia Simmer, and Jian Q. Feng Abstract Cell lineage tracing, an old technique which originated in the nineteenth century, regains popularity and relevance due to introduction of a more sensitive tomato fluorescent protein under the control of a ubiquitous promoter (Rosa 26 gene). In addition, various tissue specific CreERT2 mouse lines are widely available, making cell lineage tracing studies more specific and powerful. In this protocol, we provide a practical guide for researchers to map progeny of specific cells such as chondrocytes during development using a fluorescent reporter (tomato, red) and multiple chondrocyte Cre lines. Further, we provide valuable examples in which these tracing lines, combined with a bone reporter mouse line (2.3 Col 1a1-GFP) or costained with different immunofluorescent proteins, revealed how a chondrocyte transdifferentiates into a bone cell in vivo. Key words Cell lineage tracing, Cartilage, Bone, Chondrocyte, Growth/development

1

Introduction Cell lineage tracing and reporter analysis are powerful techniques in the study of developmental biology [1]. The pairing of a recombinase system with fluorescent markers as reporters allows us to study cell fate in vivo [2, 3]. The conventional technique commonly uses the Cre-LoxP system, wherein Cre, a recombinase enzyme, excises the STOP sequence between two loxP sites. This activates the reporter (tdTomato is favored as it encodes the brightest fluorescent protein with the strongest epifluorescence) in a specific cell lineage (Fig. 1a). In this way, the specific cell type and all descendants of those cells are permanently labeled via the fluorescent reporter. In some cases, the investigator can choose a favorable time point to activate Cre by using a drug, such as tamoxifen, causing the Cre fused to a modified form of the estrogen receptor (CreERT2) to translocate to the nucleus and induce recombination of LoxP sites ultimately

Matthew J. Hilton (ed.), Skeletal Development and Repair: Methods and Protocols, Methods in Molecular Biology, vol. 2230, https://doi.org/10.1007/978-1-0716-1028-2_18, © Springer Science+Business Media, LLC, part of Springer Nature 2021

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Fig. 1 Mechanism for cell lineage tracing and the generation of the compound mice. (a) The Cre-loxP system: Cre excises the STOP sequence between the two loxP sites, and the tdTomato protein (fluorescing red) is permanently expressed in the specific cell line. (b) Tracing embryonic cells: cross 2.3Col1a1-GFP; Rosa26tdTomato mice with Col10a1-Cre mice. Col10a1-Cre is noninducible, which reflects cell differentiation from the very beginning of Col10a1 expression (at E14.5). (c) Tracing postnatal cells: cross Aggrecan-CreERT2 (Agg-CreERT2) with 2.3Col1a1-GFP; Rosa26tdTomato mice, allowing for tamoxifen-induced activation. Activate Cre at 2 weeks of age via tamoxifen injection (Tm: tamoxifen). (d) Combining immunofluorescence with cell lineage tracing: generate Agg-CreERT2; Rosa26tdTomato mice. Activate Cre at postnatal day 3, and perform Runx2 and DMP1 immunofluorescence assays (Tm: tamoxifen).

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activating the reporter [3]. Each of these systems are described in more detail in Chapter 3. Combining these traditional designs with immunofluorescent staining makes these techniques even more powerful, allowing direct tracking of cell differentiation. Colocalization of cell lineage markers via immunofluorescence and reporter tomato signals allow the observation of the number of progeny of the founder cell, their location, and their differentiation status, if appropriate markers are selected. This provides more information than cell lineage tracing alone. In addition, the use of cell-specific markers can simplify the generation of compound mice, as there is no need to breed transgenic mice expressing various cell specific fluorescent marker (GFP, YFP, etc.), accelerating the investigation. Co application of cell lineage tracing and immunofluorescence is a powerful tool for investigating cell biology in vivo [4]. It allows for simultaneously performing immunofluorescence with two different antibodies over the tomato signal background, showing the expression patterns for two markers in one section, making it easier for the investigator to compare and analyze results. Here we describe three protocols that include (1) cell lineage tracing to examine the fate of embryonic chondrocytes in condyle formation, (2) cell lineage tracing to examine the fate of postnatal chondrocytes, and (3) a combination of cell lineage tracing with colocalization using immunofluorescence.

2 2.1

Materials Reagents

1. Tamoxifen. 2. Corn oil. 3. Ethanol. 4. Xylazine. 5. Ketaset. 6. Phosphate buffered saline (PBS). 7. Paraformaldehyde (PFA). 8. Ethylenediaminetetraacetic acid (EDTA). 9. Sucrose. 10. Optimal Cutting Temperature (OCT) compound. 11. DAPI/nonfluorescing antifade mounting solution. 12. Hyaluronidase. 13. Phosphate buffered saline with 0.1% Tween 20 (PBST). 14. Bovine serum albumin (BSA). 15. Animal serum (Goat serum for RUNX2 and DMP1 antibodies).

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16. Primary antibody for marker proteins of interest (RUNX2 and DMP1). 17. Secondary antibody. 18. Rabbit IgG for positive control. 2.2

Solutions

1. Tamoxifen solution (1 mL at 10 mg/mL): Combine 10 mg of tamoxifen powder with 0.9 mL of corn oil and 0.1 mL of ethanol (see Note 1). 2. Xylazine–Ketaset combination solution (1 mL at a 1:2 ratio): Combine 0.667 mL of Ketaset (5 mg/mL in H2O) with 0.333 mL of Xylazine (1 mg/mL in H2O) (see Note 2). 3. 4% PFA solution (pH 7.4): Combine 40 g of PFA powder with 1 L of 1 PBS. (see Note 3) 4. 10% EDTA solution (pH 7.4): Combine 100 g of EDTA powder with 1 L of 1 PBS. 5. 15% sucrose solution: Combine 150 g of sucrose powder with 1 L of 1 PBS. 6. 30% sucrose solution: Combine 300 g of sucrose powder with 1 L of 1 PBS. 7. 2 mg/mL Hyaluronidase solution (pH 5.0): Combine 2 g of Hyaluronidase powder with 1 L of 1 PBS.

3

Methods

3.1 Cell Lineage Tracing in Embryonic Chondrocytes

To illustrate the use of cell lineage tracing in embryonic chondrocytes, we initially crossed Col10a1-Cre mice with Rosa26tdTomato (B6;129S6-Gt(ROSA)26Sortm9(CAG-tdTomato)Hze/J) mice to obtain Col10a1-Cre; Rosa26tdTomato mice. We next crossed these mice with 2.3Col1a1-GFP mice to generate Col10a1-Cre; Rosa26tdTomato ; 2.3Col1a1-GFP mice for the generation of tissue sections and as subjects for these cell lineage tracing experiments (Fig. 1b). 1. Anesthetize the mice with the Ketaset–Xylazine combination solution by first removing a mouse from its cage. Next, use the left thumb and index finger to grab the skin on the back of the mouse and turn it over, exposing the abdomen. Use the right hand to hold the syringe. The optimal entry point for injection is on the left or right side of hypogastrium, avoiding the liver and bladder. Keep the syringe parallel to the hindlimbs of the mouse and inject intraperitoneally (see Note 2). Confirm the anesthetization by pinching the mouse’s ankle. If there is no reaction, the mouse is unconscious. 2. After the mouse has lost consciousness, fix the four legs of the mouse on a board to entirely expose the abdomen. Saturate the abdomen with 70% ethanol and make an incision from the

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Fig. 2 Transformation from chondrocytes to bone cells in the mandibular condyle in Col10al-Cre; 2.3Col1a1GFP; Rosa26tdTomato Compound Mice In the condylar process from 3-week-old mice, three different colors of cells can be visualized in the condylar process: pure red (chondrocyte-derived bone cells), yellow (red combined with green, indicating chondrocytederived bone cells that express the collagen 1 gene), and pure green (nonchondrocyte derived bone cells with the collagen 1 gene) [5, 6]. (This figure is adapted from Yan et al. (Jing, Y., Hinton, R. J., Chan, K. S., Feng, J. Q. Co-localization of Cell Lineage Markers and the Tomato Signal. J. Vis. Exp. (118), e54982, doi:https://doi.org/ 10.3791/54982 (2016)))

lower abdomen to the neck along the middle line. Pinch and simultaneously pull the skin to the lateral sides to reveal the peritoneal membrane. Use dissection scissors to make a longitudinal incision. Cut off and remove the front ribs to expose the heart. Puncture the left ventricle of the heart with a 22 G syringe, hold the syringe, and simultaneously cut a slot in the right atrium. Slowly inject the 4% PFA, which is perfused along the cardiovascular system while the blood flushes out of the cut from the right atrium. The volume of the PFA for perfusion is 1 mL/g. Perform this step in a Class I biosafety cabinet that is hard-ducted to the building exhaust system.

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3. Peel off the mouse’s skin and put the whole body into a 50-mL polypropylene centrifuge tube that contains 40 mL of 4% PFA to fix overnight at 4  C. 4. Use dissection scissors and #3 and #5 forceps to carefully remove the mandible and hind leg from the body and remove the muscles and tendons on the surface. Perform this step in a Class I biosafety cabinet that is hard-ducted to the building exhaust system. 5. Cut the mandible into two pieces at the distal region of the third molar. Cut the femur and tibia in the midshaft to expose the bone marrow cavity in order to accelerate decalcification. Put the part that includes the condyle and condylar process along with the hind leg into 40 mL of 10% EDTA to decalcify at 4  C for 2–4 days in a 50-mL polypropylene centrifuge tube. 6. Use 50 mL of 15% sucrose to dehydrate the condyle and hind leg overnight at 4  C in a 50-mL polypropylene centrifuge tube. 7. Use 30% sucrose to dehydrate the condyle and hind leg overnight at 4  C in a 50-mL polypropylene centrifuge tube. 8. Embed the sample with OCT along the sagittal plane on the cutting plate in the cryosection machine. Horizontally lay the condyle or the hind leg in the mounting mold. Submerge the tissue in OCT and leave it in the cryosection machine until the OCT freezes. Mount the OCT block on the cutting plate. Wait approximately 15 min before cutting to ensure that the OCT is completely frozen. 9. Cut the condyle and hind leg into 10-μm sections. Collect the sections on slides and store at 20  C. 10. Incubate the slide in a 37  C chamber to remove the water before staining. 11. Wash the slides twice with distilled water for 5 min. 12. Wipe off the water around each section. Use a hydrophobic barrier pen to circle the sections and drop DAPI or nonfluorescing antifade mounting solution into the circle. Carefully lay down the coverslip. 13. Capture fluorescent cell images using a confocal microscope at wavelengths ranging from 488 μm (green) to 561 μm (red). Take multiple stacked images at 200 Hz (dimensions of 1024  1024) using 10, 20, and 63 lenses (Fig. 2). 3.2 Cell Lineage Tracing in Postnatal Chondrocytes

To illustrate the use of cell lineage tracing in postnatal chondrocytes, we crossed Aggrecan-CreERT2 (Agg-CreERT2) mice with Rosa26tdTomato (B6;129S6-Gt(ROSA)26Sortm9(CAG-tdTomato)Hze/J) mice to obtain Agg-CreERT2; Rosa26tdTomato mice. Next, we crossed these mice with 2.3Col1a1-GFP mice to generate

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Agg-CreERT2; Rosa26tdTomato; 2.3Col1a1-GFP mice that were utilized for the following cell lineage tracing experiments (Fig. 1c). 1. Tamoxifen injection (recommend postnatal day 14): First, remove a mouse from its cage. Then, use the left thumb and index finger to grab the skin on the back of the mouse and turn it over, exposing the abdomen. Use the right hand to hold the syringe. The optimal entry point for injection is on the left or right side of hypogastrium, avoiding the liver and bladder. Keep the syringe parallel to the hind legs of the mouse and inject intraperitoneally (see Note 4). 2. Follow Subheading 3.1, steps 2 through 11 above (Fig. 3). 3.3 Combining Cell Lineage Tracing with Immunofluorescence

To illustrate the use of cell lineage tracing in postnatal chondrocytes, we crossed Aggrecan-CreERT2 (Agg-CreERT2) mice with Rosa26tdTomato (B6;129S6-Gt(ROSA)26Sortm9(CAG-tdTomato)Hze/J) mice to obtain Agg-CreERT2; Rosa26tdTomato mice (Fig. 1d). 1. Tamoxifen injection (recommend postnatal day 3): First, remove a mouse from its cage. Then, use the left thumb and index finger to grab the skin on the back of the mouse and turn it over, exposing the abdomen. Use the right hand to hold the syringe. The optimal entry point for injection is on the left or right side of hypogastrium, avoiding the liver and bladder. Keep the syringe parallel to the hind legs of the mouse and inject intraperitoneally. The dosage for injection is 75 mg tamoxifen/kg or 7.5 μL solution/g. 2. Follow Subheading 3.1, steps 2 through 11 above. 3. Wipe off the water around each section. Use a hydrophobic barrier pen to circle the sections on the slide. 4. For Immunofluorescence antigen retrieval, treat the sections with hyaluronidase in a humid chamber at 37  C for 30 min. 5. Rinse with PBST three times. 6. Prepare blocking solution that contains 3% bovine serum albumin (BSA) and 20% goat serum in 1 PBS for RUNX2 or DMP1 immunofluorescent staining. Add solution into circles to completely cover the sections (see Note 5) and incubate in a humid chamber for 1 h at room temperature. DO NOT carry out a wash step. Instead, carefully wipe away the majority of blocking solution. 7. Prepare the primary antibody solution that contains 2% goat serum in 1 PBS for RUNX2 or DMP1 immunofluorescent staining. The concentration of the primary antibody is 1:400 for RUNX2 and 1:100 for DMP1. Again add solution into circles to completely cover the sections (see Note 5). 8. Incubate slides at 4  C overnight.

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Fig. 3 Transformation from chondrocytes to bone cells (osteoblasts and osteocytes) in the mandibular condyle and long bone in Agg-CreERT2; 2.3Col1a1-GFP; Rosa26tdTomato compound mice. Cre was activated by tamoxifen on postnatal day 14, and the mice were sacrificed at 4 weeks old. There were three colored

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9. Rinse with 1 PBS three times. 10. Prepare a positive control. A rabbit IgG solution will be used as the control for immunofluorescent staining to avoid false positive results (see Note 6). The solution contains 2% goat serum in 1 PBS. The concentration of the rabbit IgG for RUNX2 control is 1:400 and for DMP1 it is 1:100. 11. Prepare the secondary antibody solutions. These contains 2% goat serum in 1 PBS for Runx2 or DMP1 immunofluorescent staining. Use the secondary antibody at a dilution of 1:500 for each. Add solution into circles to completely cover the sections (see Note 5). 12. Incubate for 2 h at room temperature. 13. Rinse with 1 PBS three times. 14. Wipe off the water around the section and drop DAPI into the circle to cover the section on the slide. Carefully lay down the cover slip. 15. Capture fluorescent cell images using a confocal microscope at wavelengths ranging from 488 μm (green) to 561 μm (red). Take multiple stacked images at 200 Hz (dimensions of 1024  1024) using 10, 20, and 63 lenses (Fig. 4).

4

Notes 1. Dosage for tamoxifen injection is typically 75 mg/kg or 7.5 μL solution/g of total mouse body weight. 2. Dosage of Ketaset–Xylazine solution for injection is typically 30 μL/g of total mouse body weight. The weights of the mice at the ages of 2 weeks, 3 weeks, and 4 weeks old are approximately 7–9 g, 11–13 g, and 16–18 g, respectively. 3. Adjust pH to 7.4 with 2 M NaOH before bringing volume fully to 1 L, and handle PFA in the hood with gloves and a facemask. 4. The dosage for injection is 75 mg tamoxifen/kg or 7.5 μL solution/g. The weight of mice at the age of 2 weeks is approximately 7–9 g.

ä Fig. 3 (continued) cells in the condylar process (a), and epiphysis and metaphysis of long bone (b, c): pure red (chondrocyte-derived bone cells, white arrows), yellow (red combined with green, indicating chondrocytederived bone cells that expressed the collagen 1 gene, white arrows), and pure green (non–chondrocyte derived bone cells with the collagen 1 gene, blue arrows). The protocol was able to provide strong evidence that chondrocytes directly transform into bone cells and contribute to the formation of the condylar process during development [5, 6] (Tm tamoxifen, C cartilage, B bone). (This figure is adapted from Yan et al. (Jing, Y., Hinton, R. J., Chan, K. S., Feng, J. Q. Co-localization of Cell Lineage Markers and the Tomato Signal. J. Vis. Exp. (118), e54982, doi:https://doi.org/10.3791/54982 (2016)))

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Fig. 4 Colocalization of two markers for osteogenic cells with the lineage-tracing background in the condylar process in 2-week-old Agg-CreERT2; Rosa26tdTomato Compound Mice. (a) Chondrocyte-derived bone cells

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5. The volume of the solution depends on the size of the section. Use 50 μL of solution for the condyle and 100 μL for the long bone. 6. The IgG control is necessary for immunohistochemical staining to avoid false-positive signals. The staining for the experimental and control groups needs to be performed simultaneously.

Acknowledgments This study was partly supported by U.S.s National Institutes of Health grants to J.Q.F. (R01DE025014 and DE025659). References 1. Kretzschmar K, Watt FM (2012) Lineage tracing. Cell 148(1–2):33–45. https://doi. org/10.1016/j.cell.2012.01.002 2. Romagnani P, Rinkevich Y, Dekel B (2015) The use of lineage tracing to study kidney injury and regeneration. Nat Rev Nephrol 11(7):420–431. https://doi.org/10.1038/nrneph.2015.67 3. Humphreys BD, DiRocco DP (2014) Lineagetracing methods and the kidney. Kidney Int 86 (3):481–488. https://doi.org/10.1038/ki. 2013.368 4. Jing Y, Hinton RJ, Chan KS, Feng JQ (2016) Co-localization of cell lineage markers and the

tomato signal. J Vis Exp (118):54982. https:// doi.org/10.3791/54982 5. Jing Y, Zhou X, Han X, Jing J, von der Mark K, Wang J, de Crombrugghe B, Hinton RJ, Feng JQ (2015) Chondrocytes directly transform into bone cells in mandibular condyle growth. J Dent Res 94(12):1668–1675. https://doi.org/10. 1177/0022034515598135 6. Hinton RJ, Jing Y, Jing J, Feng JQ (2017) Roles of chondrocytes in Endochondral bone formation and fracture repair. J Dent Res 96 (1):23–30. https://doi.org/10.1177/ 0022034516668321

ä Fig. 4 (continued) expressing Runx2 show yellow color in the nuclei (white arrows) on the surface of the trabecular bone beneath the condyle cartilage. Bone cells without Runx2 expression are pure red, and represent mature chondrocyte-derived bone cells in the bone matrix (yellow arrows). Sporadic green nuclei can be seen in either non–chondrocyte-derived bone cells or bone cells which transformed from chondrocytes before Cre activation (Tm: tamoxifen). B) Chondrocyte-derived bone cells in the bone matrix can be observed based on DMP1 staining in their cell bodies (white arrows). A few of the osteocytes are positive for DMP1 but lack red color in their cell bodies (yellow arrows). These cells are either non–chondrocyte-derived bone cells or chondrocyte-derived bone cells arising before tamoxifen injection. (This fig is adapted from Yan et al. (Jing, Y., Hinton, R. J., Chan, K. S., Feng, J. Q. Co-localization of Cell Lineage Markers and the Tomato Signal. J. Vis. Exp. (118), e54982, doi:https://doi.org/10.3791/54982 (2016)))

Chapter 19 Immunofluorescent Staining of Adult Murine Paraffin-Embedded Skeletal Tissue Neta Felsenthal and Elazar Zelzer Abstract Immunohistochemistry, or immunolabeling, is a key method for the identification of protein expression and localization. Successful detection relies on a low signal-to-noise ratio, which is affected greatly by antibody specificity as well as the staining protocol. Immunohistochemistry in the mouse is challenging, particularly in adult skeletal tissue, due to the need for long decalcification, high autofluorescence and high levels of endogenous peroxidase. Here, we describe a highly sensitive protocol for protein detection in decalcified paraffin-embedded sections from adult mouse skeletal tissue. By using four levels of amplification, this method allows for the identification of even low-abundance proteins. Key words Immunohistochemistry, Tyramide signal amplification (TSA), Horseradish peroxidase (HRP), Antigen retrieval, Protein expression, Tissue sections, Paraffin

1

Introduction Proteins are among the most fundamental components of life. They are one of the most abundant organic molecules in living organisms and they have an incredibly diverse range of functions. Structural proteins constitute the cytoskeleton as well as the extracellular matrix that holds and forms our tissues. Enzymes catalyze metabolic reaction and regulatory proteins such as transcription factors control gene expression and virtually all other inner cellular functions. Other proteins are involved in the immune system, cellular communication, synthesis, transportation and degradation of molecules, cell cycle, and more. Given the immense involvement and control of proteins over the biological process, identifying protein localization and distribution is a fundamental aspect in any research question. Immunolabeling or immunohistochemistry (IHC) allows for the identification of a protein of interest in situ, that is, in its native environment inside a cell and within a tissue. The method relies of the high affinity and binding between an antibody and an antigen,

Matthew J. Hilton (ed.), Skeletal Development and Repair: Methods and Protocols, Methods in Molecular Biology, vol. 2230, https://doi.org/10.1007/978-1-0716-1028-2_19, © Springer Science+Business Media, LLC, part of Springer Nature 2021

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which is the protein of interest. Following the initial binding between an antibody and its target, comes a stage of antibody identification and visualization. The primary antibody can be directly labeled by a fluorophore or a chromogen, or it can be identified using a secondary antibody and signal amplification [1, 2]. Different levels of signal amplification exist (Table 1 summarizes the different possibilities for detecting a primary antibody and the different levels of signal amplification), and these are constantly being improved and developed. These advances allow for visualization of even low levels of proteins as well as the detection and colocalization of multiple proteins. While some organisms are easy to stain, have specific antibodies and offer a low signal-to-noise ratio, in other organisms, like the mouse, IHC is a challenging procedure that requires extensive calibrations and modifications, depending on sample type and antibodies [3]. Adult skeletal tissue presents an even greater challenge [4]. The need for long decalcification, high autofluorescence from bone and muscle tissue and high levels of endogenous peroxidase require unique staining protocols in order to facilitate specific detection with low background, while maintaining tissue morphology. The protocol presented here has been developed as a tool to identify proteins that have low abundance in formalin-fixed, decalcified paraffin-embedded adult skeletal tissue sections. It was greatly contributed by similar protocols that were shared by many colleagues [5, 6]. In this protocol, a biotinylated secondary antibody binds to the primary antibody against the target protein. This secondary antibody is then detected using a horseradish peroxidase (HRP)-conjugated streptavidin molecule, which is incubated with tyramide signal amplification (TSA) system fluorophore to produce a fluorescent signal, thus creating a four-level amplification. This enables the detection of low-abundance proteins and transcription factors while maintaining tissue integrity. Moreover, this technique can be easily combined with other immunofluorescence protocols for multiplex labeling [7].

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Materials 1. Hot plate or slide warmer. 2. Xylene. 3. Absolute ethanol. 4. Phosphate buffered saline (PBS). 5. Parafilm. 6. 30% hydrogen peroxide (H2O2). 7. 20 mg/mL Proteinase K (PK); stock concentration. 8. 4% paraformaldehyde (PFA) in PBS.

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Table 1 Methods of detection of primary antibody Amplification level Detection method

Compatibility and Notes Fast labeling protocol, compatible with high-abundance proteins

1

Primary antibody is conjugated to a fluorophore Example: Cy3-anti-tdTomato

2

A secondary antibody conjugated to a fluorophore binds to the primary antibody FITC-donkey anti-rabbit

2+

Using HRP and AP involves an enzymatic A secondary antibody, conjugated to reaction, where substrate degradation horseradish peroxidase (HRP) or to results in a colorimetric reaction. High alkaline phosphatase (AP), binds to the substrate concentration, long primary antibody. Signal is produced by incubation time might result in applying an enzymatic substrate—Either nonspecific signal DAB or TSA (for HRP) or NBT (for AP)—And performing a colorimetric reaction HRP-anti-rabbit AP-anti-rabbit

3

A secondary antibody, conjugated to biotin, Good signal amplification, suitable for weak staining. Does not require binds to the primary antibody. The quenching of endogenous peroxidase secondary antibody is then detected using a fluorophore conjugated to avidin or streptavidin Biotin-anti-rabbit; avidin Cy5

4

A secondary antibody, conjugated to biotin, Maximum amplification currently available, compatible with binds to the primary antibody. The low-abundance proteins secondary antibody is then detected using avidin conjugated to HRP. Signal is produced by applying an enzymatic substrate—Either DAB, TSA (for HRP) or NBT (for AP)—And performing a colorimetric reaction Biotin-anti-rabbit; avidin-HRP; TSA

9. 10 mM sodium citrate buffer, pH 6. 10. Tris–NaCl–Tween (TNT) buffer: 100 mM Tris–HCl pH 7.5, 150 mM NaCl, add Triton X-100 for a final concentration of 0.1%. 11. Tris–NaCl blocking (TNB) buffer: 100 mM Tris–HCl pH 7.5, 150 mM NaCl, 0.5% blocking reagent (Roche, see Note 1). 12. Humidified chamber for slides. 13. Antibody of choice, with a compatible biotinylated secondary antibody.

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14. Streptavidin-HRP. 15. TSA amplification kit. 16. 40 ,6-diamidino-2-phenylindole (DAPI). 17. Water-based mounting medium. 18. Coverslips.

3

Methods 1. Bake slides on a hot plate at 60  C for 45 min (min)—1 h (h) (see Note 2). 2. Allow the tissue to cool down to room temperature (RT) for 5–10 min, and dewax in two washes of xylene for 5 min each. 3. Rehydrate to PBS: wash twice in 100% EtOH for 5 min each; once in 75% EtOH, for 5 min, 50% EtOH for 5 min, 25% EtOH for 5 min and wash twice in PBS for 5 min. 4. Fix slides in 4% PFA/PBS for 10 min at RT. 5. Rinse in PBS, followed by additional two washes in PBS for 5 min each. 6. Antigen retrieval: For postnatal day (P) 28 tendon-bone attachment, mix 120 μL of PK in 200 mL PBS in a Coplin jar. Incubate for 15 min at RT (see Notes 3 and 4). 7. Wash twice in PBS for 5 min each. 8. Quench endogenous peroxidase: prepare fresh 3% H2O2 in MeOH, mix well and incubate slides for 45 min (see Note 5). 9. Rinse in TNT. 10. Wash twice in TNT for 5 min each. 11. Blocking: apply 100 μL of blocking solution to each slide, place in a humid chamber and cover with a Parafilm strip. Incubate for 1 h at RT (see Note 6). 12. Gently dab excessive blocking solution from the slide. Dissolve primary antibody in TNB and apply 100 μL of diluted primary antibody to each slide. Some examples of primary antibodies and their dilutions are highlighted in Figs. 1, 2 and 3. Cover with a Parafilm strip and incubate overnight at 4  C in a humid chamber (see Notes 7 and 8). 13. The following day, wash the primary antibody three times with TNT for 5 min each. 14. Dilute biotinylated secondary antibody in TNB. Appropriate secondary antibody is selected based on primary antibody and tissue section species. Apply 100 μL per slide, cover the slides with Parafilm and incubate in a humid chamber for 1 h at RT.

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Fig. 1 Immunohistochemistry staining for ENPP1 (red) in a P28 mouse Achilles tendon-bone attachment section. Antigen retrieval with PK. Primary antibody dilution is 1:100, secondary antibody is biotin anti-goat (Jackson ImmunoResearch), dilution is 1:150. Nuclei are counterstained with DAPI (blue). Scale bar: 50 μm

15. Wash three times in TNT for 5 min each. 16. Dilute streptavidin-HRP 1:200 in TNB blocking solution. Apply 100 μL per slide, cover the slides with Parafilm and incubate in a humid chamber for 1 h at RT (see Note 9). 17. Wash three times in TNT for 5 min each. 18. Develop antibody signal: dilute Cy3/FITC/Cy5 fluorophore 1:200 in amplification reagent (supplied with the kit—TSA detection system), apply 100 μL to each slide and cover with Parafilm. Incubate in a humid chamber for 15–20 min at RT (see Notes 10 and 11). 19. Quench remaining tissue peroxidase using 3% H2O2 in TNT for 10 min. 20. Wash three times in TNT for 5 min each. 21. Counterstain the nuclei with DAPI for 5 min at RT. 22. Wash twice in PBS for 3 min each. 23. Mount slides using water-based mounting medium such as Immuno-mount and cover.

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Fig. 2 Immunohistochemistry staining for RUNX2 (red, Cell Signaling) in a P28 mouse tibia bone section. Antigen retrieval with PK. Primary antibody dilution is 1:200, secondary antibody is biotin anti-rabbit (Jackson ImmunoResearch), dilution is 1:200. Nuclei are counterstained with DAPI (blue). Scale bar: 50 μm

24. Image using fluorescent microscope and appropriate filters. Sample images are provided for ENPP1, RUNX2, and Phospho-SMAD 1/5 in Figs. 1, 2 and 3.

4

Notes 1. TNB (Blocking buffer) should be prepared in advance, as the blocking reagent takes a long time to dissolve. Make a stock reagent in a 50 mL tube, heat in a water bath at 65  C until the blocking reagent has completely dissolved, aliquot and keep frozen at 20  C. 2. Baking slides on a hot plate is essential to prevent tissues from detaching from the slide during the staining protocol. Baking time can be extended if tissue appears damaged or detaches during staining. When staining embryonic tissue, incubation can be shortened to 15 min. 3. Different antibodies may require different methods of antigen retrieval, and this should be adapted per antibody. PK concentration and incubation time should also be adapted according to section thickness and tissue age. Too much PK can result in a high background, while too little PK can result in a low signal.

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Fig. 3 Immunohistochemistry staining for Phospho-Smad 1/5 (red, Cell Signaling) in a P28 mouse articular cartilage section. No antigen retrieval is needed. Primary antibody dilution is 1:200, secondary antibody is biotin antirabbit (Jackson ImmunoResearch), dilution is 1:200. Nuclei are counterstained with DAPI (blue). Scale bar: 50 μm

4. Another common method of antigen retrieval uses 10 mM sodium citrate buffer, pH 6. Place slides in a Coplin jar containing sodium citrate buffer. Put in the microwave, hot water bath or pressure cooker, bring to boil on maximum intensity for 3–5 min; then, reduce heat and cook with light bubbling for an additional 10 min. Let the slides cool completely in citrate buffer before continuing with the protocol. In case of tissue damage while using citrate buffer for antigen retrieval, consider preheating the solution to maximum 80  C in a water bath on a hot plate, and incubate at 70–80  C for 10–15 min. Let the slides cool completely in citrate buffer before continuing with the protocol [8]. 5. Endogenous peroxidase often causes background in adult skeletal tissue. Quenching should be optimized by using 1–3% H2O2 and reducing or extending quenching time. Additional methods include incubation of the slides in 100 mM sodium azide in PBS for 45 min. Additional commercial blockers are also available. Identifying endogenous peroxidase in tissues is possible by incubating tissue sections following antigen retrieval with the development substrate from step 16.

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6. Prepare a humid chamber by soaking a piece of tissue or Kimwipe in water at the bottom of a slide box. Incubation with the blocking solution can be extended. 7. Alternatively, the primary antibody can be incubated for 1 h at RT or at 37  C; however, these conditions are possible only if the staining is successful after an overnight incubation at 4  C. The proper dilution should be calibrated for each antibody. 8. If the staining is very weak or no signal is detected when using a high antibody concentration, consider incubating the primary antibody overnight at RT. 9. Streptavidin-HRP can be diluted 1:200 to 1:1000 and should be calibrated in case of high background staining. 10. Optimization of TSA fluorophore dilution and incubation time might be necessary, depending on antibody and signal intensity required. For staining tissue that contains bone, consider using Cy5 due to high autofluorescence from bone tissue and red blood cells in bone marrow, which can interfere with imaging when using Cy3/FITC fluorophores. 11. Alternatively, TSA fluorophore can be diluted in 0.003% H2O2 in 0.1 M boric acid, pH 8.5. References 1. Shi ZR, Itzkowitz SH, Kim YS (1988) A comparison of three immunoperoxidase techniques for antigen detection in colorectal carcinoma tissues. J Histochem Cytochem 36:317–322. https://doi.org/10.1177/36.3.3278057 2. de Matos LL, Trufelli DC, de Matos MGL, da Silva Pinhal MA (2010) Immunohistochemistry as an important tool in biomarkers detection and clinical practice. Biomark Insights 5:9–20 3. Ward JM, Rehg JE (2014) Rodent immunohistochemistry. Vet Pathol 51:88–101. https://doi. org/10.1177/0300985813503571 4. Akkiraju H, Bonor J, Nohe A (2016) An improved immunostaining and imaging methodology to determine cell and protein distributions within the bone environment. J Histochem Cytochem 64:168–178. https://doi.org/10. 1369/0022155415626765 5. Liu ES, Martins JS, Zhang W, Demay MB (2018) Molecular analysis of enthesopathy in a

mouse model of hypophosphatemic rickets. Development 145:dev163519. https://doi. org/10.1242/dev.163519 6. Mathew SJ, Hansen JM, Merrell AJ et al (2011) Connective tissue fibroblasts and Tcf4 regulate myogenesis. Development 138:371–384. https://doi.org/10.1242/dev.057463 7. Stack EC, Wang C, Roman KA, Hoyt CC (2014) Multiplexed immunohistochemistry, imaging, and quantitation: a review, with an assessment of Tyramide signal amplification, multispectral imaging and multiplex analysis. Methods 70:46–58. https://doi.org/10.1016/ J.YMETH.2014.08.016 8. Jiao Y, Sun Z, Lee T et al (1999) A simple and sensitive antigen retrieval method for freefloating and slide-mounted tissue sections. J Neurosci Methods 93:149–162. https://doi. org/10.1016/S0165-0270(99)00142-9

Chapter 20 Detection of Hypoxic Regions in the Bone Microenvironment Wendi Guo and Colleen Wu Abstract Oxygen serves as a critical environmental factor essential for maintaining the physiological state of a tissue. Hypoxia, or low oxygen, triggers a cascade of events which allows for cells to adapt to low oxygen tensions and to facilitate oxygen delivery required to maintain tissue homeostasis. In the bone microenvironment (BME), vascular heterogeneity, poor perfusion rates of blood vessels, and high metabolic activity of hematopoietic cells result in the generation of a unique hypoxic landscape. Importantly, in this region, hypoxia and its downstream effectors are associated with establishing stem cell niches and regulating the differentiation of committed progenitors. Given the functional importance of the hypoxic bone niche, visualizing regions of hypoxia may provide valuable insights into the mechanisms that regulate tissue homeostasis. Here, we describe the utilization of the nitroimidazole derivative, pimonidazole, to detect hypoxic regions within the BME. Key words Hypoxia, microenvironment

1

Pimonidazole,

Immunofluorescence,

Immunohistochemistry,

Bone

Introduction During oxidative phosphorylation oxygen acts as the final electron donor to generate ATP, thereby providing energy essential for cellular function. For this reason, multicellular organisms have developed complex mechanisms to regulate and maintain oxygen homeostasis. For example, activation of the hypoxia inducible factor (HIF) signaling pathway is a hallmark response to oxygen insufficiency. Here, low oxygen tension causes the stabilization and nuclear translocation of HIF transcription factors, where they bind to specific DNA sequences to drive expression of genes involved in the cellular adaptation to hypoxic environments [1]. As physiological oxygen tension in human tissues range from 11% to 1% O2 [2], it is important to note that hypoxia is a relative term and for the purposes of this chapter, physiologic hypoxia will be defined as the capacity to induce activation of the HIF signaling pathway.

Matthew J. Hilton (ed.), Skeletal Development and Repair: Methods and Protocols, Methods in Molecular Biology, vol. 2230, https://doi.org/10.1007/978-1-0716-1028-2_20, © Springer Science+Business Media, LLC, part of Springer Nature 2021

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In the BME, live imaging of murine calvaria reveals oxygen values of 4.2% near the periosteum and 1.7% in deeper perisinusoidal regions within the marrow space [3]. Notably, hypoxia and HIF signaling influences the differentiation of multipotent mesenchymal progenitor cells [4, 5], couples osteogenesis to angiogenesis [6], regulates osteoblast function [7–9], controls proliferation and survival of chondrocytes [10], and aids in establishing the hematopoietic stem cell (HSC) niche [11]. Moreover, in genetically engineered mouse models, aberrant activation or loss of the hypoxic response is associated with pathological conditions such as polycythemia and osteoarthritis [11, 12]. For these reasons, the ability to identify cells residing in hypoxic regions will not only aid in elucidating the contribution of oxygen to the maintenance of tissue homeostasis in the BME, but may also yield potential therapeutic targets to treat bone disorders [13]. A variety of methods have been employed for mapping oxygen concentrations in vivo. Until recently the polarographic oxygen electrode, or Clark’s electrode, was the only available technique to directly assess tissue O2. In this method, a probe containing two electrodes is physically inserted into a tissue and a known magnitude of voltage is applied to the cathode where oxygen is reduced. O2 measurements can be taken because the difference in voltage between the cathode and anode is proportional to the amount of molecular oxygen reduced at the cathode [14]. While this method provides a relatively easy way to assess oxygen tension in a local environment, several disadvantages are associated with this technique. Namely, the electrode consumes oxygen during the measurement process and the invasive nature of the probe can potentially damage the microvasculature, both of which can contribute to inaccurate readings [2]. In addition, the needle electrode lacks spatial resolution, allowing only for the measurement of oxygen at a single point. Given the heterogenous oxygen distribution within the BME [3, 15], a single point measurement may fail to provide an accurate representation of the hypoxic environment and the potential cellular responses to oxygen deprivation in this tissue. In contrast to Clark’s electrode, Spencer et al. [3] developed a method which provides high spatial resolution of oxygen distribution, is relatively noninvasive, and does not disrupt the local microenvironment. Here, a metalloporphyrin-based two-photon– enhanced phosphorescent probe is injected systemically into live animals and subsequent bimolecular collisions with dissolved oxygen quench the emissive triplet state of the probe [3]. Consequently, direct measurements of local oxygen tension can be made using an established phosphorescence decay time versus oxygen concentration calibration curve. However, this technique comes with its own set of caveats. For example, traditional two-photon– microscopy objectives with working distances of 2–3 mm may not

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be suitable for imaging all tissue types, especially tissues such as murine long bones which exceed this range of measurement [16]. An alternative to direct measurements of O2 is the use of nitroimidazole derivatives such as pimonidazole. These reagents have garnered particular interest as exogenous hypoxia markers due to their chemical structures which confer sensitivity to low oxygen tensions, thereby providing information on the relative oxygenation of a tissue. Pimonidazole is reduced under conditions of hypoxia by nitroreductase enzymes to form a hydroxylamine intermediate. This intermediate subsequently forms stable and irreversible adducts to nucleophilic groups such as thiol groups in proteins or DNA [17] (Fig. 1). The oxygen dependent property of pimonidazole results from oxygen competing for the addition of the first electron to pimonidazole during the initial reduction. Based on experiments comparing oxygen microelectrode measurements of O2 and misonidazole binding [18], pimonidazole has an established tissue O2 threshold of
10 mmHg (1.3% O2) with half maximal inhibition of binding at 0.15% O2 [19]. Conversely, the binding rate of pimonidazole decreases sharply at oxygen concentrations above 1.3% O2 [18]. Importantly, in relation to other hypoxia markers, pimonidazole staining has been demonstrated to be consistent with immunostaining patterns of oxygen-regulated proteins such as carbonic anhydrase 9 and LDH-5, demonstrating specificity for hypoxic cells [20, 21]. There are some important points to note when utilizing pimonidazole. Firstly, an estimated 80% of hydroxylamine intermediates are fragmented by reaction with water, and thus, not all reductively activated intermediates bind to cellular molecules [22]; however, robust labeling can still be achieved and used to predict treatment outcomes (Fig. 2) [23, 24]. Secondly, tissues such as the liver, possess high concentrations of nitroreductase enzymes which can reduce pimonidazole in an oxygen-insensitive manner. While reverse perfusion experiments show that binding of pimonidazole in the pericentral region of the liver is not due to high concentrations of cytochrome redox enzymes, care should be taken when examining tissues that possess high concentrations of redox enzymes [19]. In these cases, secondary methods should be employed to rule out nonspecific binding of pimonidazole. While pimonidazole is unable to give an exact measurement of oxygen concentration, there are several advantages to using this nitroimidazole derivative for detecting hypoxia in vivo. First, pimonidazole is a well-established standard marker of hypoxia that is widely used and commercially available. Secondly, pimonidazole purchased through Hypoxyprobe is formulated as a hydrochloride salt with high solubility in aqueous solution, yet because pimonidazole itself is highly lipophilic with an octanol–water partition coefficient of 8.5, it can penetrate all tissues including the brain [25]. Thirdly, pimonidazole binding can be detected by either

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Fig. 1 Mechanism of action of pimonidazole. Under conditions of hypoxia, nitroreductase enzymes catalyze the reduction of pimonidazole to form nitroso and hydroxylamine intermediates. These reductively activated intermediates bind irreversibly to cellular nucleophilic groups in proteins of hypoxic cells

Fig. 2 Labeling of hypoxic regions in the BME. (a) Representative image of immunohistochemical analysis of murine tibia isolated from wild type mouse. Binding of primary antibody against pimonidazole mediated oxygen dependent adduct formation is visualized by cells which have oxidized 3,3’-Diaminobenzidine (DAB) (brown). An IgG control is utilized to demonstrate specificity of primary antibody binding. Black arrowheads denote hypertrophic chondrocytes which serve as an internal positive control. (b) Representative immunofluorescent image of a tibia isolated from a LepRCre; R26-tdTomato reporter mouse. Pimonidazole binding (green) is visualized using Hypoxyprobe rabbit antisera and an Alexa Fluor 488–conjugated secondary antibody. White arrowheads point to cells that demonstrate high colocalization (yellow) of pimonidazole and leptin receptor expressing stromal cells (red). 40 ,6-diamidino-2-phenylindole (DAPI) staining of nucleus (blue). Trabecular bone (TB). Tibias isolated from 3-month B6 mice which were injected with 60 mg/kg pimonidazole hydorchloride 45 min prior to tissue harvest

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immunofluorescence in cryopreserved tissue specimens or by peroxidase-catalyzed reaction in formalin-fixed, paraffin-embedded tissue sections. Therefore, this method can be readily and easily incorporated by laboratories to provide an accurate representation of hypoxic regions in a tissue. Importantly, pimonidazole serves as a simple and effective way to visualize the hypoxic landscape and relative oxygen distribution. In this chapter, we outline a protocol using pimonidazole to label and visualize regions of hypoxia in bone tissue.

