Osteoporosis

Osteoporosis Methods in Molecular Biology John M. Walker, Series Editor 457. Membrane Trafficking, edited by Ales Van...

Author: Jennifer J. Westendorf

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Osteoporosis

Methods in Molecular Biology John M. Walker, Series Editor 457. Membrane Trafficking, edited by Ales Vancura, 2008 456. Adipose Tissue Protocols, Second Edition, edited by Kaiping Yang, 2008 455. Osteoporosis, edited by Jennifer J. Westendorf, 2008 454. SARS- and Other Coronaviruses: Laboratory Protocols, edited by Dave Cavanagh, 2008 453. Bioinformatics, Volume 2: Structure, Function, and Applications, edited by Jonathan M. Keith, 2008 452. Bioinformatics, Volume 1: Data, Sequence Analysis, and Evolution, edited by Jonathan M. Keith, 2008 451. Plant Virology Protocols: From Viral Sequence to Protein Function, edited by Gary Foster, Elisabeth Johansen, Yiguo Hong, and Peter Nagy, 2008 450. Germline Stem Cells, edited by Steven X. Hou and Shree Ram Singh, 2008 449. Mesenchymal Stem Cells: Methods and Protocols, edited by Darwin J. Prockop, Donald G. Phinney, and Bruce A. Brunnell, 2008 448. Pharmacogenomics in Drug Discovery and Development, edited by Qing Yan, 2008 447. Alcohol: Methods and Protocols, edited by Laura E. Nagy, 2008 446. Post-translational Modification of Proteins: Tools for Functional Proteomics, Second Edition, edited by Christoph Kannicht, 2008 445. Autophagosome and Phagosome, edited by Vojo Deretic, 2008 444. Prenatal Diagnosis, edited by Sinhue Hahn and Laird G. Jackson, 2008 443. Molecular Modeling of Proteins, edited by Andreas Kukol, 2008 442. RNAi: Design and Application, edited by Sailen Barik, 2008 441. Tissue Proteomics: Pathways, Biomarkers, and Drug Discovery, edited by Brian Liu, 2008

440. Exocytosis and Endocytosis, edited by Andrei I. Ivanov, 2008 439. Genomics Protocols, Second Edition, edited by Mike Starkey and Ramnanth Elaswarapu, 2008 438. Neural Stem Cells: Methods and Protocols, Second Edition, edited by Leslie P. Weiner, 2008 437. Drug Delivery Systems, edited by Kewal K. Jain, 2008 436. Avian Influenza Virus, edited by Erica Spackman, 2008 435. Chromosomal Mutagenesis, edited by Greg Davis and Kevin J. Kayser, 2008 434. Gene Therapy Protocols: Volume 2: Design and Characterization of Gene Transfer Vectors, edited by Joseph M. LeDoux, 2008 433. Gene Therapy Protocols: Volume 1: Production and In Vivo Applications of Gene Transfer Vectors, edited by Joseph M. LeDoux, 2008 432. Organelle Proteomics, edited by Delphine Pflieger and Jean Rossier, 2008 431. Bacterial Pathogenesis: Methods and Protocols, edited by Frank DeLeo and Michael Otto, 2008 430. Hematopoietic Stem Cell Protocols, edited by Kevin D. Bunting, 2008 429. Molecular Beacons: Signalling Nucleic Acid Probes, Methods and Protocols, edited by Andreas Marx and Oliver Seitz, 2008 428. Clinical Proteomics: Methods and Protocols, edited by Antonia Vlahou, 2008 427. Plant Embryogenesis, edited by Maria Fernanda Suarez and Peter Bozhkov, 2008 426. Structural Proteomics: HighThroughput Methods, edited by Bostjan Kobe, Mitchell Guss, and Huber Thomas, 2008 425. 2D PAGE: Sample Preparation and Fractionation, Volume 2, edited by Anton Posch, 2008

Osteoporosis Methods and Protocols

Jennifer J. Westendorf Editor Mayo Clinic, Department of Orthopedic Surgery Rochester, MN, USA

Editor Jennifer J. Westendorf Mayo Clinic Department of Orthopedic Surgery Rochester, MN USA [email protected]

Series Editor John M. Walker University of Hertfordshire Hatfield, Hertz UK

ISBN: 978-1-58829-828-7 e-ISBN: 978-1-59745-104-8 DOI: 10.1007/978-1-59745-104-8 Library of Congress Control Number: 2008920309 © 2008 Humana Press, a part of Springer Science+Business Media, LLC All rights reserved. This work may not be translated or copied in whole or in part without the written permission of the publisher (Humana Press, 999 Riverview Drive, Suite 208, Totowa, NJ 07512 USA), except for brief excerpts in connection with reviews or scholarly analysis. Use in connection with any form of information storage and retrieval, electronic adaptation, computer software, or by similar or dissimilar methodology now known or hereafter developed is forbidden. The use in this publication of trade names, trademarks, service marks, and similar terms, even if they are not identified as such, is not to be taken as an expression of opinion as to whether or not they are subject to proprietary rights. While the advice and information in this book are believed to be true and accurate at the date of going to press, neither the authors nor the editors nor the publisher can accept any legal responsibility for any errors or omissions that may be made. The publisher makes no warranty, express or implied, with respect to the material contained herein. Cover Illustration: Background: Provided by Dr. Stefan Judex (Chapter 22, Fig. 2). Foreground: Provided by Dr. Ralph Muller (Chapter 19, Fig. 1). Printed on acid-free paper 9 8 7 6 5 4 3 2 1 springer.com

Preface

Osteoporosis is a disease characterized by reduced bone mass, quality, and strength, changes in skeletal micro-architecture, and increased fracture risk. An astonishing number of individuals have osteoporosis throughout the world, including 75 million people in the United States, Europe, and Japan alone (1). An estimated 30–50% of women and 15–30% of men will suffer on osteoporosis-related fracture in their lifetimes (1). Recovery time from osteoporotic fractures is lengthy, often includes terminal nursing home care, and places enormous financial burdens on society, health-care systems, businesses, families, and individuals. Thus, osteoporosis has a broad impact on human health. Osteoporosis, which literally means “porous bone,” is often referred to as the silent disease because symptoms are not noticed until a fracture occurs. A tremendous comprehensive effort has been made in the last two decades to improve diagnostic methods, increase awareness, and identify treatments for osteoporosis. Known causes of osteoporosis include aging, genetics, hormonal imbalances (e.g., estrogen deficiency, hyperparathyroidism), environmental factors (e.g., diet, nutrition, smoking and exercise), and medical drugs (e.g., corticosteroids and chemotherapy). Many advances have occurred in recent years because of novel research methodologies that allow scientists to study bone cells and tissues at the cellular and molecular levels and in a variety of models. The goal of this textbook is to provide a platform for leading scientists from around the world to share their protocols for studying bone biology. We begin with in vitro models to study bone cells and tissues. The emphasis is placed on osteoblasts (bone-forming cells) and osteoclasts (bone-resorbing cells), but it is recognized that other bone marrow–derived cells, notably adipocytes and hematopoietic cells, have major influences on the activities of osteoblasts and osteoclasts and thus contribute to the development and retention of bone mass. The second section details several in vivo rodent models that can be utilized for drug testing, tissue engineering, and studying osteoporosis in either gender. The third section contains three viral gene delivery protocols for studying the roles of specific gene products in bone cells. The fourth section includes relatively new state-of-the art molecular techniques that can be utilized to assess single genes or for global genomic analysis. Section five conv

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Preface

sists of protocols for powerful imaging techniques that allow for in situ observations of bone cells and tissues. The sixth and final section describes methods to analyze bone responses to mechanical stimuli. Many beautiful color reproductions of figures present in the textbook can be found on the compact disk (CD) that accompanies it. I hope that this Methods in Molecular Biology: Osteoporosis book will help both new and experienced experimentalists studying bone biology and osteoporosis. Of course, all potential topics could not be covered in one textbook. Thus, I refer the readers to another book in the Molecular Methods in Medicine series, Bone Research Protocols, edited by Helfrich and Ralston (2), wherein additional techniques and alternative descriptions to some of the topics covered in this volume are described. A second excellent book, Handbook of Histology Methods for Bone and Cartilage, edited by An and Martin (3), focuses on histology and histomorphometry techniques. Thank you to the authors who contributed their techniques and pointers to this manual. It was a pleasure to work with them. I learned a lot! Thank you Julia Hutchcroft at the University of Minnesota and Teresa Hoff at the Mayo Clinic for your administrative assistance. Finally, thank you John Walker and Humana Press for your patience and guidance and for giving me the opportunity to edit this textbook. Jennifer J. Westendorf, PhD

1. International Osteoporosis Foundation. www.iofbonehealth.org/facts-andstatistics.html 2. Helfrich, M. H., Ralston S. T. (2003) Molecular Methods in Medicine: Bone Research Protocols. Humana Press, Totowa, NJ. 3. An, Y. A., Martin, K. L. (2003) Handbook of Histology Methods for Bone and Cartilage. Humana Press, Totowa, NJ.

Contents

List of Color Plates ..........................................................................................

xi

Part I In Vitro Models 1

Isolation and Culture of Rodent Osteoprogenitor Cells .............................................................................. Thomas A. Owen and Lydia C. Pan

2

Osteoclast Culture and Resorption Assays ........................................... Elizabeth W. Bradley and Merry Jo Oursler

3

Assessing New Bone Formation in Neonatal Calvarial Organ Cultures ....................................................................... Khalid S. Mohammad, John M. Chirgwin, and Theresa A. Guise

4

Detection of Apoptosis of Bone Cells In Vitro ....................................... Teresita Bellido and Lilian I. Plotkin

3

19

37

51

Part II In Vivo Rodent Models 5

6

7

In Vivo Parathyroid Hormone Treatments and RNA Isolation and Analysis ............................................................ Xin Li, Ling Qin, and Nicola C. Partridge Assessment of Bone Formation Capacity Using In vivo Transplantation Assays: Procedure and Tissue Analysis ............................................................... Basem M. Abdallah, Nicholas Ditzel, and Moustapha Kassem

79

89

Ossicle and Vossicle Implant Model Systems ........................................ 101 Glenda J. Pettway and Laurie K. McCauley vii

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Contents

8

Utility of the Ovariectomized Rat as a Model for Human Osteoporosis in Drug Discovery ............................................................. 111 Yogendra P. Kharode, Michael C. Sharp, and Peter V.N. Bodine

9

Orchidectomy Models of Osteoporosis .................................................. 125 Stéphane Blouin, Hélène Libouban, Marie Françoise Moreau, and Daniel Chappard

Part III

Methods of Gene Delivery

10

Gene Delivery by Adenoviruses .............................................................. 137 Renny T. Franceschi and Chunxi Ge

11

Lentivirus Delivery of shRNA Constructs into Osteoblasts ................ 149 Marc N. Wein, Dallas C. Jones, and Laurie H. Glimcher

12

Gene Delivery by Retroviruses ............................................................... 157 Valerie Deregowski and Ernesto Canalis

Part IV Molecular and Genetic Techniques 13

Chromatin Immunoprecipitation Assays: Application of ChIP-on-Chip for Defining Dynamic Transcriptional Mechanisms in Bone Cells ........................................... 165 Margaretha van der Deen, Mohammad Q. Hassan, Jitesh Pratap, Nadiya M. Teplyuk, Daniel W. Young, Amjad Javed, Sayyed K. Zaidi, Jane B. Lian, Martin Montecino, Janet L. Stein, Gary S. Stein, and Andre J. van Wijnen

14

Identification of Transcription Factor Target Genes by ChIP Display ....................................................................................... 177 Artem Barski, Steven Pregizer, and Baruch Frenkel

15

Application of the Laser Capture Microdissection Technique for Molecular Definition of Skeletal Cell Differentiation In Vivo ............................................................................. 191 Dafna Benayahu, Rina Socher, and Irena Shur

16

Quantitative Trait Loci Mapping ........................................................... 203 Dong-Hai Xiong, Jian-Feng Liu, Yan-Fang Guo, Yan Guo, Tie-Lin Yang, Hui Jiang, Yuan Chen, Fang Yang, Robert R Recker, and Hong-Wen Deng

Contents

ix

Part V Imaging Techniques 17

In Situ Nuclear Organization of Regulatory Machinery ..................... 239 Shirwin M. Pockwinse, Sayyed K. Zaidi, Ricardo F. Medina, Rachit Bakshi, Krishna P. Kota, Syed A. Ali, Daniel W. Young, Jeffery A. Nickerson, Amjad Javed, Martin Montecino, Andre J. van Wijnen, Jane B. Lian, Janet L. Stein, and Gary S. Stein

18

Bioluminescent Imaging in Bone ............................................................ 261 Yoram Zilberman, Yossi Gafni, Gadi Pelled, Zulma Gazit, and Dan Gazit

19

Micro-Computed Tomography: A Method for the Non-Destructive Evaluation of the Three-Dimensional Structure of Biological Specimens ........................................................................... 273 Martin Stauber and Ralph Müller

20

Fourier Transform-Infrared Microspectroscopy and Microscopic Imaging ........................................................................ 293 Samuel Gourion-Arsiquaud, Paul A. West, and Adele L. Boskey

Part VI Mechanical Testing 21

Assessment of the In Vivo Adaptive Response to Mechanical Loading ............................................................................ 307 Leanne Kaye Saxon and Lance Edward Lanyon

22

Determination of Bone’s Mechanical Matrix Properties by Nanoindentation ............................................................... 323 Engin Ozcivici, Suzanne Ferreri, Yi-Xian Qin, and Stefan Judex

23

Fluid Flow Assays .................................................................................... 335 Ryan C. Riddle, Amanda F. Taylor, and Henry J. Donahue

Index .................................................................................................................. 347

List of Color Plates

The following color illustrations are printed in the insert Fig. 1.1

Fig. 1.1

Fig. 1.3

Fig. 1.4

Osteoblastic differentiation of mouse calvarial cultures. a. Calvarial cells were isolated from Col3.6-Topaz transgenic mice, which express green fluorescent protein (GFP) driven by a 3.6-kb rat type I procollagen promoter (15), and cultured under osteogenic differentiation conditions. On the indicated days after plating, cells were fixed with formalin. Using a 10× objective, the same fields were photographed using epifluorescence to visualize GFP and phase contrast optics. Nascent nodules first appeared as refractile areas where cells have begun to deposit collagen and form multilayers around day 8. Cells possessing GFP fluorescence presaged visible nodule formation and were preferentially localized within differentiating nodules. As seen in the day 14 phase contrast image, nodules grew in area and acquired a granular appearance as mineralization proceeded b. A transverse section through a mineralized nodule from a rat calvarial osteoblast culture (study day 19) that was fixed with formalin, embedded in Spurr’s resin, sectioned and stained with toluidine blue. Bright field microscopy revealed mineralized matrix (purple) on the side of the nodule next to the culture dish and a layer of cuboidal osteoblasts atop of a layer of osteoid adjacent to the culture medium Alkaline phosphatase enzyme histochemical staining during osteoblastic differentiation. Rat calvarial cells were isolated and cultured under osteogenic differentiation conditions. On the indicated days after plating, a dish was fixed and air dried. At the end of the timecourse, all dishes were stained for alkaline phosphatase enzyme activity. While alkaline phosphatase activity was already detectable at day 6 (red color), it increased in intensity with osteoblastic differentiation and the strongest staining was associated with the multilayered nodules of differentiated osteoblasts Extracellular matrix mineralization during osteoblastic differentiation. Rat calvarial or bone marrow cells were isolated and cultured under osteogenic differentiation conditions. a. On the indicated days after plating, rat calvarial cultures were von Kossa stained and viewed by phase xi

xii

Fig. 20.3

List of Color Plates

contrast optics using a 4× objective. Small mineralized nodules as well as unmineralized nodules were detectable by day 8. With time, the mineralized nodules increased in size and number. b. Unmagnified views of von Kossa (VK) stained cultures showing a differentiated rat calvarial culture (ROB) in a 12-well dish 12 days after plating (ROB) and a differentiated rat bone marrow culture in a 6-well dish 18 days after plating (BMC). c. Unmagnified view of mineralized nodules visualized by calcein incorporation. ROB and BMC cultures were stained with either calcein (left panel) or calcein blue (center and right panels) and photographed on top of an ultraviolet transilluminator Hyperspectral images obtained from normal baboon cortical bone tissue. a. A typical FTIR spectrum from a single image pixel showing frequencies of interest. b. Integrated matrix areas (1,592–1,712 cm−1) are calculated and displayed as 2D color-coded images to give an IR image of the matrix distribution. c. Integrated mineral areas (916–1,180 cm−1) and the tissue mineral distribution and (d) the carbonate (840–892 cm−1) distribution. Images of the mineral:matrix ratio (e), the carbonate:mineral ratio (f), the crystallinity ratio (1,030/1,020 cm−1) (g) and the crosslink ratio (1,660/1,690 cm−1) (h) are shown. Note: all spectra are baselined and PMMA (embedding media) corrected using ISYS software

Contributors

Basem M. Abdallah, MSc, PhD Department of Endocrinology, KMEB Laboratory, University Hospital of Odense, Denmark Syed A. Ali, MD Department of Cell Biology, University of Massachusetts Medical School, Worcester, MA Rachit Bakshi, PhD Department of Cell Biology, University of Massachusetts Medical School, Worcester, MA Artem Barski, PhD Department of Biochemistry & Molecular Biology, University of Southern California Keck School of Medicine, Los Angeles, CA Dafna Benayahu, PhD Department of Cell and Developmental Biology, Sackler School of Medicine, Tel-Aviv University, Tel-Aviv, Israel Teresita Bellido, PhD Division of Endocrinology & Metabolism and Center for Osteoporosis and Metabolic Bone Diseases, University of Arkansas for Medical Sciences, Little Rock, AR Peter V.N. Bodine, PhD Osteoporosis Research, Women’s Health and Musculoskeletal Biology, Wyeth Research, Collegeville, PA Adele L. Boskey, PhD Mineralized Tissue Research, Hospital for Special Surgery, Weill Medical College and Cornell University, New York, NY Stéphane Blouin, PhD INSERM, U 922, Université d’Angers, Angers, France xiii

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Contributors

Elizabeth Bradley Mayo Clinic Graduate School, Rochester, MN Ernesto Canalis, MD Department of Research, Saint Francis Hospital and Medical Center, Hartford, CT, and The University of Connecticut School of Medicine, Farmington, CT Daniel Chappard, MD, PhD INSERM, U 922, Université d’Angers, Angers, France Shu-Gui Chen The Key Laboratory of Biomedical Information Engineering of Ministry of Education and Institute of Molecular Genetic, School of Life Science and Technology, Xi’an Jiaotong University, Xi’an, P.R. China Yuan Chen Laboratory of Molecular and Statistical Genetics, College of Life Sciences, Hunan Normal University, Changsha, Hunan, P.R. China John M. Chirgwin, PhD Department of Internal Medicine, Division of Endocrinology, University of Virginia, Charlottesville, VA Hong-Wen Deng, PhD Departments of Orthopedic Surgery and Basic Medical Sciences, University of Missouri-Kansas City, Kansas City, MO Valerie Deregowski, PhD Department of Research, Saint Francis Hospital and Medical Center, Hartford, CT Nicholas Ditzel, MSc Department of Endocrinology, KMEB Laboratory, University Hospital of Odense, Denmark Henry J. Donahue, PhD Division of Musculoskeletal Sciences and Department of Orthopaedics and Rehabiliation, Pennsylvania State University College of Medicine, Milton S. Hershey Medical Center, Hershey, PA Suzanne Ferreri Department of Biomedical Engineering, State University of New York at Stony Brook, Stony Brook, NY Renny T. Franceschi, PhD Department of Periodontics & Oral Medicine, School of Dentistry, and Biological Chemistry, School of Medicine, University of Michigan, Ann Arbor, MI Baruch Frenkel, DMD, PhD Departments of Orthopaedic Surgery and Biochemistry & Molecular Biology, University of Southern California Keck School of Medicine, Los Angeles, CA

Contributors

Yossi Gafni, MSc, DMD Skeletal Biotechnology Laboratory, Hebrew University- Hadassah Medical Center, Jerusalem, Israel Dan Gazit, PhD, DMD Skeletal Biotechnology Laboratory, Hebrew University- Hadassah Medical Center, Jerusalem, Israel Zulma Gazit, PhD Skeletal Biotechnology Laboratory, Hebrew University- Hadassah Medical Center, Jerusalem, Israel Chunxi Ge, MD, PhD Departments of Periodontics & Oral Medicine, School of Dentistry, and Biological Chemistry, School of Medicine, University of Michigan, Ann Arbor, MI Laurie H. Glimcher, MD Harvard Medical School and Harvard University of Public Health, Boston, MA Samuel Gourion-Arsiquaud, PhD Mineralized Tissue Research, Hospital for Special Surgery, New York, NY Theresa A. Guise, MD Department of Internal Medicine, Division of Endocrinology, University of Virginia, Charlottesville, VA Yan Guo The Key Laboratory of Biomedical Information Engineering of Ministry of Education and Institute of Molecular Genetic, School of Life Science and Technology, Xi’an Jiaotong University, Xi’an, P.R. China Yan-Feng Guo The Key Laboratory of Biomedical Information Engineering of Ministry of Education and Institute of Molecular Genetic, School of Life Science and Technology, Xi’an Jiaotong University, Xi’an, P.R. China Mohammad Q. Hassan, PhD Department of Cell Biology, University of Massachusetts Medical School, Worcester, MA Amjad Javed, PhD School of Dentistry, University of Alabama at Birmingham Institute of Oral Health Research, Birmingham, AL Hui Jiang Laboratory of Molecular and Statistical Genetics, College of Life Sciences, Hunan Normal University, Changsha, Hunan, P.R. China Dallas C. Jones, PhD Harvard Medical School and Harvard University of Public Health, Boston, MA

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Contributors

Stefan Judex, PhD Department of Biomedical Engineering, State University of New York at Stony Brook, Stony Brook, NY Moustapha Kassem, MD, PhD, DSc Department of Endocrinology, KMEB Laboratory, University Hospital of Odense, Denmark Yogendra P. Kharode, MS Osteoporosis Research, Women’s Health and Musculoskeletal Biology, Wyeth Research, Collegeville, PA Krishna P. Kota, PhD Department of Cell Biology, University of Massachusetts Medical School, Worcester, MA Lance E. Lanyon, CBE, BVSc, PhD, DSc, MRCVS, FMedSci Royal Veterinary College, London, UK Xin Li, PhD Department of Periodontics & Oral Medicine, School of Dentistry, University of Michigan, Ann Arbor, MI Jane B. Lian, PhD Department of Cell Biology, University of Massachusetts Medical School, Worcester, MA Hélène Libouban, PhD INSERM, U 922, Université d’Angers, Angers, France Jian-Feng Liu Departments of Orthopedic Surgery and Basic Medical Sciences, University of Missouri-Kansas City, Kansas City, MO Laurie K. McCauley, DDS, PhD Departments of Periodontics & Oral Medicine and Pathology, University of Michigan, Ann Arbor, MI Ricardo F. Medina, PhD Department of Cell Biology, University of Massachusetts Medical School, Worcester, MA Khalid S. Mohammad, MD, PhD Department of Internal Medicine, Division of Endocrinology, University of Virginia, Charlottesville, VA Martin Montecino, PhD Departamento de Biologia Molecular, Universidad de Concepcion, Facultad de Ciencias Biologicas, Concepcion, Chile

Contributors

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Marie Françoise Moreau, PhD INSERM, U 922, Université d’Angers, Angers, France Ralph Müller, PhD Institute for Biomechanics, ETH Zürich, Zürich, Switzerland Jeffrey A. Nickerson, PhD Department of Cell Biology, University of Massachusetts Medical School, Worcester, MA Merry Jo Oursler, PhD Endocrine Research Unit, Mayo Clinic, Rochester, MN Thomas A. Owen, PhD Department of Cardiovascular, Metabolic and Endocrine Diseases, Pfizer Global Research and Development, Groton, CT Engin Ozcivici Department of Biomedical Engineering, State University of New York at Stony Brook, Stony Brook, NY Lydia C. Pan, PhD Department of Cardiovascular, Metabolic and Endocrine Diseases, Pfizer Global Research and Development, Groton, CT Nicola C. Partridge, PhD Department of Physiology and Biophysics, University of Medicine and Dentistry of New Jersey, Robert Wood Johnson Medical School, Piscataway, NJ Gadi Pelled, DMD Skeletal Biotechnology Laboratory, Hebrew University- Hadassah Medical Center, Jerusalem, Israel Glenda J. Pettway, MS Departments of Periodontics & Oral Medicine and Biomedical Engineering, University of Michigan, Ann Arbor, MI Lilian I. Plotkin, PhD Division of Endocrinology & Metabolism and Center for Osteoporosis and Metabolic Bone Diseases, University of Arkansas for Medical Sciences, Little Rock, AR Shirwin M. Pockwinse, BA Department of Cell Biology, University of Massachusetts Medical School, Worcester, MA Jitesh Pratap, PhD Department of Cell Biology, University of Massachusetts Medical School, Worcester, MA

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Contributors

Steven Pregizer Department of Biochemistry & Molecular Biology, University of Southern California Keck School of Medicine, Los Angeles, CA Ling Qin, PhD Department of Physiology and Biophysics, University of Medicine and Dentistry of New Jersey, Robert Wood Johnson Medical School, Piscataway, NJ Yi-Xian Qin, PhD Department of Biomedical Engineering, State University of New York at Stony Brook, Stony Brook, NY Robert R. Recker, MD, MACP, FACE Osteoporosis Research Center, Creighton University, Omaha, NE Ryan C. Riddle, PhD Division of Musculoskeletal Sciences and Department of Orthopaedics and Rehabilitation, Pennsylvania State University College of Medicine, Milton S. Hershey Medical Center, Hershey, PA Leanne K. Saxon, Ba Appl Sci (Hon), PhD Royal Veterinary College, London, UK Michael C. Sharp Osteoporosis Research, Women’s Health and Musculoskeletal Biology, Wyeth Research, Collegeville, PA Irena Shur, MD, PhD Department of Cell and Developmental Biology, Sackler School of Medicine, Tel-Aviv University, Tel-Aviv, Israel Rina Socher. MSc Department of Cell and Developmental Biology, Sackler School of Medicine, Tel-Aviv University, Tel-Aviv, Israel Martin Stauber, PhD Institute for Biomechanics, ETH Zürich, Zürich, Switzerland Gary S. Stein, PhD Department of Cell Biology, University of Massachusetts Medical School, Worcester, MA Janet L. Stein, PhD Department of Cell Biology, University of Massachusetts Medical School, Worcester, MA Amanda F. Taylor, PhD Division of Musculoskeletal Sciences and Department of Orthopaedics and Rehabiliation, Pennsylvania State University College of Medicine, Milton S. Hershey Medical Center, Hershey, PA

Contributors

xix

Nadiya M. Teplyuk, PhD Department of Cell Biology, University of Massachusetts Medical School, Worcester, MA Margaretha van der Deen, Dr Department of Cell Biology, University of Massachusetts Medical School, Worcester, MA André J. Van Wijnen, PhD Department of Cell Biology, University of Massachusetts Medical School, Worcester, MA Marc N. Wein, PhD Harvard Medical School and Harvard University of Public Health, Boston, MA Paul A. West, PhD Mineralized Tissue Research, Hospital for Special Surgery, New York, NY Dong-Hai Xiong Osteoporosis Research Center, Creighton University, Omaha, NE Fang Yang Laboratory of Molecular and Statistical Genetics, College of Life Sciences, Hunan Normal University, Changsha, Hunan, P.R. China Tie-Lin Yang The Key Laboratory of Biomedical Information Engineering of Ministry of Education and Institute of Molecular Genetic, School of Life Science and Technology, Xi’an Jiaotong University, Xi’an, P.R. China Daniel W. Young, PhD Novartis Institute for Biomedical Research, Cambridge, MA Sayyed K. Zaidi, PhD Department of Cell Biology, University of Massachusetts Medical School, Worcester, MA Yoram Zilberman, PhD, DMD Skeletal Biotechnology Laboratory, Hebrew University-Hadassah Medical Center, Jerusalem, Israel

Chapter 1

Isolation and Culture of Rodent Osteoprogenitor Cells Thomas A. Owen and Lydia C. Pan

Abstract Osteoblasts are the cells responsible for formation of new bone throughout life. Rats are one of the most widely studied mammalian species in skeletal biology and serve as useful models for many aspects of human skeletal physiology. The availability of genetically modified mice as research tools has greatly enabled our understanding of how specific genes contribute to the process of skeletogenesis. In order to explore the impact of biochemical, genetic, or pharmacological manipulation on bone formation, various osteogenic cell culture systems have been developed. Two of the most widely accepted rodent osteogenic culture models, using osteoprogenitor cells isolated from calvaria or bone marrow, are described in this chapter. Keywords Osteoblast, bone marrow, mineralization, alkaline phosphatase, calcein.