2

Materials

2.1 Pimonidazole Injection

1. 1 PBS. 2. Pimonidazole HCl (Hypoxyprobe Omni Kit, also contains rabbit antisera to detect adducts) stock solution: Add 10 ml of 1 PBS to 1 g of pimonidazole hydrochloride to make a stock solution at 100 mg/ml. Solution will appear yellow in color. Aliquot and store stock solution at 4  C protected from light (see Note 1). 3. Scale. 4. 25-gauge needles. 5. 1 ml syringes.

2.2

Tissue Isolation

1. Surgical dissection tools: scissors, forceps. 2. 70% ethanol. 3. Tissue cassettes. 4. Pencil.

2.3 Frozen Tissue Sample Preparation

1. 4% paraformaldehyde (PFA). 2. 15% sucrose in 1 PBS. 3. 30% sucrose in 1 PBS. 4. Embedding medium: Cryomatrix (Shandon). 5. Dry ice. 6. Tape-transfer windows (Electron Microscopy Sciences). 7. UV optical adhesive (Norland Products). 8. Roller. 9. Cryostat. 10. High-profile Biosystems).

stainless-steel

microtome

blade

(Leica

11. UV light: CryoJane Tape-Transfer System (Electron Microscopy Sciences). 12. Slide box.

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2.4 Paraffin Tissue Sample Preparation

1. 10% neutral buffered formalin. 2. 20% ethylenediaminetetraacetic acid (EDTA) in distilled H2O: add EDTA 50 g at a time using sodium hydroxide pellets to dissolve EDTA with constant stirring on a hot plate. Once all the EDTA is dissolved, wait until solution reaches room temperature and adjust pH to 7.4. Store at room temperature. 3. 70% ethanol. 4. Rocker. 5. Microtome. 6. Low-profile Biosystems).

2.5 Pimonidazole Staining and Immunofluorescent Detection

stainless-steel

microtome

blade

(Leica

1. Humidified slide chamber. 2. Hydrophobic pen. 3. Slide mailer. 4. Rabbit antisera (Hypoxyprobe Omni Kit). 5. Anti-rabbit fluorescent secondary antibody: such as goat antirabbit IgG Alexa Fluor® 488. 6. 1 PBS-Tween 20 (PBS-T): 0.1% Tween 20 in 1 PBS. 7. Blocking buffer for frozen sections: 1.5% serum of appropriate species for secondary antibody in 1 PBS-T. 8. DAPI mounting medium. 9. Clear nail polish.

2.6 Enzymatic Detection for Immunohistochemistry

1. Slide warmer. 2. Humidified slide chamber. 3. Hydrophobic pen. 4. 100% xylene. 5. Graded ethanol series: 100%, 95%, 90%, 80%, and 70%. 6. 3% hydrogen peroxide. 7. Enzyme-conjugated secondary antibody: such as anti-rabbit IgG, HRP-linked antibody. 8. Streptavidin/Biotin Blocking Kit (Vector Laboratories). 9. DAKO Protein Block Serum-Free (Agilent). 10. ImmPACT DAB Peroxidase (HRP) Substrate (Vector Laboratories). 11. Streptavidin–HRP conjugate (Calbiochem): Reconstitute 200 μg in 1 ml of distilled H2O for a stock solution of 200 μg/ml. Store at 20  C. 12. Mounting medium (Cytoseal 60 or equivalent).

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3.1 Pimonidazole Injection

1. Pimonidazole hydrochloride working stock: Dilute 100 mg/ ml stock solution 1:10 in 1 PBS to generate a 10 mg/ml working stock solution. 2. Weigh mice to be injected with pimonidazole. 3. Inject mice intraperitoneally with 60 mg/kg of pimonidazole (made in step 1) using a 25-gauge needle and syringe (see Note 2). 4. Set a timer for 45 min before sacrificing mice to harvest tissue (see Note 3).

3.2 Tissue Isolation and Sample Preparation for Frozen Sections

1. 45 min post-injection, euthanize mice by CO2 asphyxiation followed by secondary method of euthanasia approved by the institutional IACUC committee. 2. Spray down mice using 70% ethanol to wet and flatten the hair. Extract long bones, removing as much soft tissue as possible. A lobe of liver can be taken at this time for a positive control. 3. For tibias, cut off the distal end of the bone just proximal to the ankle joint to allow for maximal penetration of fixative into the bone tissue and marrow space. For femurs, cut off the proximal end of the bone just below the femoral head. 4. Place cut long bones in labeled tissue cassettes and immerse cassettes in a large plastic container or glass beaker with 4% PFA overnight (8–12 h) at 4  C (see Note 4). 5. After overnight incubation in 4% PFA, transfer cassettes into a 15% sucrose solution and incubate for 24 h at 4  C. 6. After 24-h incubation in 15% sucrose, transfer cassettes into 30% sucrose solution and incubate for 24 h at 4  C. 7. Gently remove bones from cassettes using forceps and embed bones in Cryomatrix or similar embedding medium. Set embedding resin by placing blocks on a flat surface of dry ice. Blocks can be stored at 20  C until ready to section (see Note 5). 8. Prepare labeled slides by spreading a thin layer of UV-activated optical adhesive on glass slides. Allow slides to sit at room temperature or above for 5 min to allow glue to spread evenly on the slide. Next, place slides, roller and tape windows in cryostat set to 20  C for 5 min or until cool. 9. Take 10-μm sections using a high-profile stainless-steel blade. Place adhesive side of the tape window onto the tissue block and use roller to smooth. Transfer tape with tissue section onto a prepared glass slide and use roller to smooth.

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10. Fix slide using UV light source, sample should have a frosted appearance when fixed as opposed to a relatively clear unfixed appearance. If using the CryoJane Tape-Transfer System, flash UV source four to five times with 1 min in between each flash. After UV fixation, use roller to smooth tape window again before peeling off the tape. Placed finished slide into a cooled slide box. Store slides at 20  C until ready to stain. 3.3 Sample Preparation for Paraffin Sections

1. Collect tissues of interest in labeled cassettes and immerse cassettes in a large plastic container or glass beaker with 10% neutral buffered formalin for 24 h at 4  C. 2. After 24 h in 10% neutral buffered formalin, change solution to fresh 10% neutral buffered formalin and fix tissue for an additional 24 h at 4  C. 3. After the additional 24 h of fixation, remove cassettes and place in fresh 1 PBS for 24 h at 4  C. 4. For long bones, decalcify in 20% EDTA for 72 h at room temperature on a rocker. 5. Following 72 h of decalcification in 20% EDTA, rinse cassettes three times for 1 min each with distilled H2O and place in 70% ethanol overnight at 4  C or until ready to paraffin embed. 6. Using a microtome equipped with a low-profile stainless-steel blade, take 5-μm paraffin sections to place on glass slides. Allow sections to completely dry at room temperature overnight.

3.4 Pimonidazole Staining and Detection of Hypoxic Regions by Immunofluorescence

1. Thaw slides for 30 min at room temperature. 2. Cross-link slides for an additional 10 min with a UV light source if neccessary. 3. Using a slide mailer, wash slides three times for 5 min each with 1 PBS-T. 4. Using a hydrophobic pen, carefully draw a circle around the tissue sample. 5. Place marked slides in a humidified slide chamber filled with 1 PBS. 6. Using a pipette, add blocking buffer consisting of 1.5% goat serum if using a goat-derived secondary antibody in 1 PBS-T to completely coat each sample making sure that buffer does not run off the area marked by the hydrophobic pen. Incubate for 1 h at room temperature. 7. To prepare primary antibody, dilute rabbit anti-pimonidazole antibody 1:100 in blocking buffer (see Note 6). 8. After 1h incubation with blocking buffer, pour off blocking buffer, and add primary antibody prepared in step 7. Incubate overnight in a humidified slide chamber at 4  C.

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9. After overnight incubation with primary antibody, rinse slides three times for 5 min each with 1 PBS-T. 10. Dilute secondary antibody with blocking buffer according to manufacturer’s instructions. For goat anti-rabbit IgG Alexa Fluor 488, use a 1:200 dilution. Pipette secondary antibody onto samples and incubate for 1 h at room temperature in the dark. 11. After incubation with secondary antibody, rinse slides three times for 5 min each with 1 PBS-T. 12. Wipe off hydrophobic pen markings and place one drop of DAPI mounting medium in the center of each sample. Gently place a coverslip onto each slide being careful to avoid bubbles. Clear nail polish can be used to seal the coverslip in place. 13. Slides can be stored temporarily at 4  C in the dark until ready to image with a fluorescence microscope. 3.5 Detection of Hypoxic Regions by Peroxidase-Catalyzed Reaction

1. Place slides on a 60  C slide warmer overnight prior to staining. 2. Deparaffinize sections in xylene for 2 min. Repeat 2. 3. Hydrate sections in a graded ethanol series: 100% ethanol three times for 2 min, 95% ethanol three times for 2 min, 90% ethanol three times for 2 min, 80% ethanol three times for 2 min, 70% ethanol three times for 2 min. 4. Rinse slides in distilled H2O for 5 min. 5. Immerse slides in 3% hydrogen peroxidase to quench endogenous peroxidases for 15 min at room temperature. 6. After immersion with 3% hydrogen peroxidase, rinse gently in distilled H2O for 5 min. 7. Using a hydrophobic pen, draw a circle around the tissue sample. Immerse slides in 1 PBS until ready to stain. 8. Place slides in a humidified slide chamber and block nonspecific streptavidin binding sites using the streptavidin solution from the Streptavidin/Biotin Blocking Kit. Incubate sample in streptavidin solution at room temperature for 15 min. 9. Perform a quick rinse with 1 PBS after blocking with streptavidin solution. Block nonspecific biotin binding sites using the biotin solution from the Streptavidin/Biotin Blocking Kit. Incubate sample in biotin solution at room temperature for 15 min. 10. Perform a quick rinse with 1 PBS after blocking with biotin solution. Block nonspecific antibody binding sites using DAKO Protein Block Serum-Free. Incubate sample in protein block at room temperature for 5 min. 11. To prepare primary antibody, dilute rabbit anti-pimonidazole antibody 1:100 in 1 PBS-T.

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12. Pour off excessive block, do not rinse. Coat tissue sample with anti-pimonidazole antibody. Incubate overnight in a humidified slide chamber at 4  C. 13. Next day, rinse slides three times for 5 min each with 1 PBS. 14. Apply enzyme-conjugated secondary antibody to slides diluted to the concentration specified by the manufacturer. For antirabbit IgG HRP-linked antibody (Cell Signaling Technology), use a 1:2000 dilution in 1 PBS-T. Coat samples and incubate for 1 h at room temperature. 15. Rinse slides three times for 5 min each with 1 PBS. 16. Prepare 1:200 dilution of streptavidin–HRP conjugate from 200 μg/ml stock in 1 PBS-T. Coat samples with streptavidin–HRP conjugate in a humidified slide chamber for 1 h at room temperature. 17. Rinse slides with 1 PBS after incubation with streptavidin– HRP conjugate. 18. Add 1 drop (approximately 30 μl) of ImmPACT DAB chromogen concentrate to 1 ml ImmPACT DAB diluent. Mix well. 19. Under a dissecting scope, coat tissue samples with DAB solution. Once the hypertrophic chondrocytes stain brown, quench reaction by immersing the slide in distilled H2O for 5 min (Fig. 2). Repeat for all slides utilizing the same time point (see Note 7). 20. Counterstain if desired. Coverslip with Cytoseal 60 or desired mounting medium.

4

Notes 1. The pimonidazole hydrochloride solution will precipitate when stored at 4  C over time. If precipitate has formed in the stock solution, warm to room temperature until precipitate has dissolved. 2. For a 25 g mouse, a 60 mg/kg dose would be calculated as follows:

  mg ¼ 1:5 mg of pimonidazole needed ð0:025 kg animalÞ  60 kg 1:5 mg ¼ 0:15 ml of pimonidazole working stock to be injected mg 10 stock ml

ð6 μl of pimonidazole working stock per 1 g of mouseÞ Dosages will vary depending on animal species and do not scale linearly. For rodent studies, 60 mg/kg body weight achieves a good balance of economy and effectiveness,

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however, dosages up to 400 mg/kg have been used without toxicity or changes in tissue hypoxia [26]. For other laboratory species, doses will need to be determined on an individual basis. 3. Length of time before harvest is determined based on the plasma half-life of pimonidazole which differs between species. Tissues become anoxic during harvest, thus, it is possible for residual pimonidazole to bind and to give false measures of hypoxia. In mice, pimonidazole has a half-life of 25 min, therefore, a period of 45 min represents approximately 2 half-lives of circulating pimonidazole. Accordingly, approximately ¼ of the initial concentration of pimonidazole is present at the time of harvesting. This low circulating concentration combined with rapid transfer and fixation minimizes nonspecific pimonidazole binding. Longer wait times of 3–5 half-lives can be used if high background binding is suspected. Plasma half-lives will differ depending on the animal species used so wait times will need to be adjusted accordingly. For reference, the plasma half-life for mice is 25 min, rats 45 min, dogs 90 min and humans 300 min. 4. For a small number of bones, each bone can be fixed in a 1.5 ml Eppendorf tube with 1 ml of 4% PFA. 5. For best results, section frozen tissue blocks the same day that they are embedded. Otherwise, allow blocks to completely equilibrate to cryostat temperature (20  C) before sectioning. Do not section tissue block straight from 80  C without allowing time to equilibrate (48 hours at 20  C ), as tissue may tear upon sectioning. 6. No antigen retrieval is required for the detection of oxygen dependent adduct formation. 7. Hypertrophic chondrocytes exist in hypoxic regions and stably express HIF transcription factors. As such, these cells can be used as an internal positive control to determine the time to quench DAB staining. A tissue section which was not incubated with primary antibody can be used as a negative control to determine nonspecific binding (Fig. 2). References 1. Semenza GL (2014) Oxygen sensing, hypoxiainducible factors, and disease pathophysiology. Annu Rev Pathol 9:47–71 2. Carreau A et al (2011) Why is the partial oxygen pressure of human tissues a crucial parameter? Small molecules and hypoxia. J Cell Mol Med 15(6):1239–1253 3. Spencer JA et al (2014) Direct measurement of local oxygen concentration in the bone marrow of live animals. Nature 508(7495):269–273

4. Sheehy EJ, Buckley CT, Kelly DJ (2012) Oxygen tension regulates the osteogenic, chondrogenic and endochondral phenotype of bone marrow derived mesenchymal stem cells. Biochem Biophys Res Commun 417(1):305–310 5. Wagegg M et al (2012) Hypoxia promotes osteogenesis but suppresses adipogenesis of human mesenchymal stromal cells in a hypoxia-inducible factor-1 dependent manner. PLoS One 7(9):e46483

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6. Wang Y et al (2007) The hypoxia-inducible factor alpha pathway couples angiogenesis to osteogenesis during skeletal development. J Clin Invest 117(6):1616–1626 7. Merceron C et al (2019) Hypoxia-inducible factor 2alpha is a negative regulator of osteoblastogenesis and bone mass accrual. Bone Res 7:7 8. Shomento SH et al (2010) Hypoxia-inducible factors 1alpha and 2alpha exert both distinct and overlapping functions in long bone development. J Cell Biochem 109(1):196–204 9. Regan JN et al (2014) Up-regulation of glycolytic metabolism is required for HIF1alphadriven bone formation. Proc Natl Acad Sci U S A 111(23):8673–8678 10. Schipani E et al (2001) Hypoxia in cartilage: HIF-1alpha is essential for chondrocyte growth arrest and survival. Genes Dev 15 (21):2865–2876 11. Rankin EB et al (2012) The HIF signaling pathway in osteoblasts directly modulates erythropoiesis through the production of EPO. Cell 149(1):63–74 12. Yang S et al (2010) Hypoxia-inducible factor2alpha is a catabolic regulator of osteoarthritic cartilage destruction. Nat Med 16(6):687–693 13. Wan C et al (2008) Activation of the hypoxiainducible factor-1alpha pathway accelerates bone regeneration. Proc Natl Acad Sci U S A 105(2):686–691 14. Clark LC Jr et al (1953) Continuous recording of blood oxygen tensions by polarography. J Appl Physiol 6(3):189–193 15. Kusumbe AP, Ramasamy SK, Adams RH (2014) Coupling of angiogenesis and osteogenesis by a specific vessel subtype in bone. Nature 507(7492):323–328 16. Sencan I et al (2018) Two-photon phosphorescence lifetime microscopy of retinal capillary plexus oxygenation in mice. J Biomed Opt 23 (12):1–9

17. Masaki Y et al (2016) Imaging mass spectrometry revealed the accumulation characteristics of the 2-Nitroimidazole-based agent "Pimonidazole" in hypoxia. PLoS One 11(8): e0161639 18. Gross MW et al (1995) Calibration of misonidazole labeling by simultaneous measurement of oxygen tension and labeling density in multicellular spheroids. Int J Cancer 61 (4):567–573 19. Arteel GE et al (1995) Evidence that hypoxia markers detect oxygen gradients in liver: pimonidazole and retrograde perfusion of rat liver. Br J Cancer 72(4):889–895 20. Rademakers SE et al (2011) Metabolic markers in relation to hypoxia; staining patterns and colocalization of pimonidazole, HIF-1alpha, CAIX, LDH-5, GLUT-1, MCT1 and MCT4. BMC Cancer 11:167 21. Olive PL et al (2001) Carbonic anhydrase 9 as an endogenous marker for hypoxic cells in cervical cancer. Cancer Res 61(24):8924–8929 22. Raleigh JA, Koch CJ (1990) Importance of thiols in the reductive binding of 2-nitroimidazoles to macromolecules. Biochem Pharmacol 40(11):2457–2464 23. Olive PL et al (2000) Comparison between the comet assay and pimonidazole binding for measuring tumour hypoxia. Br J Cancer 83 (11):1525–1531 24. Kaanders JH et al (2002) Pimonidazole binding and tumor vascularity predict for treatment outcome in head and neck cancer. Cancer Res 62(23):7066–7074 25. Saunders MI et al (1984) The clinical testing of Ro 03-8799--pharmacokinetics, toxicology, tissue and tumor concentrations. Int J Radiat Oncol Biol Phys 10(9):1759–1763 26. Durand RE, Raleigh JA (1998) Identification of nonproliferating but viable hypoxic tumor cells in vivo. Cancer Res 58(16):3547–3550

Chapter 21 EdU-Based Assay of Cell Proliferation and Stem Cell Quiescence in Skeletal Tissue Sections Marco Angelozzi, Charles R. de Charleroy, and Ve´ronique Lefebvre Abstract Identifying and tracking proliferating and quiescent cells in situ is an important phenotyping component of skeletal tissues in development, physiology and disease. Among all the methods that exist, which include immunostaining for cell cycle-specific proteins, the gold standards use thymidine analogs. These compounds label proliferating cells by being incorporated into de novo–synthesized genomic DNA. 5-bromo20 -deoxyuridine (BrdU) has traditionally been used for this purpose, but its detection is lengthy and requires harsh treatment of tissue sections to give access of anti-BrdU antibody to DNA. An alternative, more recently developed, uses 5-ethynyl-20 -deoxyuridine (EdU). This thymidine analog is detected by click chemistry, that is, covalent cross-linking of its ethynyl group with a fluorescent azide that is small enough to easily penetrate native tissues and reach DNA. In addition to being simple and quick, this EdU-based assay is compatible with other protocols, such as immunostaining, on the same tissue sections. We here describe an EdU-based protocol optimized to label and functionally assess actively proliferating cells as well as slowly dividing cells, including stem cells, in mouse skeletal tissues. Key words Bone, BrdU, Cartilage, Cell labeling, Cell proliferation, EdU, In situ, Skeleton, Stem cell

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Introduction Fine coordination of cell proliferation, differentiation, migration, and death is fundamental in all aspects of skeleton development and adult maintenance [1]. Namely, adequate proliferation of osteochondroprogenitors prior to differentiation is essential to form proper intramembranous and endochondral bones, and sustained proliferation of growth plate chondrocytes before terminal maturation is essential for suitable elongation of endochondral bone templates. Also, timely control of skeletal stem cell proliferation is essential in bone throughout life, as these cells serve as a source of osteoblasts. They are mainly quiescent in homeostatic conditions, but upon such signals as tissue injury they reenter the cell cycle to give rise on one hand to progeny that commit to chondrocyte and osteoblast differentiation to repair tissue damage and on the other

Matthew J. Hilton (ed.), Skeletal Development and Repair: Methods and Protocols, Methods in Molecular Biology, vol. 2230, https://doi.org/10.1007/978-1-0716-1028-2_21, © Springer Science+Business Media, LLC, part of Springer Nature 2021

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hand to cells that return to a quiescent state and maintain the stem cell pool [2]. Perturbations of the mechanisms that control cell proliferation contribute to various types of skeletal diseases, including dysplasias (e.g., achondroplasia), degenerative disease (e.g., osteoporosis), or cancer (e.g., osteosarcomas) [3, 4]. Several methods exist to detect proliferating cells in skeletal tissues and thereby answer questions on the impacts of molecular pathways, pathogenic conditions or drug treatments on skeletal cells. These methods include the immunostaining of proteins associated with cell proliferation, such as the nuclear antigen Ki-67, the proliferating cell nuclear antigen (PCNA) and the minichromosome maintenance complex component 2 (MCM2). A main limitation of these methods is that these proteins are expressed during the S, G1, and G2/M phases of mitosis and thus mark nonquiescent rather than proliferating cells. Another limitation is that they only mark cells cycling at the time of sample collection and are thus unsuited to determine cell proliferation over prolonged periods of time. Gold standard methods to accurately, sensitively and reliably assess cell proliferation in vivo consist in labeling cells newly synthesizing DNA through the use of nucleotide analogs that are injected into live animals. The first such protocol was developed in the late 1950s and used 3H-thymidine [5]. Decades later, safer and more sensitive procedures were developed thanks to the synthesis of halogenated nucleotides. The most frequently used is bromodeoxyuridine (BrdU), but chlorodeoxyuridine (CldU) and iododeoxyuridine (IdU) also exist [6, 7]. Taking advantage of the availability of distinct halogenated nucleotides and antibodies specific to each of them, a protocol was recently developed in which these compounds were sequentially injected in adult mice and used to track neural stem cells and cohorts of progenitor and differentiated cell progeny over time [8]. The same protocol was also utilized advantageously to label skeletal stem cells and their progeny in the epiphyseal growth plate of young mice [9]. While the procedures with halogenated nucleotides remain widely used, their disadvantages are that they require harsh treatment of tissue sections to allow antibody access to genomic nucleotides, and are therefore incompatible with other staining assays. A major improvement was made a decade ago with the development of 5-ethynyl-20 -deoxyuridine (EdU). This synthetic nucleotide carries an alkyne group that can be readily detected through click chemistry [10]. The reaction is a copper-catalyzed [3+3] cycloaddition. It consists in forming a stable triazole ring by covalently coupling the alkyne group present in EdU to an Alexa Fluor®–conjugated azide group. The latter compound is small enough to effectively diffuse through fixed tissues. The assay provides similar results as the traditional BrdU assay, but takes significantly less time, and by avoiding harsh tissue treatments, can be

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coupled with other types of staining [7]. Herein we provide a detailed protocol for an EdU-based assay that can be used either to detect and quantify actively proliferating cells or to track slowly dividing stem cells in the mouse skeleton system.

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Materials EdU and several other reagents described below are classified as toxins, potential mutagens or teratogens. It is therefore imperative that precautions be taken in compliance with all pertaining local regulations when working with these chemicals (see Note 1).

2.1 Labeling Reagents and Detection Kit

1. The assay described in this chapter uses the Click-iT® EdU Alexa Fluor® 488 Imaging Kit (ThermoFisher Scientific). This kit contains all the components needed to label DNA-synthesizing cells and to detect EdU incorporated into DNA. Equivalent kits with the Alexa Fluor® 555, 594, or 647 azides can be purchased from the same company if visualization in a different fluorescent channel is desired (see Note 2). 2. EdU powder can also be purchased separately and dissolved in PBS to obtain a 10 mM solution. Vortex thoroughly and store this solution at 20  C for up to 1 year. Alternatively, dissolve EdU in DMSO.

2.2

Other Materials

Other materials needed for the assay, but not provided in the kit, are as follows: 1. Secondary container to store and transport the EdU solution. This container should close tightly and protect its content from light (opaque dark-color Tupperware®). 2. Phosphate buffered saline (PBS). 3. Paraformaldehyde (PFA) aqueous solution (16% stock). 4. 0.5 M ethylenediamine tetraacetic acid (EDTA) pH 7.4. 5. Histo-Clear or equivalent nontoxic xylene substitute. 6. Ethanol at 100, 95, and 70% in distilled water. 7. 3% (w/v) bovine serum albumin (BSA) in PBS. 8. Distilled water. 9. Vectashield® Mounting Medium or equivalent with or without DAPI. 10. Coplin jars. 11. Superfrost Plus slides or other positively charged glass slides. 12. Glass coverslips. 13. Optional: Immunostaining ImmEdge™ pen).

pen

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pap

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Methods Figure 1 provides a schematic of the major steps involved in the EdU assay, from cell labeling in vivo to quantification of data. All steps are detailed below and, except where otherwise stated, are carried out at room temperature.

3.1 In Vivo Labeling of DNASynthesizing Cells

1. For mouse administration, aspirate 100 μl of EdU solution per 10 g of mouse body weight using a sterile 30G1/2 needle attached to a 1-ml syringe and inject the solution intraperitoneally. Place a treatment card on the injected mouse cage to indicate the presence of a biohazard. Generate biological replicates by using as many animals per experimental group as needed for statistical analysis. Because mice are coprophagous, only house together mice that receive EdU at the same time. 2. To label and quantify actively dividing skeletal cells, the labeling period (i.e., time between EdU injection and mouse euthanasia) is typically 30–60 min for embryos at the gestation days 10.5 (E10.5) to E13.5, 1 to 2 h for E14.5 to E18.5 fetuses, and 2–4 h for postnatal mice. To label and track stem cells, it is recommended to inject EdU several days in a row in order to label the cells during at least one division and to collect tissues no earlier than 1 week after the last EdU injection to allow the label to be diluted out among actively dividing progeny while being detectably retained in stem cells, and to move the mice to a clean cage 1 day after the last EdU injection. 3. When time is over, euthanize the mice as approved by your Institutional Animal Care and Use Committee (e.g., by CO2 asphyxiation followed by cervical dislocation or decapitation). Then, either proceed immediately for tissue collection or place the bodies on wet ice until tissue collection, but for no more than a few hours.

3.2 Preparation of Tissue Sections

1. Dissect mouse embryos or body parts containing skeletal tissues of interest and briefly rinse them in ice-cold PBS. 2. Fix samples in 4% PFA in PBS at 4  C under gentle rocking. Embryos up to E15.5 can be fixed as a whole. E16.5 fetuses and older mice should be skinned and cut in pieces (e.g., head, trunk, and limbs) to facilitate penetration of the fixative and subsequent solutions. Fixation time is 1–2 h for E11.5 to E14.5 mouse embryos, 24 h for E15.5 to E18.5 fetuses, and 48 h from birth onward. 3. Demineralize samples from newborn and older mouse at 4  C in 0.5 M EDTA, pH 7.4, under gentle rocking. Use a volume of EDTA solution that is at least 20 times larger than the sample volume, and change this solution every day.

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Fig. 1 Schematic of the major steps involved in the EdU-based assay of actively proliferating and slowly dividing/stem cells. See Subheadings 1 and 2

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Demineralization takes 1 day for a newborn mouse, 1 week for a 7-day-old mouse, and 3 weeks for an adult mouse. It is complete when the samples are soft enough to be bent easily and without fracture. 4. Rinse samples in PBS at 4  C for 3 30–60 min, and then proceed for embedding, following standard protocols. Both paraffin and frozen sections are suitable for the EdU assay. Choose either type depending on the other assays that you want to perform on the same sections and on adjacent sections. 5. Sections should be 7- to 20-μm thick. Superfrost Plus slides, which bind tissue sections more efficiently than regular glass slides, are recommended. Generate technical replicates by dedicating at least 3 non-adjacent sections per sample to the EdU assay. 6. Just before proceeding for EdU detection, air-dry frozen sections on the bench for 1 h, and then remove the embedding medium by washing the slides in PBS for 3 2 min. If you use paraffin sections, deparaffinize them by dipping the slides in Histo-Clear for 2 5 min. Then, rehydrate them in 100, 95, and 70% ethanol for 2 1 min, and wash in PBS for 3 2 min. 7. Optional: use an immunostaining pen to circle sections with a hydrophobic barrier (see Note 3), but do not let sections dry out. 3.3

EdU Detection

Perform all steps of the assay in the dark (see Note 4) and never let the slides dry out, as this creates fluorescence background. Proceed essentially as instructed in the Click-iT® EdU Alexa Fluor® 488 Imaging Kit. 1. Freshly prepare the Click-iT® reaction cocktail as instructed in the kit and apply it to each section for 30 min. 2. Wash slides with 3% BSA in PBS for 2 min, and then with PBS for 2 min (see Note 5). 3. Stain cell nuclei by incubating sections in Hoechst 33342 solution (Component G of the kit, diluted 1/2000 in 1 PBS) for 2 min. Wash the slides in PBS for 2 2 min, and mount them with Vectashield® medium (without DAPI). Alternatively, skip the incubation with Hoechst 33342 solution and directly mount the slides with Vectashield® medium containing DAPI.

3.4 Image Acquisition and Quantification

1. Photograph tissue sections assayed for EdU incorporation under epifluorescence microscopy conditions: use an excitation wavelength of 350 nm and an emission wavelength of 461 nm to detect DAPI- or Hoechst 33342-stained nuclei, and an

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Fig. 2 EdU assay in articular and growth plate cartilage of juvenile mice illustrating the differences in results obtained when one or several of EdU injections are performed and when short or extended periods of time are applied between cell labeling and tissue analysis. Mice received a single injection of EdU (on postnatal day 14) and were euthanized 2 h, 2 days, 7 days, or 12 days later, or received EdU on three consecutive days (on postnatal days 22, 23, and 24) and were euthanized 12 days after the last injection. Sagittal knee sections were stained for EdU (green signal) and with DAPI (blue signal). The top-row pictures show that articular chondrocytes have a very low rate of proliferation. Only a few cells incorporated EdU in 2 h (left). Cell doublets are seen 2, 7, and 12 days after EdU injection and their numbers are similar to those of single cells after 2 h, indicating that the cells underwent one division within 2 days after labeling and none in the next 10 days. Note that most EdU-positive cells are located in the superficial layers of articular cartilage and gave rise to progeny aligned with the articular surface (increasing the tissue surface), and that most other EdU-positive cells are located in the upper half of the tissue (non-mineralized cartilage) and proliferated in columns perpendicular to the tissue surface (increasing the tissue depth). The bottom-row pictures show that many proximal tibial growth plate chondrocytes incorporated EdU in 2 h. These actively proliferating cells were all located in the upper half of the columnar zone (CZ). The labeling of cells in all tissue layers, including the hypertrophic zone (HZ), at day 2 and the loss of this labeling by day 7 indicate that columnar chondrocytes continued to divide and that their progeny progressed through all stages of cell maturation within a few days. By day 12, EdUlabeled cells could only be detected in the resting zone (RZ) of the growth plate. These slowly cycling cells were likely growth plate chondrocyte stem cells. Note that the primary and secondary ossification centers (POC and SOC, respectively) feature bone marrow and bone cells that massively incorporated EdU within 2 h, but that only a few cells were still labeled by day 12, indicating an abundance of actively proliferating cells and a low proportion of slowly dividing cells, presumably corresponding to hematopoietic and skeletal stem cells

excitation wavelength of 495 nm combined with an emission wavelength of 519 nm to detect Alexa Fluor® 488-EdU-positive cells. EdU-positive cell nuclei will appear green and other cell nuclei blue (Fig. 2). 2. Select section areas of interest for quantification of cell proliferation. This should be done cautiously as cell proliferation rates vary between tissue types (e.g., bone marrow versus trabecular bone) and tissue regions (e.g., columnar versus

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hypertrophic chondrocyte zones in growth plate cartilage) (Fig. 2). As appropriate, divide tissues into several areas. 3. Quantify results, according to your study goals. In most cases, the percentages of EdU-positive cells should be calculated (rather than the numbers of EdU-positive cells per tissue volume). For this, divide the numbers of EdU-positive cells by the total numbers of cells present in selected tissue areas. And multiply the result by 100. Calculate averages and standard deviation of replicates, plot data on graphs, and use appropriate statistical tests to compare cell populations or experimental groups.

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Notes 1. EdU, Hoechst 33342 and DAPI are classified as toxins and potential mutagens and teratogens, as are BrdU and other nucleotide analogs and DNA-intercalating agents. Special precautions to be taken in compliance with local regulations include working in a chemical fume hood or laminar flow hood; wearing double gloves, safety goggles, and a lab coat; storing and transporting the solutions in secondary containers (secured and properly labeled); discarding contaminated disposables (syringes, needles, mouse parts, mouse cage bedding, and paper towels) in appropriate sharp or biohazard containers to be incinerated; disposing of fixative and rinse solutions in chemical waste bottles (properly labeled and stored); and cleaning dissecting tools and contaminated areas with large volumes of running tap water. Other reagents that require special precautions include paraformaldehyde and DMSO. Handle and dispose of them according to their hazardous nature and pertaining regulations. 2. Other fluorochromes conjugated to azide for the detection of EdU include Alexa Fluor® 488, 555, 594, and 647. If you wish to perform dual-labeling with BrdU and EdU, keep in mind that some anti-BrdU antibodies cross-react with EdU (Accurate), whereas others do not (Sigma) [11]. 3. Surrounding tissue sections with a hydrophobic barrier allows smaller volumes of solution to be used during incubations than when entire slides are covered with solution, or to perform distinct assays with different sections on one slide. It is recommended that such steps as PBS washes be performed by dipping slides in Coplin jars. Note that the barrier must be added after section deparaffinization because it is completely removed by Histo-Clear.

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4. Total darkness is not necessary, but avoid direct exposure to light. Turn off room lights and stay away from windows. Perform incubations in a lightproof drawer or incubator. 5. If desired, proceed with complementary immunostaining or histochemistry assays on the slides at this time. Keep the samples protected from light during all incubations.

Acknowledgments This work was supported by NIH/NIAMS grants AR68308 and AR72649 to VL. References 1. Kozhemyakina E, Lassar AB, Zelzer E (2015) A pathway to bone: signaling molecules and transcription factors involved in chondrocyte development and maturation. Development 142:817–831 2. Rumman M, Dhawan J, Kassem M (2015) Concise review: quiescence in adult stem cells: biological significance and relevance to tissue regeneration. Stem Cells 33:2903–2912 3. Yip RKH, Chan D, Cheah KSE (2019) Mechanistic insights into skeletal development gained from genetic disorders. Curr Top Dev Biol 133:343–385 4. Marie PJ (2015) Osteoblast dysfunctions in bone diseases: from cellular and molecular mechanisms to therapeutic strategies. Cell Mol Life Sci 72:1347–1361 5. Hughes WL, Bond VP, Brecher G, Cronkite EP, Painter RB, Quastler H, Sherman FG (1958) Cellular proliferation in the mouse as revealed by autoradiography with Tritiated thymidine. Proc Natl Acad Sci U S A 44:476–483 6. Gratzner HG (1982) Monoclonal antibody to 5-bromo- and 5-iododeoxyuridine: a new reagent for detection of DNA replication. Science 218:474–475

7. Mead TJ, Lefebvre V (2014) Proliferation assays (BrdU and EdU) on skeletal tissue sections. Methods Mol Biol 1130:233–243 8. Podgorny O, Peunova N, Park JH, Enikolopov G (2018) Triple S-phase labeling of dividing stem cells. Stem Cell Rep 10:615–626 9. Newton PT, Li L, Zhou B, Schweingruber C, Hovorakova M, Xie M, Sun X, Sandhow L, Artemov AV, Ivashkin E, Suter S, Dyachuk V, El Shahawy M, Gritli-Linde A, Bouderlique T, Petersen J, Mollbrink A, Lundeberg J, Enikolopov G, Qian H, Fried K, Kasper M, Hedlund E, Adameyko I, S€avendahl L, Chagin AS (2019) A radical switch in clonality reveals a stem cell niche in the epiphyseal growth plate. Nature 567:234–238 10. Salic A, Mitchison TJ (2008) A chemical method for fast and sensitive detection of DNA synthesis in vivo. Proc Natl Acad Sci U S A 105:2415–2420 11. Zeng C, Pan F, Jones LA, Lim MM, Griffin EA, Sheline YI, Mintun MA, Holtzman DM, Mach RH (2010) Evaluation of 5-ethynyl-20 -deoxyuridine staining as a sensitive and reliable method for studying cell proliferation in the adult nervous system. Brain Res 1319:21–32

Chapter 22 Whole Mount In Situ Hybridization in Murine Tissues Deepika Sharma, Matthew J. Hilton, and Courtney M. Karner Abstract Whole mount in situ hybridization is a sensitive method used to characterize the spatial and temporal expression of RNA transcripts throughout an entire tissue. This method is an excellent tool for studying gene expression during embryonic development. Here, we describe a procedure for digoxigenin labeled in situ hybridization on whole embryos. Key words Gene expression, RNA, Digoxigenin, In situ hybridization, WISH

1

Introduction In situ hybridization has been extensively used to interrogate gene expression patterns in both tissue sections and in whole organs and embryos [1]. Understanding the temporal and spatial expression of genes is a prerequisite to understand gene function. In situ hybridization is considered to be the gold standard for evaluating gene expression in developing tissues and has been used extensively to distinguish patterns of gene expression during limb patterning and skeletal development [2–10]. The developing limb is composed of distinct zones that pattern the limb and specify the many cell types of the developing skeletal system including the osteoblasts and chondrocytes. In situ hybridization has been critical to describe patterns of gene expression associated with these distinct zones and specified cell types. Indeed, while in situ hybridization is ideal for describing gene expression patterns and detecting major differences in gene expression, it is important to note that in situ hybridization is not quantitative and is not well suited to detect subtle differences in gene expression [11–13]. The underlying principle of in situ hybridization is based on nucleotide complementarity between a labeled antisense RNA probe (also known as a riboprobe) and the endogenous mRNA transcript. There are myriad possibilities to label the riboprobe. For example, the riboprobe can be labeled with 35S, or with

Matthew J. Hilton (ed.), Skeletal Development and Repair: Methods and Protocols, Methods in Molecular Biology, vol. 2230, https://doi.org/10.1007/978-1-0716-1028-2_22, © Springer Science+Business Media, LLC, part of Springer Nature 2021

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nonradioactive reagents such as digoxigenin (DIG) [14]. DIG labeled riboprobes offer increased stability and a longer shelf life as well as a safer method of in situ hybridization when compared to radioactively labeled probes and will be the topic of this chapter [15]. Specifically, we will describe a moderately high throughput protocol for whole mount in situ hybridization using mesh buckets. This protocol is useful to rapidly analyze the spatial localization of mRNA transcripts in many samples ranging from isolated limb buds to whole embryos.

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Materials All solutions and buffers are prepared using Milli-Q (ultrapure) water (see Note 1).

2.1 Common Materials

1. Rocker. 2. 10
PBS (pH 7.4–7.6): Dissolve 80 g NaCl, 2 g KCl, 14.4 g Na2HPO4, and 2.4 g KH2PO4 in 800 mL of ultrapure water. pH the solution with HCl to 7.4 and make the final volume to 1 L with ultrapure water. Autoclave and store at room temperature. 3. 1 M Tris–HCl (pH 7.4–7.6): Dissolve 121.1 g Tris–HCl in 800 mL of ultrapure water and adjust pH with HCl. Make final volume to 1 L with ultrapure water. Autoclave and store at room temperature. 4. 10
MAB (pH 7.4–7.6): Dissolve 116 g of maleic acid (Sigma M 0375), 87.6 g NaCl, 40 g NaOH in 800 mL of ultrapure water. pH the solution with HCl to 7.4 and make the final volume to 1 L with water. Store at room temperature (see Note 2). 5. 5 M NaCl: Weigh 292.2 g NaCl and dissolve in ultrapure water to a total volume of 1 L. Store at room temperature. 6. 1 M MgCl2: Weigh 203.3 g MgCl2 and dissolve in ultrapure water to a final volume of 1 L. Store at room temperature.

2.2 Linearization of Plasmid

1. 10
buffer. 2. Restriction enzyme. 3. 15 μg of Plasmid. 4. Phenol–chloroform. 5. 3 M sodium acetate. 6. 1
TE buffer: 10 mM Tris, bring to pH 8.0 with HCl, 1 mM EDTA.

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1. 1 μg of linear cDNA template of target gene (see Note 3). 2. 10
transcription buffer (Roche). 3. 0.1 M DTT (Boehringer) (see Note 4). 4. 10
NTP Labeling Mixture:10 mM ATP, CTP, GTP, 6.5 mM UTP, 3.5 mM DIG-11-UTP, pH 7.5. 5. 10 U/μL DNase I, RNase-free. 6. 20 U/μL Protector RNase Inhibitor. 7. 20 U/μL Appropriate RNA Polymerase (see Note 3).

2.4

Embryo Fixation

1. 4% paraformaldehyde (PFA) in 1
PBS: To make 500 mL, add 25 mL of 1 M NaOH to 350 mL of PBS, add 20 g of 8% PFA. Stir until PFA dissolves. Adjust the pH to 7 using 1 M HCl (approximately 25 mL). Make 50 mL aliquots and store at 20 ˚C (see Note 5). 2. Dehydration: 25%, 50%, 75%, and 100% ethanol/PBST series.

2.5 Prehybridization and Hybridization

1. Hybridization mesh buckets (Netwell 12-well Carrier Kit, Corning 3477). 2. 1
PBST: Mix 100 mL of 10
PBS with 900 mL of ultrapure water. Add 0.1% Tween-20 (1 mL) to the solution. Autoclave, filter and store at room temperature. 3. Reagents for rehydration: 75%, 50%, and 25% ethanol/PBST series. 4. 10 μg/mL Proteinase K solution: Add 6 μL of 10 mg/mL Proteinase K in 6 mL of 1
PBST. 5. 2 mg/mL glycine: Dissolve 12 mg of glycine in 6 mL of 1
PBST. 6. Postfix: Mix 12.5 mL of 4% PFA, 100 μL of 25% glutaraldehyde (0.2%), and 12.5 μL of Tween 20. 7. Hybridization oven at 70 ˚C. 8. 20
SSC (pH 7.4): Dissolve 175.3 g of NaCl and 88.2 g of Sodium Citrate in 800 mL of ultrapure water. Adjust pH with citric acid. Adjust volume to 1 L with ultrapure water. Autoclave and store at room temperature. 9. 10% SDS: Dissolve 10 g of SDS in 80 mL of ultrapure water and bring the volume up to 100 mL. Autoclave and store at room temperature. 10. Hybridization stock solution: 50% formamide (deionized), 5
SSC, pH 4.5 (use citric acid to pH), 1% SDS, 50 μg/mL yeast tRNA, and 50 μg/mL heparin. For 50 mL final volume, add 25 mL of deionized formamide, 12.5 mL of 20
SSC (pH 4.5), 5 mL of 10% SDS, 50 μL of yeast tRNA, 2.5 mg of heparin, and 7.5 mL of ultrapure water. Aliquot into 15 mL tubes and store at 80  C.

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2.6 Post Hybridization, Blocking and Antibody Incubation

1. Hybridization oven at 70 ˚C and 65 ˚C. 2. Solution I (50% Formamide, 5
SSC pH 4.5, 1% SDS): To make 50 mL, add 25 mL of deionized formamide, 12.5 mL of 20
SSC (pH 4.5), and 5 mL of 10% SDS to 7.5 mL of ultrapure water. 3. Solution II (0.5 M NaCl, 10 mM Tris–HCl pH 7.5, and 0.1% Tween 20): To make 50 mL, add 5 mL of 5 M NaCl and 500 μL of 1 M tris (pH 7.5) to 44.5 mL of ultrapure water. 4. Solution III (50% formamide, 2
SSC pH 4.5, 0.1% Tween 20): To make 50 mL, add 25 mL of deionized formamide and 5 mL of 20
SSC (pH 4.5) to 20 mL of ultrapure water. 5. Invitrogen PureLink RNase A (20 mg/mL): To make 100 μg/ mL, add 50 μL of RNase A to 10 mL of Solution II. 6. 1
MBST (100 mM maleic acid, 150 mM NaCl, 1% Tween 20): To make 500 mL, add 50 mL of 10
MAB to 450 mL of water and 500 μL of Tween 20. 7. Blocking solution: 2% Boehringer Mannheim Blocking Reagent (BMB, Sigma) in MBST. Heat briefly to dissolve BMB, cool to RT, before adding heat inactivated sheep serum, to make 10 mL, add 1 mL of sheep serum to 9 mL of 1
MBST and dissolve 20 mg of BMB blocking reagent by warming the solution to 70 ˚C and repeated vortexing. 8. Antibody solution: Dilute anti-digoxygenin AP antibody (Sigma) 1:4000 in the blocking solution.

2.7 Post Antibody Washes and Color Development

1. NTT solution: 0.15 M NaCl, 0.1 M Tris pH 7.5 and 0.1% Tween 20. To make 100 mL, add 3 mL of 5 M NaCl, 10 mL of Tris pH 7.5 and 100 μL of Tween-20 to 87 mL of ultrapure water. 2. NTTML stock solution: 100 mM NaCl, 100 mM Tris–HCl, pH 9.5, 50 mM MgCl2, 0.1% Tween 20, and 2 mM levamisole. To make 50 mL, add 1 mL of 5 M NaCl, 5 mL Tris–HCl pH 9.5, 2.5 mL of MgCl2, 25 mg of levamisole, and 50 μL of Tween 20 to 41 mL of ultrapure water (Make fresh). 3. Heavy-duty aluminum foil. 4. Developer: BM purple AP substrate (Sigma). 5. Stereo microscope and digital camera.

3

Methods

3.1 Linearization of Plasmid

1. To linearize the plasmid, mix the following reagents at room temperature: 22 μL 10
Colorless buffer, 5 μL appropriate restriction enzyme, and 15 μg plasmid (determine volume based on concentration) and fill to 200 μL with Milli-Q-water.

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2. Incubate at 37  C overnight (see Note 6). 3. Add 1 volume of phenol–chloroform to the digestion reaction and mix well. 4. Centrifuge at maximum speed for 15 min and transfer the aqueous phase into a new tube. 5. Add 1/10th the volume of 3 M sodium acetate and three times the total volume of 100% ethanol. 6. Incubate at 20  C 1 h to overnight. 7. Centrifuge at maximum speed for 10 min at 4  C, visualize the pellet, and discard the supernatant. 8. Wash the pellet with 70% ethanol twice and dry the pellet for 10 min. Resuspend the pellet in 20 μL of TE buffer and measure the concentration. 9. Linearized plasmid can be stored at 20  C. 3.2 Generation of Probes (Can Be Prepared During or Prior to Day 1)

1. To make DIG-labeled probes, mix 6.5 μL RNase-free water, 2 μL warmed 10
transcription buffer, 2 μL 0.1 M DTT, 2 μL DIG-labeled Mix, 1 μg linear DNA plasmid, 1 μL RNase inhibitor, 1 μg linear DNA plasmid, 5 μL RNA polymerase (T7, T3, or SP6). 2. Gently mix probe solution. Incubate at 37  C for 2 h either using a water bath or hybridization oven. 3. Add 2 μL of DNase I (ribonuclease-free) and mix gently. Incubate at 37  C for 20 min. 4. Add 100 μL of TE, 10 μL 4 M LiCl, and 300 μL ethanol, mix and precipitate at 20  C for 30 min to overnight (Can be stored at 20  C indefinitely). 5. Centrifuge at maximum speed at 4  C for 30 min. Wash the pellet twice with 70% ethanol and air-dry. 6. Dissolve the pellet in TE and measure the concentration. Add 1 mL of hybridization buffer or add formamide to 50%. The probe can be stored at 80  C.

3.3 Embryo Preparation

This protocol is optimized for whole embryos from embryonic stage E7.5–E12.5 as well as isolated E10.5–E12.5 mouse limb buds. 1. Embryos are harvested in 1
PBS. 2. Upon completion of dissection, embryos are placed in an Eppendorf tube containing 1 mL of 4% PFA. The embryos are fixed overnight (12 h) at 4  C with gentle rocking. 3. After fixation, embryos are washed once in PBST for 5 min with gentle rocking. At this point, the embryos can be used for in situ hybridization immediately. If so, go to Subheading 3.3,

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step 4. Alternatively, embryos can be dehydrated and stored for further use. If so, go to next step. 4. Embryos are then dehydrated into ethanol using a graded ethanol/PBST series (25%, 50%, and 75%). Allow embryos to rock gently for 5 min in each solution. 5. At this point, embryos can be stored indefinitely in 75% ethanol/PBST at 20  C. 3.4 Prehybridization and Hybridization (Day 1)

All washes during this step are performed in hybridization mesh buckets placed in a 12-well plate (see Fig. 1). Multiple whole embryos or limb buds can be sorted into individual buckets with each bucket corresponding to a specific in situ probe. 1. Rehydrate samples in a 75%, 50%, and 25% ethanol/PBST series for 5 min. Then wash twice in PBST for 5 min each. All washes are performed at room temperature. 2. Wash embryos with PBST three times for 5 min each. 3. Incubate at room temperature with 10 μg/mL proteinase K in PBST for 15–20 min depending on embryo stage (see Note 7). 4. Wash embryos in 2 mg/mL glycine in PBST for 10 min at room temperature (make this solution fresh). 5. Wash twice in PBST for 5 min each. 6. Refix embryos with 4% PFA and 0.2% glutaraldehyde in PBST for 20 min at room temperature. 7. Wash embryos with PBST three times for 5 min each. 8. Wash embryos in a 1:1 mixture of hybridization buffer/PBST solution for 10 min at room temperature with gentle rocking. 9. Add fresh hybridization buffer (at least 400 μL) and incubate for 1 h at 70  C with gentle rocking. Seal plates with parafilm. 10. Add the appropriate DIG-labeled antisense probe to individual wells (see Note 3). Incubate overnight at 70  C. Seal the plate with parafilm to ensure the container is airtight so the probe does not evaporate overnight.

3.5 Post Hybridization Washes, Blocking, and Antibody Incubation (Day 2)

1. Before washing, it is important to prewarm the indicated solutions to the proper temperature. Solution I—70  C. Solution I: Solution II mixed 1:1—70  C. Solution III—65  C. 2. Wash embryos with Solution I twice for 30 min each at 70  C. 3. Wash embryos in Solution I/II mix for 10 min at 70  C. 4. Wash embryos with Solution II twice for 5 min each at room temperature. 5. Wash embryos with 100 μg/mL RNase A in Solution II for 1 h at 37  C.

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Fig. 1 Overview of solution changes using 12-well hybridization buckets. Each solution is placed into 12-well plates. Carrier with hybridization buckets are placed in current solution for appropriate time. To change solutions, move carrier to the next plate. Hybridization bucket can accommodate multiple embryos or individual limb buds to increase the number of animals analyzed per probe

6. Wash embryos with Solution II then Solution III for 5 min each at room temperature. 7. Wash embryos with Solution III twice for 30 min at 65  C (Make up MBST during these washes). 8. Wash embryos with MBST three times for 5 min each at room temperature. 9. Incubate embryos in blocking solution for 1 h at room temperature. 10. Remove blocking solution and add blocking/antibody solution. Incubate at 4  C overnight with gently rocking (12h). 3.6 Post Antibody Washes and Color Development (Day 3)

1. Wash embryos with MBST three times for 5 min each. 2. Wash embryos with MBST for 1 h at room temperature. Repeat this step eight times or leave overnight in MBST at 4  C (see Note 8). 3. Wash with NTTML (make fresh) three times for 5 min each. 4. Cover samples with aluminum foil to protect from light. 5. Add BM Purple to each embryo and incubate at room temperature with gentle rocking (see Note 9). 6. When the color reaction is complete, wash embryos with PBST three times for 5 min each (see Note 10). 7. Post-fix embryos in 4% PFA and 0.1% glutaraldehyde overnight at 4  C. Keep the embryos covered in aluminum foil. 8. Wash embryos with PBST three times for 5 min each. 9. To clear the embryos, wash through glycerol/PBST gradient (30%, 50%, and 80%) changing every day. Embryos can be stored in 80% glycerol at 4  C. 10. Following the clearing step, embryos can be visualized and imaged using a stereoscope and digital camera (Fig. 2).

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Fig. 2 Whole mount In situ hybridization results. Expression of Sry-box factor 9 (Sox9), Collagen type II (Col2a1) or Wnt family member 5a (Wnt5a). Sox9 demarcates mesenchymal condensations while Col2a1 is expressed in more differentiated cartilage. Wnt5a is expressed in the distal mesenchyme underneath the apical ectodermal ridge

4

Notes 1. The reagents, buffers, and instruments that are used before the hybridization step should be RNase free, but DEPC-treated water is not necessary. We find that Milli-Q (ultrapure) water is sufficient. All of the common solutions used in the prehybridization and hybridization steps should be autoclaved and filtered using 0.2 μM filters. 2. Maleic acid is hard to dissolve until NaOH has been added. Start with adding NaOH and then maleic acid once it is dissolved, add the NaCl. 10
MAB can be stored at room temperature indefinitely. 3. To generate antisense riboprobes, a plasmid containing the cDNA of interest needs to be linearized. Use a unique restriction enzyme near the 50 end of the cDNA, and the appropriate RNA polymerase specific to the promoter (T3, T7 or Sp6, depending on the plasmid) at the 30 end. 4. Dilute freshly from 1 M DTT. 5. PFA is toxic, therefore all waste should be discarded in the hazardous waste.

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6. Save 1 μL to run on the get to check if plasmid is fully digested before cleanup and another 1 μL after transcription to visualize the RNA probe. 7. For this step, be very diligent in monitoring the amount of time in proteinase K. Too much time in proteinase K can result in loss of signal or tissue. More internal tissue like the notochord require longer time as compared to the AER of the limb which is very superficial. The following are the digestion times used for E7–E12.5 embryos: E7.0 ¼ 2 min. E7.5 ¼ 5 min. E8.5–E9.5 ¼ 10 min. E10.5 ¼ 15 min. E11.5 ¼ 16–17.5 min. E12.5 ¼ 18–20 min. 8. Decreased background is obtained by a second day of washing. Embryos can be left up to 3 days washing at 4  C without apparent loss of signal. It is advantageous to leave the embryos overnight in MBST at 4  C. This decreases background and allows you to start the staining reaction first thing in the morning and monitor all day rather than monitoring all night to prevent overstaining. 9. At the end of the first wash, place BM Purple at room temperature. This solution should be close to room temperature before starting to develop the embryos. 10. The color reaction will vary from probe to probe. As a general rule, the higher the gene expression in the respective tissue the faster the color reaction will be complete. It is recommended that the color reaction be monitored every 30 min using a dissecting microscope and a light source. In our experience the color reaction typically takes from 2 h to overnight to complete depending on the probe.

Acknowledgments This work was supported by NIH/NIAMS grants (A032092 and A032121) to M.J.H and (AR071967 and AR076325) to C.M.K. References 1. Gall JG, Pardue ML (1969) Formation and detection of RNA-DNA hybrid molecules in cytological preparations. Proc Natl Acad Sci U S A 63(2):378–383

2. Riddle RD, Johnson RL, Laufer E, Tabin C (1993) Sonic hedgehog mediates the polarizing activity of the ZPA. Cell 75(7):1401–1416 3. Zakany J, Duboule D (1993) Correlation of expression of Wnt-1 in developing limbs with

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abnormalities in growth and skeletal patterning. Nature 362(6420):546–549 4. Chen H, Johnson RL (1999) Dorsoventral patterning of the vertebrate limb: a process governed by multiple events. Cell Tissue Res 296(1):67–73 5. Cohn MJ, Izpisua-Belmonte JC, Abud H, Heath JK, Tickle C (1995) Fibroblast growth factors induce additional limb development from the flank of chick embryos. Cell 80 (5):739–746 6. Lettice LA, Hill RE (2005) Preaxial polydactyly: a model for defective long-range regulation in congenital abnormalities. Curr Opin Genet Dev 15(3):294–300 7. McGlinn E, van Bueren KL, Fiorenza S, Mo R, Poh AM, Forrest A et al (2005) Pax9 and Jagged1 act downstream of Gli3 in vertebrate limb development. Mech Dev 122 (11):1218–1233 8. Sheth R, Gregoire D, Dumouchel A, Scotti M, Pham JM, Nemec S et al (2013) Decoupling the function of Hox and Shh in developing limb reveals multiple inputs of Hox genes on limb growth. Development 140 (10):2130–2138

9. Tabin C, Wolpert L (2007) Rethinking the proximodistal axis of the vertebrate limb in the molecular era. Genes Dev 21 (12):1433–1442 10. Zuniga A, Haramis AP, McMahon AP, Zeller R (1999) Signal relay by BMP antagonism controls the SHH/FGF4 feedback loop in vertebrate limb buds. Nature 401(6753):598–602 11. Acloque H, Wilkinson DG, Nieto MA (2008) In situ hybridization analysis of chick embryos in whole-mount and tissue sections. Methods Cell Biol 87:169–185 12. Wilkinson DG, Nieto MA (1993) Detection of messenger RNA by in situ hybridization to tissue sections and whole mounts. Methods Enzymol 225:361–373 13. Neufeld SJ, Zhou X, Vize PD, Cobb J (2013) mRNA fluorescence in situ hybridization to determine overlapping gene expression in whole-mount mouse embryos. Dev Dyn 242 (9):1094–1100 14. Wilkinson DG (1995) RNA detection using non-radioactive in situ hybridization. Curr Opin Biotechnol 6(1):20–23 15. Komminoth P (1992) Digoxigenin as an alternative probe labeling for in situ hybridization. Diagn Mol Pathol 1(2):142–150

Part IV Primary Cell Isolations, Cultures, and Assays

Chapter 23 Bone Marrow Stromal Cell Assays: In Vitro and In Vivo Pamela G. Robey, Sergei A. Kuznetsov, Paolo Bianco*, and Mara Riminucci Abstract Populations of bone marrow stromal cells (BMSCs, also known as bone marrow–derived “mesenchymal stem cells”) contain a subset of cells that are able to recapitulate the formation of a bone/marrow organ (skeletal stem cells, SSCs). It is now apparent that cells with similar but not identical properties can be isolated from other skeletal compartments (growth plate, periosteum). The biological properties of BMSCs, and these related stem/progenitor cells, are assessed by a variety of assays, both in vitro and in vivo. Application of these assays in an appropriate fashion provide a great deal of information on the role of BMSCs, and the subset of SSCs, in health and in disease. Key words Bone marrow, Colony forming unit-fibroblast, Bone, Cartilage, Stroma, Marrow adipocytes, In vitro assays, In vivo transplantation

1

Introduction It has long been recognized, based on the work by Friedenstein and Owen, that bone marrow contains an adherent, nonhematopoietic cell that is a component of the bone marrow stroma (reviewed in [1–5]). In a series of experiments, starting with nonclonal populations of these bone marrow stromal cells (BMSCs, also known as bone marrow-derived “mesenchymal stem cells”), and subsequently with clonal populations that arise from individual colonyforming unit fibroblasts (CFU-Fs), it was demonstrated that a subset of BMSCs is multipotent. When clonal strains were transplanted in vivo, some of the clonal strains formed bone and cartilage in closed systems (diffusion chambers). When transplanted in an open system (with access to the circulation), some of the clonal strains formed bone, stroma that supports hematopoiesis, and marrow adipocytes, all of donor origin, with blood of recipient origin [6]. These experiments firmly established the multipotent nature of

*Deceased. Matthew J. Hilton (ed.), Skeletal Development and Repair: Methods and Protocols, Methods in Molecular Biology, vol. 2230, https://doi.org/10.1007/978-1-0716-1028-2_23, © Springer Science+Business Media, LLC, part of Springer Nature 2021

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a subset of BMSCs, suggesting the existence of a stem cell able to differentiate into skeletal cell phenotypes (a skeletal stem cell, SSC, [3, 7] However, it must be noted that not all CFU-Fs that establish clonal lines are multipotent based on these in vivo transplantation assays. It has also been determined that these multipotent cells arise from specialized clonogenic BMSCs that are found on the abluminal side of bone marrow sinusoids. These cells are known as mural cells or pericytes [8]. But again, not all pericytes are multipotent. Very importantly, their ability to self-renew was established by subsequent serial transplantation of phenotype-defined clonogenic cells in vivo [8, 9]. Based on these findings, it is clear that bone marrow stroma contains a stem cell by the most rigorous criteria: the ability of the progeny of a single cell to reform and support a complete organ (the bone/marrow organ), and the ability to self renew. In addition to perivascular bone marrow skeletal stem cells [8, 10–12], recent studies have also identified skeletal stem cells in other locations such as the growth plate and the periosteum [13– 17]. While skeletal stem cells derived from these different locations share many characteristics, they are not completely identical in their functionality, based on the assays that are described below. For example, cells derived from the growth plate and the periosteum do not appear to form adipocytes as shown by in vivo transplantation assays. Consequently, there is a need to precisely define the region from which a population of cells has been derived, and to encorporate the tissue of origin into the name by which the cells are called when describing the results of various experiments. The experimental proof of the existence of SSCs is based on a number of assays that require in vivo transplantation (the gold standard by which to evaluate differentiation capacity) of ex vivo expanded, clonally derived cells, or of freshly isolated cells with a particular cell surface phenotype. Nonetheless, several in vitro differentiation assays are widely used for determination of osteogenic and adipogenic differentiation but are prone to artifacts as described below. While cartilage formation was first demonstrated by in vivo transplantation of BMSCs in diffussion chambers, the current gold standard assay relies on the formation of high density cell pellets in vitro [18, 19]. Lastly, expression of markers representative of a particular cell phenotype has also been employed as a means of determining differentiation. However, expression of several markers does not faithfully predict the differentiation capacity of cells, but assessment of the pattern of expression of markers is a useful tool when studying different stages of differentiation in conjuction with in vivo assays. Bone marrow derived skeletal stem/progenitor cells (BMSCs/ SSCs) are the main focus of this chapter, but the assays described below are also pertinent to skeletal stem/progenitor cells derived from other skeletal tissue sources (growth plate and periosterum).

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BMSCs can most likely be isolated from any species in which bone marrow exists, although culture conditions often vary from one species to another. As an example, murine and human BMSCs do vary substantially, and the methods below highlight differences between establishing and characterizing cultures from these two different species. Furthermore, murine and human BMSCs are the most frequently used, based on the wealth of transgenic and knockout animal models that exhibit skeletal disorders, and from humans, both normal and with diseases. What follows below is a description of in vitro and in vivo assays for the assessment of BMSCs, and the subset of SSCs within the population, that can be applied to normal and pathological marrow from mice and from humans.

2 2.1

Materials Solutions

Unless specifically indicated, most reagents and supplies can be obtained from many different vendors. 1. Marrow collection medium (MCM): α-MEM (with 100 U/ml sodium heparin for human bone marrow aspirates). 2. Serum-containing medium (SM): αMEM, 2 mM glutamine or glutamax, 100 U/ml penicillin, 100 μg/ml streptomycin sulfate, and 20% lot-selected fetal bovine serum, NON–heat inactivated (see Note 1). 3. Hanks balanced salt solution (HBSS). 4. 100% methanol. 5. Enzymatic digestions: Trypsin–EDTA (0.05% Trypsin with 0.53 mM EDTA in HBSS), or collagenase (1 mg/ml Collagenase IV in α-MEM). 6. Osteogenic medium (OM): αMEM, 2 mM glutamine, 100 U/ ml penicillin, 100 μg/ml streptomycin sulfate and 1–20% lot-selected fetal bovine serum, NON–heat inactivated (see Note 1), supplemented with 10
8 M dexamethasone, 10
4 M L-ascorbic acid-2-phosphate, and 2–5 mM β-glycerophosphate. 7. Adipogenic medium (AM): αMEM supplemented with (1) 0.5 μM isobutylmethylxanthine, 0.5 μM hydrocortisone, and 60 μM indomethacin or (2) αMEM with 10
4 M L-ascorbic acid 2-phosphate and 10
8 to 10
7 M dexamethasone or (3) αMEM containing glutamine and penicillin/streptomycin, with 20% lot-selected rabbit serum, 10
4 L-ascorbic acid 2-phosphate and 10
8 M dexamethasone or (4) αMEM with the 0.1–10 μM of the PPARγ ligand, rosiglitazone.

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8. Chondrogenic medium (CM): DMEM with high glucose (4.5 g/L) supplemented with 10
6 M bovine insulin, 8  10
8 M human apotransferrin, 8  10
8 M bovine serum albumin, 4  10
6 M linoleic acid, 10
3 M sodium pyruvate, 10 ng/ ml rhTGFβ1, 10
7 M dexamethasone, and 2.5  10
4 M Lascorbic acid-2 phosphate. 9. Anesthesia: Combine 225 μl ketamine, 69 μl of xylazine, 75 μl of acepromazine (Sigma), and 231 μl of H2O (total volume ¼ 600 μl), use 100 μl/mouse (25 g), or 2–5% isoflurane). 10. Betadine. 11. 70% ethanol. 12. Standard histological stains: saturated methyl violet, hematoxylin and eosin (H&E), Alizarin Red S, von Kossa, Oil Red O, and Toluidine Blue. 13. Antibodies: for cell surface analysis and immunohistochemistry (numerous vendors). 2.2 Equipment and Supplies

1. Hemocytometer: For use in cell enumeration. 2. Sterile labware: Tissue culture dishes and flasks of various sizes, pippetes, centrifuge tubes (not vendor specific), cell strainers (70 μ pore size, Becton Dickinson), Scienceware® cloning cylinders (Sigma), sterilized vacuum grease. 3. Surgical equipment: Sterile scalpels, small scissors, forceps, small spatula, and autoclips (not vendor specific). 4. Scaffolds for in vivo transplantation: Ceramic particles (hydroxyapatite/tricalcium phosphate, or variations thereof, from a variety of sources) for human and mouse transplants; collagen sponges (from a variety of sources) for mouse transplants (see Note 2). 5. CO2 incubators: Set to 37 and 5% CO2 (not vendor specific). In some instances, incubators are made hypoxic (2–5% O2) (see Note 3). 6. Microscopes: Standard inverted phase contrast, dissecting microscopes, bright- and dark-field microscopes (not vendor specific). 7. Standard flow cytometry and fluorescence-activated flow cyteometry equipment and supplies (not vendor specific). 8. Standard PCR equipment, supplies, and primers (not vendor specific).

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1. Murine: Typically femora, tibiae, and humeri, collected from any strain of mice. 2. Human: Bone fragments collected as surgical waste (IRB exempt if de-identified) and bone marrow aspirates from normal volunteers and patients with skeletal diseases under Internal Review Board approved protocols for the use of human subjects in research. 3. Guinea pigs: Guinea pigs (Hartley Davis, Charles River Laboratories) are used to create irradiated bone marrow feeder cells for certain types of murine cultures.

2.4 Recipients for In Vivo Transplantation Assays

1. Autologous transplantation: For larger animals (sheep, nonhuman primates, etc.), bone marrow is aspirated, and after ex vivo expansion, BMSCs are transplanted back into the original donor with an appropriate scaffold. 2. Syngeneic transplantation: Any inbred strain of mice, rats, rabbits, guinea pig, etc. 3. Xenogeneic transplantation: Congenic immunocompromised mice of various strains such as NSG mice (NOD.Cg-Prkdcscid Il2rgtm1Wjl/SzJ, Jackson Laboratory), or SHC mice (CB17. Cg-PrkdcscidHrhr/IcrCrl, Charles River Laboratory (see Note 4).

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Methods There have been a number of modifications of the original procedure developed by Friedenstein, which relies on the rapid adherence of BMSCs to tissue culture plastic [6, 20, 21]. These include subfractionation of bone marrow single cell suspensions by density gradient centrifugation. However, this often results in a marked loss of BMSCs/SSCs (~50%, R. Merling and S. A. Kuznetsov, unpublished results) as determined by colony forming efficiency assays (see below). More recently, prospective isolation of BMSCs/ SSCs using sets of cell surface markers and FACS have been employed (reviewed in [4, 7, 8, 22, 23]). However, there is no standard in vivo assay in which UNCULTURED SORTED cells can be transplanted directly, although it has been attempted using a single mouse skeletal stem cell along with differentiated cells as a “carrier” [13]. Hence, sorted fractions of stromal cells are usually cultured prior to in vivo assays. For this reason, establishing primary cultures at clonal density (CFU-F cultures) is a practical surrogate for purification of the whole population of clonogenic stromal progenitors [8]. Lastly, it has been suggested that low oxygen tension (2–5%) modulates proliferation and differentiation of BMSCs/SSCs (see Note 3). So far, results have been variable,

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with reports of enhancing proliferation and osteogenic differentiation [24] or inhibiting proliferation while maintaining stemness [25]. Few studies have tested the ability of BMSCs/SSCs grown in low oxygen tension to form an ectopic ossicle in vivo, but it appears that cells grown in low oxygen form marrow that is laden with adipocytes, and relatively low levels of hematopoiesis ([26], Balakumaran, Kuznetsov and Robey, unpublished results). Consequently, more work is needed to determine the impact of low oxygen tension on the functionality of BMSCs/SSCs with respect to bone formation and support of hematopoiesis. On the other hand, it is generally thought that low oxygen encourages chondrogenic differentiation, based on the avascular (and thus anaerobic) nature of cartilage [27]. 3.1 Collection and Preparation of Single Cell Suspensions of Bone Marrow

1. Euthanize mice by CO2 inhalation or terminal anesthesia in compliance with institutionally approved protocols for the use of animals in research; collect femora, tibiae, and humeri aseptically; clean muscle from bone; cut the epiphyses and flush the entire bone marrow content of medullary cavities with MCM and combine flushes from all bones. For human surgical specimens; scrape trabecular bone fragments with a steel blade into MCM. For human bone marrow aspirates, collect 0.5 ml with a Jamshidi needle; mix with 5 ml of ice-cold MCM containing 100 U/ml sodium heparin. For both types of human preparations, centrifuge at 135  g for 10 min; resuspend in fresh MCM. For human specimens where yellow marrow is abundant, such as the femoral heads or patellas from elderly patients, a second centrifugation may be necessary in order to get rid of remaining fat droplets. 2. For both mouse and human specimens, pipet up and down several times; pass through needles of decreasing diameter (gauges 16 and 20) to break up aggregates; filter through a cell strainer (see Note 5); count nucleated cells with a hemocytometer. 3. Prepare guinea pig bone marrow suspensions in a similar fashion as above for mouse marrow; irradiate them with 6000 cGy to prevent proliferation of adherent guinea pig BMSCs. These live cells are used as feeder cells for murine BMSC/SSC colony forming efficiency assays.

3.2 Colony Forming Efficiency (CFE)— Enumeration of CFU-Fs

The concentration of CFU-Fs in bone marrow, as determined by the CFE assay, is a rough estimation of the number of SSCs in the BMSC population [28], expressed as the colony forming efficiency (number of BMSC colonies per 1  105 marrow nucleated cells in the original marrow cell suspension (see Note 6, Fig. 1).

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Fig. 1 Establishment of clonal and nonclonal cultures of BMSCs. Clonal cultures are essential in order to determine the multipotency of the SSC subset of cells within the population. Single cells of bone marrow are plated at clonal density, and a single CFU-F adheres, and proliferates to form a colony. When bone marrow cells are plated at high density, nonclonal BMSC cultures are generated that can be used for general biochemical analysis. When both types of cultures near confluency, they are assessed for cartilage formation by pellet cultures, or for the ability to support the formation of a bone/marrow organ upon in vivo transplantation with appropriate scaffolds

1. Plate murine cells (6–15  105 nucleated cells), or human cells (1–6  105 nucleated cells) into 25 cm2 plastic culture flasks in 5 ml of SM in either triplicate or quadruplicate. These cell densities have been chosen based on previously established colony forming efficiency (CFE) values, such that discrete BMSC colonies are formed in numbers sufficient for statistical analysis. 2. Remove unattached cells after 2–3 h and wash vigorously three times with SM. 3. Add 5 ml of SM; for murine cultures, add irradiated guinea pig feeder cells (1.0–1.5  107 nucleated cells per flask) prepared as described above. Standard serum-containing medium does not contain all of the factors needed for 100% CFE of murine BMSCs/SSCs. On the other hand, human BMSCs/SSCs do not require a feeder cell layer for optimal CFE [21]. 4. Incubate at 37 in a humidified atmosphere of 5% CO2 with air; on day 10–14, wash with HBSS; fix with methanol; stain with an aqueous solution of saturated methyl violet. 5. Count colonies containing 50 or more cells using a dissecting microscope and determine colony forming efficiency (number of colonies per 1  105 nucleated cells plated). If cultures are harvested earlier than 10 days, colonies smaller than 50 cells can be counted; however, for the 10–14 day harvesting time, 50 cells is a reasonable cut off that discriminates colonies that are actually growing from smaller “clusters” of cells that have ceased to proliferate (Fig. 2).

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Fig. 2 Colony forming efficiency assay (CFE). Bone marrow is collected from the long bones of mice, and from bone fragments with marrow or aspirates from humans. Single cell suspensions are plated at clonal density. For murine cultures, irradiated guinea pig bone marrow cells are added as feeders to optimize colony forming efficiency. After 10–14 days, colonies with greater than 50 cells are counted and the CFE is determined as the number of colonies/100,000 bone marrow nucleated cells. To date, the CFE is the closest approximation to the number of SSCs in the BMSC population 3.3 Establishment of Single Colony-Derived Strains of BMSCs

A number of studies have focused on the characterization of single colony-derived strains, prepared as described below. It is by clonal analysis and appropriate differentiation assays that the multipotent nature of the subset of BMSCs that are SSCs is established. 1. Plate murine cells (6–15  105 nucleated cells); plate cells from human surgical specimens, (0.007–3.5  103 nucleated cells/ cm2) or from aspirates (0.14–14.0  103 nucleated cells/cm2) into 150 mm diameter Petri dishes for preparation of single colony-derived strains; add 30–50 ml of SM. The low cell densities employed in this assay have been chosen to allow discrete BMSC colonies to be formed at a distance from each other such that the colonies can grow significantly without approaching each other before being isolated. Alternatively, cells can be plated by limiting dilution into 96 well microtiter plates (1 cell/well). 2. Wash vigorously with HBSS after 2–3 h, add irradiated guinea pig cells to mouse cultures as described above.

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3. After 14–16 days, visually inspect and identify well-separated colonies of perfectly round shape for cloning; wash with HBSS: surround colony to be isolated with a cloning cylinder attached to the dish with sterilized high vacuum grease. 4. Treat cells inside the cylinder with two consecutive aliquots of Trypsin/EDTA for 5–10 min each at room temperature; add cold FBS into each fraction as it is collected (final concentration 3%) to inhibit Trypsin; combine fractions and transfer to individual wells of 6-well plates containing SM. 5. Passage before cells reach confluence (~80% confluency), usually 5–10 days later, transfer consecutively to a 25 cm2 flask (second passage) and to a 75 cm2 flask (third passage). 3.4 Preparation of Multicolony-derived Strains of BMSCs

For many studies, multicolony-derived strains are sufficient, and necessary for biochemical analysis of BMSCs undergoing differentiation into various phenotypes, and changes as the result of genetic manipulation, either naturally occuring, or induced. However, multicolony-derived strains cannot be soley used to determine the nature of SSCs (in particular, their multipotency). 1. For murine cultures (see Note 7), plate approximately 6–8  107 nucleated cells per 75 cm2 flask; human surgical specimens, plate at 5  106 to 5  107 nucleated cells; from aspirates, plate at 5  106 to 20  107 nucleated cells into 75 cm2 flasks or 150 mm diameter dishes containing 30–50 ml of SM. The cell densities used for generation of BMSC multicolony-derived strains are based on our data of many years and are chosen to ensure vigorous BMSC growth starting with hundreds of colonies in each flask. When chosing these densities, multiple factors were considered, including, for mouse cultures, the stimulating effect of hematopoietic cells on BMSC proliferation, and, for human aspirates, a highly variable degree of contamination with peripheral blood in bone marrow aspirates. 2. Culture at 37 in a humidified atmosphere of 5% CO2 with air; replace medium on day 1 for human aspirates, and at day 7 for all other cultures; passage generally is performed on day 12–14 (~80% confluent). 3. Passage cultures by washing twice with HBSS; use two treatments of Trypsin/EDTA for 25–30 min (for murine) or 10–15 min (for human) at room temperature; wash with SM. If murine cultures develop significant amounts of extracellular matrix, treatment with collagenase (1 mg/ml collagenase IV in α-MEM) may be needed prior to trypsin/EDTA. 4. Add cold FBS into each fraction as it is collected (final concentration 3%) to inhibit enzymatic activity; combine fractions; pipet to break up cell aggregates; centrifuge at 135  g for

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10 min; resuspend cell pellet in fresh SM; plate murine cells at 2–10  106 cells per 75 cm2; plate human cells at 2  106 cells per 75 cm2 flask or 150 mm diameter dish; passage again when approximately 70% confluency. These cell densities ensure a fast growth of BMSCs so that in 3–5 days, maximum BMSC numbers can be collected. For mouse cultures, also take into consideration a highly variable concentration of macrophages among BMSCs. 3.5 Flow Cytometry and FluorescenceActivated Cell Sorting (FACS)

There are a number of cell surface markers that have been utilized to prospectively isolate BMSCs from other cell types. There is a long list of mouse lines (too extensive to list here) with reporters driven by promoters of interest for prospective isolation of cells by FACS, or genetically modified to eliminate a particular cell surface marker or transcription factor for flow cytometry and phenotype analysis (reviewed in [7, 22, 23, 29]). BMSCs/SSCs are negative for hematopoietic and endothelial markers, and positive for a number of markers that are commonly expressed by many connective tissue cell types. In vivo, human BMSCs are identified by expression of ALP, CD146, CD105, CD90, (reviewed in [7]), and CD271 [30, 31]. Combinations of these markers, along with STRO-1 [32], can be used to enrich clonogenic stromal cells to near purity. There are important differences in the phenotype of murine and human cells. For example, Vcam1 isolates all CFU-Fs from mouse bone marrow while CD146 isolates none [33]. Prospective isolation experiments by FACS are crucial to define the correlation between ex vivo observed properties and in vivo identity of stromal cells. What follows below is an example of detection of a single cell surface marker by flow cyteometry; however, current technology allows for the identification of up to 18 markers from the same sample depending on the instrument. The technical details of how to perform such in depth analyses are best gleaned from the manufacturer’s instructions and will not be covered here. For FACS (to obtain live cells of a defined phenotype for either ex vivo expansion or for direct transplantation), a number of schemes have been developed for murine and human SSCs (see [8, 13–17]), and the reader is referred to several reviews for the possible strategies [22, 23, 29]. However, it must be noted that due to the variability of the strategies used, and the lack of comprehensive determination of recovery of CFU-Fs after preparatory manipulations, it is not currently clear if differences between SSC populations from different skeletal compartments is due to the inherent properties of the recovered cells, or due to the loss of cell subsets during each step of manipulation. Further studies are needed to answer this question. 1. Harvest and wash the cells with PBS; adjust the cell suspension to a concentration of 1–5  106 cells/ml in ice-cold PBS, 10% FCS or BSA, 1% sodium azide (omit for viable cell sorting).