1

Introduction

Cells of the osteoblastic lineage are integral to the process of bone formation, which maintains integrity of the skeleton. The molecular events that regulate osteoblastic differentiation are frequently studied using in vitro cellular models that recapitulate events occurring in vivo but are more accessible to biochemical, pharmacological, and genetic manipulation than whole organisms. Many widely used in vitro osteoblastic cell model systems employ either karyotically normal but immortal cell lines (e.g., MC3T3) or genetically atypical cell lines derived from tumors (e.g., UMR-106, ROS 17/2.8). However, tumor origins and genetic alterations associated with the prolonged culture of cell lines can raise questions about the extent to which they truly replicate in vivo phenomena. Two laboratory species that are widely employed in biomedical research on the mammalian skeleton are rats and mice. Rats have been widely used to model many aspects of human skeletal physiology, including responses to hormone deficiencies and mechanical loading. Recent advances in genetic technologies have made mice important models for investigating the roles of individual genes in the skeleton. From: Methods in Molecular Biology, Vol. 455: Osteoporosis: Methods and Protocols Edited by Jennifer J. Westendorf © Humana Press, Totowa, NJ

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4

T.A. Owen and L.C. Pan

In vitro model systems can be effectively developed using genetically and phenotypically normal primary osteoblast precursor cells derived from either the developing calvaria of embryonic or neonatal rodents or from the bone marrow of rodents of any age. Osteogenic cultures derived from rodent calvaria undergo a series of proliferative and maturational events characterized by highly regulated changes in gene expression. Eventually, the cells form multi-layered nodules that resemble islands of bone tissue when examined histologically (1–3). The cells in these nodules are surrounded by an organized collagenous extracellular matrix onto which hydroxyapatite crystals are deposited (2–4). Similarly, osteoblastic precursor cells contained within the bone marrow undergo sequential stages of attachment, clonal growth, extracellular matrix deposition, and mineralization (5, 6). Osteogenic precursor cells derived from bone marrow are believed to represent cells at an early stage in the mesenchymal lineage, since they readily give rise not only to osteoblastic colonies, but also to adipocytic and fibroblastic colonies. In contrast, osteogenic cells isolated from the calvarium represent a more committed cell at a later stage in the lineage. The choice of experimental species and cellular model depends upon the question under investigation. Methods for preparing osteogenic precursor cells from rodents were first published more than 40 years ago (6–8) and have been refined by many investigators over time. Isolation of osteogenic cells from rodent calvariae involves sequential enzymatic digestion, typically using trypsin and collagenase, first to remove contaminating cell types such as blood cells and fibroblasts, and then to liberate the osteogenic cells from their surrounding extracellular matrix. Late-term fetal or neonatal calvaria are most often used because they provide a relatively abundant source of osteoprogenitors without the need for extensive tissue dissection. Although they originate from a different embryonic tissue than most of the rest of the skeleton, a large body of research indicates that they share many of their physiological and pharmacological responses with osteoblasts derived from other bone sites. Bone marrow yields cultures with a lower percentage of osteogenic cells than calvaria (9); however, these cells are responsible for the formation of cancellous and endocortical bone that continuously turns over and can be studied throughout the entire lifespan of the organism. With either cell source, the isolated cells proliferate and produce an organized collagenous extracellular matrix that eventually mineralizes in a manner highly reminiscent of intact bone.

2 2.1

Materials Cell Culture Media and Reagents for Rodent Calvarial Osteoblast Isolation

1. Cell digestion PBS: Cold, sterile phosphate-buffered saline, calcium and magnesium-free (PBS) with 50 µg/mL gentamicin sulfate (or 100 units/mL penicillin plus

1 Culture of Rodent Cells

2.

3.

4.

5.

6.

7.

5

100 µg/mL streptomycin) and antimycotic solution (e.g., Fungizone, 2.5 µg/mL amphotericin B in 20 µg/mL sodium deoxycholate, Invitrogen, Carlsbad, CA). Trypsin-collagenase digestion cocktail: Dissolve an amount of Collagenase P equivalent to 15 Wünsch units in 45 mL serum-free Minimal Essential Medium (MEM). When dissolved, add 5 mL of 2.5% trypsin solution in PBS to give 0.25% final trypsin concentration (see Note 1). Final collagenase activity should be 0.3 Wünsch units/mL. Sterilize by filtration through a 0.22-µm filter. Store on ice until needed, but warm to 37 °C just prior to use. Prepare fresh each day it will be used. L-Ascorbic acid: Dissolve 1 g in 20 mL of water to give 50 mg/mL final concentration (see Note 2). Sterilize by filtration through a 0.22-µm filter and freeze in 1.2-mL aliquots at −20 °C. Each aliquot is sufficient for two 500-mL bottles of culture medium and may be frozen and thawed up to three times. 0.5 M Sodium phosphate, pH 7.4: Mix 77.4 mL of 0.5 M Na2HPO4 with 22.6 mL of 0.5 M NaH2PO4. Confirm the pH (±0.1 pH unit) and sterilize by filtration through a 0.22-µm filter. Store at room temperature (see Note 3). Dexamethasone: Dissolve cell culture grade dexamethasone in 100% ethanol to a final concentration of 10−2 M. Store in small aliquots at −20 °C. Dilute 10−2 M dexamethasone 1:100 in ethanol to make a 10−4 M stock. Diluted (10−4 M) dexamethasone is stable at −20 °C for up to 6 months even after repeated freezethaw cycles. Growth medium: Supplement 500 mL of MEM with 50 mL (10%) of fetal bovine serum (FBS) (see Note 4), 0.55 mL of 50 mg/mL gentamicin sulfate (50 µg/mL final concentration). For the preparation of rat calvarial osteoblasts, add 55 µL of 10−4 M dexamethasone (10−8 M final concentration). Dexamethasone is not necessary for the growth of murine calvarial osteoblasts. Freezing medium: MEM + 30% FBS + 10% DMSO.

2.2

Rodent Calvarial Osteoblast Growth and Differentiation

1. Growth medium as in Section 2.1 Item 6 (above). 2. Differentiation medium: Supplement 500 mL of minimal essential medium, alpha modification (MEMα) with 50 mL (10%) of FBS (see Note 4), 0.55 mL of 50 mg/mL gentamicin sulfate (50 µg/mL final concentration), 0.55 mL of 50 mg/mL L-ascorbic acid (100 µg/mL final concentration including that present in the basal medium), 2.2 mL of 0.5 M sodium phosphate, pH 7.4. This yields a 3 mM total concentration of phosphate ions including what is already present in the basal medium (see Note 5). For the preparation of rat calvarial osteoblasts, add 55 µL of 10−4 M dexamethasone (10−8 M final concentration). Dexamethasone is not necessary for the differentiation of murine calvarial osteoblasts.

6

2.3


Rodent Bone Marrow Osteoblast Isolation, Growth, and Differentiation

1. Bone marrow PBS: Cold, sterile PBS with 50 µg/mL gentamicin sulfate (or 100 U/mL penicillin plus 100 µg/mL streptomycin) and antimycotic solution (e.g., Fungizone, 2.5 µg/mL amphotericin B in 20 µg/mL sodium deoxycholate, Invitrogen, Carlsbad, CA). 2. Isomet low speed saw with diamond wafering blade (Buehler, Lake Bluff, IL). 3. 10-cc syringe with flexible tubing (~5 cm) attached. The tubing diameter should allow a snug fit around the bone from which the bone marrow cells will be isolated (i.e., adult rat vs juvenile mice) and should be sterilized by immersion in 70% ethanol for at least 10 minutes. 4. L-ascorbic acid as in Section 2.1 Item 3. 5. 0.5 M sodium phosphate, pH 7.4 as in Section 2.1 Item 4. 6. Dexamethasone as in Section 2.1 Item 5. 7. Growth medium: Supplement 500 mL of MEMα with 75 mL (15%) of FBS (see Note 4) and 0.58 mL of 50 mg/mL gentamicin sulfate (50 µg/mL final concentration). 8. Differentiation medium: Supplement 500 mL of MEMα medium with 50 mL (10%) of FBS (see Note 4), 0.55 mL of 50 mg/mL gentamicin sulfate (50 µg/mL final concentration), 0.55 mL of 50 mg/mL L-ascorbic acid (100 µg/mL final concentration including that present in the basal medium), 2.2 mL of 0.5 M sodium phosphate pH 7.4 (3 mM final concentration including the phosphate present in the basal medium). For the preparation of rat calvarial osteoblasts, add 55 µL of 10−4 M dexamethasone (10−8 M final concentration). Dexamethasone is not necessary for the differentiation of murine bone marrow-derived osteoblasts (see Note 5).

2.4

Alkaline Phosphatase Enzyme Activity

1. Alkaline phosphatase lysis buffer: 150 mM Tris-HCl, pH 9.0, 0.1 mM ZnCl2, 0.1 mM MgCl2, 1% Triton X-100. Prepare by dissolving Tris base, ZnCl2, and MgCl2 in water, adjusting the pH to 9.0 with HCl, and then adding the Triton X-100 and water to the required volume. Sterilize by filtration through a 0.22 µm filter and store at room temperature. 2. Measurement of cell protein for normalization of enzyme activity: Detergentcompatible DC protein assay reagent (BioRad, Hercules, CA) or any other protein measurement method compatible with 1% Triton X-100. 3. Enzyme substrate: 1-Step PNPP (p-nitrophenyl phosphate) (Pierce, Rockford, IL) or any other detection reagent based on PNPP for colorimetric detection of absorbance at 405 nm (see Note 6). Prewarm to 37 °C prior to use.


2.5

7

Alkaline Phosphatase Histochemical Staining

1. Fixative: 100% methanol chilled to −20 °C (see Note 7). 2. 0.2 M Tris-maleate, pH 8.4: Titrate 0.5 M Tris base solution with maleic acid to the desired pH, and then adjust volume to obtain a final concentration of 0.2 M. 3. Alkaline phosphatase histochemical stain: Dissolve 10 mg of Napthol AS-MX phosphate, sodium salt, in 0.56 mL of dimethylformamide. Add 9.44 mL of water, 10 mL of 0.2 M Tris-maleate, pH 8.4, and 20 mg of Fast Red TR salt and stir until dissolved. If the solution is not completely clear, filter it (vacuum or gravity) through Whatman 3 M paper. Filtration is not usually necessary unless high resolution microscopy is being performed. Do not use if the solution has turned brownish yellow and cloudy. Prepare only as much stain as is needed for immediate use (see Note 8).

2.6

von Kossa Staining for Assessment of Mineralization

1. Fixative: 100% methanol (−20 °C) (see Note 7). 2. 5% AgNO3 in water. Store at room temperature protected from light. Solutions containing silver nitrate should be disposed of as hazardous waste.

2.7

Calcein Staining for Assessment of Mineralization

1. Fix: 100% methanol (−20 °C) (see Note 7). 2. Calcein (CAS Number: 1461-15-0) or calcein blue (CAS Number: 54375-47-2): 2.5 mg/mL in PBS. Sterilize by filtration through a 0.22-µm filter and store in the dark at 4 °C.

3 3.1

Methods Rodent Calvarial Osteoblast Isolation

1. This isolation protocol works best with rat pups ranging in age from day 21 of gestation through day 1–2 postpartum or with neonatal mice up to 7 days old. In accordance with institutional policies and procedures, euthanize pregnant dams, remove fetuses by Caesarean section and euthanize (e.g., by decapitation). For neonates, euthanize in accordance with institutional policies and procedures

8

2.

3.

4.

5.


(e.g., CO2 asphyxiation followed by decapitation). Dispose of the bodies and retain heads for dissection of the calvariae. Using sterile instruments, dissect each calvarium consisting of parietal bones plus adjoining sections of frontal and occipital bones, but exclude lateral sutures in order to minimize soft tissue contamination. This procedure may be done in a laminar flow hood to minimize the chance of bacterial or fungal contamination, but has also been performed on the benchtop without negative consequences. Cut each calvarium in half along the midline to prevent bones from stacking and sticking together during enzymatic digestion. Collect bone pieces in cell digestion PBS. Rinse bones twice with Cell Digestion PBS to eliminate as much blood as possible. Bones may be digested immediately or stored in this solution at 4 °C for up to 3 days without significant loss of cell viability (see Note 9). Transfer bones to a sterile 50-mL conical tube using sterile forceps for enzymatic digestion. a) Add pre-warmed trypsin-collagenase digestion cocktail, using 10 mL for up to 50 calvariae. Scale up volume for larger numbers of bones, but use no less than 10 mL for small numbers of bones. Fasten the tube horizontally in an orbital shaker and agitate at 120 rpm at 37 °C for 15 minutes. The cells released by this digestion (fraction I) are primarily red blood cells with a small percentage of fibroblasts. This first digest also releases loosely attached fragments of overlying tissue. Remove the tube from the orbital shaker and allow the bone fragments to settle to the bottom of the tube. Pipet off the supernatant and discard it. b) To the bone fragments, add the same volume of fresh, pre-warmed trypsincollagenase digestion cocktail as used in step 5.1. Fasten the tube horizontally in an orbital shaker and agitate at 120 rpm at 37 °C for 30 minutes. This digestion (fraction II) generally releases a large percentage of the osteogenic cell population from the calvarial fragments. Remove tube from the orbital shaker and allow the bone fragments to settle to the bottom. Using a pipet, transfer the supernatant containing cells into a clean sterile tube and add FBS to 10% (v/v) to neutralize protease activity. Keep cell suspension on ice until the next step is completed. c) To the bone fragments, add the same volume of fresh, pre-warmed trypsincollagenase digestion cocktail as used in step 5.1. Fasten the tube horizontally in an orbital shaker and agitate at 120 rpm at 37 °C for an additional 30 minutes. This digestion (fraction III) releases most of the remaining osteogenic cells from the calvarial fragments. Remove the tube from the orbital shaker and allow the bone fragments to settle to the bottom of the tube. Pipet the supernatant containing cells into a clean sterile tube and add FBS to 10% (v/v) to neutralize the activity of the digestion enzymes. Rinse the bone fragments with 10 mL of growth medium to recover any additional cells and add this rinse to the tube containing the fraction III digest.


9

6. Combine the FBS-neutralized fraction II and III supernatants and filter through a sterile 70- or 100-µm nylon mesh such as a cell strainer (BD Falcon) into a sterile 50-mL collection tube. 7. Pellet the cells by centrifugation at 250 g for 5 minutes. 8. Wash the cell pellet once with growth medium and centrifuge again as in step 7. 9. Resuspend the cell pellet in growth medium using 0.5 mL medium per calvarium. 10. Dilute a sample of cells 1:1 with 0.4% Trypan Blue in PBS and count viable cells using hemacytometer. Typical cell yields are in the range of 1.0 × 106 cells/calvarium for rats and 1.0–1.5 × 106 cells/calvarium for mice. 11. Plate cells in growth medium at a density of 20,000 cells/cm2 (rat) or 60,000 cells/cm2 (mouse). Incubate for 24 hours at 37 °C in a 5% CO2 incubator (see Note 10). 12. Detach cells using 0.25% trypsin with 1 mM EDTA, pellet by centrifugation for 5 minutes at 250 g, then resuspend cells in growth medium and count a sample of the cells. Pellet the cells again by centrifugation at 250 g for 5 minutes. Based on the cell count, resuspend the resulting pellet in freezing medium at 2 × 106 cells/mL. Freeze in cryovials at 1 mL/vial. Once frozen, transfer to liquid nitrogen for long-term storage.

3.2

Rodent Calvarial Osteoblast Growth and Differentiation

1. Thaw a vial of frozen osteoblasts (2 × 106 cells) into a T75 flask containing 20 mL of growth medium and incubate overnight. 2. The following day (day 1), cells may be trypsinized and replated for experiments as primary cultures. Alternatively, cultures may be re-fed with growth medium to remove residual DMSO and allowed to proliferate prior to subculturing (see Note 11). 3. Trypsinize the cells, determine viable cell concentration by Trypan Blue exclusion and plate these cells for the desired experiment at 20,000 cells/cm2 in growth medium. The use of culture vessels with a surface area of at least 2 cm2 per well (24-well dishes) is recommended (see Note 12). The day of plating is study day 0. Feed all wells again with growth medium on study day 2. 4. The cells should be completely confluent by study day 4. On study day 4 and every other day thereafter, feed wells with differentiation medium. After this point, care must be taken not to dislodge the cell layer when feeding. Multilayered nodules of differentiated osteoblasts first become visible as irregular, opaque foci in regions of highest cell density, typically beginning around study day 8. These nodules should be extensively mineralized by study day 14. The rate and extent of differentiation can be affected by several factors including genetics and variability in fetal bovine serum lots. Fig. 1.1a shows the appearance of differentiating calvarial cultures bearing a fluorescent transgene by phase contrast and fluorescence microscopy. Histologically, a cross-section of one of these multilayered

10


Fig. 1.1 Osteoblastic differentiation of mouse calvarial cultures. a. Calvarial cells were isolated from Col3.6-Topaz transgenic mice, which express green fluorescent protein (GFP) driven by a 3.6kb rat type I procollagen promoter (15), and cultured under osteogenic differentiation conditions. On the indicated days after plating, cells were fixed with formalin. Using a 10× objective, the same fields were photographed using epifluorescence to visualize GFP and phase contrast optics. Nascent nodules first appeared as refractile areas where cells have begun to deposit collagen and form multilayers around day 8. Cells possessing GFP fluorescence presaged visible nodule formation and were preferentially localized within differentiating nodules. As seen in the day 14 phase contrast image, nodules grew in area and acquired a granular appearance as mineralization proceeded (See Color Plates).

nodules of cells resembles that of newly formed woven bone. A layer of cuboidal osteoblasts can be seen on top of the nodule overlaying a region of dense collagenous extracellular matrix, the bottom of which is the site of active mineral deposition (Fig. 1.1b).

3.3

Rodent Bone Marrow Osteoblast Isolation, Growth and Differentiation

1. This procedure has been used to isolate total bone marrow cells capable of osteoblastic differentiation from rodents varying in age from 6 weeks to 28 months.


11

Fig. 1.1 (continued) b. A transverse section through a mineralized nodule from a rat calvarial osteoblast culture (study day 19) that was fixed with formalin, embedded in Spurr’s resin, sectioned and stained with toluidine blue. Bright field microscopy revealed mineralized matrix (purple) on the side of the nodule next to the culture dish and a layer of cuboidal osteoblasts atop of a layer of osteoid adjacent to the culture medium (See Color Plates).

In accordance with institutional policies and procedures, euthanize the rats or mice and dissect out the tibiae (rats) or femorae and tibiae (mice). Using a lowspeed Isomet saw with a diamond edged wafering blade and bone marrow PBS in the reservoir, cut off the distal end of the tibia immediately proximal to the tibiofibular junction. Remove the proximal end of the tibia including the metaphysis. Both ends of mouse femorae should be removed, leaving a cylinder of bone (see Note 13). 2. Rinse the syringe with attached tubing with PBS or medium to remove residual ethanol and fill with growth medium. Fit tubing over the distal end of the tibia and depress the plunger. Typically, the marrow cells will be extruded as an intact “plug.” This step may be performed in a laminar flow hood if desired, but can also be done aseptically on an open lab bench. 3. After all bone marrows have been extruded, carry out the remainder of the procedure in a laminar flow hood. Triturate the bone marrow using a 5- or 10-mL tissue culture pipet until it is a well dispersed cellular suspension. Adjust the volume with growth medium to approximately 5 mL per rat tibia or 1 mL per mouse bone. Remove any tissue fragments by filtration through a sterile 70- or 100-µm nylon filter such as a Cell strainer (BD Falcon) into a sterile 50-mL collection tube. 4. Remove 100 µL of the cell suspension and add to 900 µL of PBS, then further dilute with an equal volume of Trypan Blue solution. This results in a 1:20 dilution of the cells and will typically yield a convenient concentration of cells for counting. The dilution factor can be adjusted as needed. Count all cells, regardless of size. Calculate the total cell concentration, remembering to take the appropriate dilution factor into account. A typical yield of cells is in the range of 1.0 × 108 cells/tibia for adult rats and 4–5 × 107 cells from two tibiae plus two femorae of one adult mouse.

12


Fig. 1.2 Osteoblastic differentiation of rat bone marrow cultures. Rat bone marrow cells were isolated and cultured under osteogenic differentiation conditions. a. Unmagnified view of differentiated osteoblastic colonies in a 12-well dish after 18 days in culture. These mineralized colonies are visible as opaque patches in the culture dish. b. Enlarged view of a mineralized osteoblastic colony photographed under a 4× objective.

5. Plate the isolated bone marrow cells into 6-well (or larger) dishes at 1.8 × 105 cells/cm2 in growth medium (see Note 12). The day of plating is called study day 0. Feed all wells again with growth medium on study day 4. There are usually not many attached cells visible prior to study day 4. 6. On study day 7 and every third day thereafter, feed all wells with differentiation medium. Small colonies of cells can be observed by study day 7 and by study day 10, some colonies will begin to form multilayered nodules of differentiated osteoblasts, becoming opaque as a result of mineral deposition. Fig. 1.2 shows a rat marrow culture with mineralized nodules on study day 18 (see Note 14).

3.4

Alkaline Phosphatase Enzyme Activity

1. Aspirate the medium from cultures to be analyzed and rinse once with PBS. Add alkaline phosphatase lysis buffer to the cells (50 µL/cm2 of cell growth area) and incubate on a rocking platform for 30 minutes at room temperature. Assay for alkaline phosphatase enzyme activity either immediately (below) or store the samples in lysis buffer at −80 °C. 2. To assay for alkaline phosphatase enzyme activity, thaw the samples at room temperature or use freshly lysed samples. Do not subject lysates to more than one freeze-thaw cycle prior to analysis. Unless known from previous experiments, pipet various volumes of lysate in triplicate, ranging from 1 to 50 µL, into a 96-well assay plate. Add 200 µL of pre-warmed (37 °C) alkaline phosphatase substrate (PNPP) to each well and read the change in absorbance as a function of time at 405 nm (see Notes 6 and 15). 3. Determine the protein concentration of each sample using a reagent such as the DC protein assay reagent (BioRad) that is compatible with the presence of 1% Triton X-100. Use this value to normalize the amount of enzyme activity determined in step 3.4.2 and express the data as substrate converted per minute per µg protein (10).


13

Fig. 1.3 Alkaline phosphatase enzyme histochemical staining during osteoblastic differentiation. Rat calvarial cells were isolated and cultured under osteogenic differentiation conditions. On the indicated days after plating, a dish was fixed and air dried. At the end of the timecourse, all dishes were stained for alkaline phosphatase enzyme activity. While alkaline phosphatase activity was already detectable at day 6 (red color), it increased in intensity with osteoblastic differentiation and the strongest staining was associated with the multilayered nodules of differentiated osteoblasts (See Color Plates).

3.5

Alkaline Phosphatase Histochemical Staining

1. Aspirate medium from the cultures and rinse once with PBS. Fix the cells for 10 minutes at room temperature using 100% methanol (−20 °C). Following incubation, remove the fixative and rinse once in PBS (see Notes 7 and 16). 2. Remove the PBS and add a sufficient volume of freshly prepared alkaline phosphatase staining solution to cover the dish. Incubate at 37 °C for up to 20 minutes, protected from light, until the desired intensity of staining is reached. Aspirate and discard the stain, rinse with water, and air dry (9) (Fig. 1.3).

3.6

von Kossa Staining for Assessment of Mineralization

1. Aspirate medium from the cultures and rinse once with PBS. Fix the cells for 10 minutes at room temperature using cold 100% methanol (see Note 7). Remove the fixative and rinse twice with water. It is critical to rinse with water prior to staining or else unacceptable background may occur due to precipitation of the stain. Fixed dishes may be stained immediately or stored dry at room temperature for staining at a later time. 2. Stain the cells by adding 5% AgNO3 (250 µL/cm2 of cell growth area). Develop stain by exposure to ultraviolet light until the mineralized nodules become dark brown to black in color (Fig. 1.4a). Traditionally, this has been done by placing the dishes containing AgNO3 solution in bright sunlight until the desired level of staining is obtained (11); however, use of a calibrated ultraviolet source gives more controlled and reproducible results. We routinely use a high-intensity ultraviolet light box designed for cross-linking nucleic acids to membranes (UV Stratalinker 1800, Stratagene) set to deliver 2.4 × 105 µJoules/cm2. 3. Following exposure to ultraviolet light, remove the AgNO3 stain and dispose of it as hazardous waste in accordance with institutional policies. Rinse the dishes twice with water and invert to air dry (see Note 17).

14


Fig. 1.4 Extracellular matrix mineralization during osteoblastic differentiation. Rat calvarial or bone marrow cells were isolated and cultured under osteogenic differentiation conditions. a. On the indicated days after plating, rat calvarial cultures were von Kossa stained and viewed by phase contrast optics using a 4× objective. Small mineralized nodules as well as unmineralized nodules were detectable by day 8. With time, the mineralized nodules increased in size and number. b. Unmagnified views of von Kossa (VK) stained cultures showing a differentiated rat calvarial culture (ROB) in a 12-well dish 12 days after plating (ROB) and a differentiated rat bone marrow culture in a 6-well dish 18 days after plating (BMC). c. Unmagnified view of mineralized nodules visualized by calcein incorporation. ROB and BMC cultures were stained with either calcein (left panel) or calcein blue (center and right panels) and photographed on top of an ultraviolet transilluminator (See Color Plates).

4. Images of von Kossa stained nodules can be captured for quantitation by photography on a white light box using the same manual exposure settings to maintain image uniformity throughout an experiment (Fig. 1.4b). Entire plates can be easily documented using a flatbed scanner. The parameter most frequently quantified is total nodule area using image analysis software such as ImagePro (Media Cybernetics, Bethesda, MD).

3.7

Calcein Staining for Assessment of Mineralization

1. To assess mineralization using calcein, add calcein or calcein blue from the 2.5 mg/mL stock solution to cell culture medium to achieve a final concentration of 25 µg/mL (1:100 dilution) and return cultures to the incubator for 16–24 hours.


15

If quantitation of mineralized nodule area is desired, the time of exposure should be kept constant for all samples being compared. 2. To terminate staining, aspirate the medium and rinse once with PBS. Fix the cells for 10 minutes at room temperature using 100% methanol (−20 °C) or other suitable fixative (see Note 7). Following fixation with methanol, invert and air dry the plates. If an aldehyde fixative is used, rinse the cultures once in PBS, and then invert to air dry. Plates may then be stored indefinitely at room temperature. Protection from strong light is advisable, even though calcein and calcein blue are relatively resistant to photobleaching. 3. Images of calcein or calcein blue fluorescence can be captured by photographing the stained dishes on an ultraviolet transilluminator using the same manual exposure settings to maintain image uniformity throughout an experiment (Fig. 1.4C). These images are also suitable for determination of total nodule area using image analysis software such as ImagePro (Media Cybernetics).

4

Notes

1. An alternative method to prepare trypsin/collagenase digestion cocktail is to dissolve 15 Wünsch units of collagenase P in 50 mL 0.25% trypsin-1 mM EDTA since this is the trypsin-EDTA solution commonly used in cell culture. The low concentration of EDTA in this solution does not practically affect the activity of the collagenase in this application. In many publications, collagenase P concentration has been specified in mass units, for example, 0.2 mg/mL. However, we have specified the collagenase concentration by activity units to facilitate use of commercial preparations with markedly different specific activities. Recommended sources for Collagenase P are Roche and Worthington. 2. All reagents should be prepared in tissue culture quality deionized (18 MΩ) water. All chemicals should be ACS (American Chemical Society) grade or higher or qualified for tissue culture use. 3. β-Glycerol phosphate (10 mM final concentration) may be substituted for the additional sodium phosphate. A stock solution of 1 M β-glycerol phosphate can be prepared in PBS or MEMα medium, sterilized by filtration through a 0.22µm filter, divided into 6-mL aliquots (5.5 mL is added to each 500-mL bottle of differentiation medium), and stored at −20 °C. 4. Fetal bovine serum is not a defined product and varies significantly with manufacturer and individual lots. Serum lots should be tested for their ability to support osteoblast differentiation based on mineralized bone nodule formation under these culture conditions. 5. Ascorbic acid is absolutely required for collagen synthesis and is essential for osteoblasts to lay down an organized collagenous extracellular matrix and form multilayered nodules. The basal medium, MEMα contains 50 µg/mL ascorbic acid and the addition of an extra 50 µg/mL ensures that ascorbate will not be limiting for the necessary collagen production. MEMα contains 1 mM sodium phosphate so that supplementation with an additional 2 mM sodium phosphate is sufficient to allow efficient mineralization of the collagenous extracellular matrix.