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2. Add the primary labeled antibody (0.1–10 μg/ml); incubate for ~30 min at room temperature or 4  C. 3. Wash the cells 3 by centrifugation at 400  g for 5 min; resuspend in 500 μl to 1 ml of ice-cold PBS, 10% FBS or BSA, 1% sodium azide (omit for viable cell sorting). 4. Keep the cells in the dark on ice or at 4  C until analyzed. 5. Analyze using appropriate instrument settings for the analyzer or the sorter and for data acquisition software. 3.6 In Vitro Differentiation Assays

In vitro differentiation assays do not probe the inherent, native differentiation potential of cells, but only their response to chemical cues. They are prone to artifacts. Dystrophic calcification cannot be distinguished from matrix mineralization by histochemical stains [34]. In some cases, cells adsorb lipids from the serum rather than synthesize them de novo [35]. While cartilage formation was first demonstrated by in vivo transplantation of BMSCs/SSCs in diffussion chambers, more recent assays rely on the formation of high density cell pellets in vitro [18, 19], which appears to provide the appropriate 3D configuration to support cartilage formation. To date, it has been difficult to form cartilage with BMSCs in vivo in open systems due to the lack of appropriate scaffolds that inhibit vascular invasion but maintain nutrient exchange. However, one such scaffold has been recently identified [36]. While the in vitro assays are prone to artifacts, if performed properly, they provide a first glimpse at the differentiation capacity of a given population of cells.

3.6.1 In Vitro Osteogenic Differentiation Assay

1. Plate BMSCs at a density of 1.5  103 cells/cm2 in SM, switch to OM when cells just reach confluency (see Note 8). 2. Incubate cultures for up to 6 weeks with medium changes every 3 days. 3. Once calcification is visually apparent (mineral is phase bright), fix and stain with either Alizarin Red S or von Kossa (Fig. 3).

3.6.2 In Vitro Adipogenic Differentiation Assay

1. Plate BMSCs at a density of 4  103 cells/cm2 in SM and then switch to one of the AM formulations indicated above once they reach confluency. 2. Incubate cultures for up to 4 weeks with medium changes every 3 days. 3. Once fat accumulation is visually apparent, fix and stain with Oil Red O (Fig. 3).

3.6.3 In Vitro Chondrogenic Differentiation Assay

1. Centrifuge BMSCs (2.5  105) at 500  g in 15 ml polypropylene conical tubes in 5 ml of chondrogenic medium (see Note 9).

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Fig. 3 In vitro differentiaton assays. The osteogenic and adipogenic in vitro differentiation assays are highly prone to artifacts but are often used. Cells plated in SM, and then switched to OM prior to confluence will begin to calcify, as shown by alizarin red S staining. Cells plated in SM and switched to AM prior to confluence will form multilocular fat droplets within their cytoplasm as shown by staining with oil red O. On the other hand, chondrogenic differentiation is best done in vitro, by forming a high density pellet culture. If successful, chondrocytes will be seen lying in lacunae, surrounded by a matrix that stains purple with toluidine blue (metachromasia)

2. Incubate with caps partially unscrewed for 3 weeks at 37 in 5% CO2, with a medium change at 2–3 day intervals. 3. Harvest by washing with PBS, fix in 4% neutral buffered formalin for 2 h, brief demineralization with 10% EDTA in PBS, embed in paraffin, section at 5 μm, and perform histological analysis by staining with toluidine blue (cartilage matrix is stained purple) (see Note 10, Fig. 3). 3.6.4 Analysis of Gene Expression

There are numerous methods for the analysis of gene expression of cultured cells (and from cells transplanted in vivo as described below) using RT-PCR, quantitative RT-PCR, microarray profiling, RNA-seq and single cell RNA-seq which are too lengthy to describe in detail here (reviewed in [37]). Nonetheless, there is a very characteristic pattern of gene expression as BMSCs/SSCs undergo differentiation. However, measurement of expression of these markers is not a guarantee of true differentiation: expression of markers must be matched with evidence of true differentiation as based on appropriate assays (in vivo transplantation as described below for osteogenic and adipogenic differentiation, the in vitro cartilage pellet assay as described above). Some markers useful for distinguishing differentiated phenotypes are listed below (but there are many others): 1. For osteogenesis: ALPL, BGLAP, BGN, COL1A1, COL1A2, DCN, DMP-1, FGF-23, FN, IBSP, IGFBP-3, MEPE, PDPN, PHEX, SCUBE3, SOST, SP7, SPARC, SPP1, RUNX2. 2. For adipogenesis: ADIPOQ, ADIPOR1, ADIPOR2, CIDEA, CHC22, DLK1, FABP, FATP1-6, GALECTIN-12, GLUT4,

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LEPTIN, PERILIPIN-2, PGC1A, PPARG, PREF-1, BSCL2, UCP1, VSTM2A, VSTM2B, ZIC1. 3. For chondrogenesis: ACAN, ANXA6, CTSB, CD44, CD151, CHADL, CHAD, COL2A1, COL9A2, COL10A1, COL11A1, COL12A1, COMP, CRTAC1, DSPG3, FAM20B, MAT1, MAT3, MAT4, SOX5, SOX6, SOX9. 3.7 In Vivo Differentiation Assay—Formation of an Ectopic Bone/ Marrow Organ (Ossicle)

In vivo transplantation of BMSCs/SSCs has become the gold standard by which to measure their multipotential nature. Both murine and human BMSCs have the ability to form bone, myelosupportive stroma and adipocytes when transplanted subcutaneously along with an appropriate carrier (an ectopic ossicle) . However, human BMSCs form ectopic ossicles only with ceramic/hard particles, whereas murine BMSCs can do so on both ceramic-based scaffolds and in collagen sponges [38] (Fig. 4). When murine and human clonal strains, derived from a single CFU-F, were interrogated by in vivo transplantation, ~10–20% were found to be multipotent (formed a complete bone/marrow organ), ~50% form bone only, and the remainder form fibrous tissue [39, 40]. Thus, not all BMSCs, not even all CFU-Fs, are multipotent. The in vivo transplantation assay is the only assay that descrimates between cells that are multipotent, and cells that are not [4, 7, 28].

3.7.1 Ceramic Carrier Constructs (Human and Mouse BMSCs)

1. Sterilize ceramic particles by heating at 220  C overnight, aliquot 40 mg aseptically into sterile round bottomed 1 ml cryotubes. 2. Pellet BMSCs at 135  g for 10 min; resuspend in SM to the volume in mls equal to the number of transplants to be prepared. 3. Wash ceramic particles twice with SM. 4. Transfer BMSCs (1–2  106 cells in 1 ml of SM) into tubes with particles; mix and incubate at 37  C for 70–100 min with slow rotation. 5. Centrifuge particles with adherent BMSCs (135  g for 1 min); remove the supernatant. 6. Transplant as described below.

3.7.2 Collagen Sponge Constructs (Murine BMSCs)

1. Sterilize sponges if necessary; cut collagen sponges into cubes of the desired size or into any other shape; place into SM; squeeze with forceps to remove air bubbles. 2. Transfer BMSCs (1–2  106 cells/1 ml of SM) into individual 1 ml Eppendorf tube, pellet at 135  g for 10 min; discard all but 50–150 μl of the supernate (depending on the size of the sponge); resuspend the pellet.

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Fig. 4 In vivo differentiation assays. Murine BMSCs will form a bone/marrow organ (bone, hematopoiesis supportive stroma, marrow adipocytes (adipo) of donor origin, with hematopoiesis (hp) of recipient origin when transplanted in conjunction with collagen sponges (a, b), and with hydroxyapatite/tricalcium phosphate (HA/TCP, s—scaffold) (data not shown). On the other hand, human BMSCs will only form a bone/marrow organ with hard (ceramic, devitalized bone, etc.) scaffolds (c, d)

3. Blot sponges between two sheets of sterile filter paper; immediately place into freshly resuspended cells in the Eppendorf tube where the sponge expands, absorbing the cells. 4. Transplant as described below. 3.8 Surgery to Create Ectopic Ossicles

This procedure describes the use of mice as recipients. Similar procedures can be used when using other species of recipients. 1. Anesthetize the mouse; shave or use a depilatory if necessary; clean the skin with betadine and 70% ethanol; make a single 3 cm longitudinal incision with a sterile scalpel in the skin along the dorsal surface. 2. Use the tip of sterile round-tipped scissors to make a pocket for the transplant by inserting the scissor subcutaneously, open the scissors by approximately 1 cm; use a sterile spatula to insert ceramic transplants or sterile forceps to insert collagen sponge transplants (usually 4 transplants/mouse); close the incision with several autoclips or with surgical glue. 3. Harvest at various time points; fix with 4% neutral buffered formaldehyde overnight; decalcify; embed in paraffin for standard histological analyses. If histochemical staining of the

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paraffin sections is intended, it is better to perform demineralization with 10% EDTA in PBS. Its duration depends on the size of the transplants, on the amount of bone and/or of hydroxyapatite in the transplants, on the temperature (much longer at 4  C than at room temperature). Decalcification is shorter if the solution is replaced often and if shaking is performed. The most gentle demineralization can take up to 6 weeks. To be sure that demineralization is completed and no more calcified structures are left, X-rays of the transplants may be performed. 4. Determine donor origin in transplants by donor-specific in situ hybrization probes or antibodies, or by use of reporters introduced into the donor cells.

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Notes 1. It is not well recognized that culture conditions vary from one species to another [21], and that fetal bovine serum must be tested extensively to select lots that are suitable for one animal species or another. The specific lot of fetal bovine serum used is critical for determination of CFE [41]. Furthermore, it has been determined that heat inactivation can substantially reduce the ability of FBS to support colony formation and growth [41]. 2. Identification of a scaffold that is able to maintain the biological activities of BMSCs/SSCs is critical. Unfortunately, many of the commercially available scaffolds, both ceramic or otherwise, have not been found to support even bone formation very well. Generally speaking hydroxyapatite/tricalcium phosphate (60%/40%) ceramics have been useful (e.g., the mineral component of Attrax™, Nuvasive Inc., MASTERGRAFT™, Medtronic, Inc.; BioOss™, Geistlich), and other scaffolds currently under development may be even better. For murine cells, Gelfoam™, Pfizer, Inc. has shown the most consistent results to date. 3. It has been reported that the colony forming efficiency and proliferation of murine BMSCs (and possibly human BMSCs as well) is increased by growth in hypoxic (2–5% O2) conditions. However, it is not yet clear that the full biological activity of BMSCs is maintained under these growth conditions. 4. When human cells are implanted into immunocompromised mice, donor cells can be identified by either anti-human antibodies (such as antibodies against human mitochondria) or anti-human DNA sequences (such as ALU). If, however, mouse cells are implanted, donor cells of male origin can be

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identified in a female recipient by a FISH probe against mouse Y chromosome, produced by many companies. 5. Excessive pressure, both positive and negative, should be avoided while passing cell suspensions through the needles. Murine cells, in particular, are very sensitive to rapid changes in pressure. 6. Not all BMSCs are clonogenic. The cell concentrations indicated will result in densities that will allow for density independent growth of BMSCs from a single CFU-F. By clonal analysis, ~1:5 of the colonies are multipotent based on the in vivo transplantation assay [39]. Thus the colony forming efficiency is a rough estimate of the number of SSCs. 7. Rodent bone marrow stromal cells are often highly contaminated with hematopoietic cells, primarily macrophages (which can take on a BMSC-like appearance to the untrained eye). Passaging significantly reduces their presence, but does not eliminate them. Magnetic bead sorting or FACS sorting strategies have been used to eliminate the hematopoietic cells from murine BMSC cultures [33]. 8. The in vitro osteogenic assay is highly variable from one animal species to another, from one strain of mice to another, and if from different cell preparations to another. The cell layer has the propensity to roll up if it becomes superconfluent with abundant extracellular matrix and the OM is not added at the right time. Optimization may be required by adding OM at different times before or after reaching confluency, or by reducing the level of serum. 9. For the chondrogenic assay, it is extremely important to use polypropylene tubes, which prevent cell attachment to the walls of the tube. 10. In histological evaluation of pellet cultures, staining with toluidine blue is essential to determine if cartilage is formed. Bona fide chondrocytes must be seen lying in lacunae, surrounded by matrix that stains purple with toluidine blue. Although Alcian Blue Or Safranin O are often used, Alcian Blue is not specific enough (osteoid will stain lightly with Alcian Blue), and Safranin O is also used as a nuclear stain.

Acknowledgments Work in the authors’ laboratories was supported by the Division of Intramural Research of the National Institute of Dental and Craniofacial Research, a part of the Intramural Research Program of the National Institutes of Health, Department of Health and Human Services (to PGR and SAK), and the Department of Molecular Medicine, Sapienza University of Rome (to PB and MR).

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References 1. Owen M, Friedenstein AJ (1988) Stromal stem cells: marrow-derived osteogenic precursors. Ciba Found Symp 136:42–60 2. Bianco P, Riminucci M, Gronthos S, Robey PG (2001) Bone marrow stromal stem cells: nature, biology, and potential applications. Stem Cells 19:180–192 3. Bianco P, Robey PG (2004) Skeletal stem cells. In: Lanza RP (ed) Handbook of adult and fetal stem cells. Academic Press, San Diego, pp 415–424 4. Bianco P, Cao X, Frenette PS, Mao JJ, Robey PG, Simmons PJ, Wang CY (2013) The meaning, the sense and the significance: translating the science of mesenchymal stem cells into medicine. Nat Med 19:35–42 5. Bianco P, Robey PG (2015) Skeletal stem cells. Development 142:1023–1027 6. Friedenstein AJ (1980) Stromal mechanisms of bone marrow: cloning in vitro and retransplantation in vivo. Hamatol Bluttransfus 25:19–29 7. Bianco P, Kuznetsov S, Riminucci M, Robey PG (2006) Postnatal skeletal stem cells. Meth Enzymol 419:117–148 8. Sacchetti B, Funari A, Michienzi S, Di Cesare S, Piersanti S, Saggio I, Tagliafico E, Ferrari S, Robey PG, Riminucci M, Bianco P (2007) Self-renewing Osteoprogenitors in bone marrow sinusoids can organize a hematopoietic microenvironment. Cell 131:324–336 9. Sacchetti B, Funari A, Remoli C, Giannicola G, Kogler G, Liedtke S, Cossu G, Serafini M, Sampaolesi M, Tagliafico E, Tenedini E, Saggio I, Robey PG, Riminucci M, Bianco P (2016) No identical "mesenchymal stem cells" at different times and sites: human committed progenitors of distinct origin and differentiation potential are incorporated as adventitial cells in microvessels. Stem Cell Rep 6:897–913 10. Mendez-Ferrer S, Michurina TV, Ferraro F, Mazloom AR, Macarthur BD, Lira SA, Scadden DT, Ma’ayan A, Enikolopov GN, Frenette PS (2010) Mesenchymal and haematopoietic stem cells form a unique bone marrow niche. Nature 466:829–834 11. Park D, Spencer JA, Koh BI, Kobayashi T, Fujisaki J, Clemens TL, Lin CP, Kronenberg HM, Scadden DT (2012) Endogenous bone marrow MSCs are dynamic, fate-restricted participants in bone maintenance and regeneration. Cell Stem Cell 10:259–272 12. Zhou BO, Yue R, Murphy MM, Peyer JG, Morrison SJ (2014) Leptin-receptor-expressing mesenchymal stromal cells represent the

main source of bone formed by adult bone marrow. Cell Stem Cell 15:154–168 13. Chan CK, Seo EY, Chen JY, Lo D, McArdle A, Sinha R, Tevlin R, Seita J, Vincent-Tompkins J, Wearda T, Lu WJ, Senarath-Yapa K, Chung MT, Marecic O, Tran M, Yan KS, Upton R, Walmsley GG, Lee AS, Sahoo D, Kuo CJ, Weissman IL, Longaker MT (2015) Identification and specification of the mouse skeletal stem cell. Cell 160:285–298 14. Mizuhashi K, Ono W, Matsushita Y, Sakagami N, Takahashi A, Saunders TL, Nagasawa T, Kronenberg HM, Ono N (2018) Resting zone of the growth plate houses a unique class of skeletal stem cells. Nature 563:254–258 15. Chan CKF, Gulati GS, Sinha R, Tompkins JV, Lopez M, Carter AC, Ransom RC, Reinisch A, Wearda T, Murphy M, Brewer RE, Koepke LS, Marecic O, Manjunath A, Seo EY, Leavitt T, Lu WJ, Nguyen A, Conley SD, Salhotra A, Ambrosi TH, Borrelli MR, Siebel T, Chan K, Schallmoser K, Seita J, Sahoo D, Goodnough H, Bishop J, Gardner M, Majeti R, Wan DC, Goodman S, Weissman IL, Chang HY, Longaker MT (2018) Identification of the human skeletal stem cell. Cell 175:43–56 e21 16. Debnath S, Yallowitz AR, McCormick J, Lalani S, Zhang T, Xu R, Li N, Liu Y, Yang YS, Eiseman M, Shim JH, Hameed M, Healey JH, Bostrom MP, Landau DA, Greenblatt MB (2018) Discovery of a periosteal stem cell mediating intramembranous bone formation. Nature 562:133–139 17. Duchamp de Lageneste O, Julien A, AbouKhalil R, Frangi G, Carvalho C, Cagnard N, Cordier C, Conway SJ, Colnot C (2018) Periosteum contains skeletal stem cells with high bone regenerative potential controlled by Periostin. Nat Commun 9:773 18. Johnstone B, Hering TM, Caplan AI, Goldberg VM, Yoo JU (1998) In vitro chondrogenesis of bone marrow-derived mesenchymal progenitor cells. Exp Cell Res 238:265–272 19. Barry F, Boynton RE, Liu B, Murphy JM (2001) Chondrogenic differentiation of mesenchymal stem cells from bone marrow: differentiation-dependent gene expression of matrix components. Exp Cell Res 268:189–200 20. Friedenstein AJ, Latzinik NV, Gorskaya Yu F, Luria EA, Moskvina IL (1992) Bone marrow stromal colony formation requires stimulation by haemopoietic cells. Bone Miner 18:199–213

396

Pamela G. Robey et al.

21. Kuznetsov SA, Friedenstein AJ, Robey PG (1997) Factors required for bone marrow stromal fibroblast colony formation in vitro. Br J Haematol 97:561–570 22. Morrison SJ, Scadden DT (2014) The bone marrow niche for haematopoietic stem cells. Nature 505:327–334 23. Ambrosi TH, Longaker MT, Chan CKF (2019) A revised perspective of skeletal stem cell biology. Front Cell Dev Biol 7:189 24. Hung SP, Ho JH, Shih YR, Lo T, Lee OK (2012) Hypoxia promotes proliferation and osteogenic differentiation potentials of human mesenchymal stem cells. J Orthop Res 30:260–266 25. D’Ippolito G, Diabira S, Howard GA, Roos BA, Schiller PC (2006) Low oxygen tension inhibits osteogenic differentiation and enhances stemness of human MIAMI cells. Bone 39:513–522 26. Suire C, Brouard N, Hirschi K, Simmons PJ (2012) Isolation of the stromal-vascular fraction of mouse bone marrow markedly enhances the yield of clonogenic stromal progenitors. Blood 119:e86–e95 27. Markway BD, Tan GK, Brooke G, Hudson JE, Cooper-White JJ, Doran MR (2010) Enhanced chondrogenic differentiation of human bone marrow-derived mesenchymal stem cells in low oxygen environment micropellet cultures. Cell Transplant 19:29–42 28. Bianco P, Robey PG, Simmons PJ (2008) Mesenchymal stem cells: revisiting history, concepts, and assays. Cell Stem Cell 2:313–319 29. Chen KG, Johnson KR, Robey PG (2017) Mouse genetic analysis of bone marrow stem cell niches: technological pitfalls, challenges, and translational considerations. Stem Cell Rep 9:1343–1358 30. Quirici N, Soligo D, Bossolasco P, Servida F, Lumini C, Deliliers GL (2002) Isolation of bone marrow mesenchymal stem cells by antinerve growth factor receptor antibodies. Exp Hematol 30:783–791 31. Tormin A, Li O, Brune JC, Walsh S, Schutz B, Ehinger M, Ditzel N, Kassem M, Scheding S (2011) CD146 expression on primary nonhematopoietic bone marrow stem cells is correlated with in situ localization. Blood 117:5067–5077 32. Simmons PJ, Torok-Storb B (1991) Identification of stromal cell precursors in human bone

marrow by a novel monoclonal antibody, STRO-1. Blood 78:55–62 33. Chou DB, Sworder B, Bouladoux N, Roy CN, Uchida AM, Grigg M, Robey PG, Belkaid Y (2012) Stromal-derived IL-6 alters the balance of myeloerythroid progenitors during toxoplasma gondii infection. J Leukoc Biol 92:123–131 34. Bonewald LF, Harris SE, Rosser J, Dallas MR, Dallas SL, Camacho NP, Boyan B, Boskey A (2003) von Kossa staining alone is not sufficient to confirm that mineralization in vitro represents bone formation. Calcif Tissue Int 72:537–547 35. Diascro DD Jr, Vogel RL, Johnson TE, Witherup KM, Pitzenberger SM, Rutledge SJ, Prescott DJ, Rodan GA, Schmidt A (1998) High fatty acid content in rabbit serum is responsible for the differentiation of osteoblasts into adipocyte-like cells. J Bone Miner Res 13:96–106 36. Kuznetsov SA, Hailu-Lazmi A, Cherman N, de Castro LF, Robey PG, Gorodetsky R (2019) In vivo formation of stable hyaline cartilage by naive human bone marrow stromal cells with modified fibrin microbeads. Stem Cells Transl Med 8:586–592 37. Lowe R, Shirley N, Bleackley M, Dolan S, Shafee T (2017) Transcriptomics technologies. PLoS Comput Biol 13:e1005457 38. Krebsbach PH, Kuznetsov SA, Satomura K, Emmons RV, Rowe DW, Robey PG (1997) Bone formation in vivo: comparison of osteogenesis by transplanted mouse and human marrow stromal fibroblasts. Transplantation 63:1059–1069 39. Kuznetsov SA, Krebsbach PH, Satomura K, Kerr J, Riminucci M, Benayahu D, Robey PG (1997) Single-colony derived strains of human marrow stromal fibroblasts form bone after transplantation in vivo. J Bone Miner Res 12:1335–1347 40. Sworder BJ, Yoshizawa S, Mishra PJ, Cherman N, Kuznetsov SA, Merlino G, Balakumaran A, Robey PG (2015) Molecular profile of clonal strains of human skeletal stem/ progenitor cells with different potencies. Stem Cell Res 14:297–306 41. Kuznetsov SA, Mankani MH, Bianco P, Robey PG (2009) Enumeration of the colonyforming units-fibroblast from mouse and human bone marrow in normal and pathological conditions. Stem Cell Res 2:83–94

Chapter 24 Isolation and Culture of Periosteum-Derived Progenitor Cells from Mice Chinedu C. Ude, Girdhar G. Sharma, Jie Shen, and Regis J. O’Keefe Abstract This chapter describes the methods of isolation of mouse periosteal progenitor cells. There are three basic methods utilized. The bone grafting method was developed utilizing the fracture healing process to expand the progenitor populations. Bone capping methods requires enzymatic digestion and purification of cells from the native periosteum, while the Egression/Explant method requires the least manipulation with placement of cortical bone fragments with attached periosteum in a culture dish. Various cell surface antibodies have been employed over the years to characterize periosteum derived progenitor cells, but the most consistent minimal criteria was recommended by the International Society for Cellular Therapy. Confirmation of the multipotent status of these isolated cells can be achieved by differentiation into the three basic mesodermal lineages in vitro. Key words Periosteum, Periosteum Progenitor Cells, Bone Grafting, Bone Capping, Egression

1

Introduction The periosteum is an important tissue that is located on the outer surface of cortical bone and possesses mesenchymal progenitors that are involved in bone modeling and repair in the presence of appropriate signals. During development, periosteum contributes to bone growth and modeling in addition to its crucial healing role in the event of bone injury. Periosteum is present on almost all long bones, with only sesamoid bones and the intra-articular portions of bones lacking this tissue [1–3]. Histologically, the periosteum is divided into two distinct layers, an outer fibrous layer, and an inner cambium layer. The cambium is the region of the periosteum that is rich in progenitor cells. During development and growth the periosteum is thick with a robust cambium, however progressively becomes thinner with age. Thus, in adults, the periosteal vessel density and number of progenitor cells are decreased and the periosteum is present as a thin sheath enveloping the bone. This causes loss of much of the childhood osteoblastic potential [3–5],

Matthew J. Hilton (ed.), Skeletal Development and Repair: Methods and Protocols, Methods in Molecular Biology, vol. 2230, https://doi.org/10.1007/978-1-0716-1028-2_24, © Springer Science+Business Media, LLC, part of Springer Nature 2021

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however, upon stimulation by fracture or injury, progenitor cells proliferate, accumulate, and then undergo differentiation. This process is essential for normal fracture healing. In approximately 5–10% of cases, fracture healing is compromised. The isolation of periosteum cells has enabled study of the signals that are necessary to promote the proliferation and differentiation of the periosteum. Periosteal cells can differentiate into both bone and cartilage cells in vivo [1, 6–8]. Periosteum plays an important role in endochondral bone formation, which involves a cartilage intermediate that undergoes calcification and replacement by bone. Intramembranous bone formation, on the other hand, involves osteogenic differentiation directly from periosteal progenitors without the formation of a cartilage intermediate [9, 10]. Following a fracture, cells from the inner cambium layer of the periosteum proliferate to create a critical mass of progenitor cells that subsequently undergo differentiation and bone formation. In the central region of the fracture which is the most avascular, progenitor cells differentiate into cartilage tissue which forms bone through endochondral ossification. Endochondral bone repair is the most common type of facture healing and occurs in the setting of motion at the fractured site which leads to reduced vascularization and a hypoxic environment [3]. At the periphery of the fracture along the surface of the bone, where there is a rich vascular supply, progenitor cells differentiate directly into osteoblasts and bone forms through intramembranous ossification. The isolation of periosteal cells for study in vitro enables approaches to study the nature of the cell population and the signals that regulate the differentiation of these cells. Three different procedures have been developed in the mouse. One method involves the transplantation of a 4 mm segment of the midshaft of the murine femur with periosteum. This allows for the in vivo expansion of the periosteal progenitors, at which time the cells can be isolated by enzymatic digestion [11–14]. The second method involves use of agarose to cap the articular surface of murine femurs and tibias, and the isolation of periosteum derived progenitor cells using enzymatic digestion [15]. Third method involves placement of murine bone fragments into a culture dish to allow the egression of periosteal cells onto the culture dish [7]. Both the bone grafting and bone capping methods have been utilized mainly in mice, but the periosteal egression model has been used to isolate periosteal cells from mice as well as multiple other species including horse, chicken, and human [15–17]. Bone grafting and capping methods require manipulations through enzymatic digestions and centrifugations. This may induce some physiological stress to the isolated cells. On the other hand, egression is more natural as it allows viable cells to explant into the culture medium without any manipulation. In this chapter, we describe our experience with these three basic methods of periosteal cell isolation in mice and the culture

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procedures necessary to maintain and proliferate these cells through many passages (Fig. 1). Elucidating the regenerative potential of periosteal cells has been of importance in orthopaedic research. Periosteum provides a unique niche for progenitor cells and can be a source for molecular factors that modulate cell behavior [9]. Skeletal stem cells (SSCs) represent a group of native uncommitted cells, that are multipotent, capable of both self-renewal and differentiation into lineages of mesodermal origin, including cartilage, bone, and adipose. These progenitor cells were originally identified in the bone marrow stroma, but have been isolated from other tissues including: amniotic fluid, Wharton’s jelly, umbilical cord blood, adipose tissue, skin, synovial membrane, articular cartilage, compact bone, and periosteum [18–21]. Recent nomenclatures, support the naming of these cells according to the tissue in which they reside, reflecting their origin. Hence the periosteum harbors periosteum derived progenitor cells (PDPCs) that act as major players in bone development and fracture healing [8, 22–24]. Historically, diverse opinions on the markers to characterize a pure PDPC population has been debatable for researchers. However, the initial consensus on the utilization of the classic “mesenchymal stem cells” antigenic profile was proposed in agreement with the minimal criteria by the International Society for Cellular Therapy [9]. A summarized report showed that different authors have independently identified that PDPCs were either positive or negative to the following antibodies: Minimal criteria for mesenchymal progenitor cells: CD90+, CD105+, CD73+, CD45
, HLA-DR
, CD14
, CD34
; Integrins: CD29+, CD49e+; Adhesion molecules: CD31
, CD44+, CD166+, CD54+, CD146+; MHC class: HLA-ABC+; Hematopoietic markers: CD14
, CD33
, CD34
, CD45
, CD133
; and some selected additions, including pluripotent markers: MSCA-1+, CD9 +/, CD13+, STRO-1+, SSEA-4+, ScaI+, Sox2+, Oct4+, and Nanog+ [25, 26]. Colnot et al. identified a multipotent periosteal progenitor cell pool that is dependent on the expression of periostin and has high in vitro and in vivo clonogenic potential. They played important roles in fracture regeneration, nevertheless, the loss of periostin impaired their number and function [7]. Chan et al., provided a panel of surface antibodies that included CD45, CD235a, TIE2, CD31, PDPN, CD146, CD73, CD164, and CD90 to separated growth plate zone cells into distinct subpopulations, in order to identify skeletal stem cells. A combination of in vitro and in vivo assessments supported the identity of PDPN+CD146-CD73 +CD164+ fetal growth plate cells as self-renewing, multipotent skeletal stem cells [27]. In a recent work by Debnath et al., a human analogous periosteal stem cell (PSC), present in the long bones and calvarium of

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Fig. 1 Methods of periosteum Isolation. (a) Isolation through bone autografting (BG) This model recreates the events after fracture through an autograft resection and implantation surgery. A hematoma is formed, which is stabilized by the surrounding soft tissues, and cells from the inner cambium layer proliferate, differentiate and migrate nearer to the fracture site, forming a spongy callus. Five days later, through enzymatic digestion, proliferating progenitor cells are harvested from the callus. The cells liberated after digestion are passed through cell strainer and collected by centrifugation before plating in the culture dish/ well. (b) Isolation through bone capping with Agarose (BC). This model targets the direct harvest of progenitor cells that are within the periosteum layer covering the diaphysis of long bones. Long bones are dissected and epiphyses are capped with low-melting-point (LMP) agarose to prevent contamination of the harvest with chondrocytes and bone marrow cells. Through enzymatic digestion, progenitor cells are liberated. The liberated cell populations are filtered through a cell strainer and collected by centrifugation for proliferation in culture media. (c) Isolation through bone egression (EG). Isolation through egression requires no enzymatic digestion or callus formation. It allows both viable committed and uncommitted (native) cells to migrate from the periosteum onto the culture dish/well. Long bones are dissected, epiphyses are cut out and bone marrow flushed with media. The bones may be further cut into smaller sizes to increase periosteal surface contact area. The bones pieces are cultured and periosteal cells explant and proliferate within days

mice was discovered. These cells displayed clonal multipotency, selfrenewal, and sit at the apex of a differentiation hierarchy. In sets of three periosteal cell populations, which all lack THY1.2 and 6C38, and were CD49low CD51low; the cells expressing CD200 +CD105
were termed the periosteal stem cells (PSCs); the cells expressing CD200
CD105
were termed the periosteal progenitor 1 (PP1) cells; while the cells expressing CD105+CD200variable, were designated periosteal progenitor 2 (PP2) cells. It was further shown that PSCs differentiated into PP1 and PP2 in

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addition to THY1.2+ and 6C3+ cells, but neither PP1s nor PP2s produced PSCs in culture. Immunostaining confirmed the presence of these CD200+ cells in the periosteum and a subset expressing gremlin 15 and nestin1 [6]. More recently, Ambrosi et al. proposed a set of markers that broadly label human skeletal stem cell populations with variable CFU-F ability and differentiation to osteogenic, chondrogenic, adipogenic, and stromal lineages. These include CD105, CD140a, CD73, CD90, STRO-1, CD271, and CD44. They also recommended using CD271, CD90, CD51, CD44, CD146, CD106, and LepR with variations depending on the developmental stages investigated. For mouse skeletal stem cells, they proposed using cathepsin K, Sox9, MSSC, and Mx1. However, it was concluded that all bone-resident stem cells cannot be defined, other than they are likely of nonhematopoietic (CD45
) and nonendothelial (CD31
) origin [28]. Among these varied cell surface antibodies that have frequently been utilized to characterize PDPCs, CD90.2+, CD105+, Sca1+ and CD45
, CD34
have commonly been reported in the literature [14, 29–31]. A combination of these markers with CD 200+ can enrich the characterization of progenitor cells in a population of isolated periosteal cells [6]. Hence, we utilized a panel of cell surface antibodies for characterization of the isolated periosteal cells and describe staining protocols to assess a trilineage differentiation to confirm their multipotency.

2 2.1

Materials Mice

1. Sex and aged matched mice at about 6 weeks of age for bone autografting (n ¼ 5) (see Note 1). 2. Sex and aged matched mice at about 3–6 weeks of age or younger for bone capping or egression models of PDPC isolation (n ¼ 5) (see Note 1).

2.2 Surgical Instruments and Anesthesia

1. Shaving clipper. 2. Dremel cutter. 3. Spinal needle (25G). 4. Dissecting scissors-sharp point straight 4.500 . 5. Tissue forceps 4.7500 with blunt, serrated ends (2 sets). 6. Syringe needle (22G). 7. Scalpel handle #3 and surgical blade #15. 8. Forceps scissors-stainless steel hemostat 500 with blunt, serrated ends. 9. Mini plier-cutter.

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10. Suture 5-0 Ethilon 1800 . 11. Needle holder 500 . 12. Sterile gauze. 13. Autoclaved firm paper (3  2 cm). 14. 70% ethanol (EtOH). 15. Povidone–iodine prep pads. 16. Anesthesia cocktail: Combination of ketamine, xylazine, and dH2O or 2–5% isoflurane. 2.3

Cell Culture

1. Sterile Petri dish 60cm2. 2. Sterile culture dish—tissue culture treated 60cm2. 3. Sterile 6-well plates—tissue culture treated. 4. Sterile micropipette tips (500 μL). 5. Micropipette dispenser. 6. Sterile polypropylene centrifuge tube—(15 mL, 50 mL). 7. Cylindrical flask (500 mL). 8. Cell strainer (70 μm). 9. Collagenase D or P solution (prepare 1 mg/mL working stock). 10. Alpha DMEM. 11. 1 phosphate-buffered saline (PBS), pH 7.4. 12. Fetal bovine serum (FBS). 13. Gentamicin sulfate (10 mg/mL). 14. Basic fibroblast growth factor-2 (bFGF-2 50 μg/mL). 15. 70% ethanol (EtOH). 16. Complete medium (Alpha DMEM/F12—Dulbecco’s Modified Eagle Medium + 10% FBS + 0.05% gentamicin sulfate +20 ng/mL bFGF-2). 17. 0.05% trypsin–EDTA solution. 18. Agarose (prepare a working stock of 5% low melting point agarose with TE buffer and autoclave).

2.4

Flow Cytometry

1. Cells (1–5  106 cells/mL) in ice-cold PBS + 10% FBS + 1% sodium azide. 2. Polystyrene round bottom 12  75 mm2 Falcon tubes. 3. 1% Paraformaldehyde in ice-cold PBS to prevent deterioration for extended storage 4. Conjugated primary antibodies (CD 90.2-Phycoerythrin PE, CD 105-PE-CY7, Sca1-Alexa flour 488, CD 200-PerCPeFlour 710, CD 45-Pacific blue, and CD 34-APC).

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2.5 Osteogenic Differentiation

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1. StemPro Osteogenic Differentiation Kit (Gibco). 2. Alizarin Red S staining solution: 1% Alizarin Red S in 95% ethanol. 3. 4% Paraformaldehyde in 1 PBS, store at 4  C.

2.6 Chondrogenic Differentiation

1. StemPro Adipogenic Differentiation Kit (Gibco). 2. Alcian Blue staining solution (pH 2.5) (Sigma Aldrich catalog number B8438). 3. 3% acetic acid solution. 4. 0.1% Nuclear Fast Red solution. 5. 4% Paraformaldehyde in 1 PBS, store at 4  C.

2.7 Adipogenic Differentiation

1. StemPro Adipogenic Differentiation Kits (Gibco). 2. Oil Red O staining kit (Sigma Aldrich Catalog Number MAK194). 3. 4% paraformaldehyde in 1 PBS. 4. 100% and 60% isopropanol, store at 4  C.

3

Methods Three methods have been utilized for PDPC isolation. These include the bone grafting, capping of long bones with agarose, and an Egression/Explant procedure. (see Note 2). These isolations result in some variations in cell yield in our laboratory. Our standard expected cell yields are shown in Table 1.

3.1 Isolation of PDPCs Through Bone Autografting (BG)

Isolation of PDPCs occurs through the enzymatic digestion of expanded progenitor cell populations present along the 4 mm surface of the mid-shaft femur autograft 5 days after transplantation. As mentioned above, in this model, a hematoma forms after bone transplantation, which is stabilized by the surrounding soft tissues, and cells from the inner cambium layer proliferate along the surface of the 4 mm bone graft (Fig. 1a). 1. Anesthetize the mouse using 0.15 mL of ketamine/xylazine cocktail (20 μg/mL) by intraperitoneal injection before surgery. Allow approximately 3–5 min for the drug to take effect (see Note 3). 2. As soon as the mouse is sedated, pull it out from its cage and shave the areas around the right or left hind leg, to the lower back and abdomen. Clear the shaved hairs and clean the skin first with 70% EtOH and subsequently with povidone–iodine (see Note 3). 3. Place the mouse on a right or left lateral recumbent to expose the right or left femur (depending on limb of choice for

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Table 1 The range of typical PDPCs yield per mouse from the BG, BC and EG Methods Methods of PDPCs isolation

Expected cell yield per mouse in 14 days

Bone grafting

3  105 to 5  105

Bone capping

8  105 to 1  106

Egression/Explant

1  105 to 3  105

procedure). Using the forceps, pull the skin up from the muscle below and make about 1 cm incision with the scissors to expose the muscle covering the femur. 4. With two forceps, grab the muscle covering the femur and tease them apart to both ends of the incision. Slide one forceps under the exposed femur, to secure the muscles’ movement and the bone. 5. With Dremel cutter, cut about 4 mm of bone at the mid-shaft of femur. Starting from the proximal area, place the autoclaved hard paper under the bone, then to the distal part. Place the graft in a culture dish with basal DMEM medium (see Note 4). 6. Use a sterile 22G needle to burrow through the cut femur until protruding from the proximal femur in the trochanteric region near the hip joint. Insert a 25G spinal needle into the protruding 22G needle, and then remove the 25G needle. Pull the 22G needle to draw the spinal needle through the femur and pass in the 4 mm graft to the distal end of the femur and knee joint (see Note 4). 7. Secure the graft by first bending the spinal needle at the proximal head, then the tibia-femoral junction. Use the pliers to cut the extra protrusion at the knee joint, before bending. Close the muscle above the femur and the skin. Use Ethilon suture to close the incision (see Note 5). 8. Finally, give an intramuscular injection of 0.10 mL buprenorphine (0.5–1.0 mg/kg) as an analgesic to help recuperation and put mice back to the cage and monitor for recovery, suture dehiscence, and allow mouse to recover for 5 days. 9. On the fifth day post-surgery, euthanize mice using CO2 chamber or cervical dislocation. Alternatively, the mouse can be sedated with ketamine cocktail for the harvest of the graft, then euthanized with CO2 afterward. 10. Soak mice in a cylindrical flask containing 70% EtOH for 5–10 min to sterilize the skin. Alternatively, shave the surgical areas after sedation and clean with alcohol and povidone-iodine to maintain a sterile graft harvest. Cut off the bent end of the

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spinal needle holding the grafted bone at the tibia-femoral junction. 11. Dissect the grafted area and pull the spinal needle to retract from the tibia-femoral junction, to remove the bone graft. Then place the graft in a petri dish containing about 1 mL of alpha-DMEM. Wash the resected tissues a few times by changing the DMEM on the culture dish, two times, to remove red blood cells. Using the scalpel and blade, scrape the periosteum callus on the resected bone graft, from the sides and the ends (see Note 6). 12. Place the scraped tissues and about 1 mL DMEM into a 15 mL conical tube containing 10 mL of 1 mg/mL collagenase D or P (see Note 6). 13. Put the mixture in a shaking incubator to incubate at 37  C for about 45 min—1 h. Take out the digested mixture and pour it through a cell strainer into a 50 mL conical tube. Collect the filtrate and centrifuge at 400  g for 5 min (see Note 6). 14. Then resuspend with 5 mL complete culture medium and seed in a 30 cm2 culture dish or two wells of 6-well plate. 3.2 Isolation of PDPCs Through Bone Capping with Agarose (BC)

Isolation of PDPCs occurs through direct enzymatic digestion of periosteal tissues covering the diaphyseal bone after embedding epiphyses in low-melting-point (LMP) agarose (Fig. 1b). 1. Euthanize mice using CO2 chamber or cervical dislocation and dissect their hind limbs. Clear off the muscles and connective tissues on the femur and tibia under sterile conditions and place the isolated bones in a petri dish containing 2 mL DMEM. 2. Dip the articular end/epiphyses into a prewarmed 5% LMP agarose and placed them in another Petri dish containing a small amount of media to solidify the agarose (see Note 6). 3. After solidification of the agarose over about 5 min, place the bones in a 15 mL conical tube containing 10 mL of 1 mg/mL collagenase D or P. 4. Digest initially for 10 min and discarded the digests as they could contain cells from remnant muscle and connective tissue. Then proceed with a second 1 h digest to isolate PDPCs. Filter the digest solution through a cell strainer into a 50 mL conical tube. 5. Collect the filtrate and centrifuge at 400  g for 5 min. Then resuspend with the complete culture medium and seed in a 60 cm2 culture dish or four wells of a 6-well plate.