16


6. Alternative alkaline phosphate substrates may be utilized, depending on the level of sensitivity required and on the equipment available to read the assay plates. For example, a fluorescent substrate (AttoPhos®) is available from Promega (Madison, WI) and a chemiluminescent substrate (LumiPhos®) can be obtained from Pierce (Rockford, IL). Mature calvarial or bone marrow-derived osteoblast cultures produce high levels of alkaline phosphatase activity, so sensitivity is not generally an issue. As a rule, differentiated calvarial cultures will express significantly higher levels of alkaline phosphatase than will bone marrow-derived cultures. 7. Many alternative fixatives can be used prior to alkaline phosphatase histochemical staining or analysis of mineralization. Limited fixation with either 4% paraformaldehyde, made by diluting 10 mL of 32% paraformaldehyde stock (Electron Microscopy Sciences, Ft. Washington, PA) with 8 mL of 10× PBS and 62 mL of water and storing at room temperature or commercially prepared 10% formalin for 10 minutes at room temperature works well. These aldehyde fixatives are preferable in situations where organic solvents must be avoided, for example, to preserve lipid deposits for adipocyte staining. 8. Commercial alkaline phosphatase histochemical staining kits, for example, leukocyte alkaline phosphatase (kit 86-C) from Sigma Diagnostics are satisfactory and provide other colors of stained product. An analogous kit that produces a fluorescent product, ELF 97 Endogenous Phosphatase Detection Kit (Invitrogen kit E6601) can be substituted for the Napthol AS-MX phosphate and fast red in protocol 3.5.2 (12). 9. The viability of calvarial osteoblasts for several days when stored in this manner allows for shipping between laboratories or for additional analyses that may be needed to genotype or phenotype genetically modified mice. Although we have had considerable success with this option, it is possible that this step could introduce additional experimental variables related to differential cell survival. 10. All cell culture incubations are done at 37 °C in a 5% CO2 incubator. Trypan Blue exclusion does not accurately predict cell viability after the enzymatic digestion protocol, so it is advisable to culture the cells as described in step 11 to allow full recovery prior to initiating experiments. Relatively little proliferation has been observed during this first 24 hours of culture. Freshly isolated calvarial cells can be plated for experiments at this point without freezing. If this is desired, proceed directly to protocol step 3.2.3. 11. Mouse calvarial cultures show a marked loss of differentiation capacity with subculturing; therefore, it is advisable to use primary cultures if osteoblastic differentiation is the desired endpoint. In contrast, rat calvarial cells can be expanded for three to four population doublings without significant loss of the potential for mineralized nodule formation (13). 12. For experiments with osteoblastic differentiation as an endpoint, plating calvarial osteoblasts in wells 20 µL, reduce the volume of assay buffer to get a final volume of 90 µL. Do not use >30 µL of sample and maintain the final volume of the standard to the same value as the samples (i.e., if you add 25 µL of sample, add 15 µL lysis buffer to the standards). 8. Foam will incorporate oxygen in the solution and inhibit the polymerization of the gel. 9. An image analysis system able to detect chemiluminescence can be used instead of film. This method has the advantage that the images are directly stored as electronic files, avoiding the need to scan the film. However, the sensitivity is lower and some weak bands may not be detected using image analysis. 10. The normal serum use to block has to be from the species in which the secondary antibody was raised. 11. The biotin-avidin system allows the amplification of the system, obtaining a better signal. However, FITC-labeled antibodies can be used if biotinylated antibodies are not available or the signal is strong. Alternatively, an HRPlabeled secondary antibody can be used, followed by addition of a peroxidase substrate such as diaminobenzidine. 12. Do not leave with fixative for more than 10 minutes, as fluorescence will be reduced. Four percent p-formaldehyde (15 minutes at room temperature) can also be used to fix the cells. Do not use methanol or acetone, as these fixatives destroy the fluorescent proteins. 13. The amount of protein to load depends on the efficiency of transfection: The higher the efficiency, the lower the amount of total lysate that needs to be loaded in the gel. 14. Make sure you do not leave trypsin in too long, as cells will start dying. 15. GFP targeted to the nucleus is also expressed in the cytoplasm, although with lower intensity than in the nucleus. This property can be used to evaluate cell morphology by quantifying the number of cytoplasmic projections per cell. The expression of nuclear-targeted RFP (and cyan fluorescent protein, commercially available) is restricted to the nucleus and cannot be detected in the cytoplasm; therefore, they cannot be used to evaluate cell morphology.

4 Detection of Apoptosis of Bone Cells In Vitro

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19. Hill, P. A., Tumber, A., Meikle, M. C. (1997) Multiple extracellular signals promote osteoblast survival and apoptosis. Endocrinology 138, 3849–3858. 20. Plotkin, L. I., Manolagas, S. C., Bellido, T. (2002) Transduction of cell survival signals by connexin-43 hemichannels. J Biol Chem 277, 8648–8657. 21. Plotkin, L. I., Manolagas, S. C., Bellido, T. (2006) Dissociation of the pro-apoptotic effects of bisphosphonates on osteoclasts from their anti-apoptotic effects on osteoblasts/osteocytes with novel analogs. Bone 39, 443–452. 22. Weinstein, R. S., Chen, J. R., Powers, C. C., et al. (2002) Promotion of osteoclast survival and antagonism of bisphosphonate-induced osteoclast apoptosis by glucocorticoids. J Clin Invest 109, 1041–1048. 23. Kogianni, G., Mann, V., Ebetino, F., et al. (2004) Fas/CD95 is associated with glucocorticoidinduced osteocyte apoptosis. Life Sci 75, 2879–2895. 24. Ahuja, S. S., Zhao, S., Bellido, T., et al. (2003) CD40 ligand blocks apoptosis induced by tumor necrosis factor alpha, glucocorticoids, and etoposide in osteoblasts and the osteocytelike cell line murine long bone osteocyte-Y4. Endocrinology 144, 1761–1769. 25. Almeida, M., Han, L., Bellido, T., et al. (2005) Wnt proteins prevent apoptosis of both uncommitted osteoblast progenitors and differentiated osteoblasts by beta-catenin-dependent and independent signaling cascades involving Src/ERK and phosphatidylinositol 3-kinase/AKT. J Biol Chem 280, 41342–41351. 26. Plotkin, L. I., Mathov, I., Aguirre, J. I., et al. (2005) Mechanical stimulation prevents osteocyte apoptosis: requirement of integrins, Src kinases and ERKs. Am J Physiol Cell Physiol 289, C633–C643. 27. Fatokun, A. A., Stone, T. W., Smith, R. A. (2006) Hydrogen peroxide-induced oxidative stress in MC3T3-E1 cells: the effects of glutamate and protection by purines. Bone 39, 542–551. 28. Park, B. G., Yoo, C. I., Kim, H. T., et al. (2005) Role of mitogen-activated protein kinases in hydrogen peroxide-induced cell death in osteoblastic cells. Toxicology 215, 115–125. 29. Bellido, T., Huening, M., Raval-Pandya, M., et al. (2000) Calbindin-D28k is expressed in osteoblastic cells and suppresses their apoptosis by inhibiting caspase-3 activity. J Biol Chem 275, 26328–26332. 30. Liu, Y., Porta, A., Peng, X., et al. (2004) Prevention of glucocorticoid-induced apoptosis in osteocytes and osteoblasts by calbindin-D28k. J Bone Miner Res 19, 479–490. 31. Pavalko, F. M., Gerard, R. L., Ponik, S. M., et al. (2003) Fluid shear stress inhibits TNFalpha-induced apoptosis in osteoblasts: a role for fluid shear stress-induced activation of PI3kinase and inhibition of caspase-3. J Cell Physiol 194, 194–205. 32. Bakker, A., Klein-Nulend, J., Burger, E. (2004) Shear stress inhibits while disuse promotes osteocyte apoptosis. Biochem Biophys Res Commun 320, 1163–1168. 33. Kitase, Y., Jiang, J. X., Johnson, M. L., et al. (2006) The anti-apoptotic effects of mechanical strain on osteocytes are mediated by PGE2 and monocyte chemotactic protein-3 (MCP-3): selective protection by MCP-3 against glucocorticoid (GC), but not TNF-a induced apoptosis. J Bone Miner Res 21, S48. 34. Alessi, D. R., Cuenda, A., Cohen, P., et al. (1995) PD 098059 is a specific inhibitor of the activation of mitogen-activated protein kinase kinase in vitro and in vivo. J Biol Chem 270, 27489–27494. 35. Nicholson, D. W., Ali, A., Thornberry, N. A., et al. (1995) Identification and inhibition of the ICE/CED-3 protease necessary for mammalian apoptosis. Nature 376, 37–43. 36. Jilka, R. L., Weinstein, R. S., Bellido, T., et al. (1998) Osteoblast programmed cell death (apoptosis): modulation by growth factors and cytokines. J Bone Min Res 13, 793–802. 37. Frisch, S. M., Vuori, K., Ruoslahti, E., et al. (1996) Control of adhesion-dependent cell survival by focal adhesion kinase. J Cell Biol 134, 793–799. 38. Murakami, T., Yamamoto, M., Ono, K., et al. (1998) Transforming growth factor-beta1 increases mRNA levels of osteoclastogenesis inhibitory factor in osteoblastic/stromal cells and inhibits the survival of murine osteoclast-like cells. Biochem Biophys Res Commun 252, 747–752. 39. Kameda, T., Mano, H., Yuasa, T., et al. (1997) Estrogen inhibits bone resorption by directly inducing apoptosis of the bone-resorbing osteoclasts. J Exp Med 186, 489–495.

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40. Chua, C. C., Chua, B. H., Chen, Z., et al. (2003) Dexamethasone induces caspase activation in murine osteoblastic MC3T3-E1 cells. Biochim Biophys Acta 1642, 79–85. 41. Mahajan, N. P., Harrison-Shostak, D. C., Michaux, J., et al. (1999) Novel mutant green fluorescent protein protease substrates reveal the activation of specific caspases during apoptosis. Chem Biol 6, 401–409. 42. Tyas, L., Brophy, V. A., Pope, A., et al. (2000) Rapid caspase-3 activation during apoptosis revealed using fluorescence-resonance energy transfer. EMBO Rep 1, 266–270. 43. Mohr, S., McCormick, T. S., Lapetina, E. G. (1998) Macrophages resistant to endogenously generated nitric oxide-mediated apoptosis are hypersensitive to exogenously added nitric oxide donors: dichotomous apoptotic response independent of caspase 3 and reversal by the mitogen-activated protein kinase kinase (MEK) inhibitor PD 098059. Proc Natl Acad Sci U S A 95, 5045–5050. 44. O’Brien, C. A., Jia, D., Plotkin, L. I., et al. (2004) Glucocorticoids act directly on osteoblasts and osteocytes to induce their apoptosis and reduce bone formation and strength. Endocrinology 145, 1835–1841. 45. Hiroi-Furuya, E., Kameda, T., Hiura, K., et al. (1999) Etidronate (EHDP) inhibits osteoclasticbone resorption, promotes apoptosis and disrupts actin rings in isolate-mature osteoclasts. Calcif Tissue Int 64, 219–223. 46. Hughes, D. E., Wright, K. R., Uy, H. L., et al. (1995) Bisphosphonates promote apoptosis in murine osteoclasts in vitro and in vivo. J Bone Min Res 10, 1478–1487. 47. Hughes, D. E., Dai, A., Tiffee, J. C., et al. (1996) Estrogen promotes apoptosis of murine osteoclasts mediated by TGF-beta. Nat Med 2, 1132–1136. 48. Chang, C. C., Liao, Y. S., Lin, Y. L., et al. (2006) Nitric oxide protects osteoblasts from oxidative stress-induced apoptotic insults via a mitochondria-dependent mechanism. J Orthop Res 24, 1917–1925. 49. Wiren, K. M., Toombs, A. R., Semirale, A. A., et al. (2006) Osteoblast and osteocyte apoptosis associated with androgen action in bone: requirement of increased Bax/Bcl-2 ratio. Bone 38, 637–651. 50. Ho, W. P., Chen, T. L., Chiu, W. T., et al. (2005) Nitric oxide induces osteoblast apoptosis through a mitochondria-dependent pathway. Ann NY Acad Sci 1042, 460–470. 51. Kim, G. S., Hong, J. S., Kim, S. W., et al. (2003) Leptin induces apoptosis via ERK/cPLA2/ cytochrome c pathway in human bone marrow stromal cells. J Biol Chem 278, 21920–21929. 52. Oursler, M. J., Bradley, E. W., Elfering, S. L., et al. (2005) Native, not nitrated, cytochrome c and mitochondrial-derived hydrogen peroxide drive osteoclast apoptosis. Am J Physiol Cell Physiol 288, C156–C168. 53. Wu, X., McKenna, M. A., Feng, X., et al. (2003) Osteoclast apoptosis: the role of Fas in vivo and in vitro. Endocrinology 144, 5545–5555. 54. Qin, Y. J., Zhang, Z. L., Yu, L. Y., et al. (2006) A20 overexpression under control of mouse osteocalcin promoter in MC3T3-E1 cells inhibited tumor necrosis factor-alpha-induced apoptosis. Acta Pharmacol Sin 27, 1231–1237. 55. Contardi, E., Palmisano, G. L., Tazzari, P. L., et al. (2005) CTLA-4 is constitutively expressed on tumor cells and can trigger apoptosis upon ligand interaction. Int J Cancer 117, 538–550. 56. Postiglione, L., Di, D. G., Ramaglia, L., et al. (2003) Behavior of SaOS-2 cells cultured on different titanium surfaces. J Dent Res 82, 692–696. 57. Alikhani, M., Alikhani, Z., Boyd, C., et al. (2007) Advanced glycation end products stimulate osteoblast apoptosis via the MAP kinase and cytosolic apoptotic pathways. Bone 40, 345–353. 58. Kim, H. J., Zhao, H., Kitaura, H., et al. (2006) Glucocorticoids suppress bone formation via the osteoclast. J Clin Invest 116, 2152–2160. 59. Mogi, M., Togari, A. (2003) Activation of caspases is required for osteoblastic differentiation. J Biol Chem 278, 47477–47482. 60. Wang, Y.H., Liu, Y., Rowe, D.W. (2007) Effects of transient PTH on early proliferation, apoptosis, and subsequent differentiation of osteoblast in calvarial osteoblast cultures. Am J Physiol Endocrinol Metab 292, E594–E603.

Chapter 5

In Vivo Parathyroid Hormone Treatments and RNA Isolation and Analysis Xin Li, Ling Qin, and Nicola C. Partridge

Abstract Intermittent parathyroid hormone (PTH) administration increases bone formation and bone mass and is being used as a therapy for osteoporosis. By contrast, chronic hyperparathyroidism results in the metabolic bone disease osteitis fibrosa characterized by local bone resorption and peritrabecular bone marrow fibrosis in humans. The animal models that can mimic the paradoxical effects of PTH provide the basis for further study of the functions of this hormone in skeletal tissues. In both rats and mice, the anabolic effects of PTH on bone can be achieved by daily injections subcutaneously and the catabolic effects can be achieved by continuous infusion with osmotic pumps. This chapter offers detailed information, such as the dosage and preparation of PTH, using the example of treatment of rats with PTH intermittently or continuously. High-quality, RT-PCR ready RNA is required for the analysis of gene expression. For the analysis of gene expression in bone, usually long bones are used for RNA extraction. Here we describe how to extract RNAs from the metaphyseal trabecular primary spongiosa of rat femur by a method based on two commercially available kits. This protocol can be used in other tissues with slight modification of the amount of reagent used according to the tissue size. Keywords PTH, injection, infusion, RNA, bone, in vivo.

1

Introduction

Excess PTH as seen in primary hyperparathyroidism (PHPT) has been long known for its catabolic effect to promote osteoclast activity and bone resorption and considered as a cause of secondary osteoporosis (1). Later studies have shown that PTH also has an anabolic effect on the mammalian skeleton showing stimulation of new bone formation in humans and experimental animals (2–5). Intermittent PTH application is an established pharmacological principle for its capability to stimulate bone formation. Catabolic effects of PTH can be achieved by infusion with implanted ALZET osmotic pumps continuously delivering PTH. The animal models mimic PTH’s dual effects and are a basis for further investigations

From: Methods in Molecular Biology, Vol. 455: Osteoporosis: Methods and Protocols Edited by Jennifer J. Westendorf © Humana Press, Totowa, NJ

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of the molecular mechanisms underlying the effects of this hormone. Performing microarray analysis with RNA isolated from these animal models provided entire gene expression profiles associated with the anabolic or catabolic effects of the two different PTH treatments. Bone is a hard tissue and rich in degradative enzymes, which makes RNA extraction difficult. We obtained high-quality RNA from rat (and mouse) long bones using a method based on two commercially available kits. Rat femora were resected with all connective tissue removed completely. A region subjacent to the growth plate about 3 mm wide of the metaphyseal trabecular primary spongiosa was stored in RNAlater reagent (Qiagen, Valencia, CA) for RNA isolation using Tri Reagent (Sigma, St. Louis, MO). A RNA cleanup kit (Qiagen) was used to further purify the RNA after isolation to increase the quality of the RNA.

2 2.1

Materials Equipment

1. Sterile syringes, needles, scissors, hemostats, clips, sterile scalpels, forceps, and bone cutter. 2. A scale to weigh animals and a balance to weigh osmotic pumps. 3. ALZET osmotic pumps (DURECT Corp., Cupertino, CA). 4. Isoflurane anesthetic system to anesthetize rats and perform the implantation of pumps. 5. PT 10-35 Polytron homogenizer (Kinematica GA, Littau, Switzerland) to pulverize bone samples. 6. UV spectrophotometer and quartz cuvettes.

2.2 1. 2. 3. 4. 5.

Reagents

Synthetic PTH (1–34) (human) (Bachem, King of Prussia, PA). Saline. Sprague-Dawley rats. Heat-inactivated sera from rats of the same gender and age without treatments. Diethylpyrocarbonate (DEPC)–treated water: Add DEPC to distilled water to a final concentration of 0.1% and incubate at room temperature overnight. Autoclave the next day to inactivate the DEPC. 6. Sodium hydroxide (1 N). 7. Chloroform. 8. Ethanol, 75% and 96–100%.

5 PTH In Vivo and RNA Isolation

9. 10. 11. 12. 13. 14.

15. 16. 17. 18. 19.

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Isopropanol. β-Mercaptoethanol (β-ME), 100% (see Note 1). RNAlater RNA Stabilization Reagent (Qiagen, Valencia, CA). Tri Reagent (Sigma). RNeasy kit (Qiagen). 5 × MOPS buffer: 0.4 M MOPS (morpholinopropanesulfonic acid), 100 mM sodium acetate, 10 mM EDTA. The buffer is adjusted to pH 7.0 with 1 M NaOH and sterilized by autoclaving. Agarose. Formaldehyde. Ethidium bromide, 10 mg/mL. RNA denaturation buffer: 10 mL 100% deionized formamide, 3.5 mL 40% formaldehyde, 1.5 mL 5 × MOPS buffer. 10 × RNA loading buffer: 50% glycerol, 0.4% bromophenol blue, and 0.4% xylene cyanol.

Methods PTH Preparation

1. The synthetic PTH peptide is a lyophilized white powder when purchased and must be dissolved in sterile saline to make an aliquot stock (50 ng/µL), which should be stored at or below −20 °C. 2. Make vehicle solution, saline with 1% heat-inactivated sera: Collect blood from untreated animals of the same gender and age as the animals used in the experiments. Leave the blood at room temperature for at least 30 minutes before centrifuging at 250 g for 10 minutes to separate sera. Heat sera at 56 °C for 30 minutes. Dilute with saline to the final concentration of 1%. Use a 0.20 micron sterile syringe filter (Corning) to make the solution sterile. 3. Weigh and record body weight of each rat to calculate the amount of PTH used in each rat. For injections, the final dose of PTH was 8 µg/100 g (rat body weight) at a final volume of 200 µL for each injection. For infusion, the dose of PTH was 4 µg/100 g (rat body weight) each day. Add vehicle to the PTH stock solution before loading for injections and infusions.

3.2

Injections

Use a 1-mL syringe with 26G 3/8 needle to inject the rats subcutaneously into the hind region.

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3.3

Infusions

3.3.1

Loading Pump (see Notes 2–5)

1. Weigh the empty pump together with its flow moderator. 2. With the flow moderator removed, hold the pump in an upright position, and insert the filling tube through the opening at the top of the pump until it can go no further (see Note 6). 3. Use the smallest volume syringe to fill through a 0.20-µ syringe-end filter; holding the pump in an upright position, slowly push the plunger of the syringe (see Note 7). Stop filling when the solution appears at the outlet and carefully remove the tube. 4. Wipe off the excess solution and insert the flow moderator until the cap is flush with the top of the pump (see Note 8). 5. Weigh the filled pump with its flow moderator in place. The filled volume should be over 90% of the reservoir volume (see Note 9). 6. Place each loaded pump in a tube filled with sterile saline and incubate at 37 °C overnight (or at least for 6 hours) to normalize the flow speed before implantation into rats.

3.3.2

Implantation Surgery

1. Anesthetize rats by isoflurane at the induction concentration of 4.5%, then at maintenance concentration of 2% isoflurane during the surgery. 2. Make a 1-cm-long mid-scapular incision on the back of each rat. Insert a hemostat into the incision and spread the subcutaneous tissue to create a pocket for the ALZET osmotic pump. 3. Insert a filled pump into the pocket and the close the wound skin with two clips.

3.4

Long Bone Collection

Euthanize animals with CO2 immediately after the infusion has ceased or at the indicated time after injection. One or both femora can be collected for RNA extraction (6) (see Note 10). 1. Cut the skin with a scissor to expose one femur. Resect one femur with a bone cutter. Completely remove all connective tissue, including periosteum, with a scalpel. 2. Place the femur on a sterile 100-mm culture dish face up (anterior view). Remove the distal epiphysis, including the growth plate, with a sterile scalpel. Cut from the border between cartilage and cortical bone and flip the condyles off (Fig. 5.1, see Note 11).


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Fig. 5.1 Schematic diagram of longitudinal sections of rat femur showing the trabecular bone region resected for total RNA preparation and analyses. (Reprinted from Onyia, J. E., Bidwell, J., Herring, J., et al. (1995) In vivo, human parathyroid hormone fragment (hPTH 1-34) transiently stimulates immediate early response gene expression, but not proliferation, in trabecular bone cells of young rats. Bone 17, 479–484, with permission from Elsevier).

3. With a bone cutter (dip in NaOH and DEPC-treated water each for 5 seconds before use), resect a subjacent 3-mm-wide band of the metaphyseal trabecular primary spongiosa and store it in 300 µL RNAlater RNA Stabilization Reagent (Qiagen). The tissue can be stored in RNAlater Stabilization Reagent for up to 4 weeks at 2–8 °C, up to 7 days at 18 to 25 °C, or up to 1 day at 37 °C (see Note 12).

3.5

Isolation of RNA from Bone

1. Using sterile forceps (dip into DEPC-treated water before use), remove the sample from the tube into a sterile culture dish containing Kimwipes on the bottom to quickly drain the extra solution. Transfer the sample into a round bottom Falcon tube with 3 mL Tri Reagent (Sigma) inside. 2. Immerse the bottom of the tube into a beaker containing a mixture of ice/water, pulverize the sample by using a PT 10-35 Polytron homogenizer. Pulse several times for 1 minute (see Note 13). 3. Check the bottom of the tube, if it is not a fine suspension, repeat the previous step. 4. Aliquot the 3-mL suspension from the previous step into three labeled, 1.7-mL conical bottom screw-top Eppendorf tubes on ice (see Note 14). 5. Add 200 µL of chloroform per 1 mL Tri Reagent in the tube. Tightly close the tube cap. Shake vigorously for at least 15 seconds. Leave at room temperature for 2–15 minutes.

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6. Centrifuge the resulting mixture at 12,000 g for 15 minutes at 4 °C. Centrifugation separates the mixture into three phases: a red organic phase (containing protein), an interphase (containing DNA), and a colorless upper aqueous phase (containing RNA). 7. Transfer the aqueous phase to a fresh tube and add 500 µL of isopropanol per 1 mL Tri Reagent and mix. Allow the sample to stand for 5–10 minutes at room temperature. 8. Centrifuge at 12,000 g for 10 minutes at 4 °C to precipitate the RNA pellet on the side and bottom of the tube. Remove the supernatant. 9. Wash the RNA pellet by adding 1 mL (minimum) of 75% ethanol per 1 mL Tri Reagent in the tube at step 4. Vortex the sample and then centrifuge at 7,500 g for 5 minutes at 4 °C (see Note 15). Remove the ethanol and air dry the pellet.

3.6

RNA Clean-Up

1. Add 100 µL of RNase-free water to dissolve the RNA pellet in each tube from Section 3.5. 2. Follow the RNA clean-up steps as instructed in the handbook from the RNeasy kit (Qiagen). 3. Pool the RNAs from three Eppendorf tubes of the same bone sample in a single labeled Eppendorf tube.

3.7

RNA Quality and Quantity Test

1. Use 4 or 10 µL of RNA and dilute with RNase-free water to 1 mL in a quartz cuvette. Measure the absorbance at 260 and 280 nm with a spectrophotometer. One A260 unit/mL = 40 µg/mL RNA (see Note 16). 2. The 260/280 nm ratio should be calculated to ensure the absence of protein (see Note 17). 3. Gel electrophoresis is recommended to check the integrity of the RNA samples. Agarose is prepared by melting the required amount of agarose (1%) in DEPC-treated water, cooling to approximately 60 °C (hand hot) and adding 40% formaldehyde and 5 × MOPS to give 2.2 M formaldehyde and 1 × MOPS, respectively (see Note 18). 4. RNA samples are prepared by adding up to 25 µg of RNA in a maximum of 5 µL DEPC-treated water to 15 µL RNA denaturation buffer. Add 1 µL of 10 mg/mL ethidium bromide to aid in the visualization of RNA after electrophoresis. 5. Immediately prior to loading, RNA samples are heated to 55 °C for approximately 15 minutes to denature any secondary structure and cooled on ice. Add 2 µL of sterile 10 × RNA loading buffer.


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Fig. 5.2 Gel electrophoresis of RNAs extracted from rat femora. Each lane was loaded with a different amount of RNA.

6. Samples are loaded onto the gel and electrophoresis conducted at no more than 5 V/cm until the bromophenol blue dye front has migrated approximately threefourths the length of the gel. 7. Visualization of RNA is achieved by irradiation with short wave (254 nm) UV light. Two bands corresponding to the 28S and 18S ribosomal RNAs should be clearly visible (Fig. 5.2).

4

Notes

1. The chloroform used for phase separation should not contain isoamyl alcohol or other additives. 2. ALZET pumps operate because of an osmotic pressure difference between a compartment within the pump, called the salt sleeve, and the tissue environment in which the pump is implanted. The high osmolality of the salt sleeve causes water to flux into the pump through a semipermeable membrane that forms the outer surface of the pump. As the water enters the salt sleeve, it compresses the flexible reservoir, displacing the test solution from the pump at a controlled, predetermined rate. Because the compressed reservoir cannot be refilled, the pumps are designed for single use only. 3. The rate of delivery by an ALZET pump is controlled by the water permeability of the pump’s outer membrane. Thus, the delivery profile of the pump is independent of the drug formulation dispensed. The molecular weight of a compound, or its physical and chemical properties, has no bearing on its rate of delivery by ALZET pumps. 4. The volume delivery rate of ALZET pumps is fixed by the manufacturer. ALZET osmotic pumps are available with a variety of delivery rates between 0.25 and 10 µL/hour and delivery durations between 1 day and 4 weeks. Because the volume delivery rate of the pump is fixed, different dosing rates are achieved by varying the concentration of agent in the solution used to fill the pump reservoir. 5. If a pump becomes contaminated, its surface may be wiped with an aqueous solution of 70% isopropanol immediately before use.

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6. The syringe should be free of air bubbles and allow extra syringe volume for spillage. 7. Make sure the tip of the filling tube meets the bottom of the pump reservoir to avoid air bubbles. 8. A small amount of back pressure is normal due to the tight seal at the filling portal. Avoid rapid filling, which will introduce air bubbles into the reservoir. The insertion of the moderator will displace some of the solution from the filled pump. This overflow should be wiped off. 9. If the solution volume filled in the pump is < 90% of the reservoir volume, the pump should be evacuated and refilled. 10. RNA extraction should be carried out in an RNase-free environment whenever possible. Wear gloves throughout the process and frequently change them while handling samples. It is best to use RNase-free disposable plastic ware and filter-tips. 11. Although tibiae can also be used for RNA isolation, femora are preferred for greater RNA yields due to the tissue size and higher proportion of trabecular to cortical bone. Typical yields of total RNA from a rat femur sample is 40 µg or more. If RNA is extracted from the same region in mice, 1 mL TRI reagent is enough for each sample which yields about 10 µg of RNA. 12. For archived storage at −20 °C, first incubate the sample overnight in reagent (RNAlater) at 2–8 °C and then transfer to −20 °C for storage. For archived storage at −80 °C, incubate the sample overnight in reagent at 2–8 °C, and then remove the tissue from the RNAlater RNA stabilization Reagent and transfer it to −80 °C for storage. 13. The probe of the homogenizer needs to be cleaned before use. First wash with NaOH (1 N) twice. Then wash twice with DEPC-treated water. Finally, wash with 3 mL of Tri Reagent. 14. At this step, sample suspensions can be stored at −20 °C for weeks. Before proceeding to RNA isolation, allow samples to stand for 5 minutes at room temperature to ensure complete dissociation of nucleoprotein complexes. 15. This step can be skipped if immediately performing the first step in Section 3.6. Samples can be stored in ethanol at 4 °C at least 1 week and up to 1 year at −20 °C. 16. Alternatively, the absorbance can be measured with as little as 1 µL of RNA with a NanoDrop spectrophotometer (ND-1000). 17. Final preparation of RNA should have a 260/280 ratio within the range of 1.8– 2.0 to be used in realtime RT-PCR and microarray analysis. 18. Formaldehyde is extremely toxic. Gloves should be worn when preparing and handling solutions containing formaldehyde. Electrophoresis utilizing formaldehyde-containing buffers should be performed in a fume hood. Agarose/ formaldehyde gels are inherently weaker than the equivalent percentage agarose gels and rupture of the well bottom occurs easily. Care should be taken in removing the combs prior to loading of samples.