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3.3 Isolation of PDPCs Through Egression/Explant (EG)

Isolation of PDPCs via egression requires neither enzymatic digestion nor callus formation. It allows both viable committed and uncommitted (native) cells to migrate from the tissue into the culture flask (Fig. 1c). 1. See Subheading 3.2, step 1 above. 2. Cut out articular ends of the bone and growth plates and flush the bones with DMEM to remove bone marrow from within the core which would contaminate the culture. 3. Cut the bones into smaller sizes of about 4 mm to increase the periosteal surface contact to medium and cultured in prewarmed complete media. 4. Cells will migrate out of the explanted tissue within 3 days.

3.4 Periosteum Derived Progenitor Cell (PDPC) Culture

1. For the BG and BC methods (Subheadings 3.1 and 3.2), after primary PDPCs have been isolated via enzymatic digestion and plated with 2 or 3 mL of complete media (see Note 7) in a well of a 6-well plate or 60 cm2 culture dish, place the isolated cells in a hypoxic CO2 incubator and monitor every day for proliferation and expansion of periosteal cells in culture. Discard any cells in the well/dish that exhibit contamination. 2. Add 1 mL of culture media on the third and fifth day to replenish nutrients. Remove all media and wash cells on the seventh day with prewarmed 1 PBS and replace with fresh culture media. Observe the progress of the cell each day and change media every 2 days (see Note 7). 3. For the EG method, remove the bone fragments from the culture after 7 days and gently wash cells with prewarmed 1 PBS and replace with complete culture media (see Note 7). 4. In all methods, at 14 days post culture, remove the PDPCs from the culture dish with about 1 mL of 0.05% trypsin–EDTA for 3–5 min. Cells are counted after neutralizing trypsin with media and viability evaluated using a hemocytometer and through trypan blue exclusion. 5. Subculture cells up to 5 passages with low seeding densities for proliferation in T-75 cm2 culture flasks (see Note 8). When the desired cell numbers are attained, utilize the cells for in vitro experiments/analysis or alternatively the cells may be cryofrozen for later use.

3.5 Flow Cytometry Evaluation

Several cell surface markers have been utilized in the characterization of isolated PDPCs, however, in culture, it is generally agreed that PDPCs are negative for hematopoietic and endothelial markers, and positive for makers expressed by connective tissues. PDPCs could be minimally identified by CD 90.2+, Sca1+, CD 105+, CD 200+ and CD 45
, CD34
. Using direct flow

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Table 2 The range of positivity of the surface antibodies that we used to characterize PDPCs Surface antibody/marker

Range of expression (%)

CD 90.2

90–96%

Sca-1

94–99%

CD 105

48–59%

CD 200

0.9–2.0%

cytometry protocols, PDPCs can be evaluated. The ranges of the expression of various cell surface markers observed in our laboratory are shown in Table 2. 1. Trypsinize and wash the cells with 1 PBS, and adjust the cell suspension to a concentration of 1–1.5  106 cells/mL in ice-cold 1 PBS, containing 10% BSA and 1% sodium azide. 2. Add the primary conjugated antibody (0.1–10 μg/mL) and incubate for 30 min at room temperature or 4  C. 3. Wash cells three times followed by centrifugation at 400  g for 5 min and resuspend the cells in 500–1000 μL of ice-cold 1 PBS containing 10% BSA and 1% sodium azide. 4. Cells should be kept in the dark on ice or 4  C until the scheduled time for analysis on a flow cytometer. Data is analyzed using the FlowJo software (For best result, run the flow cytometry evaluation as soon as possible). 5. For extended storage (more than 12 h), resuspend the cells in 1% paraformaldehyde to prevent deterioration. 3.6 Osteogenic Differentiation (Fig. 2)

1. Trypsinize PDPCs and plate them in 3 mL normal culture medium at a high density of about 2.0  105 cells per well in a 12 well plate and allow the culture to sit for 12–24 h in a normoxic 37  C CO2 incubator. 2. Remove media and wash once with prewarmed 1 PBS, before the adding 2 mL of prewarmed complete osteogenic medium. Incubate cultures for up to 21 days with media changes every 2–3 days. 3. Increased deposition of extracellular matrix and calcification can be visualized from the tenth to 21st day, and the Alizarin Red S solution will stain very positive from the 14th day onward indicating calcification within the cultures. 4. Remove all culture media from each well and gently wash cells twice with 2 mL 1 PBS and fix the cells with 2 mL of 4% paraformaldehyde fixative solution for about 15 min or more at room temperature (see Note 9).

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Fig. 2 Trilineage differentiation (Osteogenic, Chondrogenic, and Adipogenic). Trilineage differentiation assays start with plating the cells at optimal cell density. The passage zero (P0) cells are allowed to proliferate until the required number is achieved. The osteogenic induction requires high seeding density (SD), to enable matrix formation and calcium deposition. The chondrogenic induction requires, very high SD to enable matrix and glycosaminoglycan deposition. However, the adipogenic induction requires moderate SD. After a maximum induction period of 21 days, Alizarin Red S solution shows a bright/dark orange-red stain for bone; Alcian Blue shows positive cartilage formation (blue) and Oil red O shows positivity for adipocyte formation (red)

5. Remove the fixative and wash the cells three times with 2 mL of distilled water (ddH2O). Add about 2 mL of working stock Alizarin Red S solution. Incubation at room temperature for about 10–20 min is sufficient to show a bright/dark orangered stain (depending on the extent of calcium deposition). 6. Remove the dye and wash cells three times with ddH2O, inspect the cells with a phase-contrast microscope and take images as desired. Store plates at
20  C if dye extraction is required. 3.7 Chondrogenic Differentiation (Fig. 2)

1. Trypsinize cells and plate a micromass of PDPCs in 50 μL normal culture medium at a high density of about 1.5  106 cells per well in a 12-well plate and allow the culture to sit for 2 h. Add 3 mL of culture medium and allow to sit for 12–24 h in a normoxic 37  C CO2 incubator. 2. Remove media and wash once, with prewarmed 1 PBS, before adding 2 mL of prewarmed complete chondrogenic medium. Incubate cultures for up to 21 days with media changes every 2–3 days.

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3. Morphological changes can be seen from the third day and deposition of extracellular matrix and glycosaminoglycans from 10 to 21 days. Alcian Blue solution will stain cells positively from the tenth day onward. 4. Remove all culture media from each well and gently wash cells twice with 2 mL 1 PBS and fix the cells with 2 mL of 4% paraformaldehyde fixative solution for about 15 min or more at room temperature (see Note 9). 5. Remove the fixation and wash the cells three times with ddH2O. Add about 2 mL working stock of Alcian Blue solution. Incubate at room temperature for about 30 min. Remove the dye and wash the cells three times with tap water. 6. Counterstain the cells with nuclear fast red solution for 5 min and wash three times with tap water. A positive result will show strongly acidic sulfated mucopolysaccharides (blue), nuclei (pink to red) and cytoplasm (pale pink). Inspect the cells with a phase-contrast microscope and take images as desired. Store plates at 4  C if dye extraction is required. 3.8 Adipogenic Differentiation (Fig. 2)

1. Trypsinize PDPCs and plate them in 3 mL normal culture medium at a density of about 2.0  105 cells per well or lower in a 12 well plate and allow the culture to settle and attach for 12–24 h in a normoxic 37  C CO2 incubator. 2. Remove media and wash once with a prewarmed 1 PBS, before adding 2 mL prewarmed complete adipogenic medium. Incubate cultures for up to 21 days with media changes every 2–3 days. 3. Minimal lipid accumulation can be seen from the third day, and once lipid accumulation becomes visually apparent from the 10th to 21st days, the Oil Red O solution will stain positive. 4. Remove all culture media from each well and gently wash cells twice with 2 mL 1 PBS and fix the cells with 2 mL of 4% paraformaldehyde fixative solution for about 15 min or more at room temperature (see Note 9). 5. Remove the fixative solution and wash the wells with 60% isopropanol. Let the wells dry completely, then add 2 mL of working stock of Oil Red O solution for about 10–20 min, do not disturb or shake the wells during this period. Then, remove the dye and wash the cells three times with ddH2O. Positive orange-red staining will identify lipid accumulation. Inspect the cells under a phase-contrast microscope and capture images as desired. 6. To elute and measure the optical density (OD), remove all water and let well dry. Elute Oil Red O by adding 100% isopropanol and incubate for about 10 min or more. Pipet

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the isopropanol with Oil Red O several times to ensure that all Oil Red O is in solution, transfer into 1.5 mL Eppendorf tubes. 7. Measure OD at 500 nm, 0.5 s per reading. Use 100% isopropanol as blank and isopropanol from an empty well stained as mentioned above. 3.9

4

Conclusion

From the above three methods of PDPC isolation, the BG method is more time consuming and cost intensive. This is due to the initial recovery surgery and 5-day waiting period post autograft surgery and the surgical materials (suture, needles, spinal needle and anesthetic drugs) that are used. The BC method is straight forward, but a bit cumbersome with the agarose capping. It requires maintaining the agarose solution at an average temperature to avoid solidification. The EG method seems to be the most simple and least expensive. However, in our experience it yields the fewest number of cells. In conclusion, we now favor the BC method for most common use in our laboratory.

Notes 1. BG method requires mice of an older age for the femur to withstand the vibration of a Dremel saw. Using mice younger than 6 weeks old results in breaking and cracking of bones, and the autografts tend to fail. For BC and EG methods, younger mice can be used as it has been reported that the periosteum is thicker with a more robust periosteal progenitor population in young or neonatal mice [3, 9]. 2. The instruments for bone and tissue harvest are generally the same, except for the BG method that requires the cutter, needles, sutures, and anesthetic drug/materials. For cell isolation and tissue harvest reagents, both BG and BC require collagenase, while BC also requires agarose and TE buffer for capping. The cell egression isolation method requires no tissue digestion reagents. 3. For BG method, recovery of the mouse after surgery is vital. The time for anesthesia to take effect is mouse-specific, but one should observe the mice closely during this time and ensure that the tongue is pulled out to the side of the mouth to allow for adequate respiration. To clean shaved hairs, adhesive tape can be applied to the shaved area to remove remaining hair. Skin is then sterilized with povidone-iodine solution. 4. The Dremel saw should be used at a low speed. Using high speed/higher vibration will likely damage and crush the bone. It may bleed after the first cut, hence using the autoclaved firm paper (preferably white), helps visibility, secures the bone edge,

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and protects the skin from injury during the second bone cut. It is possible to lose the graft as the vibration can flip the 4 mm bone out of sight. Once, the graft is cut, place it in a petri dish with DMEM to prevent drying. 5. Cut the protruding edge of the spinal needle and bend the remnant as close as possible to the skin. This prevents protrusion after suture and limits suture dehiscence and infection. 6. The isolation of PDPCs varies with different lots of collagenase. It is advised to test different collagenase lots for isolation efficiency, including the cell yield. Care should be taken during washing since the formed callus may detach and be lost. Extra DMEM could be used to rinse the area for maximum collection of callus. During digestion, examine the tube at intervals to ascertain if tissue has been digested, if not, one can extend the digestion. The tube can leak during shaking, hence sealing the cover with parafilm is advised. Capping prevents digestion of cells from the growth plate, bone marrow or articular surface and the media in the culture dish prevents periosteal tissue from drying. 7. Different lots of FBS should also be tested. It is advisable to add the growth factor to the culture medium just prior to use. A sequential addition to culture has been reported to be more effective than reconstitution in media for use at later times [32]. Allowing up to 7 days post culture before washing the cells ensures maximum attachment of cells. For the cell egression method, it is not advisable to prolong tissue culture with explanted cells. 8. Occasionally, an efficient cell harvest/isolation may not yield a population of cells that has continued proliferation. Some cell isolations have a good primary yield at P0, but cells will stagnate/cease to proliferate after passaging to P1. It is advised that one should watch the progress of the isolated cell population through growth kinetics and decide when to terminate a nonproliferating culture to save time and reagents. For a reference of the range of cell yield per method, see Table 1. 9. Cells can be kept in 4% paraformaldehyde for a couple of days at 4  C before staining. Wrap culture plate with parafilm to prevent drying and cover with aluminum foil.

Acknowledgments Help from all the previous and current members of O’Keefe laboratory is duly acknowledged. National Institute of Health if thanked for the funding (R01-AR069605) the research work in our laboratory.

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References 1. Bragdon BC, Bahney CS (2018) Origin of reparative stem cells in fracture healing. Curr Osteoporos Rep 16:490–503 2. Chang H, Knothe Tate ML (2012) Concise review: the periosteum: tapping into a reservoir of clinically useful progenitor cells. Stem Cells Transl Med 1:480–491 3. Dwek JR (2010) The periosteum: what is it, where is it, and what mimics it in its absence? Skelet Radiol 39:319–323 4. Allen MR, Hock JM, Burr DB (2004) Periosteum: biology, regulation, and response to osteoporosis therapies. Bone 35:1003–1012 5. Squier CA, Ghoneim S, Kremenak CR (1990) Ultrastructure of the periosteum from membrane bone. J Anat 171:233–239 6. Debnath S, Yallowitz AR, McCormick J, Lalani S, Zhang T, Xu R, Li N, Liu Y, Yang YS, Eiseman M, Shim JH, Hameed M, Healey JH, Bostrom MP, Landau DA, Greenblatt MB (2018) Discovery of a periosteal stem cell mediating intramembranous bone formation. Nature 562:133–139 7. Duchamp de Lageneste O, Julien A, AbouKhalil R, Frangi G, Carvalho C, Cagnard N, Cordier C, Conway SJ, Colnot C (2018) Periosteum contains skeletal stem cells with high bone regenerative potential controlled by Periostin. Nat Commun 9:773 8. Zhang X, Xie C, Lin AS, Ito H, Awad H, Lieberman JR, Rubery PT, Schwarz EM, O’Keefe RJ, Guldberg RE (2005) Periosteal progenitor cell fate in segmental cortical bone graft transplantations: implications for functional tissue engineering. J Bone Miner Res 20:2124–2137 9. Ferretti C, Mattioli-Belmonte M (2014) Periosteum derived stem cells for regenerative medicine proposals: boosting current knowledge. World J Stem Cells 6:266–277 10. Yukata K, Xie C, Li TF, Takahata M, Hoak D, Kondabolu S, Zhang X, Awad HA, Schwarz EM, Beck CA, Jonason JH, O’Keefe RJ (2014) Aging periosteal progenitor cells have reduced regenerative responsiveness to bone injury and to the anabolic actions of PTH 1-34 treatment. Bone 62:79–89 11. Feigenson M, Jonason JH, Shen J, Loiselle AE, Awad HA, O’Keefe RJ (2020) Inhibition of the prostaglandin EP-1 receptor in periosteum progenitor cells enhances osteoblast differentiation and fracture repair. Ann Biomed Eng 48 (3):927–939 12. Huang C, Tang M, Yehling E, Zhang X (2014) Overexpressing sonic hedgehog peptide restores periosteal bone formation in a murine

bone allograft transplantation model. Mol Ther 22:430–439 13. Wang Q, Huang C, Xue M, Zhang X (2011) Expression of endogenous BMP-2 in periosteal progenitor cells is essential for bone healing. Bone 48:524–532 14. Wang T, Zhang X, Bikle DD (2017) Osteogenic differentiation of periosteal cells during fracture healing. J Cell Physiol 232:913–921 15. van Gastel N, Torrekens S, Roberts SJ, Moermans K, Schrooten J, Carmeliet P, Luttun A, Luyten FP, Carmeliet G (2012) Engineering vascularized bone: osteogenic and proangiogenic potential of murine periosteal cells. Stem Cells 30:2460–2471 16. McDuffee LA (2012) Comparison of isolation and expansion techniques for equine osteogenic progenitor cells from periosteal tissue. Can J Vet Res 76:91–98 17. Nakahara H, Bruder SP, Haynesworth SE, Holecek JJ, Baber MA, Goldberg VM, Caplan AI (1990) Bone and cartilage formation in diffusion chambers by subcultured cells derived from the periosteum. Bone 11:181–188 18. Antonucci I, Iezzi I, Morizio E, Mastrangelo F, Pantalone A, Mattioli-Belmonte M, Gigante A, Salini V, Calabrese G, Tete S, Palka G, Stuppia L (2009) Isolation of osteogenic progenitors from human amniotic fluid using a single step culture protocol. BMC Biotechnol 9:9 19. Buzzi M, Alviano F, Campioni D, Stignani M, Melchiorri L, Rotola A, Tazzari P, Ricci F, Vaselli C, Terzi A, Pagliaro PP, Cuneo A, Lanza F, Bontadini A, Baricordi OR, Rizzo R (2012) Umbilical cord blood CD34(+)cellderived progeny produces human leukocyte antigen-G molecules with immuno-modulatory functions. Hum Immunol 73:150–155 20. Grau-Vorster M, Rodriguez L, Del MazoBarbara A, Mirabel C, Blanco M, Codinach M, Gomez SG, Querol S, GarciaLopez J, Vives J (2019) Compliance with good manufacturing practice in the assessment of immunomodulation potential of clinical grade multipotent mesenchymal stromal cells derived from Wharton’s jelly. Cell 8(5):484 21. Ude CC, Ng MH, Chen CH, Htwe O, Amaramalar NS, Hassan S, Djordjevic I, Rani RA, Ahmad J, Yahya NM, Saim AB, Idrus RB (2015) Improved functional assessment of osteoarthritic knee joint after chondrogenically induced cell treatment. Osteoarthr Cartil 23:1294–1306

Method of Isolation and Culture of Mouse Periosteum Cells 22. Lu W, Gao B, Fan J, Cheng P, Hu Y, Jie Q, Luo Z, Yang L (2019) Mesenchymal progenitors derived from different locations in long bones display diverse characteristics. Stem Cells Int 2019:5037578 23. Ochareon P, Herring SW (2007) Growing the mandible: role of the periosteum and its cells. Anat Rec (Hoboken) 290:1366–1376 24. Zhang X, Awad HA, O’Keefe RJ, Guldberg RE, Schwarz EM (2008) A perspective: engineering periosteum for structural bone graft healing. Clin Orthop Relat Res 466:1777–1787 25. Evans SF, Chang H, Knothe Tate ML (2013) Elucidating multiscale periosteal mechanobiology: a key to unlocking the smart properties and regenerative capacity of the periosteum? Tissue Eng Part B Rev 19:147–159 26. Ferretti C, Borsari V, Falconi M, Gigante A, Lazzarini R, Fini M, Di Primio R, MattioliBelmonte M (2012) Human periosteumderived stem cells for tissue engineering applications: the role of VEGF. Stem Cell Rev Rep 8:882–890 27. Chan CKF, Gulati GS, Sinha R, Tompkins JV, Lopez M, Carter AC, Ransom RC, Reinisch A, Wearda T, Murphy M, Brewer RE, Koepke LS, Marecic O, Manjunath A, Seo EY, Leavitt T, Lu WJ, Nguyen A, Conley SD, Salhotra A, Ambrosi TH, Borrelli MR, Siebel T, Chan K,

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Schallmoser K, Seita J, Sahoo D, Goodnough H, Bishop J, Gardner M, Majeti R, Wan DC, Goodman S, Weissman IL, Chang HY, Longaker MT (2018) Identification of the human skeletal stem cell. Cell 175:43–56 e21 28. Ambrosi TH, Longaker MT, Chan CKF (2019) A revised perspective of skeletal stem cell biology. Front Cell Dev Biol 7:189 29. Kim YK, Nakata H, Yamamoto M, Miyasaka M, Kasugai S, Kuroda S (2016) Osteogenic potential of mouse periosteum-derived cells sorted for CD90 in vitro and in vivo. Stem Cells Transl Med 5:227–234 30. Umrath F, Steinle H, Weber M, Wendel HP, Reinert S, Alexander D, Avci-Adali M (2019) Generation of iPSCs from jaw periosteal cells using self-replicating RNA. Int J Mol Sci 20:1648 31. Umrath F, Thomalla C, Poschel S, SchenkeLayland K, Reinert S, Alexander D (2018) Comparative study of MSCA-1 and CD146 isolated periosteal cell subpopulations. Cell Physiol Biochem 51:1193–1206 32. Ude CC, Seet WT, Sharen Aini S, Aminuddin BS, Ruszymah BHI (2018) Shelf life evaluation of clinical grade Chondrogenic induced aged adult stem cells for cartilage regeneration. Sci Rep 8:4345

Chapter 25 Isolation and Culture of Murine Primary Chondrocytes: Costal and Growth Plate Cartilage Yihan Liao, Jason T. Long, Christopher J. R. Gallo, Anthony J. Mirando, and Matthew J. Hilton Abstract Primary chondrocyte isolation and culture is a useful tool to characterize how cellular perturbations impact chondrocyte behavior and mineralization in vitro. This protocol conveys methods for isolating and culturing primary chondrocytes from costal and growth plate cartilage. Following gross dissection of the neonatal murine anterior rib cage or long bone growth plate cartilage, chondrocytes are isolated via enzymatic digestion and plated at high density. Genetic perturbation of plated primary murine chondrocytes using viral infection of Cre recombinase to excise floxed alleles and/or overexpress genes of interest are also described. Key words Cartilage, Chondrocyte, Costal, Growth plate, Cell culture, Mouse

1

Introduction Primary chondrocyte isolation and culture is an important part of in vitro studies in cartilage biology, since the isolated cells can be used to characterize important in vivo cellular properties and/or functions including: proliferation, differentiation, metabolism, migration, and matrix production. Although there are several mesenchymal cell lines regarded as chondrogenic, such as ATDC5 [1], C3H10T1/2 [2, 3], and RCJ3.1C5.18 cells [4], these lines need to be cultured in specific conditions to induce and maintain chondrogenic properties. For instance, ATDC5 requires the presence of insulin [1, 5, 6] while C3H10T1/2 necessitates BMP-2 or TGF-β1 [7]. Primary chondrocytes better maintain chondrogenic properties and are especially useful when transgenic and floxed mouse lines are available. Following the addition of treatment media or Cre adenoviruses to recombine floxed alleles, isolated murine chondrocytes can be used to directly examine how certain molecules or perturbations in signaling influence each of the

Matthew J. Hilton (ed.), Skeletal Development and Repair: Methods and Protocols, Methods in Molecular Biology, vol. 2230, https://doi.org/10.1007/978-1-0716-1028-2_25, © Springer Science+Business Media, LLC, part of Springer Nature 2021

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aforementioned cellular processes. Evaluating gene or protein expression at particular time-points aids our understanding chondrocyte behavior at the molecular level. Chondrocytes produce and maintain the cartilage extracellular matrix (ECM) by expressing Sry-box 9 (Sox9), collagen type II (Col2a1), and Aggrecan (Acan). During endochondral ossification, mesenchymal progenitor cells differentiate into chondrocytes, which then undergo the process of proliferation, exit of cell cycle, hypertrophy (expressing collagen type X, Col10a1), late stage hypertrophy (expressing matrix metalloproteinase 13, Mmp13), and apoptosis or a cell fate change [8–12]. Endochondral ossification is defined by having an intermediate cartilage scaffold that precedes bone formation [13]. Regions of cartilage remain to allow for continued bone growth—termed growth plate cartilage—and to allow for proper movement and articulation—termed articular and costal cartilages. These arise early in development. While the growth plate persists in mice, it is completely replaced by bone in humans [14]. Sternal, or costal, cartilage forms to create articulation between the ribs and sternum and between adjacent sternal segments prior to fusion [15]. Following fusion, costal cartilage is then restricted to a thin layer between the ribs and sternum. As each type of cartilage serves a separate purpose, the differences between them can be seen at the molecular level and thus need to be considered when studying development, homeostasis, and repair. For instance, costal chondrocytes likely have a distinct profile from growth plate chondrocytes, given that growth plate cartilage is weight-bearing while costal cartilage is minimally loaded. Thus, based on the aim of an experiment, the utility of different types of primary chondrocytes should be considered. Many studies have utilized primary chondrocytes from different animal models in vitro. Here, we describe a protocol for isolation and culture of murine primary chondrocytes from costal cartilage and growth plate cartilage.

2

Materials

2.1 Primary Chondrocyte Isolation

1. 70% ethanol. 2. Absorbent bench pad. 3. (2) #5 Forceps or the equivalent. 4. One curved end forceps (5/45). 5. One pair of surgical scissors. 6. #10 scalpel or equivalent. 7. Class II biosafety cabinet. 8. Cold sterile 1
phosphate-buffered saline (PBS), pH 7.4.

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9. (2) 0.2 μm filters. 10. (2) 20–30 mL syringes. 11. Penicillin and streptomycin (P/S). 12. Dulbecco’s modified Eagle’s medium (DMEM). 13. 3 mg/mL collagenase D solution: Dissolve 90 g of collagenase D (Roche) in 30 mL of DMEM with 1% penicillin–streptomycin (P/S) and then filter-sterilize through a 0.2 μm filter. 14. 2 mg/mL Pronase solution: Dissolve 30 mg of Pronase (Roche) in 15 mL of 1
PBS and then filter-sterilize through a 0.2 μm filter (see Note 1). 15. Sterile 50 mL polypropylene centrifuge tubes. 16. Sterile 10 cm petri dish. 17. 25 mL pipettes. 18. Centrifuge. 19. 37  C shaking water bath. 20. Sterile 45 μm cell strainers. 21. P2-P5 neonatal pups for growth plate/costal chondrocyte isolation (see Note 2). 2.2 Plating and Culturing of Primary Murine Chondrocytes

1. Sterile 50 mL polypropylene centrifuge tubes. 2. Hemocytometer or automated cell counter. 3. Complete culture media: DMEM, 10% fetal bovine serum, 1% P/S. 4. Tissue culture-treated polystyrene multiwell plates (12-well or 24-well).

2.3 Viral Infection of Primary Murine Costal Chondrocyte Cultures (Optional)

3

1. 10 mg/mL Polybrene stock solution (see Note 3). 2. Adenoviral particles (see Note 4). 3. Complete culture media (defined above in Subheading 2.2) minus antibiotics.

Methods

3.1 Primary Murine Costal Chondrocyte Isolation

All steps should be performed using aseptic technique in a Class II biosafety cabinet. 1. Sacrifice P2-P5 neonatal pups by decapitation or according to IACUC-approved protocols (see Note 2). 2. Sterilize torsos with 70% ethanol. 3. To isolate rib cages and sterna, make an excision just below the rib cage and another excision at the base of the neck to obtain a tubular structure. Make a parallel incision along the spinal

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Fig. 1 (a) Isolated rib cage with sternum after cutting along both sides of the spinal column. The rib cage was flattened as displayed here. (b) Isolation of growth plate chondrocytes in early postnatal mice, with a dissection just above the visible marrow cavity

column to flatten the sternum and ribs. Remove all skin and organs. Use the edges of two forceps to gently scrape as much soft tissue away from the sternum/ribs as possible over an absorbent bench pad moistened with 70% ethanol (see Note 5). Place the isolated sternum and ribs into a 50-mL conical tube with 1
PBS on ice. Repeat the isolation procedure for each neonatal pup (see Fig. 1a). 4. Wash rib cage pieces once with 1
PBS. 5. Digest the rib cages and sternal tissue in 15 mL of 2 mg/mL Pronase solution at 37  C for 1 h with constant agitation in a shaking water bath (see Note 6). 6. Wash the tissue thoroughly three times with 1
PBS, aspirating the solution between washes. 7. Digest the tissue in 15 mL of 3 mg/mL of collagenase D solution in a 50-mL conical tube laying horizontally in a 37  C humidified cell culture chamber for 1 h (place tube on a 10-cm petri dish to capture any leaking solution). Agitate the tissue every 30 min to ensure adequate digestion (see Note 7). 8. Wash the tissue three times with 1
PBS, aspirating the solution between washes. 9. Add 15 mL of fresh collagenase D solution and transfer the tissue to a 10-cm petri dish. Incubate the samples at 37  C in a humidified cell culture chamber for 4–6 h. After 2 h, pipette the solution up and down a few times with a 25-mL pipette to disaggregate cells and tissue clumps. Repeat the process every 2 h (see Note 8). 10. Filter the cell suspension with a 45 μm cell strainer into a new 50 mL conical tube. 11. Centrifuge the cell suspension at 400
g at 4  C for 5 min. 12. Plate and culture cells (see Subheading 3.3).

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1. Sacrifice P2-P5 neonatal pups by decapitation or according to IACUC-approved protocols (see Note 2). 2. Sterilize torsos with 70% ethanol. 3. To isolate the tibiae and femurs, cut each hind limb from the body just proximal to the femur head. Using a pair of scissors, additionally transect the distal end of the limb to aid in removal of skin. Remove as much soft tissue as possible with two forceps. Disarticulate the joint to clean the remaining soft tissue. Identify the edge between the primary ossification center and the growth plate (see Fig. 1b). Using forceps to stabilize the bone, make a cut at the chondro-osseous junction using #10 scalpel. Transfer the cartilage piece to 1
PBS on ice. Repeat the isolation procedure for each hind limb element. 4. Wash growth plate cartilage pieces once with 1
PBS. 5. Digest growth plate cartilage in 15 mL of 2 mg/mL Pronase solution at 37  C for 1 h with constant agitation in a shaking water bath (see Note 6). 6. Wash the growth plate cartilage thoroughly three times with 1
PBS, aspirating the solution between washes. 7. Digest the growth plate cartilage in 15 mL of 3 mg/mL of collagenase D solution in a 50-mL conical tube laying horizontally in a 37  C humidified cell culture chamber for 1 h (place tube on a 10-cm petri dish to capture any leaking solution). Agitate the tissue every 30 min to ensure adequate digestion (see Note 7). 8. Wash the digested growth plate cartilage three times with 1
PBS, aspirating the solution between washes. 9. Add 15 mL of fresh collagenase D solution and transfer digested tissue to a 10-cm petri dish. Incubate the samples at 37  C in a humidified cell culture chamber for 4–6 h. After 2 h, pipette the solution up and down a few times with a 25-mL pipette to disaggregate cells and tissue clumps. Repeat the process every 2 h (see Note 8). 10. Filter the cell suspension with a 45-μm cell strainer into a new 50-mL conical tube. 11. Centrifuge the cell suspension at 400
g at 4  C for 5 min. 12. Plate and culture cells (see Subheading 3.3).

3.3 Plating and Culturing Primary Murine Costal/Growth Plate Chondrocytes

1. Resuspend the cell pellet in 3–5 mL of complete culture media. Count the number of cells in the cell suspension using a hemocytometer or an automated cell counter. 2. Add additional media to bring the suspension to the appropriate cell density. Plate the cell suspension into 12-well tissue culture plates at 5
105 cell/well density. Place the plates in a

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Fig. 2 Bright-field microscopy images show plated costal and growth plate chondrocytes grown 1 and 3 days after seeding at a density of 500,000 cells/ well

Fig. 3 qPCR comparison of genes in plated costal and growth plate chondrocytes grown for 3 days. Chondrogenic genes include Sox9, Col2a1, Acan, and Col10a1. Note the difference in expression of Col10a1 in each population of primary chondrocytes

humidified cell culture incubator at 37  C with 5% CO2 to allow the chondrocytes to adhere to the plate (see Note 9). 3. Culture the chondrocytes to confluence in complete culture media (see Note 10). Change the media every 2 days up to desired time points. Figure 2 demonstrates the physical appearance of cultured murine costal and growth plate chondrocytes at 3 days in culture, and Fig. 3 shows qPCR quantification of standard chondrogenic genes for each sample type after 3 days in culture. 3.4 Viral Infection of Primary Murine Chondrocyte Cultures (Optional)

1. Culture the chondrocytes to roughly 70–80% confluence. 2. Prior to infection, determine the amount of adenovirus that will be needed for the desired multiplicity of infection (MOI) as follows: (a) (Number of cells in culture)
(MOI) ¼ Total plaqueforming units (pfu). (b) (Total pfu)/(virus titer in pfu/mL) ¼ mL needed for infection at desired MOI.

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3. Remove culture media. Next, add fresh media without antibiotics at half the volume normally used for the particular culture vessel. Add an appropriate amount of polybrene to the media such that the final concentration is between 5 and 10 μg/mL. Gently swirl the media to mix. Add the adenovirus directly to the media and, again, swirl to mix. Alternatively, if multiple wells/plates are to be infected with the same virus, it may be preferential to first dilute the polybrene and virus in the total amount of media needed and, subsequently, aliquot the appropriate amount per well/plate. 4. Incubate chondrocyte cultures at 37  C with 5% CO2 in an incubator approved for Biosafety Level 2 agents. Remove the virus and add complete culture media 24–48 h postinfection. If waiting for 48 h, it is best to add an additional volume of complete culture media to the culture vessel at 24 h. 5. Continue to culture the chondrocytes at 37  C with 5% CO2 until desired endpoints (see Note 10). RNA and/or protein can be isolated at each time point in order to monitor changes in gene or protein expression for factors important in regulating chondrocyte phenotype or differentiation.

4

Notes 1. To ensure Pronase is fully dissolved, prepare solution prior to harvest of tissue, mix well with constant agitation in a shaking 37  C water bath, and filter-sterilize. 2. This protocol is based on the usage of eight animals, which typically yield around 5 M cells. Solutions can be scaled to adjust for number of mice. Since the secondary ossification center forms and becomes vascularized around postnatal day 7 [8], 2- to 5-day-old neonatal pups are ideal for pure chondrocyte isolation from the ribs and limbs. 3. The addition of polybrene to the culture media for adenovirus infection is documented to enhance transduction efficiency. Polybrene is a cationic polymer thought to neutralize the charge of the cell membrane, thereby reducing repulsive forces between the virus and target cell surface. Polybrene can be purchased from vendors as a ready-to-use solution or powder to be diluted to the desired stock concentration in sterile nuclease-free water. 4. We regularly isolate primary costal chondrocytes from floxed mice and use adenovirus expressing Cre recombinase to delete a segment of DNA flanked by loxP sites in vitro. We purchase ready-to-use Ad5-CMV-Cre-GFP or Ad5-CMV-Cre and control Ad5-CMV-GFP adenoviral particles from Vector

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Development Laboratory (Baylor College of Medicine, Houston, TX). We have also had success with viruses purchased from Vector Biolabs (Philadelphia, PA). Vector Biolabs has many premade adenoviruses convenient for use in overexpression studies. 5. Using an adsorbent bench pad moistened with 70% ethanol will help reduce potential contamination of cells in culture. Complete growth media for incubation includes penicillin and streptomycin to reduce the risk of contamination. 6. Constant agitation of Pronase solution aids in removal of soft tissue. The solution will likely become cloudy following the 1-h incubation. 7. The first collagenase digestion is used to remove remaining soft tissue. It is advised to not have constant agitation for this step as it may disrupt and cause loss of some chondrocytes. 8. The second collagenase step dissociates the chondrocytes from their matrix. Use of a petri dish allows for any remaining fibroblasts to adhere to the plate, as chondrocytes do not readily attach in the time frame described. In as little as 4 h, cells can be recovered. However, it appears that maximal cell count occurs at approximately 6 h of digestion—a time frame that does not lead to cell stress or death. 9. In plating both costal and growth plate chondrocytes, 12-well plates are customarily used. The specific plating density can range between 200,000 cells and 700,000. However, we have observed that plating 500,000 cells allows for confluence to be obtained without alterations in cell morphology or fate changes. Higher density cultures may be needed for experiments such as adenoviral infections. Adjust accordingly. 10. Depending on the duration of culture, complete culture media may need to be substituted for chondrocyte maturation media (Complete culture media with 50 μg/mL ascorbic acid and 10 mM β-glycerophosphate). Chondrocyte maturation media will prevent chondrocytes from undergoing dedifferentiation. The addition of ascorbic acid to media is essential for chondrocyte maturation and subsequent cartilage ECM mineralization. Additionally, the cartilage matrix will not mineralize properly until provided a phosphate source, such as β-glycerophosphate, during extended culture periods.

Acknowledgments This work was supported by NIH/NIAMS grants: R01 grants (AR063071 and AR071722) to M.J.H.

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References 1. Atsumi T, Ikawa Y, Miwa Y, Kimata K (1990) A chondrogenic cell line derived from a differentiating culture of AT805 teratocarcinoma cells. Cell Diff Dev 30:109–116 2. Denker AE, Haas AR, Nicoll SB, Tuan RS (1999) Chondrogenic differentiation of murine C3H10T1/2 multipotential mesenchymal cells: I. Stimulation by bone morphogenetic protein-2 in high-density micromass cultures. Differentiation 64:67–76 3. Ahrens M et al (1993) Expression of human bone morphogenetic proteins-2 or -4 in murine mesenchymal progenitor C3H10T1/ 2 cells induces differentiation into distinct mesenchymal cell lineages. DNA Cell Biol 12:871–880 4. Chang W et al (1999) Calcium sensing in cultured chondrogenic RCJ3.1C5.18 cells. Endocrinology 140:1911–1919 5. Yao Y, Zhai Z, Wang Y (2014) Evaluation of insulin medium or chondrogenic medium on proliferation and chondrogenesis of ATDC5 cells. Biomed Res Int 2014:569241 6. Shukunami C et al (1997) Cellular hypertrophy and calcification of embryonal carcinomaderived chondrogenic cell line ATDC5 in vitro. J Bone Miner Res 12:1174–1188 7. Haas AR, Tuan RS (1999) Chondrogenic differentiation of murine C3H10T1/2 multipotential mesenchymal cells: II. Stimulation by bone morphogenetic protein-2 requires modulation of N-cadherin expression and function. Differentiation 64:77–89

8. Kozhemyakina E, Lassar AB, Zelzer E (2015) A pathway to bone: signaling molecules and transcription factors involved in chondrocyte development and maturation. Development 142:817–831 9. Liu C-F, Samsa WE, Zhou G, Lefebvre V (2017) Transcriptional control of chondrocyte specification and differentiation. Semin Cell Dev Biol 62:34–49 10. Yang G et al (2014) Osteogenic fate of hypertrophic chondrocytes. Cell Res 24:1266–1269 11. Yang L, Tsang KY, Tang HC, Chan D, Cheah KS (2014) Hypertrophic chondrocytes can become osteoblasts and osteocytes in endochondral bone formation. Proc Natl Acad Sci U S A 111:12097–12102 12. Zhou X et al (2014) Chondrocytes transdifferentiate into osteoblasts in endochondral bone during development, postnatal growth and fracture healing in mice. PLoS Genet 10: e1004820 13. Mackie EJ, Ahmed YA, Tatarczuch L, Chen KS, Mirams M (2008) Endochondral ossification: how cartilage is converted into bone in the developing skeleton. Int J Biochem Cell Biol 40:46–62 14. Nilsson O, Baron J (2005) Impact of growth plate senescence on catch-up growth and epiphyseal fusion. Pediatr Nephrol 20:319–322 15. Liakhovitskaia A et al (2010) The essential requirement for Runx1 in the development of the sternum. Dev Biol 340:539–546

Chapter 26 Isolation and Culture of Neonatal Mouse Calvarial Osteoblasts Madison L. Doolittle, Cheryl L. Ackert-Bicknell, and Jennifer H. Jonason Abstract This chapter describes the isolation and culture of neonatal mouse calvarial osteoblasts. This primary cell population is obtained by sequential enzymatic digestion of the calvarial bone matrix and is capable of differentiating in vitro into mature osteoblasts that deposit a collagen extracellular matrix and form mineralized bone nodules. Maturation of the cultures can be monitored by gene expression analyses and staining for the presence of alkaline phosphatase or matrix mineralization. This culture system, therefore, provides a powerful model in which to test how various experimental conditions, such as the manipulation of gene expression, may affect osteoblast maturation and/or function. Key words Mouse, Calvaria, Osteoblast, Bone, Maturation, Mineralization

1

Introduction Osteoblasts are the cells responsible for new bone formation during skeletal development, remodeling and repair. They arise from multipotent mesenchymal progenitor cells through a process governed by both systemic and local growth factors. These factors modulate signaling pathways that lead to the activation of transcription factors critical for the expression of genes required for differentiation into and throughout the osteoblast lineage (for review, [1–3]). Once committed to the lineage, the osteoblast phenotype is defined by the sequential expression of genes involved in proliferation, extracellular matrix (ECM) maturation, and mineralization [4– 6]. During the proliferative stage, immature osteoblasts begin to express and secrete type I collagen. This is the major protein product of the osteoblast and the primary building block of the organic bone matrix. As cells mature and exit the cell cycle, however, they begin to express other ECM molecules including various glycoproteins, proteoglycans, and γ-carboxylated (or gla) proteins that are involved in the subsequent maturation and mineralization of the matrix. Several of these proteins are considered established

Matthew J. Hilton (ed.), Skeletal Development and Repair: Methods and Protocols, Methods in Molecular Biology, vol. 2230, https://doi.org/10.1007/978-1-0716-1028-2_26, © Springer Science+Business Media, LLC, part of Springer Nature 2021

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“markers” of osteoblast maturation and are thought to regulate the ordered deposition and turnover of hydroxyapatite crystals within the organic bone matrix. Alkaline phosphatase (encoded by mouse Alpl), for example, is the most abundant glycoprotein in the ECM and is expressed by osteoblasts immediately upon exit from the cell cycle [6]. It is a positive regulator of bone mineral deposition and deficiency leads to hypophosphatasia in both humans and mice [7]. Other maturation “markers” include bone sialoprotein (encoded by mouse Ibsp), osteopontin (encoded by mouse Spp1), and osteocalcin (encoded by mouse Bglap). Bone sialoprotein and osteopontin are both phosphoproteins and members of the small, integrin-binding ligand, N-linked glycoprotein (SIBLING) family of proteins. Osteopontin is a potent inhibitor of bone matrix mineralization, while bone sialoprotein is a confirmed nucleator of hydroxyapatite crystals making it a positive regulator of mineralization [8–10]. Osteocalcin, a gla protein, is secreted solely by mature osteoblasts and, when carboxylated, binds strongly to hydroxyapatite crystals in the mineralized matrix [11]. Rather than playing a direct role in matrix mineralization, however, it was shown that osteocalcin is released from the bone matrix during resorption and plays an active endocrine role in regulation of glucose metabolism [12–15]. Mature osteoblasts, therefore, regulate not only bone formation but also other physiological processes by secreting proteins with endocrine properties. To date, much progress has been made in defining the molecular and cellular properties of the osteoblast phenotype through characterization of primary calvarial-derived osteoblast cultures from chicken, rat, and mouse. Primary bone cells were first successfully isolated from the frontal and parietal bones of fetal and neonatal rat calvaria by Peck et al. in 1964 [16]. Viable, alkaline phosphatase-expressing cells were released from the bone matrix by collagenase digestion; however, culture conditions did not allow for the formation of bone nodules and did not prevent the overgrowth of fibroblasts. Wong and Cohn later modified the procedure to isolate sequential populations of cells from mouse calvaria via short successive incubations with collagenase [17]. This method allows for the enrichment of cells with an osteoblastic phenotype from the third, fourth and fifth populations [18, 19]. These cells produce a type I collagen matrix, express alkaline phosphatase, and generate mineralized bone nodules containing hydroxyapatite crystals when cultured with ascorbic acid and β-glycerophosphate [20– 22]. This is still the standard isolation procedure used by most groups today and the one described in this chapter. Because these cells are able to proliferate and synthesize a mature mineralized collagen matrix in vitro, they make an appealing experimental model system for testing how a given factor, gene product or small molecule inhibitor, for example, may affect this process. While some cell lines, especially MC3T3 E1, also provide a

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good model for the study of osteoblast maturation, it is, at times, beneficial to use primary cells isolated from genetically altered mice [23]. Cells from floxed mice, for example, can be infected with virus encoding Cre recombinase to delete, in vitro, a portion of DNA flanked by loxP sites. The same cells, infected with a control virus, offer a “wild type” control for comparison in downstream applications such as the maturation assay. Over the course of maturation in vitro, typically 21–28 days, the cells will express maturation “marker” genes such as those described above and begin to build a mineralized bone matrix. The efficiency of this process can be monitored and compared among experimental groups and conclusions can be drawn with regard to how specific factors or growth conditions affect this process. It should be kept in mind, however, that this is a heterogeneous cell population comprised of osteoblasts that were likely at different stages of the maturation process upon isolation. Additionally, there are likely to be some contaminating fibroblasts or periosteal progenitor cells present in the population. Measurements of gene expression, therefore, should be interpreted as averages of expression from all cells in the population. Regardless of this heterogeneity, when cultured in appropriate conditions, the cultures as a whole are very efficient at synthesizing a mineralized bone matrix as a result of enhanced expression and secretion of osteoblastic ECM marker genes making them a suitable in vitro model of bone formation. It should also be noted that while the genes described in this chapter are those most widely accepted as “markers” of osteoblast maturation, a number of genomic screening studies have identified others [24–28]. In this chapter, we will not only describe the procedure for isolation of primary calvarial osteoblasts from neonatal mice, but will also describe the culture conditions necessary for inducing the maturation of these cells in vitro with detailed staining protocols for detection of alkaline phosphatase and mineralization of the bone matrix (Fig. 1). A brief description of the procedure for adenoviral transduction of these cells is also provided.

2

Materials

2.1 Isolation of Osteoblasts from Neonatal Mouse Calvaria

1. Neonatal mice from postnatal age (P) 2–5 days (see Note 1). 2. Dissection scissors, fine forceps (Dumont #5), and standard forceps with blunt, serrated ends. These should be cleaned well with 70% ethanol prior to use. 3. Sterile 10 cm petri dishes. 4. Sterile cell scraper. 5. Sterile specimen cup (120 ml capacity) with screw cap. Alternatively, a sterile 50 ml polypropylene centrifuge tube could be used.

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Fig. 1 In vitro maturation of mouse primary calvarial-derived osteoblasts. Osteoblasts isolated from the calvaria of P4 mice were seeded at 10,500 cells/cm2 in 6-well plates, grown to confluence and, subsequently, cultured in osteogenic media until harvest on the indicated days. Cells were then stained for the presence of alkaline phosphatase (right) or matrix mineralization (left) (a). Gene expression analyses were also performed via reverse transcriptase quantitative PCR for the indicated genes (b)

6. Sterile 70 μm cell strainers. 7. Sterile 50 ml polypropylene centrifuge tubes. 8. 75 cm2 cell culture flask, tissue-culture treated with vented cap. 9. Sterile 1
phosphate-buffered saline (PBS), pH 7.4. 10. Collagenase A solution: Dissolve Collagenase A (Roche) to a final concentration of 1 mg/ml in Opti-MEM Reduced Serum Medium (Gibco) or MEM α supplemented with 100 U/ml penicillin and 100 μg/ml streptomycin. Make fresh and filtersterilize (see Note 2). 11. Complete culture media: MEM α, no ascorbic acid (Gibco) supplemented with 10% FBS (do not heat-inactivate), 100 U/ ml penicillin, and 100 μg/ml streptomycin.

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2.2 Mouse Calvarial Osteoblast Culture and Maturation Assays

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1. 0.05% trypsin–EDTA solution. 2. Tissue-culture treated polystyrene multiwell plates (12-well and 6-well) (see Note 3). 3. Osteogenic media: Complete culture media with 50 μg/ml ascorbic acid and 10 mM β-glycerophosphate (see Note 4). Filter-sterilize. 4. 1
phosphate-buffered saline (PBS), pH 7.4. 5. 4% paraformaldehyde in 1XPBS. 6. Double distilled water (ddH2O). 7. 1-Step NBT/BCIP solution (Pierce Thermo Scientific). 8. Aluminum foil. 9. Alizarin Red S solution: 1% Alizarin Red S in ddH2O, pH 4.1–4.3 using 10% Ammonium Hydroxide, store at room temperature. 10. 5% Perchloric acid (diluted from 70% perchloric acid with dH2O—use caution as it is a strong acid) (Sigma).

2.3 Adenoviral Transduction of Mouse Calvarial Osteoblasts

1. 10 mg/ml Polybrene stock solution (see Note 5). 2. Adenoviral particles (see Note 6). 3. Complete culture media (defined above in Subheading 2.1) minus antibiotics. 4. Complete culture media.

3

Methods

3.1 Isolation of Osteoblasts from Neonatal Mouse Calvaria

1. Euthanize neonatal mice via an approved IACUC method and proceed immediately with the protocol to avoid loss of cell viability. 2. From this point forward, all steps should be carried out in a Class II biological safety cabinet using sterile technique. 3. Douse the head and upper body in 70% ethanol and decapitate using scissors. Place the head in a sterile petri dish. 4. Grasp each head with fine forceps placed ventrally and through the back of the head. Using blunt-ended forceps, peel the skin away from the top of the head toward the nasal bone revealing the calvaria. Cut along the edges of the parietal bones and place in a second petri dish with sterile PBS. 5. Using a small cell scraper, gently remove any loose connective tissue from the calvaria and transfer cleaned calvaria to a third petri dish with sterile PBS. 6. Carefully transfer the calvaria to a sterile specimen cup containing 10 ml of the 1 mg/ml Collagenase A solution, cap the cup

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tightly, and place in a 37 70–80 rpm for 20 min.



C shaking water bath set at

7. Remove the specimen cup from the water bath and spray liberally with 70% ethanol before reentering the biological safety cabinet. Discard the cells liberated during the first digest by aspirating the Collagenase A solution. Care should be taken to avoid touching the calvaria. 8. Add another 10 ml of the 1 mg/ml Collagenase A solution, cap the cup tightly, and place in the 37  C shaking water bath for another 20 min. 9. Repeat steps 7 through 8, only this time incubate for 30 min in the 37  C shaking water bath. 10. Remove the specimen cup from the water bath and spray liberally with 70% ethanol before reentering the biological safety cabinet. Collect the Collagenase A solution (digest 3) with a sterile 10 ml pipette and dispense over a 70 μm cell strainer positioned over a 50 ml conical tube. Pellet the cells by centrifugation. Remove the supernatant by aspiration and resuspend the cells in 15 ml of complete culture media. Plate all of the cells in a 75 cm2 cell culture flask with vented cap and culture at 37  C with 5% CO2. 11. Add another 10 ml of the 1 mg/ml Collagenase A solution to the specimen cup, cap the cup tightly, and place in the 37  C shaking water bath for another 30 min. 12. Repeat steps 10 and 11 two additional times to collect cells from digests 4 and 5. 13. Culture the cells to 70–80% confluence prior to passaging (see Notes 7 and 8). 3.2 Mouse Calvarial Osteoblast Cultures for Maturation Assays

1. Trypsinize and count the cells. Using complete culture media, plate the cells between 10,000 and 30,000 cells/cm2 in 6- or 12-well plates and grow to confluence (see Note 9). See Subheading 3.5 if adenoviral transduction of the cells is required. 2. Replace the media with osteogenic media and change the media every 2–3 days as needed throughout the maturation assay. 3. Harvest the cells every 2–3 days throughout the length of the maturation assay for isolation of mRNA (see Note 10) or staining as described below. At this point, all procedures can be performed on the benchtop or in a chemical fume hood as sterile conditions are no longer required.

3.3 Alkaline Phosphatase Staining of Mouse Calvarial Osteoblast Cultures

1. Wash cells once in 1X PBS (see Note 11). 2. Fix cells in 4% paraformaldehyde at room temperature for 10–15 min. Perform this step in a chemical fume hood. 3. Wash cells three times in ddH2O.

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4. Add a volume of 1-Step NBT/BCIP solution equivalent to the volume used when culturing the cells. Incubate in the dark (wrapped in foil) at room temperature on a rocking platform for 15–60 min (see Note 12). 5. Remove NBT/BCIP solution and wash cells three times in ddH2O. Allow cells to air dry and store at room temperature away from light. 6. Quantitative Alkaline Phosphatase staining can be performed in separate wells with the SensoLyte pNPP Alkaline Phosphatase Colorimetric Assay Kit per manufacturer’s instructions (AnaSpec). 3.4 Alizarin Red S Staining of Mouse Calvarial Osteoblast Cultures

1. Follow steps 1 and through 3 of Subheading 3.3. 2. Add a volume of Alizarin Red S staining solution equivalent to the volume used when culturing the cells. Incubate at room temperature on a rocking platform for 20 min. 3. Remove Alizarin Red S solution and wash cells three times in 1
PBS. Allow cells to air dry and store at room temperature away from light. 4. Make sure cell plates have fully dried and been scanned to obtain digital images. 5. Add a volume of 5% perchloric acid equivalent to the volume used when culturing the cells and place on a rocking platform for 5 min. This will destain the cells and turn the solution color from red to yellow. 6. Collect the supernatant solution from each cell culture well. 7. Create Alizarin Red standards of 0, 0.0625, 0.125, 0.5, 1, 2, and 4 mM by diluting the 1% (~42 mM) Alizarin Red S solution in 5% Perchloric Acid. 8. Measure absorbance of standards and collected samples at 405 nm in a cell plate reader and plot absorbances against Alizarin Red concentrations to obtain quantified sample concentrations.

3.5 Adenoviral Transduction of Mouse Calvarial Osteoblasts (Optional)

1. Trypsinize and plate cells in a format compatible for the desired downstream application (if carrying out a maturation assay subsequent to infection, plate cells as specified in Subheading 3.2). Culture cells to roughly 70–80% confluence. 2. Prior to infection, determine the amount of adenovirus that will be needed for the desired MOI (multiplicity of infection) (see Note 13) as follows: (a) (number of cells in culture)
(MOI) ¼ total pfu (plaque forming units)

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(b) (total pfu)/(virus titer in pfu/ml) ¼ ml needed for infection at desired MOI 3. Remove culture media and add fresh media without antibiotics at half the volume normally used for the particular culture vessel. Add an appropriate amount of Polybrene to the media such that the final concentration is between 5 and 10 μg/ml. Gently swirl the media to mix. Add the adenovirus directly to the media and, again, swirl to mix. Alternatively, if multiple wells/plates are to be infected with the same virus, it may be preferential to first dilute the Polybrene and virus in the total amount of media needed and, subsequently, aliquot the appropriate amount per well/plate. 4. Incubate cell cultures at 37  C with 5% CO2 in an incubator approved for Biosafety Level 2 agents. Remove virus and add complete culture media 24–48 h post-infection. If waiting 48 h, it is best to add an additional volume of complete culture media to the culture vessel at 24 h. 5. Continue to culture cells at 37  C with 5% CO2 until desired end points.

4

Notes 1. A litter of 6 to 8 mice should yield approximately 4
106 to 5
106 cells after 3–5 days in culture. This is sufficient for setting up a standard maturation assay with staining and isolation of mRNA as end points. 2. Another enzymatic digestion solution commonly used is 0.05% trypsin (0.25% trypsin diluted in DMEM) with 1.5 U/ml collagenase P (Roche). We have not noticed any differences in cell numbers collected following digestion with this solution or the solution containing Collagenase A. 3. We recommend using 12-well plates for staining and 6-well plates for the collection of mRNA or protein during the maturation assay. This plating format can be scaled up, if desired, but scaling down is not recommended as it is very difficult to plate the cells evenly in a culture vessel smaller than one well of a 12-well plate. Additionally, the amount of mRNA obtained from a culture less than that in one well of a 6-well plate may not be sufficient for the desired number of qPCR reactions. We have not noticed a difference among plate manufacturers with regard to their suitability for this assay. 4. The addition of ascorbic acid to the media is essential for osteoblast maturation and bone nodule formation as it promotes the synthesis and secretion of collagen [20, 29, 30]. Ascorbic acid is light sensitive once in solution, so is most effective when made fresh, but can be aliquoted and

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stored at 20  C in the dark. Additionally, nodules will not mineralize without the addition of an organic phosphate, such as β-glycerophosphate. 5. The addition of Polybrene to the culture media along with the addition of virus is documented to enhance adenovirus transduction efficiency. Polybrene is a cationic polymer thought to neutralize the charge of the cell membrane, thereby, reducing repulsive forces between the virus and target cell surface. Polybrene can be purchased from many vendors as a ready-to-use solution or as a powder that should then be diluted in sterile nuclease-free water to the desired stock concentration. 6. We regularly isolate primary osteoblasts from floxed mice and use adenovirus expressing Cre recombinase to delete, in vitro, a segment of DNA flanked by loxP sites. We purchase ready-touse Ad5-CMV-Cre-GFP or Ad5-CMV-Cre and control Ad5-CMV-GFP adenoviral particles from Vector Development Laboratory (Baylor College of Medicine, Houston, TX). We have also had success with virus purchased from Vector Biolabs (Philadelphia, PA). Vector Biolabs has many premade adenoviruses convenient for use in overexpression studies. 7. Culturing cells to this density may require anywhere from 3 to 7 days depending on the initial number of calvaria used for the isolation. During this time, the growth media should be changed every 3 days and the cells should not be grown to confluence as this may cause them to begin to mature. Morphologically, the cells should appear large and polygonal in shape with a single, large nucleus. Keep in mind that this is a heterogeneous population, however, and there may be contaminating fibroblasts or periosteal osteoprogenitors in the cultures. In our experience, though, this does not inhibit the osteogenic maturation of the cells in downstream applications. We have found that cells can be successfully passaged twice following the initial plating. 8. Contamination with bacteria or fungus is always a risk when culturing primary cells. Amphotericin B (Fungizone) can be added to the culture media (between 0.25 and 2.5 μg/ml) to prevent fungus growth. Additionally, cultures can be washed with sterile 1XPBS and replenished with fresh media 24 h after initial plating. This will remove any remaining cellular debris and reduce the risk of contamination. 9. If possible, plating at the higher density is preferred as the cells will have to undergo fewer mitotic divisions and the cultures will reach confluence within 24–72 h. Since primary osteoblasts have limited proliferative capacity, it is important that maturation of the cultures be initiated prior to onset of cellular senescence.

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10. For isolation of mRNA, we typically use the RNeasy Mini kit (Qiagen) and QIAshredder. Alternatively, when it is desired to obtain both protein and mRNA from the same cultures, we use the PARIS kit (Life Technologies) with subsequent concentration and cleanup of mRNA via the RNeasy MinElute Cleanup kit (Qiagen). When using the PARIS kit, however, it is usually necessary to break up the mineralized extracellular matrix that these cultures generate by passing the lysate through a syringe needle several times prior to adding it to the column. 11. Care should be taken to avoid touching the cell cultures during the staining procedure. Cultures at later time points will have a mineralized extracellular matrix that may easily lift from the plate while adding and aspirating liquid from the wells. 12. The length of time for staining can range from 15 to 60 min. Cultures from all time points within an experiment should be incubated with the staining solution for the same amount of time. Keep in mind that the earlier time points will express significantly less alkaline phosphatase and have little, if any, detectable mineral in the matrix. 13. MOI is the number of viral genomes per cell in the given culture. When using a particular virus for the first time, it is recommended to test a range of MOIs (e.g., 0, 50, 100, 200, 500, and 1000). Control and experimental viruses should be used at equivalent MOIs within an experiment. When calculating the volume of a given virus to be used, be sure to use the value corresponding to the titer (pfu/ml) and not the total particles/ml. The virus stock may first need to be diluted 1:10, or even 1:100, in PBS to reach a suitable working stock concentration.

Acknowledgments This work was supported by the following NIH grants: R01 AR064790 (C.L.A) and P30 AR069655 (J.H.J). References 1. Jensen ED, Gopalakrishnan R, Westendorf JJ (2010) Regulation of gene expression in osteoblasts. Biofactors 36(1):25–32; Epub 2010/ 01/21 2. Komori T (2011) Signaling networks in RUNX2-dependent bone development. J Cell

Biochem 112(3):750–755; Epub 2011/02/ 18 3. Long F (2012) Building strong bones: molecular regulation of the osteoblast lineage. Nat Rev Mol Cell Biol 13(1):27–38; Epub 2011/ 12/23

Mouse Calvarial Osteoblast Isolation 4. Aronow MA, Gerstenfeld LC, Owen TA, Tassinari MS, Stein GS, Lian JB (1990) Factors that promote progressive development of the osteoblast phenotype in cultured fetal rat calvaria cells. J Cell Physiol 143(2):213–221; Epub 1990/05/01 5. Malaval L, Liu F, Roche P, Aubin JE (1999) Kinetics of osteoprogenitor proliferation and osteoblast differentiation in vitro. J Cell Biochem 74(4):616–627; Epub 1999/08/10 6. Owen TA, Aronow M, Shalhoub V, Barone LM, Wilming L, Tassinari MS et al (1990) Progressive development of the rat osteoblast phenotype in vitro: reciprocal relationships in expression of genes associated with osteoblast proliferation and differentiation during formation of the bone extracellular matrix. J Cell Physiol 143(3):420–430; Epub 1990/06/01 7. Whyte MP (2010) Physiological role of alkaline phosphatase explored in hypophosphatasia. Ann N Y Acad Sci 1192:190–200; Epub 2010/04/16 8. Boskey AL, Maresca M, Ullrich W, Doty SB, Butler WT, Prince CW (1993) Osteopontinhydroxyapatite interactions in vitro: inhibition of hydroxyapatite formation and growth in a gelatin-gel. Bone Miner 22(2):147–159; Epub 1993/08/01 9. Boskey AL, Spevak L, Paschalis E, Doty SB, McKee MD (2002) Osteopontin deficiency increases mineral content and mineral crystallinity in mouse bone. Calcif Tissue Int 71 (2):145–154; Epub 2002/06/20 10. Hunter GK, Goldberg HA (1993) Nucleation of hydroxyapatite by bone sialoprotein. Proc Natl Acad Sci U S A 90(18):8562–8565; Epub 1993/09/15 11. Hauschka PV, Lian JB, Cole DE, Gundberg CM (1989) Osteocalcin and matrix Gla protein: vitamin K-dependent proteins in bone. Physiol Rev 69(3):990–1047; Epub 1989/ 07/01 12. Ferron M, Wei J, Yoshizawa T, Del Fattore A, DePinho RA, Teti A et al (2010) Insulin signaling in osteoblasts integrates bone remodeling and energy metabolism. Cell 142 (2):296–308; Epub 2010/07/27 13. Fulzele K, Riddle RC, DiGirolamo DJ, Cao X, Wan C, Chen D et al (2010) Insulin receptor signaling in osteoblasts regulates postnatal bone acquisition and body composition. Cell 142(2):309–319; Epub 2010/07/27 14. Wei J, Karsenty G (2015) An overview of the metabolic functions of osteocalcin. Rev Endocr Metab Disord 16(2):93–98

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15. Zoch ML, Clemens TL, Riddle RC (2016) New insights into the biology of osteocalcin. Bone 82:42–49 16. Peck WA, Birge SJ Jr, Fedak SA (1964) Bone cells: biochemical and biological studies after enzymatic isolation. Science 146 (3650):1476–1477; Epub 1964/12/11 17. Wong G, Cohn DV (1974) Separation of parathyroid hormone and calcitonin-sensitive cells from non-responsive bone cells. Nature 252 (5485):713–715; Epub 1974/12/20 18. McCarthy TL, Centrella M, Canalis E (1988) Further biochemical and molecular characterization of primary rat parietal bone cell cultures. J Bone Miner Res 3(4):401–408; Epub 1988/ 08/01 19. Wong GL, Cohn DV (1975) Target cells in bone for parathormone and calcitonin are different: enrichment for each cell type by sequential digestion of mouse calvaria and selective adhesion to polymeric surfaces. Proc Natl Acad Sci U S A 72(8):3167–3171; Epub 1975/08/01 20. Bellows CG, Aubin JE, Heersche JN, Antosz ME (1986) Mineralized bone nodules formed in vitro from enzymatically released rat calvaria cell populations. Calcif Tissue Int 38 (3):143–154; Epub 1986/03/01 21. Bhargava U, Bar-Lev M, Bellows CG, Aubin JE (1988) Ultrastructural analysis of bone nodules formed in vitro by isolated fetal rat calvaria cells. Bone 9(3):155–163; Epub 1988/01/01 22. Nefussi JR, Boy-Lefevre ML, Boulekbache H, Forest N (1985) Mineralization in vitro of matrix formed by osteoblasts isolated by collagenase digestion. Differentiation 29 (2):160–168; Epub 1985/01/01 23. Sudo H, Kodama HA, Amagai Y, Yamamoto S, Kasai S (1983) In vitro differentiation and calcification in a new clonal osteogenic cell line derived from newborn mouse calvaria. J Cell Biol 96(1):191–198; Epub 1983/01/01 24. Beck GR Jr, Zerler B, Moran E (2001) Gene array analysis of osteoblast differentiation. Cell Growth Differ 12(2):61–83; Epub 2001/03/ 13 25. Garcia T, Roman-Roman S, Jackson A, Theilhaber J, Connolly T, Spinella-Jaegle S et al (2002) Behavior of osteoblast, adipocyte, and myoblast markers in genome-wide expression analysis of mouse calvaria primary osteoblasts in vitro. Bone 31(1):205–211; Epub 2002/07/12 26. Nishikawa K, Nakashima T, Takeda S, Isogai M, Hamada M, Kimura A et al (2010)

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Madison L. Doolittle et al.

Maf promotes osteoblast differentiation in mice by mediating the age-related switch in mesenchymal cell differentiation. J Clin Invest 120(10):3455–3465; Epub 2010/09/30 27. Roman-Roman S, Garcia T, Jackson A, Theilhaber J, Rawadi G, Connolly T et al (2003) Identification of genes regulated during osteoblastic differentiation by genomewide expression analysis of mouse calvaria primary osteoblasts in vitro. Bone 32 (5):474–482; Epub 2003/05/20 28. Seth A, Lee BK, Qi S, Vary CP (2000) Coordinate expression of novel genes during

osteoblast differentiation. J Bone Miner Res 15(9):1683–1696; Epub 2000/09/08 29. Franceschi RT, Iyer BS (1992) Relationship between collagen synthesis and expression of the osteoblast phenotype in MC3T3-E1 cells. J Bone Miner Res 7(2):235–246; Epub 1992/ 02/01 30. Peterkofsky B (1991) Ascorbate requirement for hydroxylation and secretion of procollagen: relationship to inhibition of collagen synthesis in scurvy. Am J Clin Nutr 54 (6 Suppl):1135S–1140S. Epub 1991/12/01

Chapter 27 Mitochondrial Function and Metabolism of Cultured Skeletal Cells Li Tian, Clifford J. Rosen, and Anyonya R. Guntur Abstract Measuring cellular metabolism accurately is necessary to understand bioenergetic pathways in cells. The major ATP generating pathways in cells are oxidative phosphorylation and glycolysis. We have recently analyzed and published bioenergetic pathways active in osteoblasts undergoing differentiation in response to various substrates. Based on those studies, here we provide step-by-step procedures to isolate, culture, plate and run a seahorse assay for measuring cellular metabolism. Furthermore, we provide an example of oxygen consumption and extracellular acidification rate traces obtained from MC3T3E1-C4 cells using the XFe96 seahorse analyzer. One of the limitations of studying bioenergetics in bone cells is the current lack of techniques to analyze bioenergetics in vivo in live animals. There are currently techniques that have been developed using third harmonic generation to study osteocytes using three-photon microscopy along with metabolic changes using endogenous two-photon excited fluorescence. However, these sophisticated techniques are not widely available. The relative ease with which one can obtain data pertaining to metabolic parameters using the XF technology makes it a very attractive technique to utilize on a monolayer of adherent cells. Key words Calvarial osteoblasts, Bone marrow stromal cells, MC3T3E1C4 preosteoblast oxidative phosphorylation, Glycolysis, Oxygen consumption rate, Extracellular acidification rate

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Introduction Bone is a multicellular endocrine organ. Bone and the enclosed marrow space consist of a number of different cell types including chondrocytes, osteoblasts, osteocytes, osteoclasts, and marrow adipocytes. The last three decades has seen tremendous progress in identifying the genetic and signaling pathways involved in controlling proliferation and differentiation of these various cell types during skeletal development. The more recent advances using mouse genetics identified a role for the skeleton and bone marrow as an endocrine organ regulating whole body energy metabolism [1]. This now established endocrine function for bone has spurred interest in understanding how cells involved in forming and

Matthew J. Hilton (ed.), Skeletal Development and Repair: Methods and Protocols, Methods in Molecular Biology, vol. 2230, https://doi.org/10.1007/978-1-0716-1028-2_27, © Springer Science+Business Media, LLC, part of Springer Nature 2021

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maintaining bone generate ATP, which substrates are preferred and how these are processed and if these change with age and disease [2]. The current information identifying the bioenergetic pathways during differentiation of these cells is mostly based on in vitro characterization. This chapter discusses the methods we have used for culturing primary calvarial osteoblasts, bone marrow stromal cells and preosteoblast cell line MC3T3E1C4 cells and using the Agilent Seahorse XF Analyzers to measure oxidative phosphorylation and glycolysis simultaneously in the same cell populations. Primary calvarial osteoblast assays were performed on the XF24, whereas assays involving primary bone marrow stromal cells and MC3T3E1C4 cells were performed on both a XF24 and XF96 analyzer. Numerous protocols have been published describing the use of this technique and analysis for various cell types but not specifically for bone cells [3, 4]. Oxygen consumption rate (OCR) is indicative of oxidative phosphorylation and extra cellular acidification rate (ECAR) is indicative of glycolytic lactate production in combination with CO2 evolution from the tricarboxylic acid cycle. There are currently two ways you can correct for CO2 evolution; one method has been described in [5] and we have employed this method in our recent study [6]. The second method is described by Agilent and can be employed using XF glycolytic rate assay provided in a kit. This is necessary to ensure that the glycolytic rate is accurate and not contaminated with acidification from Krebs cycle. There are published studies that have utilized the Agilent seahorse technology to measure bioenergetics in different skeletal cells. Articular chondrocytes from humans with arthritis have been studied in culture and metabolic pathway changes in response to growth factors have been identified [7]. Osteoblasts have been studied extensively using this technology. In these studies, both primary bone marrow stromal cells and calvarial osteoblasts along with preosteoblast cell lines have been studied. There is evidence showing that aerobic glycolysis is a major source of ATP in a differentiated osteoblast. In contrast to this, there are studies using mesenchymal stem/progenitor cells (MSCs) demonstrating that oxidative phosphorylation is the predominant ATP producer when MSCs are provided with osteogenic cues. These differences might be accounted for due to the different source of cells used as starting material [6, 8, 9]. Evidence suggests that cells use glycolysis to sustain a cells need for biosynthesis and specifically in the osteoblast this could be because of the increased need for a differentiated osteoblast to generate extracellular matrix. It has also been postulated that there is a need for increased citrate production for proper nanocrystal structure of bone to be formed. The increase in citrate production would lead to a loss in ATP generated through the TCA cycle which is substituted by glycolysis.

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Osteocytes have been relatively less studied but one recent report studied primary osteocytes from control and aged growth hormone receptor null mice to show compromised mitochondrial function with age. A more recent study showed that the established osteocyte cell line, Ocy454, increase metabolic response to treatments of PTH and Scriptaid, an HDAC complex corepressor inhibitor [10, 11]. Osteoclast bioenergetic studies suggest that these cells are metabolically flexible and employ both oxidative phosphorylation and glycolysis at different phases of differentiation and resorption, with bone resorption relying more on glycolysis [12]. The study of primary osteoclast cultures is challenging, as the cultures do not tend to be a homogenous monolayer and this results in a variable signal for measuring OCR and ECAR. In addition, the cells are a mixture of both mononucleated and multinucleated cells and it is difficult to determine whether the signal is specific to osteoclasts. We are currently working on assays to identify ways to normalize these data and hope to provide this methodology in the future.

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Materials

2.1 General Materials, Reagents, and Kits

1. XF analysers: Agilent Seahorse XF24 and Agilent Seahorse XFe96. 2. Mito Stress test reagents (Agilent). 3. Glycolytic stress test reagents (Agilent). 4. Oligomycin. 5. 2-[2-[4-(trifluoromethoxy)phenyl]hydrazinylidene]-propanedinitrile (FCCP) 6. Antimycin. 7. Rotenone. 8. Hoechst dye. 9. Penicillin/Streptomycin (P/S). 10. Agilent Seahorse XF base media (Agilent). 11. 24-well Seahorse V7 culture plates (Agilent) 12. 96-well Seahorse V7 culture plates (Agilent) 13. Collagenase P. 14. Trypsin/EDTA (Gibco). 15. 1 PBS (Sterile) 16. Osteoblast (OB) growth media: α-MEM containing 10% v/v fetal bovine serum (FBS), 100 units/mL penicillin, 100 μg/ mL streptomycin.

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17. Osteoblast (OB) differentiation media: α-MEM containing 10% v/v fetal bovine serum (FBS), 100 units/mL penicillin, 100 μg/mL streptomycin, 8 mM β-glycerophosphate, and 50 μg/mL ascorbic acid. 18. General dissection equipment (scissors, forceps). 19. 50 mL conical tubes 20. 37  C incubator 21. Autoclave. 22. Micro centrifuge. 23. Shaker (Labquake, Barnstead). 24. BioTek Cytation 1 imaging reader. 25. Nonessential amino acids (Gibco). 26. DMEM, α-MEM (Gibco). 27. Beckman Coulter Allegra X -15R centrifuge. 28. 10 cm corning tissue culture plates. 2.2 Calvarial Osteoblast (COB) Culture Reagents

1. Animals: Mouse neonates 1–3 days of age (see Note 1). 2. COB enzyme solution: 1.5 U/mg Collagenase P; 1 PBS (pH 7.4); 8 mL trypsin–EDTA. 3. COB plating media: DMEM containing 10% FBS, 100 units/ mL penicillin, 100 μg/mL streptomycin, 250 μM nonessential amino acids.

2.3 Bone Marrow Stromal Cell (BMSC) Culture Reagents

1. Animals: Mice at 8-weeks of age (see Note 2).

2.4 MC3T3-E1 C4 Cell Culture Reagents

1. MC3T3-E1 C4 cell line.

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2. Plating Media: α-MEM, 10% FBS, 100 units/mL penicillin, 100 μg/mL streptomycin.

2. Plating Media: α-MEM, 10% FBS, 100 units/mL penicillin, 100 μg/mL streptomycin.

Methods

3.1 Isolation and Culture of Calvarial Osteoblasts for Agilent Seahorse Assay

1. To dissect calvaria from mouse neonates, first euthanize pups according to guidelines put forth in your institutional animal use and care committee (IACUC) protocol. 2. Stabilize the head with forceps and use scissors to cut away fur. Then spread scissors to peel skin. Use forceps to peel skull cap off brain and cut calvaria along sutures into a trapezoid. Next, cut trapezoid into the small pieces and hold in 1PBS on ice until all calvaria are dissected.

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3. Once all calvaria are pooled, discard 1 PBS and add 10 mL COB enzyme solution (Digestion 1) in 50 mL conical tube and set on shaker at 37  C for 15 min. 4. Discard supernatant to waste, leaving the calvarial tissue in the tube and add 10 mL COB enzyme solution to same calvarial pieces (Digestion 2). Set on shaker at 37  C for 15 min. 5. Collect Digestion 2 supernatant into a 50 mL conical tube and add 10 mL plating media and save at 37  C. 6. Add 10 mL COB enzyme solution to same calvarial pieces (Digestion 3). Set on shaker at 37  C for 15 min. Again, pool the digested supernatant and add an extra 10 mL of plating media and store at 37  C. 7. Add 10 mL COB enzyme solution to same calvarial pieces (Digestion 4). Set on shaker at 37  C for 15 min. 8. Collect all of the supernatant and combine with saved solutions from Digests 2 and 3 in a 50 mL conical tube. Total volume should be approximately 50 mL. 9. Centrifuge the cells at ~1200  g for 5 min at 4  C. 10. Pour off supernatant and resuspend pellet in 5 mL of plating media. 11. Using a syringe with an 18-gauge needle, mix thoroughly and filter through a 70 μM cell strainer. 12. Count the cell number and plate COBs in a 10 cm dish, once the cells are confluent trypsinize the cells and plate for experiments. 13. Plates 50,000 COBs per well in 100 μL of α-MEM medium in Agilent seahorse cell culture microplate (24-well plate). Ensure no trypsin is in cell suspension by spinning down cells and resuspending in complete medium. Seed cells by resting the pipette tip at an angle just below the circular rim at the top of each well and ejecting cell suspension slowly; pipet media only into designated “blank” wells. 14. Rock plate gently after seeding cells and place plate in the hood for 1 h with the hood closed and fan/UV light off in order to allow cells to adhere. 15. Add 150 μL of α-MEM medium to each well and move plate to 37  C CO2 incubator. 16. When cells reach 100% confluency, initiate differentiation (Next day of plating). 17. Replace media with OB differentiation media, change media every other day and continue until day 7, 14, or 21 days. Proceed with Seahorse experiment on the eighth, 15th, or 22nd day of culture (see Note 3) and Agilent seahorse protocol 4.

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3.2 Isolation and Culture of BMSCs for Agilent Seahorse Assay

1. To dissect the femur and tibia from 8-week-old mice for BMSC collection, first euthanize the animals according to guidelines put forth in your institutional animal use and care committee (IACUC) protocol. 2. Remove all skin and muscle tissue from the hindlimb and place the cleaned femur and tibia into sterile cold 1 PBS in a 50 mL conical tube. Process all bones in the tissue culture hood for marrow collection. 3. Place 100 μL of plating media into 1.5 mL microcentrifuge tubes. 4. Cut the end of 200 μL pipette tips so that they fit in the micro centrifuge tubes. Insert the tips inside the tubes and ensure that the lids can close. 5. Using scissors, cut both ends of femurs and tibiae and place them in Petri dishes containing cold sterile 1 PBS. 6. Place cut bones inside the cut pipette tips within the micro centrifuge tubes, usually 2 femurs and 2 tibiae from a single mouse in one tube. 7. Perform a quick spin within a microcentrifuge (~15,680  g for 15 s at room temperature) to obtain marrow from the femur and tibia. 8. Once centrifugation is complete, flush out the marrow from the end of the pipette tip as much as possible and discard the inserted pipette tips. 9. Combine and transfer all samples of the same test group into a 15 mL conical tube. Add 10 mL of plating media to the sample and pipet it up and down a couple of times. 10. Plate all cells coming from the femurs and tibiae of one group into a T150 flask (or cells of one mouse on a 10 cm petri dish) and culture in 37  C CO2 incubator. 11. After 48 h of culture, remove the culture media and wash the flask with 1 PBS gently as not to disturb the attached cells. 12. Remove 1 PBS and add the 0.25% trypsin/EDTA solution and incubate for 3–5 min. Quench the trypsin by adding plating media. 13. Calculate the number of cells needed for plating. Centrifuge the required volume of cell suspension at 1200  g for 5 min at room temperature. 14. Remove the media and resuspend the cell pellet in the volume of culture media needed for plating (see Note 4). 15. Seed 5000 BMSCs per well in 80 μL of α-MEM media (96-well plate) and set the wells in the four corners of the plate as the blank well with media only.

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16. Shake the plate carefully and guarantee the cell distribution evenly in the well. 17. Place the plate in the culture hood for 1 h with the hood closed along with the fan/UV light off. 18. Add 100 μL α-MEM medium to each well, and put the put the plate into the 37  C CO2 incubator. 19. Once the cells are confluent, remove α-MEM medium and switch cells to 180 μL of OB differentiation media. 20. Change OB differentiation media every other day until day 7 or day 14. 21. Upon completion of differentiation, proceed with the seahorse experiment as outlined below. 3.3 Culturing of MC3T3-E1 C4 Cells for Agilent Seahorse Assay

1. Thaw the MC3T3-E1 C4 cell and plate in a 10 cm Peri dish. 2. Culture in a 37  C in a CO2 incubator until cells are confluent. 3. Rinse cells gently with 1 PBS. 4. Remove 1 PBS, add a 0.25% trypsin–EDTA solution, and place in incubator for 3–5 min. 5. Quench the trypsin by adding plating media and count the cells. 6. Seed 50,000 (24 well plate in 200 μL) or 1500 MC3T3-E1 C4 cells per well cells (96 well plate in 180 μL) in α-MEM media and reserve the wells in the four corners of plate as the blank wells (media only). 7. Shake the plate carefully to guarantee the cells are distributed evenly across all wells. 8. Put the plate in the hood for 1 h with the hood closed along with the fan/UV light off in order to allow for cell adherence. 9. Add 100 μL α-MEM media to each well, and put the put the plate into a 37  C CO2 incubator. 10. Culture cells until confluent. If OB differentiation is to be performed, switch cells to OB differentiation media, change the differentiation media every other day until day 7. 11. Set up the Agilent Seahorse experiment for a 24- or 96-well plate and follow the protocol as outlined in Subheading 3.4 If using cell number for normalization (see Note 5).