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References 1. Bauer, W., Aub, J., Albright, F. (1929) Studies in calcium phosphorus metabolism: study of bone trabeculae as ready available reserve supply of calcium. J Exp Med 49, 145–162. 2. Burr, D. B., Hirano, T., Turner, C. H., et al. (2000) Intermittently administered human parathyroid hormone (1-34) treatment increases intracortical bone turnover and porosity without reducing bone strength in the humerus of ovariectomized cynomolgus monkeys. J Bone Miner Res 16, 157–165. 3. Neer, R. M., Arnaud, C. D., Zanchetta, J. R., et al. (2001) Effect of parathyroid hormone (1-34) on fractures and bone mineral density in postmenopausal women with osteoporosis. NEJM 344, 1434–1441. 4. Dempster, D. W., Cosman, F., Kurland, E. S., et al. (2001) Effects of daily treatment with parathyroid hormone on bone microarchitecture and turnover in patients with osteoporosis: a paired biopsy study. J Bone Miner Res 16, 1846–1853. 5. Brommage, R., Hotchkiss, C. E., Lees, C. J., et al. (1999) Daily treatment with human recombinant parathyroid hormone-(1-34), LY333334, for 1 year increases bone mass in ovariectomized monkeys. J Clin Endocrinol Metab 84, 3757–3763. 6. Onyia, J. E., Bidwell, J., Herring, J., et al. (1995) In vivo, human parathyroid hormone fragment (hPTH 1-34) transiently stimulates immediate early response gene expression, but not proliferation, in trabecular bone cells of young rats. Bone 17, 479–484. 7. Alzet Guide to Pump Use (Alzet). 8. Protocol for Sigma’s TRI Reagent (Sigma). 9. RNase Mini Handbook (Qiagen). 10. Li, X. (2006) Parathyroid hormone’s dual effects (anabolic versus catabolic) on osteoblast gene expression. Doctoral dissertation. University of Medicine and Dentistry of New Jersey, Newark, NJ.

Chapter 6

Assessment of Bone Formation Capacity Using In vivo Transplantation Assays: Procedure and Tissue Analysis Basem M. Abdallah, Nicholas Ditzel, and Moustapha Kassem

Abstract In vivo assessment of bone formation (osteogenesis) potential by isolated cells is an important method for analysis of cells and factors controlling bone formation. Currently, cell implantation mixed with hydroxyapatite/tricalcium phosphate in an open system (subcutaneous implantation) in immunodeficient mice is the standard method for in vivo assessment of bone formation capacity of a particular cell type. The method is easy to perform and provides reproducible results. Assessment of the donor origin of tissue formation is possible, especially in the case of human-to-mouse transplantation, by employing human specific antibodies or in situ hybridization using human specific Alu-repeat probes. Recently, several methods have been developed to quantitate the newly formed bone using histomorphometric methods or using non-invasive imaging methods. This chapter describes the use of in vivo transplantation methods in testing bone formation potential of human mesenchymal stem cells. Keywords Mesenchymal stem cell, in vivo bone, heterotropic bone, osteoblast, hydroxyapatite.

1

Introduction

Assessment of osteogenic potential of cells has traditionally been examined in vitro using a combination of molecular biology and cell biology based assays including gene expression of osteoblast specific genes and proteins as well as in vitro mineralization assays (1). However, the expression of these markers does not directly correlate with in vivo bone formation capacity (2). Also, the in vitro environment does not usually reflect the complex in vivo conditions, and thus may be a limiting factor when studying the cellular and molecular control of in vivo bone formation. For these reasons, assessment of bone formation capacity of cells in vivo is an important assay to be employed by researchers interested in understanding cellular and molecular control of bone formation. From: Methods in Molecular Biology, Vol. 455: Osteoporosis: Methods and Protocols Edited by Jennifer J. Westendorf © Humana Press, Totowa, NJ

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Several methods have been employed to assess the in vivo bone formation capacity of cells. Traditionally, implanting cells in diffusion chambers has been employed by several investigators (3). Diffusion chambers are small plastic tubes bound with semi-permeable membranes that can be loaded with cells and then implanted intraperitoneally in immunodeficient mice. Bone and cartilage are usually formed inside the tubes after 20 days. The advantage of diffusion chambers is the ease of identification of donor cells because they are physically separated from recipient cells. However, diffusion chambers are small and difficult to be loaded with cells and the assay has a low success rate. They also cause discomfort for the animals. Alternatively, transplantation of cells mixed with a carrier, hydroxyapatite/tricalcium phosphate (HA/TCP), has been very successful in supporting osteogenesis (4). The advantage of this method is the ease of the procedure, its reproducibility and the possibility of quantitating the bone formed. This assay has been used to test for the stem cell potential of mesenchymal stem cells and quantitate the effects of factors on bone formation (5).

2 2.1

Materials Human Mesenchymal Stem Cell Culture

1. Heparin (100 U/mL). 2. Lymphoprep. 3. Standard growth medium: Modified essential medium (MEM), 10% (v/v) fetal calf serum (FCS), 1% penicillin/streptomycin. Store at 4 °C, for 1 month.

2.2

Loading Cells with HA/TCP Vehicles

1. Hydroxyapatite/tricalcium phosphate (HA/TCP) ceramic powder (Zimmer Scandinavia, Hørsholm, Denmark).

2.3

Transplantation Procedures

1. Non-obese diabetic/severe combined immunodeficient NOD/SCID (NOD/LtSzPrkdcscid) mouse recipients (8-week-old females, ca. 20 g) (6) (see Note 1). Mouse breeding pairs are originally obtained from Jackson Laboratories (Bar Harbor, ME). 2. Ketaminol (Ketamine) Store at 4 °C. 3. Rompun (Xylazine). Store at 4 °C.

6 In Vivo Bone Formation Assay

2.4

Fixation and Embedding of Implants

2.4.1

Plastic and Paraffin Embedding

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1. Methyl methacrylate and glycol methacrylate Technovit 9100 kit (Heraeus, Kulzer GmbH, Wehrheim, Germany). 2. Heavy duty microtome Leica SM2500. 3. Neutral-buffered formaldehyde. 4. Formic acid solution: 0.4 M formic acid, 0.5 M sodium formate.

2.5

Histological Analysis of the Tissues Within the Implants

2.5.1

Hematoxylin/Eosin (H&E) Staining

1. Xylene. 2. Ethanol: 95%, 93%, 77%. 3. Hematoxylin/eosin stain: To prepare 1 L of hematoxylin staining solution, dissolve 1 g of hematoxylin (Merck 15938) in 1 L of distilled water. Add 0.2 g of sodium iodate (Merck 6525) and 50 g of potassium aluminium sulfate (Merck 1047). When this is dissolved, add 50 g of chloral hydrate (Ph Eur) and 1 g of citric acid (Merck 244). To prepare 1 L of eosin staining solution, make an eosin stock solution of 2.1% (w/v) eosin in distilled water. Mix 250 mL of the 2.1% eosin stock solution with 750 mL distilled water and add 150 µL acetic acid (100%). 2.5.2 1. 2. 3. 4.

Goldner’s Trichrome Stain

2-Methoxyethyl-acetate. Ethanol: 99%, 96%. 1% Acetic acid. Weigert Hematoxylin: Prepared fresh by mixing two solutions, A + B 1:3 (v/v). Solution A: Dissolve hematoxylin 1% (w/v) in 99% ethanol. Solution B: Dissolve ferrichloride • 6H2O 2.5% (w/v) and ferrosulfate • 7H2O 4.5% (w/v) in 98 mL distilled water and then mix with 2 mL concentrated hydrochloric acid. 5. Ponceau/Fuchsin Solution: Dissolve 3 g Ponceau 2 R (Merck) and 1.5 g Fuchsin acid (Merck) in 300 mL distilled water and mix with 0.6 mL 100% acetic acid, pH 3.1. Store in dark place at room temperature for 3 months. 6. Orange G Staining Solution: Dissolve 6 g Orange G (Merck 15925) and 12 g Phosphorwolframic acid (Merck 583) in 300 mL distilled water, pH 2.1. Store in dark place at room temperature for 3 months. 7. Fast green FCF solution: Mix 0.6 g Fast Green FCF (Sigma F-7252) and 300 mL distilled water. Then mix with 0.6 mL 100% acetic acid. Store in a dark place at room temperature for 3 months.

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2.5.3

1. 2. 3. 4.

5. 6. 7. 8. 9.

Identifying the Origin of Heterotropic Bone Formation (Immuno-Histochemical Staining)

H2O2 (hydrogen peroxide) 1.5% in TBS buffer. TBS: 100 mL of 0.05 M Tris-HCl, pH 7.4, and 900 mL of 0.15 M NaCl. Normal goat serum. Primary Antibodies: Polyclonal rabbit anti-human Collagen type I antibody Coll I (LF-67), polyclonal rabbit anti-human osteonectin (ON) (LF-37) antibody (7, 8), monoclonal mouse anti-human osteopontin (OPN) (NCL-O-Pontin) (Novocastra, UK, Newcastle upon Tyne) and mouse IgG (negative control). Secondary antibodies: biotinylated goat anti-mouse (Dako, Glostrup, Denmark) and biotinylated goat anti-rabbit (Dako). Peroxidase-conjugated streptavidin. 3-Amino-9-ethyl carbazole. Mayers hematoxylin. Glycergel.

3 3.1

Methods Human Bone Marrow Mesenchymal Stem Cell Cultures

1. Bone marrow is obtained by aspiration from the iliac crest of healthy volunteers (five females 24–30 years old) after infiltration of the area with local anesthetic (Lidocaine, 10 mg/mL) as described previously (9). 2. Aspirate 10 mL of bone marrow and mix with 10 mL of heparin (100 U/mL). Isolate the low-density mononuclear cells by centrifugation over a Lymphoprep density gradient (density = 1.077 ± 0.001 g/cm3) at 1,050 g for 25 minutes at room temperature. Wash the cells once in standard growth medium. 3. Count the viable cells after methylene blue staining in a Burker-Turk counting chamber and seed round 1 × 107 hMSC per T-75 flask. 4. Incubate the cultures in a humidified atmosphere of 5% CO2 at 37 °C and feed the cells by completely replacing the medium once a week until 80–90% confluence.

3.2

Loading Cells with HA/TCP Vehicles

1. Wet the hydroxyapatite/tricalcium phosphate (HA/TCP) powder (40 mg) with 100 µL of standard growth medium to avoid static electricity and transfer it into 1 mL syringe (Codan Medical, Denmark) as shown in Fig. 6.1A. 2. Trypsinize the cultured hMSC cells and add around 5 × 105 cells (in 200 µL medium) carefully on the top of HA/TCP granules in 1 ml syringe. Incubate


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overnight at 37 °C in 5% CO2. Use the HA/TCP granules without any cells as a control (see Note 2). 3. Prepare four samples of hMSC loaded onto HA/TCP granules (as mentioned in step 2) per each donor to produce four implants. 4. For dose-response experiments, the number of hMSC loaded onto HA/TCP granules are as follows: 30 × 105, 15 ×105, 5 × 105, 0.75 × 105, and 0.1 × 105 cells. 5. To examine the effect of different growth factors on the BFC of hMSC, cells are induced with different growth factors in culture for 4 days, then trypsinized and loaded onto the HA/TCP granules for transplantation (5).

3.3

Transplantation Procedures

Transplant the HA/TCP loaded without (vehicle alone as control) or with hMSC into immuno-deficient NOD/SCID (NOD/LtSz-Prkdcscid) mouse recipients (8-week-old females, ca. 20 g) as described previously (11) and as shown in Fig. 6.1B–F). 1. Perform the operations under anesthesia, which is achieved by intra-peritoneal injection of ketaminol (100 mg/kg ketamine) and Rompun (10 mg/kg xylazine). 2. Shave the mouse and disinfect the skin with 70% ethanol prior to operation.

Fig. 6.1 Step-by-step transplantation procedure of hMSC cells with HA/TCP in NOD/LtSzPrkdcscid mice. a. Prepare the cells for implantation by loading them onto HA/TCP granules in 1-mL syringes as shown and described in the Methods section. b. Shave the mouse and clean the area with ethanol. Make an incision 10- to 12-mm long through the skin only. c. Insert a sterile 1-mL syringe into the incision to form a subcutaneous pocket approximately 3 cm deep. Prepare the first implant by removing the medium above the HA/TCP (leave about 1 mm of medium). Insert the syringe with the implant into the pre-formed pocket. Inject the implant by pushing the piston. d. Withdraw the syringe slowly. e. Suture the incision and perform the procedure on the other side of the mouse. f. Arrows indicate the positions of the implants under the skin at the end of the transplantation procedure.

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3. Make two mid-longitudinal skin incisions of about 1 cm in length on the dorsal surface of each mouse, and make subcutaneous pouches by blunt dissection. 4. Place one HA/TCP implant with or without cells into each pouch (two implants per mouse). Close the incisions with surgical sewing and then keep the operated mouse in separate cage until the end of the experiment. 5. Repeat steps 2–5 on another mouse. 6. Keep the implants in mice for 8 weeks.

3.4

Fixation and Embedding of Implants

3.4.1

Plastic Embedding (for Non-Decalcified Sections)

1. Sacrifice the mice 8 weeks post-transplantation by cervical dislocation. Remove the implants and fix them in 70% ethanol. 2. Use the Technovit 9100 Methyl Methacrylate kit according to the manufacturer’s instructions to obtain plastic blocks with mineralized implants ready for sectioning (see Note 3). 3. Use a heavy duty microtome (e.g., Leica SM2500, Leica Microsystems) to cut three serial sections (7.5-µm thick) close to the block surface and repeat it every 100 µm depth through the implant to obtain a total of nine sections per implant at different levels.

3.4.2

Paraffin Embedding (for Decalcified Sections)

1. Sacrifice the mice at 8 weeks post-transplantation by cervical dislocation. Remove the implants and fix them in 4% neutral buffered formaldehyde for 24 hours. 2. Decalcify the implants by immersing them in formic acid solution for 3 days. 3. Use the standard paraffin embedding procedure previously described by other investigators (12–14) to prepare decalcified sections for hematoxylin/eosin (H&E) or immunohistochemical staining.

3.5

Histological Analysis of the Tissues Within the Implants

3.5.1

Hematoxylin/Eosin (H&E) Staining

1. Deparaffinize the sections by submerging in the following series of solutions: xylene for 10 minutes, 99% ethanol for 5 minutes, 93% ethanol for 2 minutes, and 77% ethanol for 2 minutes. 2. Rinse in tap water. 3. Stain with the standard protocol for H&E (Fig. 6.2A).


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Fig. 6.2 Histological analysis of heterotropic bone formation. a. Histological analysis of heterotropic bone in either decalcified paraffin-embedded implants (a, b, and c) or non-decalcified plastic-embedded implants (d, e, and f) harvested after 8 weeks of subcutaneous transplantation of hMSC with HA/TCP in immunodeficient mice. The paraffin sections were stained with H&E, and plastic sections were stained with Goldener’s Trichrome. hMSC cells formed Osteoid (O), trabecular bone (B), within the HA/TCP granules (H). Note that only the Goldener’s Trichrome staining could distinguish between osteoid and mineralized bone. Sections show the advantage of using the in vivo bone assay to compare the capacity of different hMSC clones to differentiate into bone in vivo. Sections were scored as, non-bone forming cells (a, d), cells with moderate bone formation (b, e) and cells with high bone formation (c, f) (magnification a–f ×10). b. Identifying the origin of heterotropic bone formation in the implants by immuno-staining using specific antihuman Collagen type I, osteonectin and osteopontin. Arrows show positive stained bone matrix and osteocytes (magnification a–c × 20).

3.5.2

Goldner’s Trichrome Stain

1. Stain the tissue sections for 30 minutes in 2-methoxyethyl-acetate followed by hydration in 99% ethanol, 96% ethanol, and water for 5 minutes each (see Note 4). 2. Perform the counter-staining with Weigert-Hematoxylin consisting of solution A and solution B 1:3 (v/v) for 5 minutes. 3. Wash the tissue sections for 10 minutes in water and stain with Ponceau/fuchsin solution for 15 minutes. 4. Rinse the sections in 1% acetic acid (2 × 1 minute) and stain with orange G solution for 7 minutes. Sections are rinsed again in 1% acetic acid (2 × 1 minute) followed by staining in Fast green for 20 minutes. 5. Perform a final wash in 1% acetic acid (2 × 1 minute). Dehydrate the tissue sections in 96% ethanol followed by 99% ethanol and mount with glass coverslips (Fig. 6.1A, d–e).

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3.5.3


Identifying the Origin of Heterotropic Bone Formation (Immuno-Histochemical Staining)

To identify the donor origin of the osteogenic tissue, immuno-histochemical staining with human-specific antibodies against osteonectin (ON), osteopontin (OPN) and collagen type I (Col I) was employed as described previously (11). Another method to demonstrate the human origin of the formed bone is in situ hybridization using human specific alu-repeats probes (15). 1. Immerse the tissue sections (from paraffin embedded implants) in 1% acetic acid for 10 minutes and treat with H2O2 in TBS (20 minutes) to abolish endogenous peroxidase. 2. After washing in TBS for 5 minutes, incubate the sections in 10% normal goat serum (NGS). 3. Incubate the tissue sections with primary antibodies overnight at 4 °C (see Note 5), wash with TNT buffer and incubate with secondary antibodies for 1 hour and wash again with TNT buffer. 4. Immuno-reactivity is detected by incubation with peroxidase conjugated streptavidin for 1 hour and developed for 30 minutes with 3-amino-9-ethyl carbazole (AEC). 5. Counter-stain the sections with Mayers Hematoxylin for 30 seconds, and mount in glycergel (Fig. 6.2B).

3.6

Quantifying the Amount of Bone Tissue Formed in the Implants

The following three quantitative methods can be independently performed on an identical set of sections or the whole implant (only for micro-CT method).

3.6.1

Visual Scoring Method (Based on Pixels Counting Method)

This simple and fast analysis method can be used to semi-quantitatively estimate the amount of bone formation using either the plastic or the paraffin-embedded sections as described previously (16, 17). 1. Scan serial sections of paraffin or plastic embedded implant (every 10th section) using Leica microscope linked through a camera Leica DFC 480 to a Leica Dellcompatible computer using the Image-Pro Plus 5.1 image analysis program. 2. Determine the total bone area for each section by selecting a color range that highlighted bone (as bone pixels) based on eosin-stained tissue (in case of paraffin-embedded sections) or Goldner’s Trichrome-stained tissue (in case of plastic-embedded sections) and import into Adobe Photoshop (Adobe Systems, Mountain View, CA).


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3. Estimate the bone area by dividing the total number of bone pixels by the total number of ceramic pixels based on color. 4. Values of serial sections through the implant are summed and then divided by the number of sections to obtain an average section score.

3.6.2

Bone Histomorphometric Analysis

This is a more accurate method and is based on the Cavalieri principle (see Note 6). 1. Serial sections (every 10th section) from each implant are cut and stained with Goldner’s trichrome stain as described. The amount of bone formed is estimated by point counting according to Cavalieri’s principle using CAST software (Olympus, Denmark, Ballerup) as described by our group (11). 2. To quantify the amount of bone tissue formed, two sets, each with the first section placed randomly in the implant, of five successive histological sections stained with Goldner’s Trichrome stain with a fixed intra-distance (75 µm) are prepared and analyzed from each implant according to Cavalierie’s Principle, which allows an unbiased volume estimate of an irregular three-dimensional (3D) structure from uniform random sections (18). 3. The amount of bone formed is determined in each section as a percentage of bone volume per total volume (BV/TV) by point counting using a grid as described previously (19). All counts are performed without knowledge of how many cells are implanted or the nature of donor cells. Intra-observer and interobserver variation for bone quantification are 9% and 17%, respectively.

3.6.3

Micro Computer Tomography

This is a newly developed procedure by our group and it may provide an easy and quick method for quantitation of in vivo formed bone. However, the major limitation is that micro-CT scanning technology is unable to reliably distinguish between the implanted hydroxyapatite/tricalcium phosphate granulates and newly formed mineralized bone. Nevertheless, the micro-CT analysis can provide data on parameters relating bone volume to total implant volume (BV/TV). It is essential that all the implants compared in a given data set were made with the same type of hydroxyapatite granulate because the interpretation of the data assumes equal mean hydroxyapatite particle size and equal distance between particles. If these criteria are met, the contribution of bone-forming cells to increases in the volume of bone in the implant can be measured as an increase in the BV/TV, when compared with control implants made without cells. 1. Carefully lift whole implants out of the fixation solution and place them in a sample-tube supplied with the micro-CT scanner (choose the tube with the smallest possible diameter) (see Note 7). Place the implant in an upright position to minimize the number of image slices required to enclose the entire implant.

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Fig. 6.3 Evaluation of heterotropic bone formation by 3D micro-CT analysis. The micro-CT analysis was performed on the whole implants obtained from the transplantation of HA/TCP granules alone (control) or with hMSC clones (with different capacity for in vivo bone formation) using the viva CT 40-system, which gives a cubic voxel size of 12*12*12 µm3 in the 3D reconstructions.

2. Perform the scanning procedure at high resolution using for example, the viva CT 40-system (Scanco Medical AG, Zürich, Switzerland) that gives a cubic voxel size of 12*12*12 µm3 in the 3D reconstructions. 3. Evaluate the scans by drawing a “region of interest” around the external surface of each implant. Perform 3D evaluation using the µCT Evaluation Program V5.0 (Scanco Medical AG, Zürich, Switzerland). As an example, the 3D micro-CT reconstruction images and parameters obtained from the scanning of two different implants with low and high bone formation are compared with an implant without cells and presented in Fig. 6.3.

4

Notes

1. Handle all the mice under sterile conditions and maintain in a separate NODSCID facility. All animal experiments presented in this study were carried out with permission from the Danish Experimental Animal Inspectorate. 2. As a model of hMSC, our immortalized hMSC named hMSC-TERT cell line (1, 10) were used through the whole in vivo procedure. hMSC-TERT cells are grown in standard growth medium.


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3. The Technovit 9100 kit is an embedding medium system based on methyl methacrylate (MMA). It is designed for the embedding of mineralized tissues with extensive possibilities of staining for light microscopy. 4. Negative controls for histochemical staining protocols are tissue sections from a mouse femur and a human bone biopsy processed in a similar way. 5. As a negative control, mouse IgG (Dako) is used as a primary antibody. 6. Cavalierie’s Principle sums up the areas of object cross-sections and multiplies the section thickness according to the formula: Vreference = T*a*ΣP(treference), where T is the section thickness, a is the area associated with the grid point, and ΣP(treference) is the number of points hitting the profile area. 7. Micro-CT analysis should be done on the whole fixed implant before decalcification or any histological evaluation.

References 1. Abdallah, B. M., Haack-Sorensen, M., Burns, J. S., et al. (2005) Maintenance of differentiation potential of human bone marrow mesenchymal stem cells immortalized by human telomerase reverse transcriptase gene in despite of extensive proliferation. Biochem Biophys Res Commun 326, 527–538. 2. Kuznetsov, S. A., Krebsbach, P. H., Satomura, K., et al. (1997) Single-colony derived strains of human marrow stromal fibroblasts form bone after transplantation in vivo. J Bone Miner Res 12, 1335–1347. 3. Bab, I., Ashton, B. A., Gazit, D., et al. (1986) Kinetics and differentiation of marrow stromal cells in diffusion chambers in vivo. J Cell Sci 84, 139–151. 4. Krebsbach, P. H., Kuznetsov, S. A., Satomura, K., et al. (1997) Bone formation in vivo: comparison of osteogenesis by transplanted mouse and human marrow stromal fibroblasts. Transplantation 63, 1059–1069. 5. Kratchmarova, I., Blagoev, B., Haack-Sorensen, M., et al. (2005) Mechanism of divergent growth factor effects in mesenchymal stem cell differentiation. Science 308, 1472–1477. 6. Shultz, L. D., Schweitzer, P. A., Christianson, S. W., et al. (1995) Multiple defects in innate and adaptive immunologic function in NOD/LtSz-scid mice. J Immunol 154, 180–191. 7. Fisher, L. W., Robey, P. G., Tuross, N., et al. (1987) The Mr 24,000 phosphoprotein from developing bone is the NH2-terminal propeptide of the alpha 1 chain of type I collagen. J Biol Chem 262, 13457–13463. 8. Fisher, L. W., Hawkins, G. R., Tuross, N., et al. (1987) Purification and partial characterization of small proteoglycans I and II, bone sialoproteins I and II, and osteonectin from the mineral compartment of developing human bone. J Biol Chem 262, 9702–9708. 9. Kassem, M., Mosekilde, L., Eriksen, E. F. (1993) 1,25-dihydroxyvitamin D3 potentiates fluoride-stimulated collagen type I production in cultures of human bone marrow stromal osteoblast-like cells. J Bone Miner Res 8, 1453–1458. 10. Simonsen, J. L., Rosada, C., Serakinci, N., et al. (2002) Telomerase expression extends the proliferative life-span and maintains the osteogenic potential of human bone marrow stromal cells. Nat Biotechnol 20, 592–596. 11. Stenderup, K., Rosada, C., Justesen, J., et al. (2004) Aged human bone marrow stromal cells maintaining bone forming capacity in vivo evaluated using an improved method of visualization. Biogerontology 5, 107–118. 12. Bruder, S. P., Kurth, A. A., Shea, M., et al. (1998) Bone regeneration by implantation of purified, culture-expanded human mesenchymal stem cells. J Orthop Res 16, 155–162.

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13. Kuznetsov, S. A., Mankani, M. H., Robey, P. G. (2000) Effect of serum on human bone marrow stromal cells: ex vivo expansion and in vivo bone formation. Transplantation 70, 1780–1787. 14. Mankani, M. H., Krebsbach, P. H., Satomura, K., et al. (2001) Pedicled bone flap formation using transplanted bone marrow stromal cells. Arch Surg 136, 263–270. 15. Hatano, H., Tokunaga, K., Ogose, A., et al. (1998) Origin of bone-forming cells in human osteosarcomas transplanted into nude mice—which cells produce bone, human or mouse? J Pathol 185, 204–211. 16. Dennis, J. E., Konstantakos, E. K., Arm, D., et al. (1998) In vivo osteogenesis assay: a rapid method for quantitative analysis. Biomaterials 19, 1323–1328. 17. Kaigler, D., Wang, Z., Horger, K., et al. (2006) VEGF scaffolds enhance angiogenesis and bone regeneration in irradiated osseous defects. J Bone Miner Res 21, 735–744. 18. Gundersen, H. J., Bendtsen, T. F., Korbo, L., et al. (1988) Some new, simple and efficient stereological methods and their use in pathological research and diagnosis. APMIS 96, 379–394. 19. Kerndrup, G., Pallesen, G., Melsen, F., et al. (1980) Histomorphometrical determination of bone marrow cellularity in iliac crest biopsies. Scand J Haematol 24, 110–114.

Chapter 7

Ossicle and Vossicle Implant Model Systems Glenda J. Pettway and Laurie K. McCauley

Abstract Bone regeneration and repair is a goal of many skeletal therapies and numerous agents positively or negatively impact these processes. New therapeutic agents and effective model systems are continually sought to identify agents and characterize their mechanisms of action are in constant demand. In addition, investigations of tumor cell–bone interaction in the skeletal metastatic microenvironment require well-defined and readily orchestrated models. This chapter describes a novel ectopic ossicle model and a vossicle modification that can be used to provide focused and rapid feedback of bone growth and bone–cellular interactions. The ossicle model is a bone marrow stromal cell (BMSC)–based model and the vossicle model is a neonatal vertebral bone transplant model. These models offer opportunities to mix and compare mesenchymal (donor derived) and hematopoietic elements (host derived). Multiple implants can be placed in one mouse to facilitate various outcome analyses, such as histomorphometry, micro-CT, gene expression studies, and cell tracking using markers such as luciferase, in response to pharmacological or genetic manipulation. Implants can also be combined with other cell types, such as cancer cells to evaluate the bone–tumor microenvironment. Keywords Bone marrow stromal cell, mesenchymal stem cell, parathyroid hormone, bone implant, bone growth, bone regeneration, ectopic ossicle.

1

Introduction

There are a wide variety of agents that positively or negatively impact bone growth and regeneration and there is a great deal of interest to better characterize their actions in bone and identify new therapeutic agents for tissue repair and engineering. In vitro model systems using osteoblastic cells have been beneficial to identify transcriptional mediators and agents with direct actions on cells of the osteoblast lineage, but many agents have disparate actions in vitro vs. in vivo. Often the From: Methods in Molecular Biology, Vol. 455: Osteoporosis: Methods and Protocols Edited by Jennifer J. Westendorf © Humana Press, Totowa, NJ

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in vivo model systems, such as the mouse ovariectomy model, take a long time to orchestrate and evaluate. Model systems in which agents could be tested in a relatively short time period and with discrete and readily measurable endpoints would be beneficial. Our laboratory has focused on investigations of the anabolic actions of parathyroid hormone (PTH) in bone and the impact of tumor-derived PTH-related protein (PTHrP) in skeletal metastasis. PTH is currently the only FDA-approved anabolic agent for osteoporosis treatment in the United States. PTH is also being investigated for potential use in enhancing fracture healing (1, 2), the promotion of implant integration (3), and in tissue engineering applications. The use of PTH in healing osseous defects may be addressed through the use of bone regeneration models, such as the cell-based ectopic ossicle model system presented here and well characterized by other groups (4, 5). This bone-engineered model involves the implantation of bone marrow stromal cells (BMSCs) in immunocompromised mice (Fig. 7.1) to generate ossicles containing cortical and trabecular bone and a hematopoietic marrow. The mesenchymal components of this model are from the donor and the hematopoietic elements are from the host. This model system offers the advantage that multiple distinct bone ossicles can be generated in a single host and used for various endpoint analyses, such as histomorphometry, immunohistochemistry, micro-computed tomography (micro-CT), bioluminescent imaging (BLI), and gene expression studies. Another benefit of this model is that implants can be placed and/or removed from the same mouse in a temporal fashion. This model facilitates the ability to compare the effects of pharmaceutical agents on developing

Fig. 7.1 Schematic of ectopic ossicle model. Tissue engineered bone is generated from transplanted bone marrow stromal cells (BMSCs). This model system can be used to characterize the effects of pharmacologic agents, such as parathyroid hormone (PTH), on bone. (Reproduced from Schneider, A., Taboas, J. M., McCauley, L. K., et al. (2003) Skeletal homeostasis in tissue-engineered bone. J Orthop Res 21:859–864 with permission of John Wiley & Sons, Inc.)