3.4 Agilent Seahorse Experiment

1. Prepare XF Assay Media by supplementing XF base media with 25 mM glucose and 10 mM pyruvate pH 7.4 (see Note 6). 2. Wash cells by removing 200 μL of media and adding 500 μL XF Assay Media. Remove another 500 μL media and add a final 500 μL XF Assay Media.

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Fig. 1 (a) OCR and (b) ECAR profiles of VEH and DFP (1 mM) treated MC3T3-E1 C4 osteoblasts after Mito Stress test using a mixture of substrates. The presence of Deferiprone (iron chelator) blocked a substantial amount of OCR suggesting that osteoblasts ability to generate ATP through oxidative phosphorylation is impaired. The increase observed in ECAR suggests that the cells have increased glycolysis to compensate for the ATP deficit created by DFP treatment

3. Place cells in non-CO2 incubator at 37 for 1 h prior to start of assay. 4. Use the Seahorse XF Extracellular Flux Assay Kit, and fill each well of the utility plate with 200 μL Agilent Seahorse XF Calibrant (PH 7.4), put the sensor cartridge into the utility plate, and keep the sensors submerging the calibrant media (see Note 7). Place the kit in a non-CO2 37  C incubator at least 3 h or overnight. 5. Warm the XF DMEM Seahorse media (PH 7.4) in a sterile bottle and add Seahorse XF substances (XF glucose, pyruvate, and glutamine solutions) for the XF DMEM Seahorse media; 6. Prepare reagents for assay (injection volume of 75 μL for each reagent per well) from 2.5 mM Seahorse stock solutions: 1.2 μM oligomycin, 0.56 μM FCCP, 0.96 μM antimycin A, and Rotenone. Dissolve these reagents in the above XF DMEM Agilent Seahorse media (see Note 8).

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7. Load the reagents to the injection ports within the sensor cartridge. Use the Wave controller software to set up the required template. 8. After the sensor cartridge (hydrated and loaded with reagents) is ready, place the sensor cartridge on the machine for calibration. 9. Once the sensor cartridge calibration is complete, load your cell plate on the XF Analyzer, and use the Wave controller software template following the command dialogs to measure OCR and ECAR (see Note 9). 10. We have included data from an experimental run on the XF96 using MC3T3E1-C4 cells treated with a mitophagy inducer (Deferiprone) in Fig. 1 to illustrate some of the above points. 11. Cells were plated and cultured as described in Subheading 3.3. The day before the assay, cells were treated with Vehicle (Methanol) or Deferiprone (1 mM) overnight. 12. The day of the assay the cells were processed for the XFe96 Mito Stress test as described in 4. After the completion of the assay the wells were imaged in a BioTek Cytation 1. This data was used for normalization. 13. Oxygen consumption rate and extracellular acidification rates are shown in Fig. 1. The data can be processed for a number of different parameters as described in our recent publication [6] using different assay kits provided by Agilent (see Note 10).

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Notes 1. 5–8 pups is sufficient per group. 2. 3 mice is sufficient per group 3. We have performed the osteoblast differentiation assay for 14 and 21d with media changes daily or every other day. The major issue we come across with differentiation is that the cells sometimes in some of the wells curl up and are not optimal for the seahorse assay. To overcome this we recommend using as many technical replicates as possible for each group. 4. A more detailed protocol has been published in [13]. 5. At the end of the XF analyzer run, we utilize protein content as measured by Bradford assay, BCA assay or cell number as imaged using a fluorescent dye for normalization. This is necessary for comparing different populations of cells that have undergone different treatments, which might result in differences in cell proliferation or number. We have recently started using cell number for normalization using a BioTek Cytation 1 instrument that is dedicated with the XF96 instrument for in

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situ cell counting. A number of technical briefs are available on the Agilent website that give detailed protocols to perform normalization. The injection of a cell-permeable Hoechst dye labels the nuclei, and these fluorescently labeled nuclei can be imaged and counted on the BioTek Cytation. The process is very straightforward and user-friendly; more information is available in the technical protocol provided by Agilent at (https://www.agilent.com/en/products/cell-analysis/sea horse-xf-imaging-normalization-solution). 6. Ensure that the XF media PH is accurate. This is critical for the ECAR measurements. Agilent now makes media available commercially that is ready to use. 7. A couple of things to keep in mind while incubating the cartridge for calibration overnight at 37  C. Add sufficient amount of calibrant to avoid evaporation. Before starting the calibration on the XF machine check that, the orientation of the wells is in the right order. Check the cells under the microscope to ensure that the cells are intact and viable. 8. Prepare the compounds for assay (injection volume of 20 μL for each reagent per well): 2 μM oligomycin, 2 μM FCCP, 1 μM antimycin A, and 1 μM rotenone, separately (For XF96). Concentrations of the inhibitors should be empirically determined in case other cell types are utilized. The cells can also be pretreated for various times with any compound/s of interest or they can be injected through the ports before the mitochondrial inhibitors, to study both long term and acute treatment effects on cells. 9. The assay protocols for the seahorse analyzers are available with the instrument manuals and at the following websites (https:// www.agilent.com/en/products/cell-analysis/seahorseanalyzers/seahorse-xfe96-analyzer and https://www.agilent. com/en/products/cell-analysis/seahorse-analyzers/seahorsexfe24-analyzer). For the XF24 we generally use the following cycles (Mix 3 min Wait 2 min and then measure for 3 min), for the XFe96 we typically use (Mix 3 min, wait 0 min and measure 3 min). 10. Currently Agilent seahorse provides a number of different test kits to measure ATP production rates (https://www.agilent. com/en/products/cell-analysis/seahorse-xf-atp) and glycolytic rates (https://www.agilent.com/en/products/cell-analy sis/seahorse-xfe-consumables/kits-reagents-media/seahorsexf-glycolytic-rate-assay-kit), which are useful. We have used a different procedure to measure ATP production and glycolytic rates in our recent publication [6].

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Acknowledgments This work was funded by NIGMS to A.R.G. through P20GM121301, Phase I: Mesenchymal and Neural Regulation of Metabolic Networks, Lucy Liaw, PhD, Program Director. Disclosure Statement: The authors have nothing to disclose References 1. Karsenty G, Oury F (2012) Biology without walls: the novel endocrinology of bone. Annu Rev Physiol 74(1):87–105. https://doi.org/ 10.1146/annurev-physiol-020911-153233 2. Lee WC, Guntur AR, Long F, Rosen CJ (2017) Energy metabolism of the osteoblast: implications for osteoporosis. Endocr Rev 38 (3):255–266. https://doi.org/10.1210/er. 2017-00064 3. Zhang J, Zhang Q (2019) Using seahorse machine to measure OCR and ECAR in cancer cells. Methods Mol Biol (Clifton, NJ) 1928:353–363. Epub 2019/02/07. https:// doi.org/10.1007/978-1-4939-9027-6_18 4. Divakaruni AS, Paradyse A, Ferrick DA, Murphy AN, Jastroch M (2014) Analysis and interpretation of microplate-based oxygen consumption and pH data. Methods Enzymol 547:309–354. Epub 2014/11/25. https:// doi.org/10.1016/b978-0-12-801415-8. 00016-3 5. Mookerjee SA, Goncalves RLS, Gerencser AA, Nicholls DG, Brand MD (2015) The contributions of respiration and glycolysis to extracellular acid production. Biochim Biophys Acta 1847(2):171–181. Epub 2014/12/03. https://doi.org/10.1016/j.bbabio.2014.10. 005 6. Guntur AR, Gerencser AA, Le PT, DeMambro VE, Bornstein SA, Mookerjee SA, Maridas DE, Clemmons DE, Brand MD, Rosen CJ (2018) Osteoblast-like MC3T3-E1 cells prefer glycolysis for ATP production but adipocyte-like 3T3-L1 cells prefer oxidative phosphorylation. J Bone Miner Res 33(6):1052–1065. https:// doi.org/10.1002/jbmr.3390 7. Wang C, Silverman RM, Shen J, O’Keefe RJ (2018) Distinct metabolic programs induced by TGF-beta1 and BMP2 in human articular chondrocytes with osteoarthritis. J Orthop

Translat 12:66–73. https://doi.org/10. 1016/j.jot.2017.12.004 8. Guntur AR, Le PT, Farber CR, Rosen CJ (2014) Bioenergetics during calvarial osteoblast differentiation reflect strain differences in bone mass. Endocrinology 155 (5):1589–1595. https://doi.org/10.1210/ en.2013-1974 9. Shum LC, White NS, Mills BN, Bentley KL, Eliseev RA (2016) Energy metabolism in mesenchymal stem cells during osteogenic differentiation. Stem Cells Dev 25(2):114–122. https://doi.org/10.1089/scd.2015.0193 10. Liu Z, Solesio ME, Schaffler MB, FrikhaBenayed D, Rosen CJ, Werner H, Kopchick JJ, Pavlov EV, Abramov AY, Yakar S (2019) Mitochondrial function is compromised in cortical bone osteocytes of Long-lived growth hormone receptor null mice. J Bone Miner Res Off J Am Soc Bone Miner Res 34 (1):106–122. Epub 2018/09/15. https:// doi.org/10.1002/jbmr.3573 11. Sun N, Uda Y, Azab E, Kochen A, Santos R, Shi C, Kobayashi T, Wein MN, Divieti PP (2019) Effects of histone deacetylase inhibitor Scriptaid and parathyroid hormone on osteocyte functions and metabolism. J Biol Chem 294(25):9722–9733. https://doi.org/10. 1074/jbc.RA118.007312 12. Arnett TR, Orriss IR (2018) Metabolic properties of the osteoclast. Bone 115:25–30. Epub 2017/12/25. https://doi.org/10.1016/j. bone.2017.12.021 13. Maridas DE, Rendina-Ruedy E, Le PT, Rosen CJ (2018) Isolation, culture, and differentiation of bone marrow stromal cells and osteoclast progenitors from mice. J Vis Exp (131):56750. https://doi.org/10.3791/ 56750

Chapter 28 Radiolabeled Amino Acid Uptake Assays in Primary Bone Cells and Bone Explants Leyao Shen and Courtney M. Karner Abstract Radiolabeled amino acid uptake assays are a highly sensitive method used to characterize the uptake of amino acids by cells or tissues in culture. This method is an excellent tool to quantify changes in amino acid consumption that are associated with states of cellular differentiation and/or disease. The methods presented here can be adapted to measure the transport of all amino acids and can be applied to cultured cells and bone explants. Key words Tritium, 3H, Radiolabeled isotopes, Amino acid uptake, Primary cells, Bone explants

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Introduction Amino acids are critical regulators of cellular function and activity. In addition to direct incorporation into polypeptide chains, amino acid metabolism provides critical intermediate metabolites that function as enzymatic cofactors, epigenetic regulators, TCA cycle intermediates, contribute to de novo amino acid synthesis and regulate cellular redox status. Many studies demonstrate the importance of amino acid metabolism in various cellular functions including cell proliferation, differentiation and pluripotency maintenance [1–8]. Osteoblasts are the principle bone forming cell whose primary function is to produce and secrete Type 1 Collagen and other proteins that comprise the bone matrix. A constant supply of amino acids is required to sustain high rates of protein synthesis associated with bone anabolism. To meet this requirement, osteoblasts must maximize the production or acquisition of amino acids. Indeed, recent reports have linked the regulation of amino acid uptake and metabolism to osteoblast function and bone formation [7–10]. The major sources of cellular amino acids are extracellular environment, intracellular protein degradation and de novo synthesis. The methods described here will focus on amino acid uptake

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from the extracellular milieu. The most common methods to evaluate amino acid uptake utilize either radiolabeled (e.g., 3H or 14C) or heavy isotope labeled (e.g., 13C) amino acids [11, 12]. Heavy isotope assays are time-consuming but provides a more thorough analysis of amino acid metabolism after uptake. By comparison, radiolabeled amino acid uptake assays are fast, sensitive and cheap but are less informative about downstream metabolism [13, 14]. Here, we describe two modified protocols for analyzing radiolabeled amino acid uptake using L-[3,4-3H]-proline in both cultured primary cells and in bones ex vivo. Importantly, these protocols are easily adaptable allowing for the evaluation of uptake of other radiolabeled amino acids in bone cells both in vitro and ex vivo. Finally, we include important troubleshooting information pertinent to working with radioactivity.

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Materials 1. Primary osteoblasts and bones isolated from mice. 2. Scalpels. 3. 1 mL syringes 4. Injection needles G30. 5. Dissection scissors. 6. 12-well/24-well tissue culture plate 7. Sterile 1 Phosphate-buffered saline (PBS), pH 7.4. 8. 37  C cell incubator 9. Krebs Ringers HEPES (KRH), pH 8.0: (120 mM NaCl; 5 mM KCl; 2 mM CaCl2; 1 mM MgCl2; NaHCO3; 5 mM HEPES; 1 mM D-Glucose). Adjust pH utilizing 1 M NaOH and 1 M HCl. 10. L-[2,3-3H]-Proline; >97%; 250 μCi (see Note 1). 11. Protection shield. 12. 1% sodium dodecyl sulfate (SDS) buffer (w/v). 13. Radioimmunoprecipitation assay (RIPA) buffer: [150 mM NaCl; 5 mM EDTA; 50 mM Tris (pH 8.0); 0.5% NP40 (v/v); 0.5% DOX (w/v); 0.1% SDS (w/v)]. 14. 1.5 mL Eppendorf tubes 15. Ice. 16. Ultima Gold™ (Scintillation solution). 17. Glass Scintillation vials. 18. Beckman LS6500 Scintillation counter or equivalent. 19. Sonicator.

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20. Radioactivity decontaminant. 21. Geiger counter (see Note 2).

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3.1 Amino Acid Uptake in Primary Osteoblasts

1. Isolate calvarial osteoblasts according to Chap. 26 (see Note 3). 2. Seed 1  105 primary calvarial cells on 12-well tissue culture plates in α-MEM containing 15% FBS. Seed cells in duplicate plate to allow for appropriate normalization in the final step (Subheading 3.1, step 13). Place the plate in a humidified cell culture incubator at 37  C with 5% CO2. 3. Culture the cells for 2–3 days until confluent. 4. Prewarm 1 PBS and KRH to 37  C. 5. Wash cells two times with 1 PBS, pH 7.4. 6. Wash cells once with KRH. 7. Perform all the experiments associated with radiation behind the protection shield. 8. Make 4 μCi/mL L-[2,3-3H]-Proline working media by diluting 4 μL of [1 μCi μL 1] L-[2,3-3H]-Proline stock in 1 mL KRH (see Note 4). 9. Incubate cells with 4 μCi/mL L-[2,3-3H]-Proline for 5 min. 10. Remove the radioactive medium. Wash the cells three times briefly with ice-cold KRH. Discard all the washes in the radioactive liquid waste container (see Note 1). 11. Lyse cells with 1 mL 1% SDS. Pipette up and down for 10 times. Transfer cell lysates to 1.5 mL Eppendorf tubes. Discard cell culture plates and pipette tips in radioactive solid waste container (see Note 1). 12. Centrifuge at > 9500  g for 10 min. Transfer supernatants to scintillation vials containing 8 mL scintillation solution. Discard tubes and pipette tips in radioactive solid waste container (see Note 1). 13. Read radioactivity in counts per minute (cpm) using Scintillation counter. Discard scintillation vials in radioactive glass waste container (see Note 1). 14. Trypsinize extra plate (from Subheading 3.1, step 2) and quantify cells using a hemocytometer. Normalize the cpm to cell number (see Note 5). 15. Spray the cell culture hood, instruments and bench with radioactivity decontaminant. Perform wipe tests to eliminate the possibility of radioactive contamination.

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3.2 Amino Acid Uptake in Bone Explants (See Figs. 1, 2, and 3)

1. Prewarm KRH to 37  C. 2. Dissect both humeri from each mouse. Remove all extemporaneous tissues using a scalpel. Remove the epiphyses from the bone. 3. Flush out marrow from the bone and weigh the bone shafts to normalize for the amino acid uptake from Subheading 3.2, step 15 (see Note 6). 4. Boil one humerus in 1 PBS on heat block at 100  C for 10 min to decellularize the bone as control (see Note 7). 5. Equilibrate humeri in KRH for 30 min in the cell culture incubator at 37  C. 6. Perform all the experiments associated with radiation behind the protection shield. 7. Make 4 μCi/mL L-[2,3-3H]-proline working media by diluting 4 μL of [1 μCi μL 1] L-[2,3-3H]-proline stock in 1 mL KRH. 8. Incubate both the experimental and boiled humeri with 4 μCi/ mL L-[2,3-3H]-proline for up to 60 min in the cell culture incubator at 37  C. The actual incubation time should be determined empirically (see Note 8). 9. Remove radioactive medium. Terminate the reaction by washing humeri three times using ice-cold KRH. Discard all the washes in the radioactive liquid waste container (see Note 1). 10. Transfer each bone into 1.5 mL Eppendorf tube. Add 500 μL RIPA buffer. Discard culture plates and pipette tips in radioactive solid waste container (see Note 1). 11. Homogenize the humeri by chopping 100 times with scissors in 1.5 mL Eppendorf tubes. 12. Sonicate bone homogenates (Amplitude: 35%, Pulse 1 s, Duration: 10 s) (see Note 9). 13. Clarify the lysate by centrifugation at > 9500  g for 10 min. Transfer 200 μL of the supernatant to scintillation vials containing 8 mL scintillation solution. Discard tubes and pipette tips in radioactive solid waste container (see Note 1). 14. Read radioactivity (cpm) using Scintillation counter. Discard scintillation vials in radioactive glass waste container (see Note 1). 15. Divide the radioactivity from Subheading 3.2, step 14 by the bone weight from Subheading 3.2, step 3 in order to normalize amino acid uptake with different bone sizes from different animals. 16. Spray the cell culture hood, instruments, and bench with radioactivity decontaminant. Perform wipe tests to eliminate the possibility of radioactive contamination.

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Fig. 1 L-[2,3-3H]-proline uptake assays in calvarial osteoblasts (cOBs). Example of proline uptake assay in cOBs treated with the amino acid analog 2-(methylamino)-isobutyric acid (MeAIB). MEAIB is a competitive inhibitor of System A amino acid transporters known to transport proline [15]. cOBs were cultured with labeled proline for 5 min in the presence of water or 5 μM MeAIB

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Notes 1. For the use of radioactive materials, please refer to Office of Radiation Safety at your home institution before conducting any radiation-associated experiments. 2. Tritium is a very low energy beta emitter and even large amounts of this isotope pose no external dose hazard to exposed persons. Geiger counters are not sensitive to tritium radiation. However, we recommend using the Geiger counter as a courtesy to alert people in the area that you are working with radiation. If you are working with isotopes with stronger radioactivity (e.g., 14C and 35S), use of the Geiger counter is required. 3. This protocol can be applied to cell lines including ST2, ATDC5, and MC3T3 or other primary skeletal cells like chondrocytes, bone marrow stromal cells, and bone marrow macrophages. 4. Other radioactive isotope labels (e.g., appropriate and detectable.

14

C and

35

S) are also

5. Results can also be normalized by DNA content for cells in vitro. This is recommended for mineralizing cells that are difficult to trypsinize and quantify. 6. Flush out the bone marrow by centrifugation for adult mice. In young mice or embryonic bones, it is better to flush out bone marrow using a syringe and 30G needle.

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Fig. 2 Evaluating radiolabeled amino acid uptake assay in bones ex vivo. (a) Schematic overview of amino acid uptake assay in humeri cultured ex vivo. (b) Sample preparation for amino acid uptake in humeri ex vivo. (i) Image of a

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Fig. 3 Ex vivo L-[2,3-3H]-Proline uptake assays in neonatal humeri. (a) L-[2,3-3H]-Proline uptake assays performed over 90 min in (b) live and boiled humeri isolated from 3 day old C57BL/6 mice. Proline uptake increases linearly for the first 60 min and then plateaus. (c) L-[2,3-3H]-Proline uptake assay in humeri performed in the presence or absence of 5 μM MeAIB

ä Fig. 2 (continued) humerus with extemporaneous tissues removed. (ii) Image of a humerus after epiphyses are removed. (iii) Image of a humerus cut in the middle to ease the removal of marrow using a syringe or centrifugation

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7. The contralateral bone is boiled as a control because radioactive amino acids can adsorb to bone matrix independent of facilitated transport by bone cells. 8. For adult bone (older than 1 month), consider longer incubation in radioactive medium. 9. Sonication of small sample volumes can result in foaming. If this occurs, centrifuge the sample and let rest for 10 min to remove bubbles.

Acknowledgments This work was supported by NIH/NIAMS grant AR071967 to C. M.K. References 1. Karner CM, Long F (2017) Wnt signaling and cellular metabolism in osteoblasts. Cell Mol Life Sci 74(9):1649–1657 2. Nelsen CJ, Rickheim DG, Tucker MM, McKenzie TJ, Hansen LK, Pestell RG et al (2003) Amino acids regulate hepatocyte proliferation through modulation of cyclin D1 expression. J Biol Chem 278 (28):25853–25858 3. Krall AS, Xu S, Graeber TG, Braas D, Christofk HR (2016) Asparagine promotes cancer cell proliferation through use as an amino acid exchange factor. Nat Commun 7:11457 4. Green CR, Wallace M, Divakaruni AS, Phillips SA, Murphy AN, Ciaraldi TP et al (2016) Branched-chain amino acid catabolism fuels adipocyte differentiation and lipogenesis. Nat Chem Biol 12(1):15–21 5. Shiraki N, Shiraki Y, Tsuyama T, Obata F, Miura M, Nagae G et al (2014) Methionine metabolism regulates maintenance and differentiation of human pluripotent stem cells. Cell Metab 19(5):780–794 6. Comes S, Gagliardi M, Laprano N, Fico A, Cimmino A, Palamidessi A et al (2013) L-Proline induces a mesenchymal-like invasive program in embryonic stem cells by remodeling H3K9 and H3K36 methylation. Stem Cell Rep 1(4):307–321 7. Karner CM, Esen E, Okunade AL, Patterson BW, Long F (2015) Increased glutamine catabolism mediates bone anabolism in response to WNT signaling. J Clin Invest 125(2):551–562 8. Yu Y, Newman H, Shen L, Sharma D, Hu G, Mirando AJ et al (2019) Glutamine metabolism regulates proliferation and lineage

allocation in skeletal stem cells. Cell Metab 29 (4):966–978. e4 9. Rached MT, Kode A, Xu L, Yoshikawa Y, Paik JH, Depinho RA et al (2010) FoxO1 is a positive regulator of bone formation by favoring protein synthesis and resistance to oxidative stress in osteoblasts. Cell Metab 11 (2):147–160 10. Elefteriou F, Benson MD, Sowa H, Starbuck M, Liu X, Ron D et al (2006) ATF4 mediation of NF1 functions in osteoblast reveals a nutritional basis for congenital skeletal dysplasiae. Cell Metab 4(6):441–451 11. Hahn TJ, Downing SJ, Phang JM (1969) Amino acid transport in adult diaphyseal bone: contrast with amino acid transport mechanisms in fetal membranous bone. Biochim Biophys Acta 183(1):194–203 12. Rosenbusch JP, Flanagan B, Nichols G Jr (1967) Active transport of amino acids into bone cells. Biochim Biophys Acta 135 (4):732–740 13. Maleknia SD, Johnson R (2011) Mass spectrometry of amino acids and proteins. In: Hughes AB (ed) Amino acids, peptides and proteins in organic chemistry: analysis and function of amino acids and peptides. WileyVCH, Weinheim, pp 1–50 14. Rennie MJ (1999) An introduction to the use of tracers in nutrition and metabolism. Proc Nutr Soc 58(4):935–944 15. Yee JA (1988) Effect of parathyroid hormone on amino acid transport by cultured neonatal mouse calvarial bone cells. J Bone Miner Res 3 (2):211–218

Chapter 29 RANKL-Based Osteoclastogenic Assay from Murine Bone Marrow Cells Zhenqiang Yao, Lianping Xing, and Brendan F. Boyce Abstract The osteoclast is the unique type of cell that resorbs bone in vivo and it is required for normal skeletal development and postnatal homeostasis. Osteoclast deficiency impairs skeletal development during embryogenesis and results in osteopetrosis and impaired tooth eruption. In contrast, excessive osteoclast formation in adults results in bone loss in a number of conditions, including osteoporosis, rheumatoid arthritis, and metastatic bone disease. Osteoclasts are derived from monocytes/macrophages; they can be generated in vitro by treatment of these precursor cells with macrophage colony stimulating factor (M-CSF) and receptor activator of NF-κB ligand (RANKL). This chapter describes procedures for generating osteoclasts from mouse bone marrow cells in vitro using M-CSF and RANKL and assessing their ability to form resorption lacunae on thin bone slices. Key words Osteoclasts, Culture, M-CSF, RANKL, Mouse, Bone marrow

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Introduction Osteoclasts, the multinucleated cells that resorb bone, are derived from precursors in the monocyte/macrophage lineage and participate in the essential biological processes of bone modeling and remodeling during embryonic development and post-natal growth. Increased osteoclast activity is responsible for pathological bone loss and destruction in many diseases, such as osteoporosis, rheumatoid arthritis, and cancer cell metastasis to bone. Although osteoclasts had been identified in tissue sections and characterized in ex vivo organ cultures [1] and their origin from hematopoietic precursors had been established [2], the first papers describing osteoclast generation from precursors in vitro were published by Allen et al. in 1981 [3, 4]. More efficient methods to generate osteoclasts in vitro were reported later by Ibbotson et al. in 1984 [5] and Roodman et al. in 1985 [5, 6] in Dr. Greg Mundy’s group. In these early osteoclastogenic assays, primary bone marrow cells were cultured in medium containing 1,25-dihydroxy-vitamin

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D3 for 1–2 weeks to form multinucleated cells with several osteoclast characteristics, including positive staining for tartrate-resistant acid phosphatase (TRAP) and formation of resorption pits on bone slices. The establishment of an osteoclastogenic assay greatly improved understanding of how osteoclasts formed and were activated in response to cytokines and growth factors as these became available as pure recombinant proteins. However, a major limitation of the assay was consistency of results because the extent of osteoclastogenesis largely depended on the source of the serum, 1,25-dihydroxy-vitamin D3, and the culture medium used. The discovery of receptor activator of nuclear factor kappa-B ligand (RANKL) in 1997 [7, 8] as an essential factor for osteoclast formation along with macrophage-colony stimulating factor (M-CSF) dramatically improved the success and efficiency of osteoclastogenic assays. Unlike 1,25-dihydroxy-vitamin D3 which promotes osteoclastogenesis by stimulating RANKL production by stromal cells, RANKL directly binds to its receptor, RANK, on precursor cells to promote osteoclast differentiation. RANKL can induce osteoclast formation from RANK-expressing cells from multiple sources, including bone marrow, blood, spleen, liver, and lymph nodes. In this chapter, we will describe detailed protocols for RANKLmediated osteoclast formation from mouse bone marrow cells, along with TRAP staining and bone resorption assays, two commonly used criteria for osteoclast identification and functional analysis, respectively. There are other osteoclast assays, such as cocultures of osteoclast precursors with various types of cells that produce RANKL, especially osteoblastic cells. Some cell lines also can give rise to osteoclast-like cells in the presence of RANKL, such as the RAW264.7 cell line. However, the RANKL-based osteoclastogenic assay using primary bone marrow cells is the most commonly used assay to generate osteoclasts in vitro. Protocols for other osteoclastogenic assays have been described [9]. The RANKL-based osteoclastogenic assay utilizes soluble forms of RANKL and monocyte colony stimulating factor (M-CSF), both of which are essential for proliferation and survival of monocytes into osteoclast precursors. M-CSF also induces expression of RANK on these precursors and RANKL completes their differentiation into osteoclasts as well as osteoclast activation and survival, which are also supported by M-CSF.

2 2.1

Materials Instruments

1. Autoclaved scissors. 2. Autoclaved fine forceps. 3. 10 ml disposable syringes

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4. 21 gauge needles 5. 70% ethanol. 2.2

Reagents

1. Alpha minimal essential medium (α-MEM). 2. Fetal bovine serum (FBS) (see Note 1). 3. Penicillin/Streptomycin (P/S) solution 10,000 U/ml. 4. Phosphate buffered saline (PBS). 5. RBC lysis buffer. 6. 10% neutral buffered formalin phosphate 7. 0.5% toluidine blue in PBS 8. 96 well culture plate 9. 10 cm petri dish 10. 15 ml tube 11. Recombinant murine RANKL (R&D). Make 10 μg/μl of stock solution and store in aliquots at
80  C. 12. Recombinant human M-CSF (R&D). Make 30 μg/μl of stock solution and store in aliquots at
80  C (see Note 2). 13. Bone slices: Individual bone slices are prepared from the cortices of bovine long bones obtained from a local slaughterhouse. Muscles and other soft tissues are scraped from the cortices, which are then cut into longitudinal blocks ~1  2  4 cm using an EXAKT Pathology Saw. Uncut or cut bone blocks can be stored at
20  C in a closed container. Blocks are mounted into the vice on a Buehler IsoMet Low Speed Cutting Machine and 300 μm thick slices are cut. Approximately 4.7 mm (3/16 in.) diameter circular discs are punched from these slices using a McGill Hole punch. Bone slices are washed with distilled water 4 times, placed in 70% ethanol for 30 min at room temperature, washed with autoclaved 1 PBS 4 times, dried in a sterilized incubator hood, and stored at
20  C in a closed container. The discs are placed into the wells of 96-well plates for osteoclast resorption assays.

2.3 Solutions and Media

1. TRAP buffer: Combine 9.2 g sodium acetate anhydrous, 11.4 g of L-(+)tartaric acid, 950 ml of distilled water, and 2.8 ml of glacial acetic acid. Dissolve and adjust pH to 4.7–5.0 with 5 M sodium hydroxide (NaOH) to increase or glacial acetic acid to decrease pH. Bring total volume to 1 l with distilled water. 2. 5 M NaOH (for pH adjustment): Combine 50 g NaOH pellets and 250 ml of distilled water.

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3. Naphthol AS-BI Phosphate Substrate (store for 3 weeks at 4  C) (Solution #2): Combine 5 mg of Naphthol AS-BI Phosphate and 250 μl ethylene glycol monoethyl ether. 4. TRAP Solution (50 ml, freshly made): Combine 30 mg Fast Red Violet LB salt in 50 ml of TRAP buffer and mix well (Solution #1). Add 250 μl of Solution #2. Mix Solution #1 and Solution #2 and keep in the dark at 4  C. 5. Washing medium (50 ml of α-MEM-2% FBS-P/S): Combine 49 ml of α MEM, 1.0 ml of FBS, and 0.5 ml P/S. 6. Osteoclast culture medium (50 ml of α-MEM-10% FBS-P/S): Combine 45 ml of α-MEM, 5.0 ml of FBS, 0.5 ml of P/S, 0.5 ml of nonessential amino acid (NEAA), and 0.5 ml of L-glutamine (see Note 3).

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3.1 Preparation of Bone Marrow Cells

1. Harvest bone marrow cells from long bones of 2–4-month-old mice (although any age of mice older than 1 month can be used). In fact, any tissue containing monocytes/macrophages can be used to culture osteoclasts (see Note 4). 2. Sacrifice mice by carbon dioxide asphyxiation, followed by cervical dislocation to ensure death, according to a protocol approved by the Institutional Animal Care and Use Committee. 3. Sterilize the mouse by immersing it whole in 50 ml of 70% ethanol for 2–5 min. 4. Remove the femora and tibiae of both hind limbs using sterilized dissecting scissors and forceps by cutting through the skin surrounding the hip joint. Tear the skin posteriorly toward the feet to remove it. Disarticulate the hip joint and remove the muscle surrounding the femur and tibia. Place clean bones in a 10 cm petri dish. 5. Cut open both ends of each femur and tibia to expose the marrow cavity. Flush out bone marrow with 10 ml washing medium using a 21 gauge needle. Pass the cells through the needle twice to make single cell suspensions. Collect the cells with 10 ml αMEM with 2% FBS into a 15-ml tube (see Note 5). 6. Centrifuge the tube at 1000  g for 5 min at room temperature. Discard the supernatant. 7. Resuspend the cell pellet in 2 ml of washing buffer, add 8 ml of RBC lysis buffer and mix thoroughly. Incubate the cells for 10 min at room temperature. 8. Spin down and discard the supernatant.

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9. Resuspend the cells with 10 ml wash buffer and repeat step 7. 10. Resuspend the cells in 10 ml of osteoclast culture medium and mix thoroughly. 11. Mix 10 μl of cell suspension with 90 μl of osteoclast culture medium. Count cell numbers in a hemocytometer: Cell number/ml ¼ cell count (# of cells in 4 squares of a hemocytometer)  10  104. 3.2

Cell Culture

1. On day 0, seed bone marrow cells at 4–6  104 cells in 200 μl of osteoclast medium per well in a 96-well plate with 5 ng/ml MCSF for 2 days (see Note 6). 2. On day 2, remove 100 μl medium from the culture and replace with 100 μl of freshly made osteoclast culture medium containing 5 ng/ml M-CSF and 10 ng/ml RANKL for 2 days. An alternative is to add RANKL at day 0 (see Notes 7 and 8). 3. On day 4, observe the cells under an inverted microscope. In general, osteoclasts begin to form after 2 days (or 4 days if RANKL is given on day 0) of RANKL treatment [10]. To assess early stages of osteoclast formation, the culture can be stopped and fixed with 10% neutral formalin. To observe later stages of osteoclast formation, replace half the medium with freshly made osteoclast culture medium containing 5 ng/ml M-CSF and 10 ng/ml RANKL at day 4, stop the culture by discarding the culture medium and fix the cells with 10% neutral formalin on the next day (day 5). Large multinucleated cells can be seen under an inverted microscopy at day 4 to 5 (Fig. 1a) (see Note 9). 4. Remove medium and add 200 μl of 10% formalin for 20 min at room temperature. Remove formalin and wash cells thoroughly with water (see Note 10).

3.3 Staining Cells for TRAP Activity

1. Add 60 μl of TRAP solution for 10–30 min at room temperature. To speed up the staining, the plate can be heated to 37  C. Check under microscope for large red/purple multinucleated cells to decide when to stop staining. Examples of TRAP+ osteoclasts are demonstrated in Fig. 1b. 2. Wash the cells thoroughly with water. 3. Counter stain the cells with Mayer’s Hematoxylin for 30 s followed by 0.5% Ammonia Water for 30 s. 4. Wash the cells thoroughly with water. 5. Air dry.

3.4

Bone Resorption

1. Place bone slices in the wells of a 96 well dish in 200 μl of α-MEM-10% FBS-P/S for 2 h to overnight in a cell culture incubator at 37  C.

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Fig. 1 Osteoclast culture assays. Bone marrow cells were isolated from C57BL/6 mice and were cultured with M-CSF for 2 days and followed by M-CSF and RANKL for an additional 4–8 days. (a) The cells were cultured on a plastic culture dish and observed under an inverted light microscope. The image shows large multinucleated osteoclasts. The border of an osteoclast is outlined by red arrows. Mononucleated cells are indicated by green arrows. (b) Cells were cultured on a plastic culture dish and subjected to TRAP staining. TRAP+ osteoclasts are indicated by green arrows. (c) Cells were cultured on a bone slice and subjected to toluidine blue staining after removal of cells. Resorption pits are indicated by red arrows

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2. Remove medium, seed 4–6  104 cells on top of the bone slices. Follow Subheading 3.3 above to culture osteoclasts with 5 ng/ml M-CSF and 10 ng/ml RANKL. 3. Remove half the culture medium and replace with new M-CSF and RANKL every other day. Examine for osteoclast formation under an inverted microscope every day. After 4–5 days, large multinucleated cells will have formed. Stop the culture and fix the cells on the bone slices 4 days after large osteoclasts are seen under the microscope, as described in Subheading 3.3 above (see Notes 11 and 12). 4. Stain the bone slices in the 96-well plates for TRAP activity, as described in Subheading 3.4, but without Mayer’s Hematoxylin counter staining. 5. Quantify osteoclast numbers. 6. Dip bone slices in 0.5% toluidine blue solution for a few seconds to stain the pits and brush the bone slice with a toothbrush with water to remove the cells. 7. Dip bone slices in 0.5% toluidine blue solution again for a few seconds and wipe off the staining solution with a piece of paper. 8. Air dry. 9. Examine resorption pits by turning the slices upside down and viewing them under an inverted microscope. Examples of bone resorption pits are demonstrated in Fig. 1c.

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Notes 1. We typically test different lots of FBS from different suppliers in the osteoclast assay to find the best lot of FBS that supports the maximum number of osteoclasts in standard culture conditions. We then purchase a large amount of the best lot of FBS (2000–3000 ml) and store it at
80  C. 2. We have found that recombinant human M-CSF induces more osteoclasts than recombinant murine M-CSF. Conditioned medium from M-CSF-expressing cells can be used to replace recombinant M-CSF. We use 1:50 dilution of the M-CSFconditioned medium [11]. 3. We make a small amount of osteoclast culture medium with M-CSF or/and RANKL. 4. We have demonstrated that cells from peripheral blood [12], spleen [11], liver and popliteal lymph nodes from adult C57/B6 mice can differentiate to mature osteoclasts using this protocol. If spleen (some osteopetrotic mice do not have bone marrow) is used to culture osteoclasts, mesh the spleen in

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a cell strainer [11]. Seed 2  105 spleen cells (after lysis of red blood cells) in each well of a 96-well plate. 5. We obtain approximately 5  107 total cells per mouse including both legs. 6. We seed different densities of cells to determine the optimal cell numbers at the beginning of each new project, for example when using cells from a new transgenic or knockout mouse. 7. Be careful. We add medium very gently because a high force from addition of medium can disturb cells on the bottom of the well. A vigorous pipetting technique can result in fewer osteoclasts, uneven cell distribution, and huge well-to-well variation. 8. Different amounts of RANKL (1–10 ng/ml) and M-CSF (3–30 ng/ml) can be used to study the responses of cells to osteoclastogenic cytokines, for example to determine if they act synergistically. 9. Typically, osteoclasts are formed 1–2 days after the second RANKL addition. 10. Plates can be stored in
20  C (discard formalin without washing) for months before TRAP staining. 11. Observe osteoclasts daily under an inverted microscope after 4–5 days of culture to determine if there are osteoclasts present on the plastic around the bone slices. These can be seen as large multinucleated cells (Fig. 1a). If there are osteoclasts, wait for another 2–3 days before stopping the experiment to allow time for resorption pits to form. 12. TRAP staining can be done beforehand.