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bone versus endogenous host bone. Furthermore, this model can be used to compare mesenchymal elements from mice with certain genetic mutations with hematopoietic elements from normal mice or visa versa. The ectopic ossicle model system has been effectively used to study skeletal responses to PTH (6, 7) and may also be used to evaluate the effects of other biological agents on both modeling and remodeling bone. A modification of this model is the vossicle model (8), in which instead of implanting bone marrow stromal cells, intact vertebral bodies from neonatal mice are utilized. This modification has the advantage of being more rapid because the step of expanding cells ex vivo is eliminated. A limitation is that there is carryover of hematopoietic elements from the donor with the vertebral implants. So although the vossicles are composed mostly of hematopoietic elements from the host, there is contamination from the donor. An additional feature of this model is that cells such as tumor cells can be co-implanted with the vertebral bodies to simulate a tumor–bone microenvironment.

2 2.1

Materials Bone Marrow Stromal Cell Isolation and Culture

1. Four- to eight-week-old donor mice for bone marrow stromal cell (BMSC) isolation. 2. Surgical Instruments: Forceps, scissors, scalpel, #15 blades, 22- and 25-gauge needles, 1-mL syringes. 3. Growth Medium: Alpha-modified minimum essential medium (α-MEM) supplemented with 20% fetal bovine serum (FBS, Hyclone, Logan, UT), 1% penicillin/streptomycin sulfate/glutamine (PSG), and 10 nM dexamethasone. 4. Hanks’ balanced salt solution (HBSS). 5. Solution of trypsin (0.25%) and ethylenediamine tetraacetic acid (EDTA). 6. 70% Ethanol.

2.2

Surgical Implantation of Bone Marrow Stromal Cells

1. Gelatin sponges (Gelfoam®, Sullivan-Schein, a Henry Schein Company, Melville, NY). 2. No. 3 Whatman filter paper. 3. Surgical Instruments: forceps, scissors, surgical wound clips (Autoclip, 9 mm, Becton Dickinson, Sparks, MD). 4. Anesthesia: 1.5% isoflurane/air anesthesia or 90 mg/kg ketamine and 5 mg/kg xylazine.

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5. Four- to six-week-old male nude mice (NIH III Nude, Charles River Laboratories, Wilmington, MA) (see Note 1).

2.3 1. 2. 3. 4. 5.

Vertebral Disk (Vossicle) Isolation

Neonatal mice (d4 postpartum). 70% EtOH. Surgical instruments: forceps, scissors, scalpel, #15 blades. 60-mm Petri dishes. Phosphate buffered saline (PBS).

2.4

Surgical Implantation of Vossicles

1. Surgical Instruments: forceps, scissors, surgical wound clips (Autoclip, 9 mm, Becton Dickinson, Sparks, MD). 2. Hamilton syringe (if co-implantation with other cells is desired). 3. 70% EtOH. 4. Anesthesia: 1.5% isofluorane/air anesthesia or 90 mg/kg ketamine and 5 mg/kg xylazine. 5. Four- to six-week-old male athymic nude mice (Athymic Nu/Nu, Harlan Laboratories, Madison, WI).

3 3.1

Methods Bone Marrow Stromal Cell Isolation

1. Sacrifice mice that will be used for BMSC isolation (see Note 2). 2. Wash each mouse in 70% ethanol to remove loose fur and disinfect. 3. Remove hind limbs and forelimbs from each mouse and place in HBSS containing 2% PSG. The PSG is used to reduce contamination. 4. In a sterile environment, dissect muscle from femurs, tibiae, and humeri and cut epiphyses (see Note 3). 5. Insert a 1-mL syringe with 25-gauge needle filled with growth medium into marrow cavity of bones and flush out bone marrow. 6. After collecting bone marrow from all bones, aspirate up and down in cell suspension with a 22-gauge needle to break apart clusters of marrow. 7. Add 15 mL of growth medium to a T75 flask followed by cell suspension.

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8. Incubate cells at 37 °C in an atmosphere of 100% humidity and 5% carbon dioxide until a confluent adherent cell layer is observed (see Note 4). 9. When approaching confluency, harvest BMSCs using two washes with HBSS followed by trypsinization with trypsin/EDTA at 37 °C for 10 minutes (see Note 5). 10. Perform a 1:2 split of cells and maintain culture until a confluent adherent layer is observed (see Note 6).

3.2

Surgical Implantation of Bone Marrow Stromal Cells

1. In a sterile environment, place Gelfoam in a 100-mm dish and cut desired number of Gelfoam scaffolds (approximately 5 × 5 × 5 mm cubes) using a scalpel (see Note 7). 2. Saturate Gelfoam cubes with growth medium and place in 37° incubator. 3. Harvest BMSCs for implantation using two washes with HBSS followed by trypsinization with trypsin/EDTA (see Note 8). 4. Pool together BMSCs from all donor mice and count cells. 5. Add volume of growth medium required to obtain 2–3 × 106 cells per 1 mL of cell suspension. 6. Add 1-mL of cell suspension to Eppendorf tubes and centrifuge at 210 g for 5 minutes. 7. While cells are being centrifuged, place presoaked Gelfoam in between two pieces of sterile Whatman filter paper to remove excess growth medium. 8. Aspirate medium from cell pellet. 9. Incorporate BMSCs into Gelfoam using capillary action by placing one Gelfoam scaffold/Eppendorf tube (containing 2–3 × 106 BMSCs) and gently rubbing Gelfoam to bottom of tube where it will absorb the cells. 10. Anesthetize host mice with isoflurane/air anesthesia or with intraperitoneal injection of ketamine/xylazine. 11. Clean the dorsal surface of mice using 70% ethanol and make a small (approximately 1 cm long) vertical incision at hind limbs and forelimbs level. 12. Create bilateral subcutaneous pouches by dissecting skin with blunt scissors (see Note 9). 13. Place one BMSC implant in each “quadrant” of the animal’s back (see Note 10). 14. Close wounds using surgical wound clips. 15. Monitor the recovery of mice and remove clips 2 weeks post-surgery. 16. Treat BMSC recipient mice with desired pharmacologic agent and harvest implants at the end of treatment regime (see Note 11). 17. Perform appropriate endpoint analyses, such as radiographic analysis (see Fig. 7.2A), microCT analysis (see Fig. 7.2B), and histomorphometry using paraffin (see Fig. 7.2C) embedded and/or undecalcified plastic (see Fig. 7.2D) sections. The effects of treatment regime on proliferation of implanted BMSCs

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Fig. 7.2 PTH stimulation of bone formation in ectopic tissue-engineered bone. Representative (a) microradiographic images, (b) micro-computed tomography (microCT), (c) paraffin embedded H&E stained histological sections, and (d) undecalcified plastic embedded tetrachrome stained histological sections of ossicles containing trabecular (T) and cortical (C) bone. Ossicles were treated with PTH or vehicle for 3 weeks, with treatment initiated 1 week after implanting BMSCs. Note the qualitative increase in radiopacity and trabecular bone in implants treated with PTH versus implants treated with vehicle (0.9 % saline). Reconstructed microCT images were generated using a microCT 40 system (Scanco USA, Inc., Wayne, PA) at a resolution of 6 µm.

may also be evaluated using bioluminescence imaging (BLI) (see Fig. 7.3). For this application, BMSCs from mice expressing luciferase are used to generate ossicles and then tracked in vivo with BLI (see Note 12).

3.3

Vossicle Isolation

1. Sacrifice mice that will be used for vertebral isolation (see Note 13). 2. Wash back of mouse with 70% ethanol. 3. In a sterile environment, dissect skin and muscle from vertebrae. Isolate lumbar vertebrae by cutting just under where the ribs attach to the spine and cut just above the pelvis. 4. Transfer vertebrae to a 60-mm dish containing PBS. 5. Carefully dissect the vertebrae with a #15 surgical blade and cut to separate single vertebral disks (vossicles) (see Note 14). 6. Transfer isolated vossicles into fresh PBS in a new 60-mm dish until implantation (see Note 15).

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Baseline (D0)

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b

Baseline (D0)

D28 High

Low Vehicle

PTH

Fig. 7.3 Bioluminescent images of ossicles derived from BMSCs isolated from mice that constitutively express luciferase. Prior to imaging mice were injected by intraperitoneal (ip) injections with 100 µL of 40 mg/mL luciferin dissolved in sterile PBS. Bioluminescent images were acquired 12 minutes after injection under isofluorane/air anesthesia on a cooled CCD IVIS system equipped with a 50-mm lens (Xenogen Corp., Alameda, CA) and coupled to a data-acquisition PC running LivingImage Software (Xenogen Corp.). Luciferase activity was similar at (a) baseline (i.e., day of BMSC implantation), and markedly increased at (b) D28 in implants exposed to PTH for 21 days, with treatment initiated 7 days after implanting BMSCs.

3.4

Surgical Implantation of Vossicles

1. In a sterile environment, anesthetize host mice with isoflurane/air anesthesia or an intraperitoneal injection of ketamine/xylazine. 2. Disinfect the dorsal surface of mice using 70% ethanol and make a small (approximately 1 cm long) vertical incision at hind limbs and forelimbs level (see Note 16). 3. Create bilateral subcutaneous pouches by dissecting skin with blunt scissors. Place one vertebral disc (vossicle) implant in each “quadrant” of the animal’s back. If co-implantation studies are desired, inject cells into vertebral bodies just prior to implantation (see Notes 10 and 17). 4. Close wounds using surgical wound clips. 5. Monitor the recovery of mice and remove clips 2 weeks post-surgery. 6. Treat vossicle recipient mice with the desired pharmacological agent and harvest implants at the end of treatment regime (see Note 18). 7. Perform appropriate endpoint analyses, such as radiographic and histomorphometric analysis (see Fig. 7.4) or other analyses described for ossicles (see Note 19). The effects of treatment regimes on proliferation of implanted vossicles may also be evaluated using BLI where either the vossicle is tracked (using vertebral bodies from luciferase expressing mice) or the tumor is tracked (using luciferase tagged tumor cells).

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a

b

c

d

f

Woven bone Tumor

Fig. 7.4 Vossicle implants. Microradiographic images of vossicles 3 weeks after implantation in a vehicle treated mouse (a), or mouse treated with PTH for 3 weeks (b). Photomicrograph of vossicle from a vehicle treated mouse (c) or mouse treated with PTH for 3 weeks (d). Vossicles can also be co-implanted with tumor cells. (e) Control sham injected without cells, or (f) ACE-1 prostate cancer cells (10,000) cells injected to vertebral bodies just prior to implantation and implants removed 4 weeks later. ACE-1 cells formed tumors and stimulated an “osteoblastic” woven bone phenotype.

4

Notes

1. Implants of BMSCs can also be placed back into the same strain of mice if it is an inbred strain of mice. 2. Surgical instruments used to isolate and implant BMSCs should be autoclaved prior to use to prevent contamination. Bone marrow stromal cells isolated from the long bones of two to three mice can be plated to one T75 flask. Therefore, two to three mice can be sacrificed at a time for BMSC harvest. 3. Ends of bones should be cut so as to preserve as much of the epiphyses as possible since the BMSCs are enriched in the trabecular bone area. Hence, a minimal amount of bone should be removed to enable access with a needle.

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4. BMSCs isolated from the long bones of two to three mice plated in one T75 flask usually reach confluency after approximately 7 days in culture. 5. The adherent BMSCs may be difficult to detach from flask. It may be necessary to gently tap the flask to aid in detaching cells. A higher concentration (i.e., 0.5%) of trypsin/EDTA and/or a cell scraper may also be used. 6. BMSCs are implanted at passage 2. Cells should be passaged when they are approximately 90% confluent. If cells reach 100% confluency they may begin to differentiate into osteoblasts, which could compromise bone formation when implanted in vivo. At passage 2, approximately 6–8 × 106 cells may be obtained from bone marrow harvested from the long bones of one 4- to 6-week-old mouse. 7. Sterile forceps can be used to hold Gelfoam in place while cutting with scalpel. 8. When preparing BMSCs for implantation, it is important to work as quickly as possible. The capacity of cells to form bone is reduced if they remain at room temperature for a long period of time. 9. It is important that mouse remains in a lateral position during surgery to place implants securely in pouches created on the backs of mice. If implants are not placed far enough laterally, they may migrate back to midline and coalesce with other implants and/or reside too close to the surgical wound. 10. A total of four implants may be placed in each mouse. 11. The ideal time to initiate injections of the anabolic agent PTH (40–80 µg/kg per day) is 1 week after implantation. Administer PTH daily for 3 weeks. 12. If BMSCs are taken from mice expressing luciferase (e.g., B6; C3-Tg TettTALuc mice from Jackson Laboratory, Bar Harbor, ME) bioluminescent imaging can be used to monitor ossicle growth as a reflection of increased numbers of BMSCs in the implants (see Fig. 7.3). 13. Surgical instruments used to isolate and implant BMSCs should be autoclaved prior to use to prevent contamination. 14. Carefully but firmly hold vertebrae with forceps when cutting single disks. One mouse donor results in approximately six vossicles. 15. Donor and host mice should be coordinated so that vossicle implantation is done just after vossicle isolation. 16. It is important that mouse remains in a lateral position during surgery to place implants securely in pouches created on the backs on mice. 17. For applications of investigating tumor/bone interface, tumor cells can be co-implanted with vossicles (see Fig. 7.4F). Just prior to implantation (see step 3) cells can be injected into the vossicle implant using a Hamilton syringe in a volume of 3–5 µL. Typically 5,000–10,000 tumor cells are used, but this varies widely depending on the growth characteristics of the cell line and the optimal numbers should be determined for each cell line. Depending on the volume of cells injected, two vertebral bodies may be used for one vossicle implant. 18. Mice bearing vossicle implants can have administration of pharmacological agent the next day after implantation. 19. Often cartilage elements are noted in the vossicles upon histological evaluation. They are typically restricted in size and inactive in appearance and can easily be omitted from calculation of bone area or other osseous parameters.

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Acknowledgments The authors acknowledge the following individuals who have contributed to the development and/or characterization of these models: Amy Koh, Paul Krebsbach, Abraham Schneider, Ana Mattos, and Jinhui Liao. Rashesh Kapadia (Scanco) is acknowledged for the micro-CT images of the ossicle model, as is Thomas Rosol for providing the ACE-1 prostate cancer cells. This work was supported by the National Institutes of Health DK53904 and CA93900.

References 1. Andreassen, T. T., Ejersted, C., Oxlund, H. (1999) Intermittent parathyroid hormone (1-34) treatment enhances callus formation and mechanical strength of healing rat fractures. J Bone Miner Res 14, 960–968. 2. Holzer, G., Majeska, R. J., Lundy, M. W., et al. (1999) Parathyroid hormone enhances fracture healing. A preliminary report. Clin Orthop 366, 258–263. 3. Skriptiz, R., Aspenberg, P. (2001) Implant fixation enhanced by intermittent treatment with parathyroid hormone. J Bone Jt Surg Br 83, 437–440. 4. Kuznetsov, S. A., Friedenstein, A. J., Robey, P. G. (1997) Factors required for bone marrow stromal fibroblast colony formation in vitro. Br J Haematol 97, 437–440. 5. Krebsbach, P. H., Kuznetsov, S. A., Satomura, K., et al. (1997) Bone formation in vivo: comparison of osteogenesis by transplanted mouse and human marrow stromal fibroblasts. Transplantation 63, 1059–1069. 6. Schneider, A., Taboas, J. M., McCauley, L. K., et al. (2003) Skeletal homeostasis in tissueengineered bone. J Orthop Res 21:859–864. 7. Pettway, G. J., Schneider, A., Koh, A. J., et al. (2005) Anabolic actions of PTH (1-34): use of a novel tissue engineering model to investigate temporal effects on bone. Bone 36, 959–970. 8. Koh, A. J., Demiralp, B., Neiva, K., et al. (2005) Cells of the osteoclast lineage as mediators of the anabolic actions of parathyroid hormone in bone. Endocrinology 146, 4584–4596.

Chapter 8

Utility of the Ovariectomized Rat as a Model for Human Osteoporosis in Drug Discovery Yogendra P. Kharode, Michael C. Sharp, and Peter V.N. Bodine

Abstract Ovariectomy-induced osteopenia in the rat produces skeletal responses similar to that in a post-menopausal woman. In the ovariectomized (ovx) rat, high bone turnover, and subsequent bone loss, like in the human post-menopausal condition, can be prevented by estrogen replacement. Because of the striking resemblance of skeletal responses in humans and rats in the state of estrogen deficiency, the ovx rat is considered to be a gold standard model for evaluating drugs for prevention and reversal of osteoporosis. This chapter describes the procedure for performing ovariectomy on the rat and the utility of the ovx rat model we have utilized over the last two decades in our laboratory. Keywords Ovariectomy, bone mineral density, peripheral dual x-ray absorptiometry, peripheral quantitative computed tomography, micro-CT.

1

Introduction

Osteoporosis is a disease characterized by reduction in bone mass, micro-architectural deterioration, and a decrease in biomechanical competence of appendicular and axial skeleton resulting in increased incidence of fractures (1). A variety of physiological and pharmacological conditions are known to cause osteoporosis—of which, the most prevalent is post-menopausal osteoporosis. Estrogen deficiency, as a consequence of menopause, dramatically increases bone turnover (2, 3)—with bone resorption far exceeding bone formation. Estrogen receptors are present in both osteoblasts (4, 5) and osteoclasts (6). Estrogen also promotes development of osteoblasts, increases their proliferation and increases a number of important osteoblast proteins like insulin-like growth factor-1, transforming growth factor-β, and bone morphogenic protein-6 (7–9). At high concentrations, estrogen increases histomorphometric indices of bone formation possibly via increasing synthesis of osteoblastic growth factors. Estrogen restricts osteoclast recruitment and suppresses bone resorptive activities mediated by stromal or monocytic cellular cytokines (IL-1, IL-6, TNF) and RANKL (10, 11). Estrogen also inhibits osteoclast differentiation and recruitment From: Methods in Molecular Biology, Vol. 455: Osteoporosis: Methods and Protocols Edited by Jennifer J. Westendorf © Humana Press, Totowa, NJ

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by increasing osteoblastic TGFβ (8). Moreover, estrogen increases the cellular level of nitric oxide, which is a potent inhibitor of bone resorption (12). In addition to decreased osteoblastic activity and increased osteoclastic activities, decreased intestinal calcium absorption (13) and increased urinary calcium excretion (14) contribute to the excessive bone loss in the estrogen-deficient state. Estrogen therapy (ET)—although associated with endometrial and breast cancer risk in a limited population—has been utilized and proven to be effective in prevention of postmenopausal bone loss and the incidence of fractures (15–18). Moreover, because estrogen deficiency results in an increase in bone turnover, mainly favoring an increase in bone resorption, a number of non-hormonal therapies also have been used to counteract the deleterious effects in both humans and rats. These non-hormonal agents act by attenuation of bone resorption (19, 20), enhancement of bone formation (21, 22) or dual-bone anti-resorptive and osteogenic-actions (23, 24). The rat skeleton has many similarities to that of humans with the exception of continuous modeling and longitudinal bone growth in the rat (25). The skeletal responses to depletion and repletion of estrogen (in the estrogen deficient state), as well as the pharmacological effects of known agents on the rat skeleton, are comparable with that in humans; thus, making the rat the species of choice for pre-clinical evaluation of new drugs as potential future therapies for human metabolic bone disorders such as osteoporosis.

1.1

Ovariectomy-Induced Osteopenia Model

The striking resemblance of the ovx rat to humans with respect to estrogen deficiency related pathophysiology—i.e., increase in bone turnover, bone loss—osteopenia (26–29) and prevention of the same by estrogen replacement (30–32) make the ovx rat a gold standard model of human osteoporosis. Since the ovx model effectively demonstrated the skeletal response of known agents such as conjugated equine estrogen (31) and parathyroid hormone (33, 34) in a fashion similar to that in postmenopausal women, the ovx rat model has been an animal model of choice for evaluation of new therapies for both prevention (8, 35–41) and treatment (21, 22, 42–45) of osteoporosis. In fact, the Food and Drug Administration also has recommended the utilization of the ovx model for pre-clinical evaluation of anti-osteoporotic drugs (46).

2 2.1

Materials Surgical Procedure

1. Surgical Instruments and Supplies: Clippers, scissors, disposable cauterizers, blunt forceps, tissue forceps, hemostats, scalpels #10 and #15 blades, 3-0 suture, gauze packs, syringe/needle (25 G), surgical apparel.

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2. 3. 4. 5. 6.

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Betadine. Ethyl alcohol. Ketamine (Bristol Laboratories, Syracuse, NY). Xylazine (Bristol Laboratories). Acepromazine (Aveco, Ft. Dodge, IA).

2.2

Evaluation of Agents in OVX Models

1. Ketamine, xylazine, and acepromazine as listed in Section 2.1. 2. Syringe/needle (25 G) and surgical apparel. 3. Ear-tags.

2.3

BMD Measurements Using Peripheral Dual X-Ray Energy Absorptiometer

1. Peripheral dual x-ray energy absorptiometer (pDXA), Sabre (Norland Corp., Ft. Atkins, WI).

2.4

BMD Measurements Using Peripheral Quantitative Computed Tomography

1. XCT Research SA+ (pQCT; Stratec Medizintechnik, Pforzheim, Germany).

2.5

Histology

1. Automated tissue processor (Shandon Hypercenter XP, Pittsburgh, PA). 2. Microtome (Leica RM2065, Heidelberg, Germany). 3. Bioquant Osteo image analysis system (Bioquant Image Analysis Corp., Nashville, TN). 4. Nikon E800 microscope (Nikon Corp., Melville, NY). 5. Methyl methacrylate (MMA) (Aldrich, St. Louis, MO). 6. Silane-coated positively charged slides (Surgipath Snowcoat X-tra Micro Slides, Richmond, IL).

2.6

Bio-Mechanical Testing

1. Instron 5543 with Instron 1 kN. load cell (Instron Corp., Canton, MA). 2. Isomet diamond blade wafer saw (Buehler Ltd., Lake Bluff, IL).

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Biochemical Analysis of Serum/Urine Parameter

1. Hitachi 911 analyzer. 2. Biomarker (osteocalcin, CTX, PYD) assay kits from suppliers.

3

Methods

Rats from a wide range of age groups (8 weeks to 15 months) and a variety of treatment periods are used by different investigators for examining the effects of agents in the ovx model. While use of older animals for the ovx model is attractive because of lower bone modeling and steady bone turnover rate, with careful study design and proper controls, the ovx model using young adult rats also can provide consistent, reproducible, and interpretable results (see Notes 1 and 2).

3.1

Ovariectomy Procedure

Bilateral ovariectomy (or sham operation) may be performed as follows: 1. Anesthetize rats with an intraperitoneal injection of 45 mg/kg ketamine, 8.5 mg/ kg, xylazine, and 1.5 mg/kg acepromazine, shave the fur over the dorsal lumbar area, disinfect the skin with Betadine followed by an alcohol rinse. 2. Make a 2-cm skin incision along the dorsal midline (just caudal to the last rib) and through the abdominal musculature (Fig. 8.1). 3. Gently grasp the ovarian fat pad using forceps, expose and remove the ovary. Use cautery to control any bleeding (Fig. 8.2).

Fig. 8.1 Incision site for ovariectomy in rats.


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Fig. 8.2 Anatomy of the urogenital system of the female rat.

4. Stitch the muscle with 3-0 absorbable sutures and use stainless steel wound clips (two to three each side) to close the skin incision. The wound clips may be removed in 7 to 10 days post-surgery. 5. Sham operation may be performed using above steps but without removing the ovaries. Surgery takes about 10 minutes per animal. Instruments are sterilized between animals using a glass bead sterilizer.

3.2

Prevention Protocol

Bone anti-resorptive agents like estrogens, bisphosphonates, selective estrogen receptor modulators (SERMs), and others can be evaluated for their bone sparing activities using the protocol listed in the following. We have utilized the prevention protocol for evaluation of agents such as CEE (31) and SERMs (40) following 5 or 6 weeks of treatment in ovx rats. 1. Weigh and ear-tag ovx and sham operated rats 2 days after surgery and divide them randomly in groups of 8 to 10 rats/group so that the average body weights for each ovx group is similar. Provide casein diet and water ad libitum. 2. Initiate daily treatment and continue the regimen for 6 weeks. Administer 10 mg/kg calcein, sc 2 and 9 days before necropsy (see Note 3). 3. During the study, at the desired time point, collect urine overnight using metabolism cages. 4. Prior to necropsy, evaluate bone mineral density using pDXA and/or pQCT as described in the following sections.

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5. Two hours after the last day’s treatment, euthanize animals by over exposure to CO2. Collect tibiae, femurs, spine, blood, and uteri. Other tissues may also be collected as required by the study design. 6. Place tibia in 40% ethyl alcohol (EtOH), store femurs and spine at −20 °C. Separate serum/plasma as needed and store as required by assay conditions. Excised bones can be analyzed for skeletal parameters by techniques listed in the following sections.

3.3

Evaluation of Agents in OVX Rats with Established Osteopenia

Ovx produces significant trabecular bone loss in long bones of rats in 3 to 4 weeks after the surgery. Osteogenic agents like PTH reverse the ovx-induced bone loss. The length of treatment period usually depends on the severity of osteopenia, e.g., if the osteopenia development period (post-ovx) is longer prior to initiation of the osteogenic regimen, the reversal of osteopenia typically requires longer treatment period. We have observed that the 3 weeks post-ovx osteopenia development period is adequate for establishing a significant trabecular bone loss in mature Sprague-Dawley rats and the lost bone may be completely restored by daily sc administration of 10 µg/kg per day, hPTH for 3 weeks (43). Maintenance of therapeutic effects of hPTH by anti-catabolic agents like estrogen and SERMs may also be evaluated using this protocol (47). However, longer durations of osteopenia are also utilized to establish a lower remodeling turnover model. The variation of this protocol from the prevention protocol are listed below: 1. Weigh and ear-tag ovx and sham operated rats 2 days after surgery and divide them in groups of 8 to 10 rats/group. Provide casein diet and water ad libitum. 2. Four weeks after ovx, evaluate base line bone mineral density using pDXA and/ or pQCT as described in the following. 3. Initiate daily treatment and continue for 4 weeks. Administer 10 mg/kg calcein, sc 2 and 9 days before necropsy (see Note 3). 4. Prior to necropsy, evaluate bone mineral density using pDXA, and/or pQCT as described in the following. 5. Two hour after the last day’s treatment, euthanize animals by overexposure to CO2. Collect tibiae, femurs, spine, blood, and uteri. 6. Place tibia in 40% EtOH, store femurs, and spine at −20 °C. Separate serum/ plasma as needed and store as required by assay conditions. For evaluation of skeletal parameters (in vivo and in excised tissues) and other parameters, we have utilized the following techniques in our laboratory.


3.4

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Evaluation of Areal BMD of 4th Lumbar Vertebra (L4) Using pDXA

BMD of the fourth lumbar vertebra (L4) may be evaluated in anesthetized rats using a pDXA. The dual energy x-ray absorptiometry (pDXA) measurements in the ovx (or sham operated) rat is performed as follows: 1. Fifteen minutes prior to DXA measurements, the anesthetized rat is placed on the Plexiglas window under the scanner perpendicular to the path of scanner aligning the spine straight in the center of Y axis. 2. A preliminary scout scan (6 × 6 cm) is performed at a scan speed of 40 mm/ second with a scan resolution of 1.0 × 1.0 mm to determine the region of interest in L4. 3. A measurement-scan (4 × 4 cm) at a scan speed of 5 mm/second and a resolution of 0.2 × 0.2 mm is taken for precise BMD measurement of L4. 4. The software allows the operator to define the total region of L4 from the images generated by the computer following the scan. The BMD is computed by the software as a function of attenuation of the dual beam (28 and 48 KeV) x-ray generated by the source underneath the subject and the detector traveling along the defined area above the subject. 5. The data for BMD values (expressed in g/cm2) and individual scans may be stored and later subjected to the statistical analysis (see Note 4).

3.5

Evaluation of Volumetric BMD of Proximal Tibia Using pQCT

Total and trabecular density of the proximal tibia can be evaluated in anesthetized rats using an XCT Research SA+ (see Note 5). 1. The right hind limb is passed through a polycarbonate tube with a diameter of 25 mm and taped to an acrylic frame with the ankle joint at a 90° angle and the knee joint at 180°. The polycarbonate tube is affixed to a sliding platform that maintained it perpendicular to the aperture of the pQCT. The platform is adjusted so that the distal end of the femur and the proximal end of the tibia would be in the scanning field. 2. A two-dimensional scout view is run for a length of 10 mm and a line resolution of 0.2 mm. After the scout view is displayed on the monitor, the proximal end of the tibia is located. The pQCT scan is initiated 3.4 mm distal from this point. The pQCT scan of 0.5 mm thickness with a voxel (three-dimensional pixel) size of 0.07 mm is obtained by 360 projections through the slice. 3. After the pQCT scan is completed, the image is displayed on the monitor. A region of interest including the tibia, but excluding the fibula is outlined. The soft tissue is automatically removed using an iterative algorithm.

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4. The density of the entire scanned bone (total density) is reported in mg/cm3. The outer 55% of the bone is peeled away in a concentric spiral. The density of the remaining bone (trabecular density) is reported in mg/cm3. 5. The results are saved in a database file for statistical analysis and the original scan are saved in the hard disk.

3.6

Micro-Computed Tomography

Three-dimensional volumetric analysis of the excised proximal tibia can be performed using Scanco µCT 40 (48, 49). 1. A scout view of the tibia is performed to include the proximal end extending approximately 7 mm distally and parallel to the growth plate. 2. A lead reference line for the CT measurement is placed at the proximal end of the tibia. Measurement begins at this point and continued distally for 350 scan slices (20 µm thickness). 3. A region of interest (ROI) distal to the growth plate is selected for analysis. 4. Images are reconstructed into three-dimensional arrays with an isotropic voxel size of 20 µm. The resulting gray-scale images are segmented using a constrained 3D Gaussian filter (σ = 1.0, support = 0.8), and bone structure is extracted using a fixed threshold (normally around 20% of maximal grayscale value; see Note 6). The trabecular bone within the proximal tibia is identified using manually drawn contouring algorithms on the region of interest (primary and secondary spongiosa). 5. Morphometric variables are computed within the volume of interest from the binary images. Morphometric analysis is performed by a direct, three-dimensional technique using Scanco software. Morphometric parameters included in the analysis are bone volume fraction (BV/TV, %), connectivity density (Conn.D.), trabecular number (Tb.N., mm−1), trabecular thickness (Tb.Th. µm) and trabecular spacing (Tb.Sp., µm) (see Notes 6, 7, and 8).

3.7

Histology

3.7.1

Evaluation of Histomorphometric Parameters in Proximal Tibia Samples

1. The proximal tibia is prepared for tissue processing, following necropsy, by cutting longitudinally through the anterior eminence with a scalpel to provide both a stable base for orientation and to allow rapid penetration of fixative into the cancellous bone. The diaphysis is then cross sectioned 1.5 cm from end of the proximal metaphysis and the resulting sample is placed into a labeled cassette and fixed in cold, approximately 5–8 °C, 40% EtOH for 48 hours for preservation


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of cellular adhesion to bone surfaces before being placed into cold 70% EtOH for subsequent processing and storage. 2. The metaphysis is placed in an automated tissue processor (Shandon Hypercenter XP) and infiltrated with MMA. The MMA infiltrated specimen is embedded with the cut surface of the anterior eminence placed face down along the bottom of a glass vial containing MMA polymerization mixture. The open vials are then put under vacuum for 3 hours and nitrogen is run over samples before capping. Complete polymerization by placing samples in a water bath at 37 °C overnight. 3. Samples are rough ground to expose the longitudinal midline of the metaphysis from which thin and thick sections are taken on a microtome (Leica RM2065) containing a tungsten carbide knife. Sections are collected on positively charged glass slides with thick sections cut to 10 µm for observation of dynamic indices and thin sections cut to 5 µm for subsequent staining and evaluation of static parameters (see Note 9).

3.7.2

Acquisition of Dynamic and Static Histomorphometric Measures in Rat Proximal Tibia

Dynamic bone indices are evaluated from fluorescein (calcein) labeled proximal tibia sectioned at 10 µm on a Bioquant Osteo image analysis system. 1. Cancellous bone in the region of secondary spongiosa, which begins approximately 450 µm below the endochondral plate, is examined with a Nikon E800 microscope using plan fluor objectives for mineralized trabecular surface for a total length of 1,500 µm into the diaphysis taking care not to include cortical bone. 2. Fluorescence emission of the calcein labels is achieved by exciting in the blue spectrum at a wavelength of 465–495 nm to reduce background signal. Mineral apposition rate (MAR) is determined by measuring the inter-label distance at 20 µm intervals between the mineralized calcein double labels and calculating the mean inter-label distance divided by the inter-label period. The resulting primary histomorphometric index is reported as µm/day. To account for error, in measuring the inter-label distance, due to oblique orientation of mineralizing surfaces the mean inter-label distance is modified by a divisor of 1.2 to improve accuracy of the measure (50, 51). 3. Total mineralized surface in the region of the secondary spongiosa is measured to include individual single label length and double label length to determine the mineralizing surface (MS = sum double label surface + ½ sum single label surface) as well as the total bone surface to derive the bone formation rate (BFR). BFR = MAR × (MS/BS) expressed in mm3/mm2 per day 4. Static parameter measures (bone volume, osteoid volume, osteoblast, and osteoclast number and surface) are taken from 5-µm sections, which are stained to reveal cellular detail and distinguish structural components using a modified Goldner’s trichrome or 0.1% toluidine at pH 3.5.

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Bio-Mechanical Testing: Compressive Strength of Vertebrae

The compressive strength of fifth lumbar vertebra (L5) is determined using Instron 5543 equipped with a 1,000 Newton (N) load cell as follows (43). 1. Spine is thawed at room temperature and L5is dissected. 2. Using a low speed Isomet diamond blade wafer saw (Buehler Ltd., Lake Bluff, IL), remove all processes. Two coplanar cuts perpendicular to the cephalocaudal axis 4.9 mm apart are made. This produces a uniform sample for compression. 3. The sample is placed in a fixture with a piston moving parallel to the cephalocaudal axis compressing the sample. 4. Data are transmitted to Instron Merlin software, which produces a load-deformation curve. Maximum load is calculated from the curve and expressed as Newtons (N).

3.9

Biochemical Parameters

Serum bone formation parameter (osteocalcin) and bone resorption parameter (CTX, cross-lap) as well as urinary pyridinoline cross-links (PYD, a bone resorption parameter) may be measured using commercially available kits and the procedures described by the manufacturers of the assay kits. Serum cholesterol may be analyzed on a Hitachi 911 instrument using Boehringer Mannheim reagents (for cholesterol). The cholesterol is measured via o-quinone imine dye (which is formed following enzymatic reactions with cholesterol) and is analyzed photometrically at 505 nm at 37 °C and expressed as mg/dL (40).

3.10

Closing Remarks

Many similarities in skeletal effects of estrogen deficiency between rat and human allow the utility of ovx rat as a gold standard model of human osteoporosis. The limitations of the rat model, however, are: (1) continuous modeling through out life, and (2) minimal cortical effects of ovx (49). Nevertheless, the ovx rat model can be used for evaluation of agents for both prevention and treatment of osteoporosis. The ovx rat model with proper study design and use of densitometry, µCT, histomorphometry, bio-mechanical testing, and bio-chemical markers may allow investigators to collect interpretable data on test compounds that can possibly help in predicting clinical effects of the compounds in humans with osteoporosis.


4

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Notes

1. In our ovx studies, we have routinely utilized 10 to 12 weeks old Sprague Dawley rats weighing 225 to 250 g. 2. Alternatively, ovx and sham operated rats may be purchased from suppliers such as Taconic Farms (Germantown, NY) or Charles River (Wilmington, MA). 3. Other chromophores such as tetracycline, demeclocycline, or xylenol orange may also be used to label the bone. The interval between the two labels may be changed as well. 4. We use n = 8 to 10 rats/group for evaluation of skeletal effects in rats. 5. Areal BMD of tibia may also be analyzed using the preceding procedure with proper positioning of the rat and defining the region of interest from the image. 6. This method may be utilized using other similar pQCT equipment such as XCT Research M or XCT 960M (43). 7. Threshold value must be set prior to analyzing the first sample of the study and should be kept the same for entire study. 8. Bone histomorphometry parameters are accepted from standardized nomenclature by Parfitt et al. (52). 9. Micro-CT evaluation of vertebrae samples may be performed using the same technique (described for tibia) by scanning the whole vertebrae and analyzing the region between the two end plates. Acknowledgment The authors wish to dedicate this article to our late director, Shunichi Harada, who devoted his professional life to osteoporosis research. We also thank Fred Bex and Barry Komm for their guidance, direction, and support of the Wyeth osteoporosis research group. Finally, they are grateful to many talented scientists whose excellent technical support over the last two decades have been instrumental in developing and implementing animal models and analytical methods in their laboratory: special thanks to Joseph Tamasi, James Morzolf, Michelle French, Stephani Smoluk, Andrea Scarcia, Vanessa Dell, Mellisa Wasco, Paula Green, Sally Selim, Colleen Millgan, and Jennifer Pirrello as well as the BioResources staff at Wyeth Research for their valuable assistance.

References 1. Consensus Development Conference. (1993) Diagnosis, prophylaxis and treatment of osteoporosis. Am J Med 94, 646–650. 2. Heaney, R. P., Recker, R. R., Saville, P. D. (1978) Menopausal changes in bone remodeling. J Lab Clin Med 92, 964–970. 3. Garnero, P., Sornayrendu, E., Chapau, M. C., et al. (1996) Increased bone turnover in late postmenopausal woman is a major determinant of osteoporosis. J Bone Miner Res 11, 337–349. 4. Komm, B. S., Terpening, C. M., Benz, D. J., et al. (1988) Estrogen binding receptor mRNA, and biologic response in osteoblast-like osteosarcoma cells. Science 241, 81–84.

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5. Ericksen, E. F., Colvard, D. S., Bergn N. J., et al. (1988) Evidence of estrogen receptors in normal human osteoblast-like cells. Science 241, 84–86. 6. Oursler, M., Osdoby, P., Pyfferon, J., et al. (1991) Avian osteoclasts as estrogen target cells. Proc Natl Acad Sci USA 88, 6613–6617. 7. Ernst, M., Heath, J. K., Rodan, G. A. (1989) Estradiol effects on proliferation, messanger ribonucleic acid for collagen and insulin-like growth factor-1, and parathyroid hormone-stimulated adenylate cyclase activity in osteoblastic cells from calvariae and long bones. Endocrinology 125, 825–833. 8. Yang, N. N., Bryant, H. U., Hardikar, S., et al. (1996) Estrogen and raloxifene stimulate transforming growth factor-beta-3 gene expression in rat bone: a potential mechanism for estrogenor raloxifene-mediated bone maintenance. Endocrinology 137, 2075–2084. 9. Rickard, D. J., Hofbaur, L. C., Bonde, S. K., et al. (1998) Bone morphogenic protein-6-production in human osteoblast-like cell lines: selective regulation by estrogen. J Clin Invest 101, 413–422. 10. Girasolle, G., Jilka, R. L., Passeri, G., et al. (1992) Marrow-derived stromal cells and osteoblasts in vitro: a potential mechanism for the anti-osteoporotic effects of estrogens. J Clin Invest 89, 883–891. 11. Rogers, A., Estell, R. (1998) Effects of estrogen therapy of postmenopausal women on cytokines measured in peripheral blood. J Bone Miner Res 13, 1577–1586. 12. Armour, C. E., Ralston, S. H. (1998) Estrogen upregulates endothelial constitutive nitric oxide synthase expression in human osteoblast-like cells. Endocrinology 139, 799–802. 13. Arjandi, B. H., Sahil, M. A., Herbert, D. C., et al. (1993) Evidence for estrogen receptorlinked calcium transport in the intestine. Bone Miner 21, 63–74. 14. Heaney, R. P., Recker, R. R., Saville, P. D. (1978) Menopausal changes in calcium balance performance. J Lab Clin Med 92, 953–963. 15. Recker, R. R., Saville, P. D., Heaney, R. P. (1977) Effects of estrogen and calcium carbonate on bone loss in postmenopausal women. Ann Int Med 87, 649–655. 15. Gallagher, J. C., Kable, W. T., Goldgar, D. (1991) Effects of progestin therapy on cortical and trabecular bone: comparison with estrogen. Am J Med 90, 171–178. 15. Vedi, S., Compston, J. E. (1996) The effect of long-term hormone replacement therapy on bone remodeling in postmenopausal women. Bone 19, 535–539. 16. Lindsay, R., Cosman, F. (2006) Effects of estrogen intervention on the skeleton, in (Favus, M. J., ed.), Primer on the Metabolic Bone Disease and Disorders of Mineral Metabolism, 5th ed. The American Society for Bone and Mineral Research, Washington, DC. 17. Kalu, D. N., Liu, C. C., Salerno, E., et al. (1991) Skeletal response of ovariectomized rats to low and high doses of 17-beta estradiol. Bone Miner 14, 175–187. 18. Plosker, G. L., Goa, K. L. (1994) Clodronate: a review of its pharmacological properties and therapeutic efficacy in resorptive bone disease. Drugs 47, 945–982. 19. Plum, L. A., Fitzpatrick, L. A., Ma, X., et al. (2006) 2MD, a new anabolic agent for osteoporosis treatment. Osteoporosis Int 17, 704–715. 20. Dempster, D. W., Cosman, F., Parisien, M., et al. (2006) Anabolic actions of parathyroid hormone on bone. Endocr Rev 14, 690–709. 21. Shen, V., Dempster, D. W., Mellish, R. W. E., et al. (1992) Effects of combined and separate intermittent administration of low-dose human parathyroid hormone fragment (1-34) and 17β-estradiol on bone histomorphometry in ovariectomized rats with established osteopenia. Calcif Tiss Int 50, 214–220. 22. Wronski, T. J., Yen, C. F., Scott, K. S. (1991) Estrogen and diphosphonate treatment provide long-term protection against osteopenia in ovariectomized rats. J Bone Miner Res 6, 387–394. 23. Jee, W. S. S., Yao, W. (2001) Overview: animal models of osteopenia and osteoporosis. J Musculoskel Neuron Interact 1, 193–207. 24. Wronski, T. J., Lowery, P. L., Walsh, C. C., et al. (1985) Skeletal alterations in ovariectomized rats. Calcif Tissue Int 37, 324–328. 25. Wronski, T. J., Walsh, C. C., Ignaszewski, L. A. (1986) Histologic evidence for osteopenia and increased bone turnover in ovariectomized rats. Bone 7, 119–123.


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Chapter 9

Orchidectomy Models of Osteoporosis Stéphane Blouin, Hélène Libouban, Marie Françoise Moreau, and Daniel Chappard

Abstract Long considered a disease of post-menopausal women, osteoporosis is increasingly being recognized among the growing population of elderly men. Androgen deficiency may be associated with an increase of bone resorption in elderly men and so, with remodeling imbalance and fracture risk. It is firmly established that androgen withdrawal induced by orchidectomy (ORX) results in decreased bone mass in animal models especially in rodents. The mature rat is the model of choice. Skeletal effects of ORX in rats have been studied at the tissular and cellular level. It induces a decrease of BMD and BV/TV with microarchitecture alterations due to an increased bone remodeling. The present chapter focuses on the ORX surgery in rats and mice. Keywords Orchidectomy, osteoporosis, rat, mice, DEXA, histomorphometry.

1

Introduction

Long considered a disease of post-menopausal women, osteoporosis is increasingly being recognized among the growing population of elderly men. There is increasing evidence for a relationship between age-related endocrine changes and osteoporosis in men. Androgen deficiency may be associated with an increase of bone resorption in elderly men and so, with remodeling imbalance and fracture risk (1). The Food and Drug Administration (FDA) and World Health Organization (WHO) have recommended the ovariectomized (OVX) rat for postmenopausal osteoporosis research (2); the orchidectomized rat (ORX) has been proposed to simulate male osteoporosis due to hypogonadism (3–15) and to evaluate injectable biomaterials (16). ORX mice were also used in few studies (17–19). This model is interesting to evidence regulation pathway in transgenic mice (20, 21). In this introduction, we will discuss the major results obtained in the rat model. The most important drawback of the rat skeleton is its lifelong growth; however, the modeling influence can be minimized by using mature animals with preponderant bone remodeling (from 4-month-old rats) (5, 9, 22). Moreover, analysis should From: Methods in Molecular Biology, Vol. 455: Osteoporosis: Methods and Protocols Edited by Jennifer J. Westendorf © Humana Press, Totowa, NJ

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be performed on the secondary spongiosa to avoid the remaining modeling activity in the primary spongiosa. Skeletal effects of ORX have been studied at the tissular and cellular level by several techniques: dual energy X-ray absorptiometry (DEXA), histomorphometry, serum dosage, and X-ray microtomograph. By DEXA, a decrease of the femoral bone mineral density (BMD) in the ORX rat was evident between 1 to 4 months postsurgery (5) but histomorphometric evaluation revealed the trabecular bone loss more rapidly and with greater sensitivity. Decreases in BV/TV in tibia were found between 2 and 3 weeks post-ORX (5, 9, 10, 23), whereas DEXA found differences after 8 weeks (24, 25). Histomorphometry showed decreased trabecular bone volume (BV/TV) of 40% in OVX mice, whereas DEXA indicated a decrease of BMD of 5% (26). ORX also induced a decrease of vertebral BV/TV between 2 and 3 weeks (5, 23) and cortical bone loss associated with increased endosteal porosity (15, 22). Histomorphometry allows measurement of microarchitecture and bone remodeling parameters. ORX induces a decrease of trabecular number and an increase of trabecular separation without alteration in trabecular thickness (9, 23, 25). Fractal dimension of the trabecular network most efficiently describes trabecular alterations because it is the earliest altered parameter (2 weeks) (25). Trabecular bone loss due to ORX is associated with increased bone remodeling. In adult rat, the osteoclast number and the osteoblastic surfaces increased early after the androgen deprivation (27). Increased bone remodeling was found 1 month post ORX; this increase was found to be transient and occurred before the trabecular bone loss observed 4 month after ORX (28). The bone remodeling can be measured by the dosage of markers evaluating resorption (TRAcP, deoxypyridinoline, and pyridinoline) or formation (osteocalcin) (23, 29–31). The mineralization rate can be affected transiently by ORX (5, 9) or not (28). In cortical bone, ORX induces a decrease of periosteal bone formation (5). The combination of different osteopenic factors has also been proposed to obtain severe bone loss in the rat. Iwamoto et al. studied the conjoined effects of ORX and sciatic neurectomy in adult rats. The association of both factors was associated with a more pronounced decrease of trabecular bone mass and increase of cortical porosity than with neurectomy alone (6, 7). Massive bone loss was obtained in a study combining the effects of ORX (acting on the whole skeleton) and paralysis induced by injection of botulinum toxin, acting on a single hind limb (31). The association of both factors led to considerable bone loss with marked alterations in bone microarchitecture (marked disconnection of trabeculae). The cumulative effect of disuse and ORX induced a peak of serum TRAcP higher than ORX alone (31). This chapter focuses on the ORX surgery in rats and mice.

2

Materials

Use of trade names is for identification purposes only and does not imply endorsement.

9 Orchidectomy Models of Osteoporosis

2.1

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Anesthesia

Anesthesia of rats or mice is performed with one of the following agents (see Note 1).

2.1.1

Isoflurane

Isoflurane is the method of choice for rodent anesthesia. It is safe and very easy to use. Induction and awakening are rapid. Gas waste must be scavenged properly. The following material is needed: 1. An induction chamber in order to progressively put to sleep a conscious animal. 2. An oxygen flowmeter to regulate flow of the oxygen bottle so that the outlet pressure is compatible with its use. 3. A conical mask adaptable to the animal’s snout to maintain animals under anesthesia. 4. A gas vaporizer filled with isoflurane. It ensures a stable concentration of the gas mixture and provides a continuous flow of anesthetic. 5. HALOSORB cartridges (Minerve, Esternay, France), which are activated carbon cartridges used to trap residual anesthesia particles protect the operator against residual anesthetic.

2.1.2

Xylazine–Ketamine

1. Rompun containing the active ingredient xylazine, solution 2% (Bayer Pharma, Puteaux, France) (20 mg xylazine/mL). It is currently used in veterinary medicine for sedation, analgesia and muscle relaxation. 2. Ketalar containing the active ingredient ketamine chlorohydrate (Parke-Davis, Courbervoie, France) (50 mg ketamine/mL). Ketamine is a powerful anesthetic used primarily in veterinary medicine for anesthesia of farm animals during routine surgery. 3. Syringes of 1 or 2 mL according to animal weights. 4. Hypodermic needles 26 G.

2.2

Animal Preparation

1. An infrared lamp to prevent heat loss (see Note 2). 2. An electric clipper or a depilatory cream to remove hair/fur from the area of surgery. 3. A weight scale to adapt the volume of anesthetic to the weight of each animal.

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Surgery (Fig. 9.1A) A beaker to immerse surgery instrument into disinfectant solution. Ethanol 70%. Michel clips for the wound closure (see Note 3). Betadine, which contains a 10% povidone-iodine solution to spray the surgical site.

Fig. 9.1 Methodology of orchidectomy. (a) Surgical equipment, (b) induction equipment of isoflurane anesthesia, (c) anesthesia maintenance of the rat using the conical mask, (d) site of incision (after hair/fur removal), (e) skin and muscle incision; testis is visible, (f) cauda epididymis, testis, caput epididymis, vas deferens and testicular blood vessels are extracted from the scrotum sac, (g) testicular blood vessels are visible after careful disconnection of epididymis and fatty tissue from testis, (h) ligature around blood vessels, (i) removal of testis 2 mm under the node.


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5. 6. 7. 8.

Surgical blunt forceps. Michel clip applying forceps. Fine straight scissors. Vicryl (polyglactin 910) suture, which is a synthetic absorbable braided sterile surgical suture (Ethicon, Johnson and Johnson, Somerville, NJ). The suture gauge should be 2-0 for rats and 4-0 for mice. 9. Sterile gauze. 10. Lab coat. 11. Sterile gloves.

3

Methods

3.1

Anesthesia

3.1.1

Isoflurane

1. 2. 3. 4.

Place the animal in the induction chamber (see Note 4) (see Fig. 9.1B). Adjust the flowmeter on the oxygen bottle to 3 L/minute. Adjust the isoflurane vaporizer to 5%. Within approximately one minute for mice and 2 minutes for rats, the animal will become anesthetized. Initially, respiratory rate will increase and then decrease. First the animal is agitated then losses its stability. After that, it falls asleep and the animal is unreacting to noise. A simple test is to knock on the induction chamber. 5. When the animal is anesthetized, it is removed from the induction chamber and placed on the surgical area with the snout in the conical mask. Prevent heat loss until the animal recovers by using the infrared lamp (see Fig. 9.1C). 6. For maintenance, adjust the isoflurane vaporizer to 3%.

3.1.2

Xylazine–Ketamine

1. If the animal is anesthetized with a combination of xylazine and ketamine, dosage must be adapted to the weight of the animal. 2. Prepare a syringe with the proper mixture of ketamine (100 mg/kg) and xylazine (10 mg/kg). For mice, a dilution of ketamine (1:10) and xylazine (1:20) should be prepared due to small weight of these animals (20 g on average). 3. The anesthetic mixture is injected intraperitoneally with a 26G needle (see Note 5). 4. Repeat when necessary, half a dose at a time (approx. every 30 minutes). 5. Prevent heat loss until the animal recovers by using the infrared lamp.

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Monitoring of Anesthesia Depth

The anesthesia depth must be checked to ensure that the animal is not too lightly anesthetized (i.e., that it does not experience pain or regain consciousness), or too deep (since vital functions can be compromised, leading to death). One can pinch the foot pad, tail, or ear of the animal to assess its anesthetic state. Any reaction from the animal indicates that it is too lightly anesthetized. Respiratory pattern and frequency give an indication of anesthesia depth as well. A decreasing respiratory rate is an indicator of increasing depth of anesthesia.

3.2

Animal Preparation

1. When the animal is properly anesthetized (reduced respiratory rate, lack of spontaneous movement to foot pad pinch), position it in dorsal recumbence. 2. Shave the hair/fur surrounding the scrotal sacs. 3. On the shaved area, apply a Betadine scrub using a sterile compress, followed by a final soaking with Betadine solution.

3.3

Surgery

3.3.1

Preparation of the Surgical Area, the Instruments, and the Operator

1. Aseptic surgical techniques must be used for this procedure. The operator must wear a lab coat (or surgical gown) and sterile gloves. 2. The surgical area should be located in a laboratory zone that is not heavily traveled. 3. Hard surface (table tops or equipment) must be disinfected with ethanol 70%. The contact time should be several minutes. 4. All instruments must be pre-sterilized by acceptable methods, such as steam sterilization. When performing surgery on more than one animal, effective sterilization can best be achieved by pre-sterilization of multiple sets of instruments. Dipping instruments in 70% alcohol between surgeries does not achieve sterility (>30 hours of contact time required) and is not an acceptable method. 3.3.2

The Surgical Procedure

Two incisions are made to separately reach each testis. Each incision is done with a scissor. Both sides are performed separately one after another from the incision to the suturing. 1. Check that the testes are localized within the scrotal sacs (see Note 6) (see Fig. 9.1D).


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2. Make an incision through the skin on the ventral side of the scrotum (about 1 cm for rats and 2 mm for mice). 3. Cut the cremaster muscles with a small incision (see Fig. 9.1E). 4. Localize the testicular fat pad and pull it through the incision using a blunt forceps. 5. Pull out the cauda epididymis together with the testis, followed by the caput epididymis, the vas deferens, and the testicular blood vessels (see Fig. 9.1F). 6. Disconnect the cauda epididymis from the testis. 7. Disconnect carefully the caput epididymis embedded in fat from the testis and avoid cutting blood vessels (see Fig. 9.1G). 8. Perform a single ligature (two or three nodes and cut with 1 mm of free end suture) around the blood vessels (see Fig. 9.1H). 9. Remove the testis by cutting with scissors and make sure no bleeding occurs (see Fig. 9.1I). 10. Replace the remaining pieces of the vas deferens, the fat, and the blood vessels back in the scrotal sac with the blunt forceps. 11. Join the lips of the wound by using the blunt forceps and close the skin with Michel clips (about three to four clips per incision) by using applying forceps. 12. Repeat steps 1 to 8 for the other testis. 13. After both testes have been removed (see Note 7), return the animal to its cage (see Note 8) and monitor anesthesia recovery as described in the following.

3.4

Surgical Recovery

1. One must prevent heat loss until the animal is ambulatory. 2. With isoflurane anesthesia, the animal is ambulatory within few minutes after the end of surgery. With xylazine/ketamine anesthesia, the following clinical parameters must be monitored at a minimum of 15-minute intervals until the animal is ambulatory: — Respiratory rate — Movement — Ability to maintain sternal recumbency. It is estimated that animals will recover within 1–3 hours post-surgery (see Note 9). 3. During the first week after surgery, infection signs and clip elimination should be monitored by checking on each animal. Clips are eliminated by animals 2 or 3 weeks after surgery.

4

Notes

1. Isoflurane and xylazine/ketamine are the most commonly used anesthetic compounds. However, pentobarbital or Avertin can also be used according to each laboratory’s practice.

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2. Hypothermia is the most commonly overlooked complication in rodent surgery and can result in a prolonged recovery period or death. To protect the animal from hypothermia, avoid metal surfaces during surgery. Animal could be wrapped in a towel to preserve body temperature. A supplemental heat source should be provided in the pre-, intra-, and post-operative periods (water re-circulating heating blanket or thermal packs can be used). 3. Clips are more suitable than suture material because animals can more easily damage the suture. 4. Animals waiting for surgery should be kept at a visual and olfactory distance from other animals undergoing surgery to minimize pre-operative stress. 5. Animals injected with a combination of xylazine and ketamine should be kept in a quiet environment to avoid anesthesia failure. 6. Sometimes testes are not localized within scrotal sacs and a massage of the peritoneal cavity should be performed from the top to the bottom of the abdomen until testes reach their normal position in the scrotal sacs. 7. For an experienced operator, the expected time for ORX is about 15 to 20 minutes. 8. Each animal under recovery should be placed in a separate cage to avoid injury by cage mates. 9. Animals may crawl into the corners of the recovery cage and bedding can sometimes block the airway. Acknowledgments The authors are greatly indebted to P. Legras and J. Roux (SCAHU) for their help with animal care.

References 1. Boonen, S., Vanderschueren, D., Geusens, P., et al. (1997) Age associated endocrine deficiencies as potential determinants of femoral neck (type II) osteoporotic fracture occurrence in elderly men. Int J Androl 20, 134–143. 2. Thompson, D. D., Simmons, H. A., Pirie, C. M., et al. (1995) FDA guidelines and animal models for osteoporosis. Bone 17, 125S–133S. 3. Audran, M., Chappard, D., Legrand, E., et al. (2001) Bone microarchitecture and bone fragility in men: DXA and histomorphometry in humans and in the orchidectomized rat model. Calcif Tissue Int 69, 214–217. 4. Fournier, P., Boissier, S., Filleur, S., et al. (2002) Bisphosphonates inhibit angiogenesis in vitro and testosterone-stimulated vascular regrowth in the ventral prostate in castrated rats. Cancer Res 62, 6538–6544. 5. Gunness, M., Orwoll, E. (1995) Early induction of alterations in cancellous and cortical bone histology after orchiectomy in mature rats. J Bone Miner Res 10, 1735–1744. 6. Iwamoto, J., Takeda, T., Katsumata, T., et al. (2002) Effect of etidronate on bone in orchidectomized and sciatic neurectomized adult rats. Bone 30, 360–367. 7. Iwamoto, J., Yeh, J. K., Takeda, T. (2003) Effect of vitamin K2 on cortical and cancellous bones in orchidectomized and/or sciatic neurectomized rats. J Bone Miner Res 18, 776–783. 8. Lerouxel, E., Libouban, H., Moreau, M. F., et al. (2004) Mandibular bone loss in an animal model of male osteoporosis (orchidectomized rat): a radiographic and densitometric study. Osteoporos Int 15, 814–819.