Acknowledgments This work was supported by the following NIH Grants: AR043510 RO1 from NIAMS; AG059775 RO1 and AG049994 from NIA. References 1. Holtrop ME, Raisz LG, Simmons HA (1974) The effects of parathyroid hormone, colchicine, and calcitonin on the ultrastructure and the activity of osteoclasts in organ culture. J Cell Biol 60:346–355 2. Ash P, Loutit JF, Townsend KM (1980) Osteoclasts derived from haematopoietic stem cells. Nature 283:669–670 3. Testa NG, Allen TD, Lajtha LG, Onions D, Jarret O (1981) Generation of osteoclasts in vitro. J Cell Sci 47:127–137

4. Allen TD, Testa NG, Suda T, Schor SL, Onions D, Jarrett O, Boyde A (1981) The production of putative osteoclasts in tissue culture - ultrastructure, formation and behavior. Scan Electron Microsc 347–354 5. Ibbotson KJ, Roodman GD, McManus LM, Mundy GR (1984) Identification and characterization of osteoclast-like cells and their progenitors in cultures of feline marrow mononuclear cells. J Cell Biol 99:471–480 6. Roodman GD, Ibbotson KJ, MacDonald BR, Kuehl TJ, Mundy GR (1985)

Osteoclast Cultures From Mouse Bone Marrow 1,25-Dihydroxyvitamin D3 causes formation of multinucleated cells with several osteoclast characteristics in cultures of primate marrow. Proc Natl Acad Sci U S A 82:8213–8217 7. Lacey DL, Timms E, Tan HL, Kelley MJ, Dunstan CR, Burgess T, Elliott R, Colombero A, Elliott G, Scully S, Hsu H, Sullivan J, Hawkins N, Davy E, Capparelli C, Eli A, Qian YX, Kaufman S, Sarosi I, Shalhoub V, Senaldi G, Guo J, Delaney J, Boyle WJ (1998) Osteoprotegerin ligand is a cytokine that regulates osteoclast differentiation and activation. Cell 93:165–176 8. Wong BR, Rho J, Arron J, Robinson E, Orlinick J, Chao M, Kalachikov S, Cayani E, Bartlett FS 3rd, Frankel WN, Lee SY, Choi Y (1997) TRANCE is a novel ligand of the tumor necrosis factor receptor family that activates c-Jun N-terminal kinase in T cells. J Biol Chem 272:25190–25194 9. Bradley EW, Oursler MJ (2008) Osteoclast culture and resorption assays. Methods Mol Biol 455:19–35

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10. Yao Z, Xing L, Boyce BF (2009) NF-kappaB p100 limits TNF-induced bone resorption in mice by a TRAF3-dependent mechanism. J Clin Invest 119:3024–3034 11. Yamashita T, Yao Z, Li F, Zhang Q, Badell IR, Schwarz EM, Takeshita S, Wagner EF, Noda M, Matsuo K, Xing L, Boyce BF (2007) NF-kappaB p50 and p52 regulate receptor activator of NF-kappaB ligand (RANKL) and tumor necrosis factor-induced osteoclast precursor differentiation by activating c-Fos and NFATc1. J Biol Chem 282:18245–18253 12. Yao Z, Li P, Zhang Q, Schwarz EM, Keng P, Arbini A, Boyce BF, Xing L (2006) Tumor necrosis factor-alpha increases circulating osteoclast precursor numbers by promoting their proliferation and differentiation in the bone marrow through up-regulation of c-Fms expression. J Biol Chem 281:11846–11855

Chapter 30 Hematopoietic Stem Cell Cultures and Assays Benjamin J. Frisch Abstract The adult hematopoietic system is repopulated in its entirety from a rare cell type known as hematopoietic stem cells (HSCs) that reside in the marrow space throughout the skeletal system. Here we describe the isolation and identification of HSCs both phenotypically and functionally. Key words Hematopoietic stem cell, HSC, HSPC, Flow cytometric analysis, FACS, CFC, LTC-IC

1

Introduction A single HSC is capable of repopulating the entire hematopoietic system [1]. This requires both unlimited self-renewal as well as the ability to differentiate into every type of hematopoietic cell. In mammals, hematopoiesis occurs, and HSCs reside, in the marrow cavity of the skeletal system. In humans, direct evaluation of HSC function is very limited. As a surrogate, in vitro assays have been developed to functionally evaluate immature hematopoietic cells [2]. Multiple methods of in vitro analysis have been developed for the identification and quantification of immature hematopoietic cells. The fastest method, flow cytometric analysis, is the only method that can prospectively identify and isolate HSCs; however, it is also the one that provides no functional data. It has however been strongly correlated with repopulating ability and is therefore widely accepted in the field [3, 4]. Colony-forming cell assays (CFCs) are in vitro functional assays that represent the second most rapid method of identifying hematopoietic progenitors. These provide some limited functional analysis, as the ability to form a multilineage colony requires both the ability to differentiate as well as some limited self-renewal. Much more time consuming assays, cobblestone area-forming cells (CAFC) and long-term culture initiating cells (LTC-IC) are used to represent the most primitive hematopoietic cell population that can be functionally assayed

Matthew J. Hilton (ed.), Skeletal Development and Repair: Methods and Protocols, Methods in Molecular Biology, vol. 2230, https://doi.org/10.1007/978-1-0716-1028-2_30, © Springer Science+Business Media, LLC, part of Springer Nature 2021

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in vitro [5]. Both of these assays require more extensive self-renewal capacity than the CFC assay. To date however the ability to serially repopulate myeloablated recipients is the only true way to determine both unlimited self-renewal, and multipotentiality (Fig. 1).

2

Materials

2.1 Immunophenotypic Identification by Flow Cytometric Analysis

1. Antibodies: CD3e, B220, CD11b, GR1, TER119, Flt3, CD48, c-Kit, Sca-1, CD150 (see Notes 1 and 2). 2. FACS staining buffer: 1
phosphate buffered saline (PBS) and 2% fetal bovine serum (FBS). 3. Red blood cell (RBC) lysis buffer: 156 mM NH4Cl, 127 μM EDTA, and 12 mM NaHCO3. 4. Vital dye to distinguish live cells from dead/dying cells (e.g., 40 ,6-diamidino-2-phenylindole [DAPI]). 5. Flow cytometer.

2.2 Colony Forming Cell Assays

1. Biosafety cabinet. 2. 5% CO2 incubator set to 37  C with humidity 95% 3. 35 mm petri dishes that have not been coated for tissue culture 4. 100 mm petri dishes 5. Sterile syringes. 6. Sterile blunt end 16-gauge needle. 7. Semisolid methylcellulose media: Multiple formulations are commercially available (see Table 1)(see Note 3). 8. Complete Iscove’s Minimum Essential Medium (IMEM). IMEM should be supplemented with 2% (by volume) FBS. 9. Red blood cell (RBC) lysis buffer: 156 mM NH4Cl, 127 μM EDTA, and 12 mM NaHCO3. 10. Recombinant CSF-1, SCF, IL-1, and IL-3.

2.3

Coculture Assays

1. Biosafety cabinet. 2. 5% CO2 incubator set to 37  C with humidity 95% 3. 5% CO2 incubator set to 33  C with humidity 95% 4. 96 well flat-bottomed tissue culture plates 5. M2-10B4 cells or primary bone marrow stromal cells. 6. 35 mm petri dishes that have not been coated for tissue culture 7. H5100 cell culture medium (Stem Cell Technologies). 8. 103 M Hydrocortisone sodium hemisuccinate in A-MEM (prepared weekly)

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Fig. 1 Flow cytometric identification of hematopoietic stem and progenitor cell populations. An example gating strategy for the quantification of phenotypic populations of Lineage/Sca1+/ckit+ (LSK) cells, multipotent progenitor 2 (MPP2) cells, multipotent progenitor 3 (MPP3) cells, multipotent progenitor 4 (MPP4) cells, short-term hematopoietic stem cells (ST-HSC), and long-term hematopoietic stem cells (LT-HSC). TOPRO is used as a viability dye in this example. Live cells are negative for TOPRO

9. H4435 methylcellulose containing medium (Stem Cell Technologies). 10. 0.25% trypsin

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Table 1 Cell # per dish, media type, and culture time for various CFC cultures started with total mononuclear bone marrow cells from either mice or humans Culture type

Cell # (per 35 mm dish)

Media

Culture time (days)

CFU-E

2
10

4

M3434/H4434

12

BFU-E

2
10

4

M3434/H4434

7–10

CFU-GM

2
104

M3434/H4434

12

CFU-GEMM

2
10

4

M3434/H4434

12

CFU Pre-B

5
10

4

M3630

7

HPP

4
10

5

M3231/H4535

28

Media is commercially available from Stem Cell Technologies, Vancouver, BC. Media listed as M is for mouse cultures, media listed as H is for human cultures

11. X-ray or gamma-radiation source. 12. All materials listed for CFC assays. 13. Light microscope capable of phase contrast imaging. 14. L-Calc™ software (Stem Cell Technologies cat#28600). 15. Recombinant stem cell factor (SCF), Flt3-Ligand (Flt3-L), and interleukin 11 (IL-11) (PeproTech). 2.4 Competitive Repopulation

1. FACS staining buffer: 1X PBS and 2% FBS. 2. 0.5 mL insulin syringes with 29G beveled needles 3. X-ray or gamma-radiation source. 4. Recipient mice: For mouse bone marrow cells the most commonly used strain is C57bl/6 mice. These mice are available with 2 different alleles of CD45 that are distinguishable by flow-cytometric analysis. CD45.2: WT C57bl/6 mice and CD45.1 (Jackson Labs, Bar Harbor, ME: Strain # 002014). 5. Antibodies for analysis of engraftment can be obtained from multiple commercial sources including eBioscience and BD Biosciences.

3

Methods

3.1 Immunophenotypic Identification by Flow Cytometric Analysis

Putative HSCs can be identified by the expression (or lack thereof) of specific cell surface antigens. This allows for rapid quantification of populations of cells that are otherwise costly and timeconsuming to assay. Moreover, most assays quantify HSCs post facto, therefore flow cytometry is the only method allowing for prospective identification of HSCs and is therefore necessary for any effort attempting to isolate HSCs. The caveat to flow

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cytometric analysis (FACS) is that it is not a functional assay. The cell surface antigen phenotypes reported have been well documented to correlate to a population of cells that is capable of repopulating the hematopoietic system of a myeloablated recipient [1, 3, 4, 6–9]. 1. An appropriate source of cells must be obtained. Depending on the details of each particular experiment the source may vary. Typical sources are cells of the marrow, spleen, or peripheral blood. 2. RBCs should be removed. One recommended method is by the use of RBC lysis buffer. Cells should be suspended in RBC lysis buffer at a concentration of approximately 1
107 cells per mL. They should be incubated at room temperature for 5 min and then promptly washed with an equal volume of FACS staining buffer. 3. Following the removal of RBCs up to 1
107 mononuclear cells should be resuspended in 100 μL FACS staining buffer. 4. An appropriate amount of each antibody should be added to the cell suspensions. Typical amounts are 0.02–0.2 μg per test, though it is recommended to titrate each antibody to determine an optimum amount. 5. Putative long-term repopulating HSCs are identified by the following surface antigens in mice. They are negative for CD3e, B220, CD11b, GR1, TER119, Flt3, and CD48. They are positive for c-Kit, Sca-1, and CD150. 6. Putative short-term repopulating HSCs are identified by the following surface antigens in mice. They are negative for CD3e, B220, CD11b, GR1, TER119, CD48, and CD150. They are positive for c-Kit and Sca-1. 7. Putative multipotent progenitors (MPPs) can be subdivided into three distinct populations whose differentiation is skewed toward specific lineages. These are identified by the following surface antigens in mice [10]. (a) MPP2 are skewed toward the myeloid, erythroid, and megakaryocyte lineages: They are negative for CD3e, B220, CD11b, GR1, TER119, and Flt3. They are positive for c-Kit, Sca-1, CD48, and CD150. (b) MPP3 are skewed toward the myeloid and megakaryocyte lineages: They are negative for CD3e, B220, CD11b, GR1, TER119, Flt3, and CD150. They are positive for c-Kit, Sca-1, and CD48. (c) MPP4 are skewed toward the Lymphoid lineages: They are negative for CD3e, B220, CD11b, GR1, TER119, and CD150. They are positive for c-Kit, Sca-1, Flt3, and CD48.

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8. Putative HSCs in humans are enriched by selecting for the population that is CD34 positive and CD38 negative (see Note 1). 3.2 Colony Forming Cell Assays

Hematopoietic progenitor cell frequencies can be determined by performing colony forming cell assays [11]. Using these assays progenitors such as colony forming unit erythrocytic (CFU-E), blast forming unit erythrocytic (BFU-E), colony forming unit granulocytic and monocytic (CFU-GM), colony forming unit granulocytic, erythrocytic, monocytic, and megakaryocytic (CFU-GEMM), and highly proliferative progenitors (HPP) can be quantified. 1. An appropriate source of cells must be obtained. Depending on the details of each particular experiment the source may vary. Typical sources are cells of the marrow, spleen, or peripheral blood. 2. Red blood cells should be removed. One recommended method is by the use of RBC lysis buffer. Cells should be suspended in RBC lysis buffer at a concentration of approximately 1
107 cells per mL. They should be incubated at room temperature for 5 min and then promptly washed with an equal volume of complete IMEM. 3. Cells should be resuspended in complete IMEM at 10
the final desired concentration. If performing mouse HPP cultures media should be supplemented with 250 ng/mL of recombinant CSF-1 and 50 ng/mL of recombinant SCF, IL-1, and IL-3. 4. For each individual sample a 15 mL conical tube should be filled with 3 mL of methylcellulose containing media if cultures are to be performed in duplicate. Add 4 mL if they are to be performed in triplicate (see Notes 3 and 4). 5. Add the stock solution of cells to the methylcellulose containing media and vortex thoroughly. Add 300 μL if cultures are to be performed in duplicate. Add 400 μL if cultures are to be performed in triplicate. 6. Allow tubes to rest for 5 min at room temperature. 7. Using a blunt end 16 gauge needle attached to a syringe dispense 1.1 mL of cell containing methylcellulose media into each 35 mm dish. 8. Evenly distribute the methylcellulose containing media across the bottom of the dish. 9. Place 2 35 mm sample containing dishes along with 1 35 mm dish containing 3 mL of sterile ultrapure water into 1 100 mm dish and place in the 37 incubator.

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10. After the indicated amount of time for the culture type analyze the cultures by counting the number of colonies present per dish (see Table 1). 3.3

Coculture Assays

HSCs require a specific microenvironment or niche to properly regulate their function. Therefore, since there is no in vitro model that faithfully recapitulates the HSC niche yet developed, in order to evaluate numbers of hematopoietic progenitors that are less mature than those found in CFC assays, a coculture assay is required. There are two cocultures most commonly used to measure the frequency of cell types that act as surrogates for HSCs; these are cobblestone area forming cells (CAFC), and long-term culture initiating cells (LTC-IC) [12–14] (see Note 5). Both use a stromal cell feeder layer to support immature hematopoietic cells however the readout is different. These cultures are somewhat controversial as they do not measure the true functional potential of HSCs to repopulate a marrow, and variations in procedures and stromal cell layers can result in different outcomes from different laboratory environments. The feeder layer used can be either primary bone marrow stromal cells, or a cell line. One well established cell line used for these cultures is M2-10B4 cells. Additionally shorter duration cocultures have been developed primarily for the analysis of HSC supportive capacity of microenvironmental populations. These assays typically use primary stromal cell populations as a feeder layer and sorted murine hematopoietic stem and progenitor cells [15]. 1. Prepare an appropriate stromal layer. If you are using M2-10B4 cells: Inoculate flat-bottomed 96-well tissue culture plates with M2-10B4 cells and maintain at 37  C. When cells have reached confluency irradiate the plates with a 20GY dose of either X-ray or gamma radiation. This serves the purpose of preventing over-proliferation of the feeder layer, while still allowing support for hematopoietic progenitors. 2. Suspend test cell population in H5100 supplemented 1:100 with 103 hydrocortisone sodium hemisuccinate. For bone marrow cells an initial concentration of 8
105 mononuclear cells per mL should be used. If performing a limiting dilution analysis it is recommended that 4–8 different concentrations be used with 12 wells/concentration. 3. 100 μL of cell suspension should be added to each well. 4. Cultures should be maintained at 33  C for 5 weeks with a half media change performed each week in which the nonadherent cells removed with the media are discarded. Aseptic technique must be strictly followed for media changes.

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5. Following 5 weeks in culture CAFCs can be counted. Cobblestone areas are identified by phase contrast microscopy. Cobblestone areas appear as dark-centered circles as they are located below the stromal layer [12]. 6. In order to measure LTC-ICs the cultured cells must be removed from the 96 well plates. Remove the nonadherent cells from the plates with the cells from each well being placed into separate tubes. Trypsin should be used to release the adherent cells from the wells. Following trypsinization adherent cells should be added to the same tubes as the nonadherent cells. CFCs should be set up using the cell population of each individual well of the 96 well plates to seed a 35 mm dish. The CFC protocol should be followed for the setup of these cultures with the use of H4435 methylcellulose containing media. After 18–20 days in culture, dishes should be scored as positive if they contain at least 1 BFU-E, CFU-GM, or CFU-GEMM. If no colonies are present the dish should be scored as negative. 7. If a limiting dilution was performed the number of CAFCs or LTC-ICs can be calculated based on the number of positive wells per dilution and using L-CALC™ software. 3.4 Short-Term Coculture

1. Prepare an appropriate stromal layer. Typically for short-term cocultures the capacity of the stromal layer to support HSCs is what is being investigated, and as such the stromal layer is variable. Rather than in LTC-IC or CAFC assays in which the stromal layer remains constant and the HSCs added to it is variable (see Note 6). 2. Following establishment of an appropriate stromal layer, typically 7–14 days of culture, prepare the desired population of hematopoietic stem and progenitor cells for coculture, typically Lin/Sca-1+/c-Kit+ (LSK) cells sorted from the bone marrow, as described in Subheading 3.1. LSK cells should be added at a concentration of 250 cells/mL, or 500 cells per well of a 6-well culture plate in 2 mL of media. Media should be A-MEM supplemented with 25 ng/mL of SCF, Flt3-L, and IL-11. 3. Following 4 days in culture cells are collected and analyzed. Analysis can include phenotypic definition by flow cytometric analysis as described in Subheading 3.1, colony forming assays as described in Subheading 3.2, or competitive repopulation to define HSC function as described in Subheading 3.5.

3.5 Competitive Repopulation

The best method by which to measure functional HSCs is by their ability to serially transplant myeloablated recipient mice [16– 18]. In mice, congenic strains result in little to no tissue rejection. To transplant human HSCs, however, immune compromised strains of mice must be used to prevent tissue rejection. These

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experiments are timely and costly, leading to the use of the previously described methods as screening tools and competitive repopulation as functional validation. 1. One day before transplantation recipient mice should be irradiated with a dose of 5GY from either an X-ray or gammaradiation source. The recipient mice should be CD45.2 if donor cells are CD45.1 or the recipient mice should be CD45.1 if donor cells are CD45.2 (see Note 7). 2. An appropriate source of donor cells must be obtained. In mice, bone marrow cells are used almost exclusively. 3. An appropriate source of competitor marrow cells must be obtained. Competitor cells should be CD45.2 if donor cells are CD45.1 or the competitor cells should be CD45.1 if donor cells are CD45.2. 4. Donor cells and competitor cells should be resuspended in FACS staining buffer at an appropriate ratio. A donor–competitor ratio of 1:1 or 1:2 is commonly used. Donor and competitor cell mixtures should be resuspended at an appropriate cell concentration so that the total injection volume does not exceed 200 μL. The minimum number of competitor cells transplanted to ensure survival of recipient mice is 1
105. Example: If donor cells are CD45.2 and a ratio of 1:2 donor– competitor is desired then 5x104 CD45.2 donor cells and 1
105 CD45.1 competitor cells will be combined and resuspended in 100–200 μL of FACS staining buffer. 5. Recipient mice should be irradiated a second time with a dose of 5GY 24 h following the first dose of radiation. The total dose of radiation should be 10GY to achieve a lethal dose. 6. Immediately following the second dose of radiation the donor and competitor cell mixture should be intravenously injected. This can be achieved by injecting directly into the lateral tail veins, or by injecting into the retroorbital sinus. If injecting by tail vein warming recipient mice with a heat lamp and using an appropriate restraining cone or tube is highly recommended. If injecting by retroorbital sinus an anesthetic or sedative must be used to avoid permanent damage to the eye (see Note 8). 7. Following transplantation the level of engraftment can be determined by flow cytometric analysis of the peripheral blood using the appropriate CD45.1 and CD45.2 antibodies. Analysis should be performed for short-term engraftment, 4–12 weeks, and long-term engraftment, greater than 16 weeks. Multilineage engraftment of donor cells should also be assessed using the following antibodies: CD3e for T-cell lineage, B220 for B-cell lineage, and CD11b for myeloid lineage.

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8. To truly measure long-term engraftment secondary transplantations should be performed. Marrow from primary recipients should be obtained and injected into secondary recipients that have been lethally irradiated as previously described. Secondary recipients should be of the same strain as primary recipients. Marrow cells obtained should not be transplanted with competitor cells. At least 1
105 whole marrow cells from the original donors should be transplanted. Successful multilineage engraftment of a secondary recipient is the gold standard for determination of a functional long-term repopulating HSC.

4

Notes 1. Proper panel design is critical to the success of flow cytometric analysis. Each antibody should be titrated to ensure optimum concentrations are being used to maximize the signal to background ratio. 2. All conjugated antibodies are prone to contamination, and degradation following exposure to room temperature and/or light (particularly tandem dyes such as PE-Cy7). Therefore, it is good practice to use all conjugated antibodies on ice, in a biosafety cabinet with the lights off. 3. To limit the number of freeze–thaw cycles, methylcellulose containing media should be aliquoted into 3 or 4 mL volumes depending on whether you are performing your assay in duplicate or triplicate respectively. 4. Methylcellulose containing media should be thawed at room temperature the day of use, or at 4  C overnight. 5. Bacterial or fungal contamination can become an issue in LTC-IC assays as the cultures are long and the media used contains no antibiotic or antifungal treatment. Therefore, the strictest aseptic technique must be maintained throughout the culture. 6. For the establishment of stromal cell feeder layers culture under hypoxic conditions, 2% O2 may be preferred depending on the cell type being used. For example, primary MSCs will become established more quickly under hypoxic conditions. 7. When irradiating mice it is recommended to immobilize them to prevent variable doses of radiation that can lead to variable engraftment rates. 8. Both tail-vein injections and injections into the retroorbital sinus require considerable skill. It is highly recommended that they be practiced prior to the use of any experimental animals.

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Acknowledgments This work was supported by NIH grants UL1 TR002001 Pilot Award to BJF. References 1. Osawa M et al (1996) Long-term lymphohematopoietic reconstitution by a single CD34low/negative hematopoietic stem cell. Science 273(5272):242–245 2. Purton LE, Scadden DT (2007) Limiting factors in murine hematopoietic stem cell assays. Cell Stem Cell 1(3):263–270 3. Matsuzaki Y et al (2004) Unexpectedly efficient homing capacity of purified murine hematopoietic stem cells. Immunity 20 (1):87–93 4. Yilmaz OH, Kiel MJ, Morrison SJ (2006) SLAM family markers are conserved among hematopoietic stem cells from old and reconstituted mice and markedly increase their purity. Blood 107(3):924–930 5. van Os R, Kamminga LM, de Haan G (2004) Stem cell assays: something old, something new, something borrowed. Stem Cells 22 (7):1181–1190 6. Okada S et al (1992) In vivo and in vitro stem cell function of c-kit- and Sca-1-positive murine hematopoietic cells. Blood 80 (12):3044–3050 7. Spangrude GJ, Heimfeld S, Weissman IL (1988) Purification and characterization of mouse hematopoietic stem cells. Science 241 (4861):58–62 8. Morrison SJ, Weissman IL (1994) The longterm repopulating subset of hematopoietic stem cells is deterministic and isolatable by phenotype. Immunity 1(8):661–673 9. Morrison SJ et al (1997) Identification of a lineage of multipotent hematopoietic progenitors. Development 124(10):1929–1939 10. Pietras EM et al (2015) Functionally distinct subsets of lineage-biased multipotent

progenitors control blood production in Normal and regenerative conditions. Cell Stem Cell 17(1):35–46 11. Bradley TR, Metcalf D (1966) The growth of mouse bone marrow cells in vitro. Aust J Exp Biol Med Sci 44(3):287–299 12. Ploemacher RE et al (1989) An in vitro limiting-dilution assay of long-term repopulating hematopoietic stem cells in the mouse. Blood 74(8):2755–2763 13. Ploemacher RE et al (1991) Use of limitingdilution type long-term marrow cultures in frequency analysis of marrow-repopulating and spleen colony-forming hematopoietic stem cells in the mouse. Blood 78(10):2527–2533 14. Sutherland HJ et al (1989) Characterization and partial purification of human marrow cells capable of initiating long-term hematopoiesis in vitro. Blood 74(5):1563–1570 15. Frisch BJ et al (2019) Aged marrow macrophages expand platelet-biased hematopoietic stem cells via Interleukin1B. JCI Insight 5: e124213 16. Ford CE et al (1956) Cytological identification of radiation-chimaeras. Nature 177 (4506):452–454 17. McCulloch EA, Till JE (1960) The radiation sensitivity of normal mouse bone marrow cells, determined by quantitative marrow transplantation into irradiated mice. Radiat Res 13:115–125 18. Szilvassy SJ et al (1990) Quantitative assay for totipotent reconstituting hematopoietic stem cells by a competitive repopulation strategy. Proc Natl Acad Sci U S A 87(22):8736–8740

INDEX A Adipocytes ............................................48, 379, 380, 384, 391, 392, 408, 437 Aggrecan..........................................................4, 117, 283, 326, 329, 331 Alcian Blue.................................................... 33, 121, 127, 131, 286–288, 295, 296, 298, 300, 394, 403, 408, 409 Alizarin Red ................................................ 218, 234, 238, 247, 272, 284, 382, 389, 390, 403, 408, 429, 431 Alkaline phosphatase (AP/ALP) ......................6, 52, 117, 123, 130, 135, 234, 248, 283, 285, 370, 388, 426–428, 430, 434 Analgesia buprenorphine............................ 69, 78, 99, 152, 404 Anesthesia isoflurane .............................................. 65, 94, 95, 99, 100, 106, 107, 382, 402 ketamine ........................................... 65, 78, 100, 101, 152, 156, 157, 382, 402–404 xylazine .....................................................65, 382, 402 Angiogenesis........................................111, 151–154, 346 Antibody primary.................................................. 126, 156, 160, 163, 164, 221, 226, 310, 316, 320, 331, 338, 340, 341, 344, 348, 352, 353, 355, 402 secondary .............................................. 121, 126, 156, 160, 163, 221, 226, 328, 333, 338–341, 343, 348, 350, 353, 354 Apoptosis ................................................................ 44, 117 Ascorbic acid .............................................. 142, 144, 422, 426, 428, 432, 440

B Beta-galactosidase .................................48, 120, 131, 285 Beta-glycerophosphate............................... 144, 381, 422, 429, 433, 440 Biomechanical tests rigidity............................................................... 29, 201 stiffness ............................................................. 29, 201 strength...................................................................... 29 torsion........................................28, 29, 79–81, 84, 85 toughness................................................................... 29 Biotin ...................................................341, 343, 350, 353

Bone calvaria ............................................... 4, 426, 429, 438 cortical ............................................12, 20, 21, 46, 69, 152, 154, 155, 158, 162, 170, 181, 184, 186, 187, 190, 192, 194, 203, 219, 234, 285, 290, 293, 307, 397 femur........................................ 81, 84, 163, 203, 253, 289, 290, 330, 351, 404 fibula ............................................................... 171, 253 formation ........................................ 4–6, 8, 10, 18–20, 29, 63, 70, 71, 76, 117, 151, 284, 305, 384, 393, 398, 425–427, 449 marrow....................................... 9, 49, 106, 117, 151, 152, 226, 233, 279, 285, 290, 297, 312, 318, 319, 330, 363, 379–394, 399, 400, 406, 411, 437, 438, 440, 453, 457–464, 469, 473–475 mineralization........................................ 304, 426, 427 patella....................................................................... 293 remodeling ................................10, 11, 303–320, 457 tibia .................................................94, 171, 203, 253, 290, 342, 404 trabecular .......................................... 21, 94, 117, 170, 173, 184, 186, 187, 190, 234, 285, 290, 296, 335, 348, 363, 384 vertebra ........................................................................ 4 Bone marrow stromal cells (BMSC) ...................... 46, 49, 117, 379–394, 438, 440, 441, 453, 468, 473 Bone morphogenetic proteins (BMPs)................. 5, 9, 76 Bone sialoprotein (BSP) ............................................... 426 Bromodeoxyuridine (BrdU)....................... 142, 358, 363

C Calcium.......................................... 76, 81, 142, 217, 233, 255, 283, 284, 304, 408 Callus bone .................................................18, 21, 23, 29, 34 cartilage............................................18, 25, 27, 29, 32 Cartilage articular ............................................. 8, 91–94, 97, 99, 283, 290, 296, 297, 343, 363, 399 degeneration........................................................ 92–94 fibrillation .................................................................. 91 formation ...................................................4, 116, 380, 385, 389, 408

Matthew J. Hilton (ed.), Skeletal Development and Repair: Methods and Protocols, Methods in Molecular Biology, vol. 2230, https://doi.org/10.1007/978-1-0716-1028-2, © Springer Science+Business Media, LLC, part of Springer Nature 2021

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AND

REPAIR: METHODS

AND

Cartilage (cont.) growth plate ......................................... 4, 8, 283, 285, 290, 293, 296, 297, 364, 415–422 mineralization................................................. 144, 422 Cell culture ...........................................50, 119, 120, 140, 144, 146, 402, 418, 419, 428, 430–432, 434, 440, 451, 452, 461, 467–475 Chondrocyte differentiation....................... 5–7, 116, 139, 140, 142 hypertrophy ........................................ 5, 6, 8, 10, 141, 348, 354, 355, 364 proliferation ................. 5–7, 139, 141, 346, 357, 363 Chondrogenesis ..........................................................7, 20 Collagens type I .................................................. 9, 116, 425, 426 type II ....................................... 4, 117, 283, 374, 416 type IX ......................................................................... 4 type X..................................................... 117, 283, 416 type XI ......................................................................... 4 Colony-forming unit fibroblasts (CFU-Fs)....... 379, 383, 385, 391, 394, 401 Cryofilm...................................................... 261, 265–269, 272–275, 278–280 Cryostat ..............................................221, 226, 267–269, 271, 272, 274, 286, 295, 318, 349, 351, 355

D Decalcification ethylenediaminetetraacetic acid (EDTA) .............. 155, 220, 284, 297, 300, 306, 307, 327, 350, 359 formic acid ............................................ 284, 286, 297, 299, 300, 307, 318 Dentin matrix acidic phosphoprotein (DMP1) ........... 48, 49, 304, 326, 328, 331, 333, 335 Differentiation.......................................... 4–9, 17, 18, 33, 41, 48, 103, 116, 117, 119, 139–142, 326, 327, 346, 357, 380, 383, 384, 386, 389–392, 398–401, 407–410, 415, 421, 422, 425, 437–441, 443, 445, 449, 458, 471 Dissection ...............................................34, 80, 119, 120, 123, 124, 129, 145, 147, 156, 174, 175, 191, 233, 235, 238, 239, 252, 285, 300, 319, 329, 349, 371, 418, 427, 440, 450

E EdU ...................................................................... 357–365 Embedding ...........................................29, 175, 176, 178, 179, 191, 218, 220, 223, 224, 231–236, 239–240, 249, 252, 256, 264–266, 274, 278, 279, 284–286, 289, 290, 292, 301, 311, 316, 318, 349, 351, 362, 404 Endochondral............................................. 4–6, 8, 19, 20, 22, 29, 64, 70, 71, 139, 357, 398

PROTOCOLS Ethanol (EtOH)................................ 106, 107, 121, 122, 124, 126–128, 152, 155, 156, 159, 161, 174, 176, 224, 232, 233, 235, 237–248, 250, 252, 253, 265, 268, 269, 271, 285–288, 290, 291, 295, 297–299, 306, 311, 316–319, 327, 328, 338, 340, 349–359, 362, 369, 371, 372, 382, 392, 402–404, 416–419, 422, 427, 429, 430, 459, 460 Extracellular matrix (ECM)............................4, 151, 283, 284, 337, 387, 394, 407, 409, 416, 425, 434, 438

F Fibroblast growth factor (FGF) ............... 5, 7, 8, 76, 402 Fibrous Tissue (FT) ............................................... 21, 391 Fixative 10% neutral buffered formalin (NBF) .................. 284, 285, 306, 310, 316, 350, 352, 390, 459 4% paraformaldehyde (PFA)......................... 120, 155, 174, 176, 220, 265, 284, 349, 369, 403, 407, 409, 411 Fluorescein .................................122, 123, 129, 132, 135 Fluorescence-activated cell sorting (FACS)......... 46, 383, 388, 389, 394, 468, 469, 471, 475 Forceps........................................... 64, 78, 80–84, 94, 95, 97, 106, 119, 125, 126, 144–146, 152, 155–157, 162, 174, 206, 207, 235, 242, 285, 286, 288, 309, 313, 330, 351, 382, 391, 392, 401, 404, 416, 418, 427, 429, 440, 458, 460 Fracture closed ................................................................... 63–71 open ........................................................................... 64 stabilized ..............................................................63, 67

G Graft.................................................................75–87, 109, 152, 156–158, 160, 162, 403–405, 411

H Hematopoietic stem cell (HSC) competitive repopulation...................... 469, 474, 475 long-term HSC (LT-HSC)........................... 467, 469, 471, 473, 475 short-term HSC (ST-HSC).......................... 469, 471, 474, 475 Hind limb ........................................................80, 83, 108, 111, 125, 266, 279, 328, 441 Histology demineralized ................................ 283–301, 311, 312 paraffin and frozen .................................................. 362 undecalcified frozen ................................................ 260 undecalcified plastic ................................................ 234 Histomorphometry ..........................................25, 99, 101

SKELETAL DEVELOPMENT Hybridization ....................................................12, 18, 29, 33–35, 99, 118, 120, 122, 128, 129, 132, 135, 140, 260, 269, 284, 300, 306, 367–375

I Immunofluorescence .......................................... 118, 121, 126, 133, 135, 217, 326, 327, 331, 333, 338, 347, 352, 353 Immunohistochemistry (IHC)............................... 29, 34, 99, 219, 259, 260, 269, 275, 306, 308, 310, 316, 317, 337, 341–343, 350, 382 Indian hedgehog (Ihh) ................................................. 6–8 In situ hybridization whole mount ..........................................118, 367–375 Intramembranous.................................... 4, 5, 8, 357, 398

J Joints.......................................................4, 5, 7, 8, 33, 68, 80, 91–93, 96–101, 141, 145, 147, 208, 213, 222, 283, 289, 290, 293, 351, 404, 419, 460

K Kidney.........................................151, 154, 157, 158, 162

L Lac-Z ..................................................................... 52, 118, 120, 125–126, 131, 133 Laser microdissection (LMD) ............................ 259–261, 267, 270, 271 Lead chromate................................................................. 27 Light-sheet microscopy ....................................... 218, 225 Lineage tracing..................................................9, 52, 118, 121, 127, 131, 325–335

M Macrophage Colony-Stimulating Factor (M-CSF)...... 10, 458, 459, 461–464 Matrix Metalloproteinase 13 (MMP13)...................6, 92, 305, 308, 310, 316, 416 Media .........................................119, 140, 231, 269, 284, 309, 404, 415, 428, 439, 451, 459, 460, 468 Medial collatoral ligament (MCL) ...........................93, 98 Meniscus ..............................................92–94, 97, 98, 290 Mesenchymal condensation......................................5, 374 Mesenchymal stem cell (MSCs) .....................48, 76, 379, 399, 438 Mesenchyme................................................ 117, 131, 374 Methyl methacrylate (MMA) ............................. 233–236, 238–241, 244, 245, 247, 249, 250, 253, 284 Microarray ........................................................ 33–35, 390 Microcomputed tomography (MicroCT) ............. 29, 85, 169–197, 208, 306

AND

REPAIR: METHODS

AND

PROTOCOLS Index 481

Microtome frozen sections......................................................... 295 paraffin sections ....................................................... 352 Microtomy .................................................. 155, 159, 231, 232, 234, 236, 241, 249, 250, 253, 255, 266, 284–286, 290, 294, 295, 311, 316, 349, 350, 352 Mineralization .............................................. 6, 20, 25, 40, 142, 144, 191, 284, 304, 305, 389, 422, 426–428 Mouse adult ...................................... 152, 300, 358, 362, 453 conditional mutant........................................... 45, 132 Cre ..................................................... 39–52, 326, 328 embryo.....................................................39, 131, 133, 141, 145, 156, 360 floxed ........................................40, 42–45, 47–49, 52, 132, 415, 421, 427, 433 postnatal ...................................................... 9, 40, 141, 331, 332, 363, 418, 427 reporter ........................................................ 40, 45, 46, 48–50, 52, 132, 348, 388 transgenic.............................................. 40, 42, 46, 49, 105, 106, 132, 139, 327, 415, 464

O Organ culture ...................... 22, 115–135, 140, 142, 457 Ossicle ...........................................................384, 391–393 Osteoarthritis ....................................12, 33, 91–101, 346 Osteoblast differentiation.........................................6–9, 357, 444 proliferation .............................. 6, 346, 357, 425, 449 Osteocalcin (Bglap)....................................................... 426 Osteoclast differentiation................................................. 439, 458 proliferation ............................................................. 458 Osteocytes ..............................................9, 10, 44, 46, 48, 49, 51, 117, 303–320, 332, 335, 437, 439 Osteopontin (Spp1) ..................................................6, 426 Osteotomy .................................................................64, 82 Osterix (Osx/Sp7) ............................................................ 8

P Parathyroid hormone related-peptide (PTHrP) . 5–7, 10, 305 PECAM ...............................................154, 156, 159, 163 Perichondrium (PC) ...................... 8, 140, 147, 154, 162 Phosphate buffered saline (PBS).......................... 78, 120, 144, 152, 176, 205, 219, 239, 285, 305, 327, 349, 359, 368, 388, 402, 415, 428, 439, 450, 459, 468 Proliferation.......................................... 4–7, 44, 116, 139, 141, 142, 346, 357–365, 384, 387, 393, 398, 400, 406, 411, 425, 437, 444, 449

SKELETAL DEVELOPMENT

482 Index

AND

REPAIR: METHODS

AND

Protein .........................................5, 18, 40, 76, 105, 123, 140, 151, 218, 248, 283, 306, 325, 337, 347, 358, 425, 444, 449, 458

R Radiography ............. 21–24, 35, 64, 65, 83, 85, 92, 235 Rat.........................................................23, 34, 48, 64, 65, 68, 70, 92, 140, 156, 160, 192, 266, 273, 279, 281, 426 Real-time/quantitative PCR (RT-PCR/qPCR) .......... 45, 305, 306, 308, 309, 314, 315, 320, 390, 420, 428, 432 Receptor activated nuclear factor kappa-B ligand (RANKL) ....................................... 6, 10, 457–464 Renal capsule ........................................................ 151–154 Ribonucleases (RNase) ....................................... 122, 260, 319, 369–372, 374 Ribonucleic acid (RNA) isolation ............................................. 33, 99, 312, 421 mRNA....................................................18–20, 33, 34, 140, 142, 260, 306, 312, 367, 368, 430, 432, 434 Riboprobes ................................................. 122, 128, 130, 135, 367, 369, 374 Runt-related transcription factor 2 (Runx2) ..............7, 8, 92, 326, 327, 331, 333, 335, 342, 390

S Scaffold ....................................................... 25, 76, 77, 83, 123, 131, 132, 217, 382, 383, 389, 391–393 Scalpel ...................................................64, 79, 82, 95, 96, 98, 106, 108, 119, 124, 175, 235, 286, 382, 392, 401, 405, 416, 419 Scissors ..................................................64, 78, 80, 82, 83, 94, 95, 98, 106, 120, 124, 125, 133, 152, 155, 158, 174, 175, 235, 285, 288, 289, 309, 329, 330, 349, 382, 392, 404, 416, 419, 427, 429, 440, 441, 450, 452, 458, 460 Sectioning ........................................................29, 99, 140, 147, 163, 217, 218, 231–256, 268, 284–286, 290, 292–295, 300, 307, 309, 355 Skeletal development .......................................... 3–12, 17, 151–154, 367, 425, 437 Skeletal repair allograft...................................................................... 76 autograft .................................................................... 75 femur fracture............................................................ 24 tendon repair .......................................................4, 116 Skeletal staining Alcian Blue/Hematoxylin/Orange G ......... 286, 287, 295–297, 300 Alizarin Red S....................... 234, 247, 390, 403, 431 alkaline phosphatase.............................. 248, 284, 430

PROTOCOLS Picrosirius Red ..............................288, 296, 298, 299 Safranin-O/Fast Green..............................32, 33, 296 tartrate-resistant acid phosphatase (TRAP).......... 234, 247, 248, 279, 284, 288, 299, 300, 310, 317, 320, 458, 462, 464 trichrome (Goldner’s)............................................. 284 Von Kossa ...................................................... 234, 236, 243–245, 248, 256, 284, 382, 389 Skeletal stem cell (SSC) ....................................10, 18, 51, 142, 357, 358, 363, 380, 383, 399, 401 Skeleton ..................................................4, 10, 12, 17, 40, 105–111, 116, 141, 199, 283, 319, 357, 359, 437 Sry-box factor 9 (Sox9) ........................................ 5, 7, 10, 116, 118, 374, 391, 401, 416, 420 Stromal .......................................................................... 438 Surgery..................................................25, 65, 68, 70, 71, 87, 92–94, 96, 100, 101, 107–111, 162, 252, 392, 393, 400, 403, 410 Suture....................................................67, 69, 79, 82, 83, 95, 97, 99, 106, 108, 110, 111, 155, 158, 159, 255, 402, 404, 410, 411, 440

T Tissue clearing .............................................. 52, 118, 121, 126, 218, 227, 309, 319 Transfection................................................................... 139 Transplantation .................................... 76, 106, 151–154, 380, 382, 383, 385, 388–391, 394, 398, 403, 475, 476

V Vascular endothelial growth factor (VEGF) ..............6, 76 Vascularization..................................................... 141, 151, 152, 155, 159, 162, 398 Vessel....................................... 5, 6, 9, 18, 22, 27, 28, 64, 111, 133, 134, 151, 152, 154, 159, 160, 162, 218, 397, 421, 432

W Whole mount ............................................. 118, 120–122, 125–128, 132–135, 219, 367–375

X X-gal staining................................................................. 118 X-ray............................................... 22, 23, 25, 29, 65, 68, 69, 71, 79, 81, 83, 85, 169, 170, 172–174, 192, 393, 469, 470, 473, 475 Xylene ................................................ 155, 159, 233, 234, 238, 243–247, 253, 256, 268, 286, 295, 297–299, 306, 311, 316, 317, 338, 340, 350, 353, 359