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9. Li, M., Jee, W. S., Ke, H. Z., et al. (1995) Prostaglandin E2 administration prevents bone loss induced by orchidectomy in rats. J Bone Miner Res 10, 66–73. 10. Libouban, H., Moreau, M. F., Legrand, E., et al. (2002) Comparison of histomorphometric descriptors of bone architecture with dual-energy X-ray absorptiometry for assessing bone loss in the orchidectomized rat. Osteoporos Int 13, 422–428. 11. Moreau, M. F., Libouban, H., Legrand, E., et al. (2001) Lean, fat and bone masses are influenced by orchidectomy in the rat. A densitometric X-ray absorptiometric study. J Musculoskel Neuron Interact 1, 209–213. 12. Peng, Z., Tuukkanen, J., Zhang, H., et al. (1994) The mechanical strength of bone in different rat models of experimental osteoporosis. Bone 15, 523–532. 13. Vanderschueren, D., Vandenput, L., Boonen, S., et al. (2000) An aged rat model of partial androgen deficiency: prevention of both loss of bone and lean body mass by low-dose androgen replacement. Endocrinology 141, 1642–1647. 14. Verhas, M., Schoutens, A., L’hermite-Baleriaux, M., et al. (1986) The effect of orchidectomy on bone metabolism in aging rats. Calcif Tissue Int 39, 74–77. 15. Wink, C. S., Felts, W. J. (1980) Effects of castration on the bone structure of male rats: a model of osteoporosis. Calcif Tissue Int 32, 77–82. 16. Blouin, S., Moreau, M. F., Weiss, P., et al. (2006) Evaluation of an injectable bone substitute (betaTCP/hydroxyapatite/hydroxy-propyl-methyl-cellulose) in severely osteopenic and aged rats. J Biomed Mater Res A 78, 570–580. 17. Erben, R. G. (2001) Skeletal effects of androgen withdrawal. J Musculoskelet Neuronal Interact 1, 225–233. 18. Most, W., Van Der Wee-Pals, L., Ederveen, A., et al. (1997) Ovariectomy and orchidectomy induce a transient increase in the osteoclastogenic potential of bone marrow cells in the mouse. Bone 20, 27–30. 19. Weinstein, R. S., Jia, D., Powers, C. C., et al. (2004) The skeletal effects of glucocorticoid excess override those of orchidectomy in mice. Endocrinology 145, 1980–1987. 20. Bellido, T., Jilka, R. L., Boyce, B. F., et al. (1995) Regulation of interleukin-6, osteoclastogenesis, and bone mass by androgens. The role of the androgen receptor. J Clin Invest 95, 2886–2895. 21. Vandenput, L., Ederveen, A. G., Erben, R. G., et al. (2001) Testosterone prevents orchidectomy-induced bone loss in estrogen receptor-alpha knockout mice. Biochem Biophys Res Commun 285, 70–76. 22. Danielsen, C. C., Mosekilde, L., Andreasen, T. T. (1992) Long-term effect of orchidectomy on cortical bone from rat femur: bone mass and mechanical properties. Calcif Tissue Int 50, 169–174. 23. Erben, R. G., Eberle, J., Stahr, K., et al. (2000) Androgen deficiency induces high turnover osteopenia in aged male rats: a sequential histomorphometric study. J Bone Miner Res 15, 1085–1098. 24. Libouban, H., Moreau, M. F., Baslé, M. F., et al. (2001) Comparison insight dual X-ray absorptiometry (DXA), histomorphometry, ash weight and morphometric indices for bone evaluation in an animal model of male osteoporosis (the orchidectomized rat). Calcif Tissue Int 68, 31–37. 25. Libouban, H., Moreau, M. F., Legrand, E., et al. (2002) Bone Architecture Measured by Fractal Dimension and Connectivity Indices Is More Precociously Altered Than Mineral Content in the Orchidectomized Rat. Birkhauser Press, Cambridge, MA. 26. Rosen, H. N., Tollin, S., Balena, R., et al. (1995) Differentiating between orchidectomized rats and controls using measurements of trabecular bone density: a comparison among DXA, histomorphometry, and peripheral quantitative computerized tomography. Calcif Tissue Int 57, 35–39. 27. Wakley, G. K., Schutte, H. D., Hannon, K. S., et al. (1991) Androgen treatment prevents loss of cancellous bone in the orchidectomized rat. J Bone Miner Res 6, 325–330. 28. Vanderschueren, D., Van Herck, E., Suiker, A. M. H., et al. (1992) Bone and mineral metabolism in the adult guinea pig: long-term effects of estrogen and androgen deficiency. J Bone Miner Res 7, 1407–1415.

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29. Rubinacci, A., Villa, I., Sibilia, V., et al. (1998) Responsiveness of urinary markers of bone resorption to orchiectomy and clodronate treatment in mature rats: a comparative study. Eur J Endocrinol 138, 120–127. 30. Vanderschueren, D., Jans, I., Van-Herck, E., et al. (1994) Time-related increase of biochemical markers of bone turnover in androgen-deficient male rats. Bone Miner 26, 123–131. 31. Blouin, S., Gallois, Y., Moreau, M. F., et al. (2006) Disuse and orchidectomy have additional effects on bone loss in the aged male rat. Osteoporos Int 18, 85–92.

Chapter 10

Gene Delivery by Adenoviruses Renny T. Franceschi and Chunxi Ge

Abstract Adenoviruses have a number advantages as gene delivery vectors, including ability to transduce a wide variety of non-dividing and dividing cells with high efficiency, relative ease of construction, and ability to be purified as high-titer viral stocks. These characteristics make adenoviruses particularly attractive for overexpressing specific genes in vitro and for evaluating in vivo biological activity in animal models. In this chapter, procedures will be described for adenovirus construction and virus delivery to in vivo sites for measurement of osteogenic activity. Keywords Adenovirus, gene therapy, bone, regeneration, bone morphogenetic protein, osteogenesis.

1

Introduction

Adenoviruses are among the most widely used gene therapy vectors. They have highly evolved mechanisms for efficient delivery of DNA to cells, are relatively easy to construct and amplify and, unlike retroviruses, are not dependent on cell replication for infection. Adenoviruses infect cells by binding of the viral fiber capsid protein by coxsackievirus and adenovirus receptor (CAR) and binding of the viral penton base by av integrins on the cell surface. The broad distribution of these receptors explains why adenoviruses can be used to infect such a wide range of cell types (1, 2). After infection, adenoviruses do not normally integrate into the host genome and, instead, remain in the nucleus as an episome that is gradually diluted and degraded as cells divide (3). Most gene therapy studies conducted to date used first-generation adenoviruses. These vectors have been genetically modified via deletion of the E1 gene to be replication incompetent, but still contain most of the viral genome, including genes encoding the major coat proteins. Because of this, cells infected with first-generation adenovirus vectors secrete viral proteins and elicit an immune response in the host that eventually result in clearance of transduced cells from the body (4). The in vivo From: Methods in Molecular Biology, Vol. 455: Osteoporosis: Methods and Protocols Edited by Jennifer J. Westendorf © Humana Press, Totowa, NJ

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half-life of transduced cells can vary from a few days up to several weeks, depending on the degree to which the site of implantation is accessible by the immune system. For example, we measured in vivo luminescence after subcutaneous implantation of fibroblasts previously transduced with an adenovirus expressing firefly luciferase and observed decay of luciferase to baseline activity within 1 week (5). Safety issues related to the immunogenicity of virally transduced cells have severely restricted the use of first-generation adenoviruses for human applications (6). Because of these problems, second-generation adenovirus-based vectors have been developed that lack genes for most or all viral proteins (7, 8). These vectors have the advantage of being able to package up to 30 kb of foreign DNA. However, they can only be propagated in the presence of helper viruses that contain the missing viral genes necessary to form a viable capsid. In spite of their limitations, first-generation adenovirus vectors have been extremely useful for expressing specific genes in cell culture with greater efficiency and duration than traditional plasmid-based systems as well as for evaluating the effectiveness of regenerative factors in animal models. In the mineralized tissues field, adenoviruses have been extensively used to evaluate the in vivo biological activity of a wide range of molecules, including members of the BMP/TGF-b family; growth factors including FGF2, IGF-1, and PDGF; and osteogenic transcription factors such as RUNX2 (9). In addition, adenovirus expression was used to explore cooperative interactions between groups of regenerative factors such as combinations of BMPs (5, 10) or combinations of angiogenic and differentiation-promoting signals (11). Lastly, adenovirus expression was used to suppress the activity of inflammatory cytokines in rheumatoid arthritis by overexpressing decoy receptors to the interleukin 1 receptor (12). There is no doubt that adenovirus expression will continue to be the method of choice for the initial in vivo evaluation of new regenerative molecules. For this reason, this chapter focuses on the construction and use of first-generation adenovirus vectors. The approaches described in this chapter were specifically developed for bone regeneration studies in the author’s laboratory using virally expressed BMPs and osteogenic transcription factors. However, these approaches can be used to express a variety of molecules in vitro or in vivo.

2 2.1

Materials Adenovirus Production

1. HEK 293T Cells. This cell line is used for packaging and amplification of adenovirus stocks and is available from several commercial sources, such as Stratagene(La Jolla, CA) (AD-293 cells). 2. HEK 293T Cell Growth Medium: Dulbecco’s modified Eagle’s medium, 4.5 g/L glucose, 110 mg/L Na pyruvate, 4.0 mM L-glutamine, 10% heat-inactivated fetal bovine serum.

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3. Phosphate Buffered Saline (PBS): 137 mM NaCl, 2.6 mM KCl, 10 mM Na2HPO4, 1.8 mM KH2PO4, adjust pH to 7.4 with HCl. 4. Virus Storage Buffer: 2.5% glycerol, 25 mM NaCl, 20 mM Tris-HCl, pH 8.0. 5. 5% Agarose in PBS: Dissolve 5 g SeaPlaque GTG agarose (BioWhittaker Molecular Applications, Cambrex Corp., East Rutherford, NJ) in 100 mL of sterile PBS and autoclave. Store in 10-mL aliquots at 4 °C in sterile 50-mL conical tubes. 6. Adeno-X Virus Purification Kits from BD Biosciences (San Jose, CA).

2.2

Ex Vivo Transduction and In Vivo Transplantation

1. Implant Lysis Buffer: 10 mM Tris-HCl, pH 7.4, 0.2 % NP40, 2.0 mM PMSF (add from 100× ethanol stock immediately before use). 2. Gelatin sponges (Gelfoam, Upohn, Kalamazoo, MI) or type I collagen hydrogels (BD Biosciences, Bedford, MA). 3. Sterile filter paper. 4. Immunodeficient mice (N: NIH-bg-nu-xid mice, Charles River Labs, Boston, MA). 5. Ketamine. 6. Xylazine. 7. 7-mm diameter trephine drill (Fine Science Tools, Foster City, CA). 8. 4-0 absorbable chromic gut (Ethicon, Inc., Somerville, NJ). 9. BLK cells. 10. Polytron Tissue Disruptor. 11. TCA: 15% and 30% in water. 12. Sigma Diagnostics kit for total calcium and phosphate.

3 3.1

Methods Vector Construction (see Note 1)

A number of kits and custom services are available for adenovirus construction (e.g., Stratagene; MP Biomedicals, Solon, OH). For this reason, specific protocols and reagents are not included here. The general strategy for making adenovirus vectors involves initial subcloning of the cDNA of interest into a shuttle vector that can either be recombined with the virus genome in E. coli as is used in the AdEasy method (13) or directly in a HEK 293T helper cell line containing CRE recombinase (14). In both methods, the use of homologous recombination greatly improves the efficiency of vector production. Commercially available shuttle vectors contain the strong cytomegalovirus promoter to drive expression of the desired gene and are also available with cDNA cloning sites 5' to an internal ribosome entry site followed by a green fluorescent protein marker gene (Stratagene). This is very useful for

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marking virally transduced cells. The Ad5 serotype vectors most commonly used have deletions in the E1 gene necessary for DNA replication. For this reason, these viruses cannot replicate except in helper cells such as HEK 293T cells that have been stably transfected with E1 (commercially available from Stratagene as Ad293T cells).

3.2

Plaque Purification

The presence of the E1 gene in HEK293T cells introduces the possibility that replication-competent viruses can be generated via recombination during virus amplification. Although the frequency of recombination is very low, the percentage of replication-competent particles increases with each amplification. For this reason, the primary virus stock for each new adenovirus made should be used for all subsequent amplifications. To assure the homogeneity of this stock, it is recommended that it be plaque purified as follows: 1. Plate HEK293T cells at a density of 5 × 105/well of a 6-well tissue culture plate. 2. After an overnight incubation, prepare a series of 1:10 dilutions of the primary viral lysate to give final dilutions from 10−4 to 10−7. 3. Add 1 mL of each dilution to a separate well of the 6-well plate of HEK293T cells. 4. Incubate for 2 hours with gentle rocking to uniformly distribute virus. 5. Melt a 10-mL aliquot of agarose by placing in boiling water and cool to 45 °C. 6. Add 30 mL growth medium, previously equilibrated at 37 °C, and mix (final agarose concentration is 1.25%). 7. Remove medium from virally transduced cells and gently pipette 3 mL agarose solution into each well. Incubate for 1 to 2 weeks. Plaques appear as white spots. 8. Use a sterile Pasteur pipet to core out well-isolated plaques and transfer each to 250 µL growth medium in a sterile centrifuge tube. After a 24-hour incubation, virus can be used to reinfect HEK293T cells for amplification and purification.

3.3

Adenovirus Amplification and Large-Scale Purification

For large-scale adenovirus purification, we routinely use Adeno-X Virus Purification Kits from BD Biosciences that use proprietary filter technology to obtain pure virus. Alternatively, virus can be purified by CsCl ultracentrifugation. 1. Infect ten 150-cm dishes of HEK293T cells (50–90% confluent) with adenovirus stock at 5–10 pfu/cell (approx. 2 × 107 pfu virus/dish). After a 2- to 5-day incubation period, cytopathic effects of viral infection are complete (2–7 days). Cells generally are not lysed, but detach from plates and appear as floating clusters in the medium. 2. Transfer cells and medium to 50-mL conical tubes. Cells that remain attached to the culture dishes can be dislodged by gentle pipetting.


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3. Centrifuge at 1,500 g for 5 minutes to obtain a cell pellet and save the supernatant. 4. Resuspend cells in 25 mL sterile of 100 mM Tris-HCl and disrupt cells with three consecutive freeze-thaw cycles. 5. Centrifuge at 1,500 g for 5 minutes to remove cell debris. Mix the cell extract with supernatant from step 3 and filter using a sterile 0.45 µ bottle top filter. 6. Virus is then purified from the filtered solution using reagents and protocols included in the Adeno-X kit. Purified virus is transferred to virus storage buffer and kept at −70 °C until use (see Note 2).

3.4

Determining Virus Titer

This method can be used to determine actual plaque forming units of a virus stock (15). It is critical that virus titer be very carefully controlled in experiments. Use of approximate methods for determining titer such as light scattering should be avoided. 1. Prepare a 96-well plate of HEK293T cells (105 cells/well) for each adenovirus preparation to be titered and let cells incubate overnight. 2. Prepare eight serial 1:10 dilutions of virus stock in growth medium containing 2% FBS. 3. For each dilution, add 100 µL to each of 10 wells. 4. After culturing for 10 days, count the number of wells showing a cytopathic effect of the virus (CPE). Make sure the lowest dilution shows 100% infection and the highest dilution has no CPE. 5. For each dilution, calculate the ratio of CPE-positive wells to total wells. Titer(T) is calculated using the following equation: T = 101+d(S – 0.5) where d = Log10 of dilution (i.e., 1 for a 1:10 dilution), S = sum of the ratios (CPE wells/total wells) for all eight dilutions. Sample calculation:

Dilution

Ratio (CPE wells/total wells)

10−1 10−2 10−3 10−4 10−5 10−6 10−7 10−8

10/10 = 1 10/10 = 1 10/10 = 1 10/10 = 1 6/10 = 0.6 2/10 = 0.2 0/10 = 0 0/10 = 0

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In this example, 100% of wells at the 10−4 dilution have a positive CPE and 0% of wells at the 10−8 dilution have a positive CPE. Therefore, d = 1, S = 1 + 1+ 1+ 1 + 0.6 + 0.2 + 0 + 0 = 4.8 Titer = 101 + 1 (4.8 – 0.5) = 105.3 pfu/100 µl or 106.3/mL = 2 × 106 pfu/mL.

3.5

Ex Vivo and In Vivo Transduction Methods

Adenovirus vectors can either be used to transduce cells in tissue culture for subsequent in vitro analysis or in vivo implantation (ex vivo transduction) or can be directly applied to in vivo sites (in vivo transduction). The same basic procedure can be used to transduce a variety of cell types. We used this procedure with primary human and mouse dermal and gingival fibroblasts, human and mouse marrow stromal cells, and mouse and rat primary calvarial osteoblast cultures. The following mouse cell lines have also been successfully used: ST2 marrow stromal cells, MC3T3-E1 preosteoblasts, BLK fibroblasts, C3H10T1/2 mesenchymal cells, and C2C12 myoblasts.

3.5.1

Ex Vivo Transduction

1. Plate cells at the desired density in their preferred growth medium. 2. After a 24-hour attachment period, add adenovirus at the desired multiplicity of infection (MOI, which reflects the number of pfu virus added per cell) in serumfree medium (see Note 1). 3. After 4 hours, add FBS to a final concentration of 2% and grow cells for an additional 24 hours. 4. Transfer cells to the desired growth or differentiation medium and feed every 2 days for the duration of the experiment or trypsinize and implant into animals on a suitable carrier (see Note 3).

3.5.2

In Vivo Cell Implantation

We routinely use either gelatin sponges or type I collagen hydrogels as substrates for in vivo cell implantation (16) (see Note 4). 1. Twenty-four hours after virus transduction, cells are trypsinized and suspended in complete medium at a concentration of 1 × 107 cells/mL. 2. For loading cells onto gelatin sponges, the sponge (precut into 0.4 cm2 pieces) is pre-wetted with compete medium and air bubbles are removed by placing the sponge between two pieces of sterile filter paper and applying gentle pressure. Sponges are then seeded with 0.5 to 1 × 106 cells by capillary action. 3. For type I collagen hydrogels, the desired number of cells is mixed with type I collagen solution and medium to give a final collagen concentration of 2 mg/mL.


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A 200-µL aliquot of the cell/collagen suspension is pipetted into a microtiter well to form a gel cylinder (see Notes 5 and 6).

3.5.2.1

In Vivo Implantation of Virally Transduced Cells

Two in vivo sites are routinely used to measure the osteogenic activity of virally transduced cells. Subcutaneous implants allow detection of osteogenic activity in the absence of endogenous bone cells and their precursors, whereas cranial implants allow detection of activity in an intramembranous bone microenvironment.

3.5.2.2

Subcutaneous Implants

Cells seeded on either Gelfoam sponges or collagen hydrogels are implanted on the backs of immunodeficient mice as follows. 1. Mice are anesthetized with ketamine (75 mg/kg) and xylazine (10 mg/kg) ip. 2. Two midline longitudinal skin incisions of approximately 1 cm in length are made on the dorsal surface of each mouse and a subcutaneous pocket is formed on each flank by blunt dissection. 3. A single implant is placed in each pocket with up to four implants per animal. 4. Incisions are closed with surgical staples.

3.5.2.3

Implanting Cells into Critical-Sized Calvarial Defects

1. Mice are anesthetized and an 0.8- to 1.0-cm incision is made in the scalp. 2. The cranial vault is exposed and a defect is created using a 7-mm diameter trephine drill, taking care not to injure the underlying dura. 3. Cells are seeded onto a precut Gelfoam sponge that is placed over the defect and secured in place with 4-0 absorbable chromic gut. In both in vivo models, mice are sacrificed after 3 to 6 weeks and bone formation is assessed using both histological and biochemical analysis as described (see Section. 3.4.3.).

3.5.3

In Vivo Transduction

Adenovirus vectors can also be directly administered to in vivo sites. In our experience, this does not generally give as reproducible results largely because of variations in efficiency of viral transduction of host cells in vivo. Both the subcutaneous and cranial models described in the preceding have been examined. Although virus can be directly injected into either site, we obtain better results when the virus is

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immobilized on a suitable carrier such as a collagen hydrogel as described in Section 3.4.1. In this case, 1 to 2 × 109 pfu adenovirus is administered in a 10- to 20-µL volume of collagen gel.

3.5.4

Standardized System for Quantifying Osteogenic Activity of Virally Expressed Osteogenic Factors

We developed a standardized system to objectively compare the in vivo osteogenic activity of different regenerative factors using adenovirus-based expression in subcutaneous implants (17). This system uses a clonal mouse fibroblast cell line (BLK cells) derived from C57BL6 mice that is transduced ex vivo with adenoviruses expressing the desired regenerative factors. Because cells are syngeneic with this mouse line, they can be implanted into mice with a minimal immune response. To further avoid immune responses elicited by virus coat proteins, this method can also be used with immunodeficient lines as in Section 3.5.1. However, in our experience the use of C57BL6 hosts gives highly reproducible data on the in vivo osteogenic activity of regenerative factors while sparing the expense of conducting experiments in immunodeficient mice. Adenovirus expression of osteogenic factors such as BMPs in this system will produce a complete ectopic bone organ including cortical and trabecular bone and marrow. Figure 10.1 shows an example of how this assay is used to compare the osteogenic activity of adenovirus expressing BMP2, BMP7, or a BMP2/7 combination. Of interest, our studies showed that combined adenoviral expression of BMP2 and 7 had approximately four times the in vivo osteogenic activity of BMP2 or BMP7 alone (5). These results were explained by the formation of BMP2/7 heterodimers having enhanced biological activity. 1. Titered adenovirus stocks encoding each regenerative molecule to be compared are used to transduce BLK cells. Normally, BLK cells are transduced with increasing titers of virus over a range of MOI from 50 to 300. 2. To ensure that each factor is expressed at similar levels, cell layers and media (for secreted molecules) are harvested and regenerative factor levels are measured by western blotting, ELISA, or another quantitative method. 3. BLK cells expressing comparable levels of each factor are implanted on the backs of mice as described in Section 3.5.1. 4. After 4 weeks, mice are sacrificed by CO2 inhalation and implants are removed for biochemical analysis and/or histology.

3.5.5

Biochemical Analysis

When comparing a large number of samples, it is easiest to use the simplest index of new bone formation, accumulation of ectopic mineral, as a means of assessing osteogenic activity.


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Fig. 10.1 In vivo osteogenic activity of AdBMP combinations. BLK fibroblasts were transduced with the indicated adenoviruses at an MOI of 100 each and implanted into C57BL/6 mice, as described in Methods. After 4 weeks, implants were harvested for determination of (a) ALP activity, (b) calcium, or (c) phosphate. Significant difference: *p < 0.05, **p < 0.01, ***p < 0.001. Histology of the implants is shown in panels D-G; (d) AdLacZ, (e) AdBMP-2, (f) AdBMP-7, or (g) AdBMP-2/7 (original magnification 4× for all sections). (Reprinted from Zhao, M., Zhao, Z., Koh, J.-T., et al. (2005). Combinatorial gene therapy for bone regeneration: cooperative interactions between adenovirus vectors expressing bone morphogenetic proteins 2, 4 and 7. J Cell Biochem 95, 1–16, with permission.)

1. Implants are homogenized in 1-mL lysis buffer with a Polytron Tissue Disruptor. Insoluble material is removed by centrifugation for 5 minutes in a microfuge. A 100-µL aliquot is removed for measurement of alkaline phosphatase (18) and the remainder of the sample is mixed with an equal volume of 30% TCA. 2. After overnight incubation at 4 °C, samples are centrifuged in a microfuge to remove precipitated protein and nucleic acids. Calcium and phosphate are solubilized by the 15% TCA. Supernatants are used to measure total calcium (Sigma Diagnostics, La Jolla, CA) and phosphate (19). Pellets can be used to measure total DNA or protein for normalization as necessary.

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3.5.6

Histological Analysis

Immediately after removal, implants are fixed at 4 °C in 10% neutral-buffered formalin for 24 hours and then demineralized in 10% formic acid for 2 days. Samples are then rinsed and stored in 70% ethanol until paraffin embedding. After sectioning, samples are de-paraffinized, hydrated, and stained with hematoxylin and eosin.

4

Notes

1. Safety Issues. Adenovirus vectors are derived from infectious human respiratory viruses and are theoretically capable of recombining with these viruses to form infectious particles. All procedures associated with adenovirus production and use must be conducted under NIH Biosafety Level 2 conditions (see http://www. cdc.gov/od/ohs/biosfty/bmbl4/bmbl4s3.htm). Furthermore, construction of vectors expressing potentially harmful genes such as oncogenes should be done with extreme caution. Researchers are advised to consult with the Biosafety Office of their institution before commencing adenovirus work. 2. Adenoviruses are denatured by freeze-thawing. It is recommended that stocks not be thawed more than two times. 3. Large variations are observed between cell types in terms of the efficiency of adenovirus transduction and expression of recombinant proteins. When beginning studies with a new virus preparation, it is important to determine the optimal titer of virus stock for expression of the protein of interest in the specific cell line being transduced. This can be accomplished using western blotting or ELISAs. 4. In our experience with a variety of cell lines and primary cultures, adenovirus expression normally persists for 1 to 2 weeks in cell culture and for approximately 1 week after subcutaneous in vivo implantation of cells (5). However, these values vary between different cell types and are dependent on factors such as proliferation rate and stability of the expressed protein and must be determined on a case-by-case basis. 5. Virally transduced cells can also be implanted in a wide variety of synthetic tissue engineering scaffolds based on a variety of chemistries, including poly-lactate/ polyglycolate polymers, alginates and ceramics (20). 6. We routinely cast gels in microtiter wells mounted on glass slides (Lab-Tek Chamber Slides, Nalge Nunc International, Rochester, NY). After the hydrogel is formed, the gel cylinder can be easily removed from the well by removing the well from the slide and gently extruding the gel with a small spatula. Acknowledgments Work cited from the authors’ laboratory was supported by NIH grants DE13386 and DE11723.


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References 1. Bergelson, J. M., Cunningham, J. A., Droguett, G., et al. (1997). Isolation of a common receptor for Coxsackie B viruses and adenoviruses 2 and 5. Science 275, 1320–1323. 2. Neumann, R., Chroboczek, J., Jacrot, B. (1988). Determination of the nucleotide sequence for the penton-base gene of human adenovirus type 5. Gene 69, 153–157. 3. Oligino, T. J., Yao, Q., Ghivizzani, S. C., et al. (2000). Vector systems for gene transfer to joints. Clin Orthop 379, S17–30. 4. Mahr, J. A., Gooding, L. R. (1999). Immune evasion by adenoviruses. Immunol Rev 168, 121–130. 5. Zhao, M., Zhao, Z., Koh, J.-T., et al. (2005). Combinatorial gene therapy for bone regeneration: cooperative interactions between adenovirus vectors expressing bone morphogenetic proteins 2, 4 and 7. J Cell Biochem 95, 1–16. 6. Yang, Y., Li, Q., Ertl, H. C., et al. (1995). Cellular and humoral immune responses to viral antigens create barriers to lung-directed gene therapy with recombinant adenoviruses. J Virol 69, 2004–2015. 7. Armentano, D., Zabner, J., Sacks, C., et al. (1997). Effect of the E4 region on the persistence of transgene expression from adenovirus vectors. J Virol 71, 2408–2416. 8. Hartigan-O’Connor, D., Amalfitano, A., Chamberlain, J. S. (1999). Improved production of gutted adenovirus in cells expressing adenovirus preterminal protein and DNA polymerase. J Virol 73, 7835–7841. 9. Franceschi, R. T. (2005). Biological approaches to bone regeneration by gene therapy. J Dent Res 84, 1093–1103. 10. Zhu, W., Rawlins, B. A., Boachie-Adjei, O., et al. (2004). Combined bone morphogenetic protein-2 and -7 gene transfer enhances osteoblastic differentiation and spine fusion in a rodent model. J Bone Miner Res 19, 2021–2032. 11. Peng, H., Wright, V., Usas, A., et al. (2002). Synergistic enhancement of bone formation and healing by stem cell-expressed VEGF and bone morphogenetic protein-4. J Clin Invest 110, 751–759. 12. Roessler, B. J., Allen, E. D., Wilson, J. M., et al. (1993). Adenoviral-mediated gene transfer to rabbit synovium in vivo. J Clin Invest 92, 1085–1092. 13. He, T. C., Zhou, S., da Costa, L. T., et al. (1998). A simplified system for generating recombinant adenoviruses. Proc Natl Acad Sci USA 95, 2509–2514. 14. Hardy, S., Kitamura, M., Harris-Stansil, T., et al. (1997). Construction of adenovirus vectors through Cre-lox recombination. J Virol 71, 1842–1849. 15. Doherty, R. (1964) Animal virus titration techniques, in (Harris, R., ed.), Techniques in Experimental Virology. Academic Press, New York. 16. Krebsbach, P. H., Gu, K., Franceschi, R. T., et al. (2000). Gene therapy-directed osteogenesis: BMP-7-transduced human fibroblasts form bone in vivo [in process citation]. Hum Gene Ther 11, 1201 –1210. 17. Franceschi, R. T., Yang, S., Rutherford, R. B., et al. (2004). Gene therapy approaches for bone regeneration. Cells Tissues Organs 176, 95–108. 18. Manolagas, S. C., Burton, D. W., Deftos, L. J. (1981). 1,25-Dihydroxyvitamin D stimulates the alkaline phosphatase activity of osteoblast-like cells. J Biol Chem 256, 7115–7117. 19. Heinonen, J. K., Lahti, R. J. (1981). A new and convenient colorimetric determination of inorganic orthophosphate and its application to the assay of inorganic pyrophosphatase. Anal Biochem 113, 313–317. 20. Alsberg, E., Hill, E. E., Mooney, D. J. (2001). Craniofacial tissue engineering. Crit Rev Oral Biol Med 12, 64–75.

Chapter 11

Lentivirus Delivery of shRNA Constructs into Osteoblasts Marc N. Wein, Dallas C. Jones, and Laurie H. Glimcher

Abstract Osteoblasts are the sole cell responsible for bone formation in vivo (1). Although genetic techniques have been extremely valuable to study the functions of certain genes in these cells in vivo, this approach is time consuming and expensive. An alternative loss-of-function approach that has been validated in many mammalian systems is shRNA-mediated gene silencing. This chapter describes methodology designed to introduce shRNA constructs into primary murine osteoblasts ex vivo in order to quickly assess the function of genes in osteoblast differentiation and extracellular matrix mineralization. Both the production of shRNA-expressing lentiviruses and the infection of calvarial osteoblasts with these lentiviruses are detailed. Keywords Osteoblasts, shRNA, lentivirus, extracellular matrix mineralization, 293T cells, NIH 3T3 cells.

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Introduction

RNA interference (RNAi) is a rapid and powerful tool to reduce levels of target gene expression in a sequence-specific fashion. RNAi was first demonstrated to function in model systems such as A. thaliana, D. melanogaster, and C. elegans (2). RNAi also suppresses gene expression in mammalian cells, although it is technically difficult to introduce synthetic small interfering (si) RNA oligonucleotides into many cell types grown in vitro. An alternative approach is to transduce mammalian cells with viruses expressing short hairpin (sh) RNAs that will subsequently generate siRNAs within cells (3, 4). A variety of viral vectors expressing shRNA sequences have been developed. Lentiviruses are particularly powerful as they can be used to infect dividing and non-dividing cells. Additionally, lentiviruses are able to infect the vast majority of cell lines and primary cells, making lentiviral-mediated shRNA expression a powerful resource for loss-of-function studies in mammalian systems. As lentiviruses can be further engineered to express drug-resistance genes (e.g., a puromycin resistance marker, as used here), it is straightforward and convenient to infect cells From: Methods in Molecular Biology, Vol. 455: Osteoporosis: Methods and Protocols Edited by Jennifer J. Westendorf © Humana Press, Totowa, NJ

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with lentiviruses, add puromycin to kill all non-infected cells, and then analyze the shRNA-expressing infected cells. Previously we used lentivirus (LV)-mediated shRNA delivery to study the function of a family of E3 ubiquitin ligases in the regulation of extracellular matrix mineralization by murine calvarial-derived osteoblasts in vitro (5). This chapter provides detailed methods for the generation of lentiviruses, the propagation of murine calvarial osteoblasts in vitro, and the infection of calvarial osteoblasts with lentiviruses.

2 2.1

Materials Production of Lentiviruses in 293T Cells

1. HEK 293T cell line can be acquired from the American Type Culture Collection (ATCC, Manassas, VA). 2. DMEM-10%: 900 mL Dulbecco’s Modified Essential Medium (DMEM), 10 mL penicillin/streptomycin, 10 mL L-glutamine, 10 mL 1 M Hepes pH 8.0, 100 mL fetal bovine serum (FBS, Hyclone, Ogden, UT). 3. Effectene Transfection Reagent (Qiagen, 301427, Valencia, CA). 4. 0.45 µM filters (Costar, 8112, Corning, NY).

2.2

Determination of Viral Titer in NIH3T3 Cells

1. The murine fibroblastic NIH3T3 cell line can be acquired from the ATCC. 2. Polybrene (hexadimethrine bromide) dissolved in sterile water at a stock concentration of 8 mg/mL and kept at 4 °C. 3. Puromycin dihydrochloride dissolved in sterile water at 2 mg/mL and kept at −20 °C.

2.3

Isolation of Calvarial Osteoblasts

1. At least six neonatal mice 2–5 days old (see Note 1 for more details). 2. 70% Ethanol to sterilize mice and surgical instruments. 3. Ice cold Dulbecco phosphate-buffered saline (DPBS) dispensed into sterile sixwell tissue culture dish (Falcon, 35-3046, Franklin Lakes, NJ). 4. Digestion solution: 50 mL Alpha Minimum Essential Medium (AMEM), 50 mg collagenase type II, 100 mg Dispase II (Roche, Germany), 500 µL penicillin/ streptomycin. Filter sterilize prior to use. 5. Osteoblast differentiation medium (OBDM-10%): 500 mL AMEM, 5 mL penicillin/streptomycin, 5 mL non-essential amino acids, 5 mL L-glutamine, 5 mL

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1 M Hepes pH 8.0, 50 mL fetal bovine serum, 25 mg L-ascorbic acid, 540 mg Glycerol 2 phosphate disodium salt hydrate. Filter sterilize prior to use. 6. 70-µm cell strainer (BD Falcon, 352350, Bedford, MA).

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Methods

Production of shRNA-expressing lentiviruses is first accomplished by transfecting viral components into packaging 293T cells. LV-containing cell supernatant is collected and filtered. Rough estimates of viral titer are then determined by infecting murine NIH3T3 cells with viruses. Finally, we describe how to isolate calvarial-derived osteoblasts from neonatal mice, and how to infect these cells with shRNA-expressing LV in vitro.

3.1

Production of Lentiviruses in 293T Cells

1. HEK 293T cells are passaged in DMEM-10% at 37 °C until ready to use. The day before transfection, cells are harvested by trypsinization, counted, and plated at a density of 2 × 106 cells in 10 mL DMEM-10% in a 10-cm dish. Plate out one dish for each virus to be produced. 2. The next day, the cells should appear 25–35% confluent and are ready for transfection with Effectene. At this time, allow DMEM-10% medium to equilibrate to room temperature. In a tissue culture hood, prepare one sterile 1.5 mL Eppendorf tube for each transfection (see Note 1). In each tube, add 300 µL buffer EC, then add 2 µg LV cDNA, 2 µg psPAX2 (gag/rev), and 200 ng pMD2. G (VSV-G) plasmid DNA to buffer EC (see Note 2). Add 34 µL enhancer solution to each tube, vortex gently for 1–3 seconds, and incubate tubes at room temperature for 5 minutes. 3. Add 80 µL Effectene solution to each tube, vortex gently for 1–3 seconds to mix, and incubate tubes at room temperature for 5 minutes. 4. Aspirate medium from adherent HEK 293T cells and gently replace with 10 mL fresh DMEM-10%. Take care to slowly pipet this fresh medium along the edge of the plate so as to not disturb the cells. 5. To each transfection tube, add 500 µL DMEM-10% and mix by gently vortexing as above. Add this mix drop wise to each plate of HEK 293T cells and gently swirl plate to mix. Place cells back in 37 °C incubator overnight. At this point, cells should be treated with standard BSL2 safety precautions. 6. The next day, aspirate off medium into bleach-containing solution. Add 10 mL fresh DMEM-10% (pre-equilibrated to room temperature) to each plate and place plates back at 37 °C. 7. The next day, harvest HEK 293T cell LV-containing supernatant. Pipette off 10 mL medium from each plate and filter through 0.45 µm filter into 10 sterile 1.5 mL Eppendorf tubes. Label LV-containing Eppendorf tubes and freeze at −80 °C until further use.

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Determination of Viral Titer in NIH3T3 Cells

1. NIH3T3 cells are passaged in DMEM-10% at 37 °C until ready for use. The day before titer determination, collect cells by trypsinization, count, and plate at a density of 3 × 104 cells in 1 mL DMEM-10% in each well of a 24-well plate. Plate enough cells for four wells per LV to be tested. 2. The next day, cells should appear 25–35% confluent and are ready for LV infection. Thaw LV stocks on ice. For each LV to be tested, prepare four sterile 1.5-mL Eppendorf tubes with 100 µl DMEM-10% plus 2 µg polybrene. To these tubes, add 0-, 1-, 10-, or 100-µL LV stock. Add this LV/medium/polybrene solution to each well in a drop wise fashion and place the cells back at 37 °C. 3. The next day, aspirate medium and replace with 1 mL DMEM-10% plus 2 µg/ mL puromycin. As described, puromycin at this concentration will effectively kill all uninfected cells. 4. Forty-eight hours later, there should be obvious signs of cell death in uninfected, puromycin-treated wells (see Note 3). At this point, collect cells by trypsinization and determine cell viability by trypan blue staining. The ratio of trypan blue-negative/trypan blue positive cells should indicate the relative cell survival in each well. Alternatively, cell viability can be easily determined with flow cytometry using the forward scatter/side scatter profiles (see Note 4). 5. Wells infected with increasing amounts of LV should show decreased numbers of dead (trypan blue–positive) cells. If, for example, cells infected with 10 µL LV show 50% survival under these conditions, the LV titer of this stock is calculated as follows: 10 µL LV infected approximately 60,000 NIH3T3 cells (assuming doubling time of 24 hours); therefore, 1 µL LV can infect 6 × 103 cells, and 1 mL LV can infect 6 × 106 cells. Determine viral titer for each desired LV, as it is important to subsequently infect osteoblasts with comparable numbers of infectious particles for each gene to be silenced (see Note 5).

3.3

Isolation of Murine Calvarial Osteoblasts

1. At least six P2–P5 neonatal mice are sacrificed by decapitation (see Note 6). Dissection tools are kept sterile by submersion in 70% ethanol, and mice are sprayed with 70% ethanol prior to decapitation. 2. Remove skin over the skull, expose calvarial bones, cut coronally just anterior to suture overlying cerebellum and just posterior to suture at rostral end of cortex. Cut sagittally along the lateral edge of skull (near the ear) to free the overlying calvarial bones from underlying brain tissue. Place calvarial bones in sterile DPBS kept cold on ice in a six-well tissue culture dish. 3. After removing calvarial bones from each mouse, carefully dissect away soft tissue and sutures from bones to avoid contaminating fibroblasts.


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4. Place dissected calvariae in 4-mL digestion solution at the bottom of a 50 mL Falcon tube. Incubate at 37 °C with shaking 275 rpm for 10 minutes. Remove digestion solution by pipetting, wash bones with 5 mL serum-free AMEM, add 4 mL fresh digestion solution and incubate again for 10 minutes at 37 °C on a rotating shaker set at 275 rpm. Wash again with 5 mL serum-free AMEM. Discard solution from these first two digestions and washes (see Note 7). 5. Add fresh 4 mL digestion solution and incubate for 10 minutes at 37 °C with shaking at 275 rpm. Aspirate off digestion solution and place in 14 mL OBDM10% (see Note 8) in a 50-mL Falcon tube. Repeat three times such that four digestion fractions are collected and pooled in 14 mL OBDM for a total of 30 mL solution. 6. Filter 30 mL solution collected in step 5 through a 70-µm cell strainer. Spin down cells at 365 g for 5 minutes at 4 °C (cell pellet will be barely visible). 7. Wash cells with 10 mL OBDM-10% and spin down again. For every two mice initially sacrificed, resuspend pellet in 3 mL OBDM-10% and plate in six-well tissue culture dish. For example, if eight mice were initially sacrificed, resuspend pellet in 12 mL OBDM-10% and plate in four wells of a six-well plate. Place cells at 37 °C. 8. Three days later, cells should appear in adherent colonies. Aspirate medium and replace with 3 mL fresh OBDM-10% pre-equilibrated to room temperature. 9. Two to three days later (5–6 days after dissection) cells should be confluent in dishes. Collect by trypsinization and count. Replate cells at a density of 3 × 104/ mL in OBDM-10% for LV infection. For example, if cells are to be infected in six-well plates (recommended for subsequent analysis of gene expression by RNA isolation), 9 × 104 cells are plated in 3 mL OBDM-10% per well.

3.4

Infection of Calvarial Osteoblasts with shRNA-Expressing Lentiviruses

1. The day after osteoblasts are plated at 3 × 104 cells/mL (step 9), cells should be 50–60% confluent. Aspirate off medium and replace with OBDM-10% containing 2 µg/mL polybrene. 2. Dilute LVs into 100 µl OBDM-10% in sterile 1.5 mL Eppendorf tubes such that the MOI is approximately 1–1.5 (see Note 9). 3. Add LV/medium mix drop wise to cells and mix by gently swirling. Place cells back at 37 °C. It is crucial to leave at least one well uninfected with LV (“mock infected”) to assess the efficiency of subsequent puromycin-induced selection. 4. The next day, add puromycin to cells to reach final concentration of 2 µg/mL. Dilute puromycin into OBDM-10% prior to adding to cells. Mix by gently swirling and place back at 37 °C. 5. Two to four days later, the mock-infected cells should be completely killed by puromycin. At this point, the medium may be changed to OBDM-10% without

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puromycin. The medium should be subsequently changed every 2–3 days to promote robust extracellular matrix production and mineralization. 6. To confirm shRNA-mediated silencing (knock-down), we typically harvest cells 5–7 days after puromycin-induced killing of mock-infected cells. Knock-down is interrogated by protein immunoblotting when possible or real-time quantitative PCR on cDNA prepared from control or gene-specific LV-shRNA cultures using gene-specific primers. 7. To investigate the effects of shRNA-mediated gene silencing on osteoblast function, we typically perform standard alkaline phosphatase stains on day 10–14 cultures to assess osteoblast differentiation and von Kossa stains on day 14–21 cultures to assess matrix mineralization.

4

Notes

1. It is essential to prepare control LV for subsequent experiments. For this purpose, we typically use LV expressing shRNA against either GFP or luciferase, genes normally not expressed in osteoblasts. 2. The empty pLKO.1 lentiviral vector as well as pLKO.1 vectors expressing specific human or mouse shRNA can be purchased from Open Biosystems (Huntsville, AL) or Sigma-Aldrich. The psPAX2 (gag/rev) and pMD2.G (VSV-G) plasmid necessary for generation of viral particles in HEK 293T cells can be acquired from Addgene.org. 3. In the uninfected, puromycin-treated wells, dying cells appear rounded and become non-adherent. In contrast, cells infected with lentivirus expressing a puromycin resistance gene adhere to the tissue culture plate and maintain their fibroblastic morphology 4. For analysis of cell death by flow cytometry, it is necessary to have a control group of cells that have not been treated with puromycin. The amount of viable cells in this population should be >90%, as determined by trypan blue exclusion on the hemocytometer. This group of untreated cells can then be utilized to generate the “live-cell” gate on the forward scatter/side scatterplot. Run each sample on the flow cytometer and collect the same number of total events with each sample. The percent of live cells in each sample can then be determined by using the “live-cell” gate that was generated with the control cells. Utilizing this method should result in no viable cells being present in the uninfected, puromycintreated group. In addition, cells can be stained with propidium-iodide prior to analysis. Viable cells can then be measured utilizing the FL2 channel on the flow cytometer, which detects dying cells that are propidium-iodide positive. 5. Given that the differentiation and function of osteoblasts can be profoundly influenced by cell density, it is important to infect cells with lentiviruses that are of a similar titer. This is crucial to ensure that each experimental condition will have a comparable cell density following puromycin selection.


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6. A relatively inexpensive and dependable source of neonatal mice must be determined. We have successfully ordered late gestational pregnant female mice from Taconic of Balb/c and C57BL/6 genetic backgrounds. Pregnant females are shipped late in gestation, and typically give birth to litters of more than six pups within 2–4 days of arrival. Other investigators (6) have found that calvarial osteoblasts are best cultured from P2-6 mice. It is important to consistently use mice of the same age to reduce inter-experimental variability. 7. The first two fractions contain mainly fibroblasts and should be discarded such that the final pooled digestions contain a relatively homogenous population of osteoblasts. 8. All collagenase/Dispase digestions must be performed in serum-free medium because serum contains anti-proteases that will inactivate these enzymes. Digestions are then stopped by collecting digestion solutions and placing them in serum-containing medium kept on ice. 9. Determine titer of virus as described in Section 3.2. For example, if virus is determined to contain 6 × 103 infectious units per 1 µL and it is desired to infect 9 × 104 cells, use 15 µL virus to achieve an MOI of approximately 1. We have found that infecting at an MOI >10 leads to significant cytotoxicity and an MOI 95%. The dissociation peak should also be checked. There are several microarray platforms available for ChIP-on-chip analysis, including microarrays from NimbleGen Systems, Inc. (Madison, WI), Agilent (Santa Clara, CA), and Affymetrix (Santa Clara, CA). Our protocol focuses on the platform developed by NimbleGen because of its ease of use. NimbleGen provides comprehensive services for DNA labeling, DNA hybridization, and data analysis of ChIP-on-chip experiments. Other platforms require array equipment that is usually available in institutional genomics facilities. The concentration of ChIP-enriched DNA is usually low and needs to be amplified to yield enough DNA for hybridization to a gene microarray. One method for amplification is ligation-mediated PCR (LM-PCR) and another method is whole genome amplification (11). Both methods are based on ligation of linkers and linear amplification. The shearing of DNA should be very well optimized and samples should not contain residual large genomic DNA. A properly sheared DNA sample will permit linear amplification of ChIP-enriched DNA, reduces background and avoids skewing of data due to disproportional amplification of DNA. Ideally, ratios of ChIP-enriched and non-enriched samples (i.e., “input” samples) are the same before and after amplification and samples need to be validated with standard quantitative PCR for positive and negative control genes. Note that DNA concentrations should be measured before amplification. DNA concentrations may need to be diluted for qPCR to obtain concentrations optimal for amplification. The amplified ChIP DNA and amplified total genomic DNA are labeled with the fluorophores Cy-5 and Cy-3, respectively, and are co-hybridized to arrays. Labeling efficiency can be measured using a NanoDrop spectrophotometer. The type of fluorophore may differ between array platforms and manufacturers. Companies provide tiling arrays that are specially designed for ChIP-on-chip. ChIP arrays are easier to design compared to conventional microarrays used for determining mRNA levels. One advantage of tiled arrays over amplicon-based probe arrays is that a positive signal can span over several probes, called the “neighbor effect.” DNA fragments of 15–70 nucleotides are designed to span or “tile” across genomic target regions. The tile density determines the resolution of the array and can be restricted to known promoter regions of genes but also whole genome arrays are available. With respect to DNA shearing, shorter fragments require more closely tiled probes than longer fragments so that real positive signals are not missed (12). Some companies also provide customized arrays.

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12. All statistical methods are based on “peak searches,” but computational approaches differ and several publications propose newer and better methods to identify positive “true” signals. A simple method of analysis is to identify peaks above a certain user-defined threshold. One method called joint binding deconvolution (JBD) uses DNA sequence information (protein-DNA binding sites) to identify peaks (13). Another approach to distinguish specific signal from noise is called algorithm for capturing microarray enrichment (ACME) (12). ACME uses a sliding window approach and it is based on the assumption that the enriched signal must be positive in one direction (one-tailed) and that the signal must be represented by multiple probes (the neighbor effect). Although there are many different ways to compute ChIP-on-chip data, independent multiple replicates are necessary to identify true signals. A typical peak search for a Runx2 target gene is depicted in Fig. 13.3.

References 1. Ren, B., Robert, F., Wyrick, J. J., et al. (2000). Genome-wide location and function of DNA binding proteins. Science 290, 2306–2309. 2. Pratap, J., Javed, A., Languino, L. R., et al. (2005). The Runx2 osteogenic transcription factor regulates matrix metalloproteinase 9 in bone metastatic cancer cells and controls cell invasion. Mol Cell Biol 25, 8581–8591. 3. Javed, A., Barnes, G. L., Pratap, J., et al. (2005). Impaired intranuclear trafficking of Runx2 (AML3/CBFA1) transcription factors in breast cancer cells inhibits osteolysis in vivo. Proc Natl Acad Sci USA 102, 1454–1459. 4. Blyth, K., Cameron, E. R., Neil, J. C. (2005). The runx genes: gain or loss of function in cancer. Nat Rev Cancer 5, 376–387. 5. Young, D. W., Hassan, M. Q., Yang, X.-Q., et al. (2007). Mitotic retention of gene expression patterns by the cell fate determining transcription factor Runx2. Proc Natl Acad Sci USA 104, 3189–3194. 6. Young, D. W., Hassan, M. Q., Pratap, J., et al. (2007). Mitotic occupancy and lineage-specific transcriptional control of rRNA genes by Runx2. Nature 445, 442–446. 7. Galindo, M., Pratap, J., Young, D. W., et al. (2005). The bone-specific expression of RUNX2 oscillates during the cell cycle to support a G1 related anti-proliferative function in osteoblasts. J Biol Chem 280, 20274–20285. 8. Shen, J., Hovhannisyan, H., Lian, J. B., et al. (2003). Transcriptional induction of the osteocalcin gene during osteoblast differentiation involves acetylation of histones H3 and H4. Mol Endocrinol 17, 743–756. 9. Shen, J., Montecino, M. A., Lian, J. B., et al. (2002). Histone acetylation in vivo at the osteocalcin locus is functionally linked to vitamin D dependent, bone tissue-specific transcription. J Biol Chem 277, 20284–20292. 10. Hassan, M. Q., Javed, A., Morasso, M. I., et al. (2004). Dlx3 transcriptional regulation of osteoblast differentiation: temporal recruitment of Msx2, Dlx3, and Dlx5 homeodomain proteins to chromatin of the osteocalcin gene. Mol Cell Biol 24, 9248–9261. 11. O’Geen, H., Nicolet, C. M., Blahnik, K., et al. (2006). Comparison of sample preparation methods for ChIP-chip assays. Biotechniques 41, 577–580. 12. Scacheri, P. C., Crawford, G. E., Davis, S. (2006). Statistics for ChIP-chip and DNase hypersensitivity experiments on NimbleGen arrays. Methods Enzymol 411, 270–282. 13. Qi, Y., Rolfe, A., MacIsaac, K. D., et al. (2006). High-resolution computational models of genome binding events. Nat Biotechnol 24, 963–970.

Chapter 14

Identification of Transcription Factor Target Genes by ChIP Display Artem Barski, Steven Pregizer, and Baruch Frenkel

Abstract Transcription factors play pivotal roles in the control of cell growth and differentiation in health and disease. In the post-genomic era, it has become possible to locate the regions occupied by transcription factors throughout the genome, leading to better understanding of their mechanism of action and the genes that they regulate. All methods for transcription factor location analysis utilize chromatin immunoprecipitation (ChIP). Although ChIP was initially used to test whether a protein binds to a candidate promoter in living cells, newly developed methods allow the unbiased identification of novel targets of transcription factors. This chapter describes ChIP Display, an affordable method for transcription factor location analysis. Despite being relatively low throughput compared with alternative methods such as ChIP-chip and ChIP-SAGE, ChIP Display provides even small molecular biology laboratories with the opportunity to discover novel targets of any transcription factor, for which high-quality antibodies are available. Keywords Transcription factor location analysis, transcription factor target genes, protein-DNA interaction in vivo, chromatin immunoprecipitation, genomics.

1

Introduction

Transcription factors play pivotal roles in cell biology, animal physiology, and disease processes. The ongoing quest to understand their function includes the search for genes whose expression they regulate. Target genes for transcription factors can be identified by expression studies, although these are fraught with problems (see Note 1). A complementary group of approaches is based on the physical interaction of transcription factors with cis-acting regulatory elements, and the assumption that such elements are frequently located close to the genes that they regulate. Most of these approaches begin with chromatin immunoprecipitation (ChIP), in which DNA fragments bound by a transcription factor of interest in living cells are immunoprecipitated with antibodies against that protein. Once identified, these fragments can be mapped to the genome, and nearby genes can be tested for their regulation by the transcription factor of interest. From: Methods in Molecular Biology, Vol. 455: Osteoporosis: Methods and Protocols Edited by Jennifer J. Westendorf © Humana Press, Totowa, NJ

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The ChIP-based methods for transcription factor target discovery are challenged by the overwhelming excess of non-specifically precipitated fragments (Fig. 14.1A). Each of these methods takes a different path to identifying those fragments that were specifically immunoprecipitated. Of these methods, ChIP Display (CD) (1) is one of the least high throughput but offers relative simplicity and can be performed in a typical molecular biology laboratory without sophisticated equipment or complicated statistical analyses (see Note 2). To overcome the high background problem, CD effectively concentrates fragments representing each target while scattering the remaining DNA. Targets are concentrated via restriction digestion (see Fig. 14.1C): All fragments representing a certain target now have the same size, allowing one to resolve them as a single band on a gel. Scattering of the non-specifically precipitated fragments is achieved by dividing the total pool of restriction fragments into families based on the identity of nucleotides at the ends of such fragments (see Fig. 14.1E–G). Because all restriction fragments representing each given target have the same nucleotides at the ends, they remain in the same family and the signal is not eroded. In contrast, the other fragments, mostly background, are scattered into many families. The CD protocol is rather simple (see Fig. 14.1). Following ChIP, immunoprecipitated DNA is dephosphorylated by shrimp alkaline phosphatase (SAP) to prevent ligation of linkers to DNA ends generated by sonication (see Fig. 14.1B). SAP is then heat-inactivated and the DNA is digested with a restriction enzyme. We use AvaII, whose recognition sequence, GGWCC (W = A or T), can be expected approximately every 500 bp in a random sequence. This is followed by ligationmediated PCR (LM-PCR) using various primer combinations (see Fig. 14.1E). Up to 36 combinations of eight nested primers are employed to amplify fragments belonging to one family at a time (see Fig. 1F,G). Each such primer contains either A or T at the +3 position of the AvaII site and one nested nucleotide, A, T, G, or C, at the 3′ end (Fig. 14.1F). Amplified fragments from two to three independent immunoprecipitates and two to three control precipitates are resolved by PAGE. Bands enriched in the ChIP lanes are considered candidate targets and are excised from the gel for further characterization. Using CD, our lab has identified dozens of targets for both Runx2 and the androgen receptor (1–4).

2

Materials

2.1 1. 2. 3. 4. 5. 6.

Digestion and Linker Ligation

Short linker oligo: 5′-TTCGCGGCCGCAC-3′. Long A linker oligo: 5′-GACGTGCGGCCGCGAA-3′. Long T linker oligo: 5′-GTCGTGCGGCCGCGAA-3′. 10× React1 and 10× React2 buffers (Invitrogen, Carlsbad, CA). Shrimp alkaline phosphatase (SAP), 1 U/µL. AvaII, 10 U/µL.

a

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5kb

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Ligate linkers 5kb

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A AvaII G G T C C N +1+2 +4+5+6 +6 +3 C

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Fig. 14.1 Principles of ChIP Display (CD). See color figure on the disc accompanying this book. a. Precipitated DNA fragments (green, specific; black, non-specific) are aligned with the genome. Graph (red) describes the representation of each nucleotide in the immunoprecipitate. The 1-Mb area shown contains two hypothetical targets, #1 and #2. b. Magnification of the two regions of interest from panel (a). Precipitated DNA fragments are treated with shrimp alkaline phosphatase (SAP) to prevent linker ligation to DNA ends generated during sonication. Crossed circles depict dephosphorylated end. (c) DNA is digested with AvaII. d. Linkers are ligated to the ends of AvaII fragments. Also shown are the nested primers. Note that different primers will amplify targets #1 and #2 (see color plate). e. PCR products amplified in three reactions: Left: Target #1 is amplified in a reaction with a single primer (family X in panel G). Middle: Target #2 is amplified with two different primers (family V in panel G); note that target #1 is amplified here again; Right: Most of the PCR reactions will amplify neither target #1 nor target #2; however, they will amplify targets from other loci. f. Linkers and nested primers. Positions +1 to +6 are defined at the top. Nucleotides at positions +3 and +6 (highlighted) are used to segregate AvaII fragments into families. Linkers (blue) contain a 1:1 A:T mixture (W) at position +3 and a C at position +1 to destroy the AvaII sites. In addition, this panel shows one of the eight PCR nested primers, the one with T at position +3 and C at position +6, which would amplify the family indicated by X in panel G. g. Thirty-six families of fragments, each amplified using one or two of eight nested primers. White squares correspond to a single primer and shaded squares correspond to two different primers. (Reproduced from Barski, A., Frenkel, B. (2004) Nucleic Acids Res 32, e104, with permission from Oxford University Press.)

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7. T4 DNA ligase, 400 U/µL. 8. 10× T4 ligase buffer. 9. Enzymatic reaction cleanup kit (Qiagen, Valencia, CA).

2.2 1. 2. 3. 4. 5.

PCR Amplification

Taq DNA polymerase, 5 U/µL. 10× PCR Reaction Buffer, –MgCl2. MgCl2, 50 mM. dNTP mix (Invitrogen), 10 mM each. Up to eight PCR primers: 5′-CGGCCGCACGWCCN-3′, where W is A or T and N is A, G, C, or T (see Note 2).

2.3

PAGE

1. 2. 3. 4.

40% polyacrylamide solution [29:1 (w/w) acrylamide/bisacrylamide]. 10% (w/v) ammonium persulfate (APS) in water (store ≤1 month at 4 °C). TEMED solution (N,N,N′,N′-tetramethylethylenediamine; store at 4 °C). 10× TBE Electrophoresis Buffer: pH 8.0: 890 mM Tris, 890 mM boric acid, and 20 mM EDTA. 5. 5× Sample Loading Buffer: 10% Ficoll 400, 50 mM disodium EDTA, pH 8.0, 0.5% sodium dodecyl sulfate, 0.25% bromphenol blue, 0.25% xylene cyanol. 6. Gel loading pipet tips. 7. 1-kb DNA ladder.

2.4 1. 2. 3. 4. 5. 6. 7. 8.

Target Identification

Spin-X Centrifuge Tube Filters (Corning, Corning, NY). HaeIII, HinfI, MspI, RsaI, 10 U/µL each. Standard/High Melt Agarose. Gel extraction kit (Qiagen). BigDye Terminator 3.1 Cycle Sequencing Kit (Applied Biosystems, Foster City, CA). AutoSeq G-50 Microcentrifuge Columns (Amersham). Hi-Di™ Formamide (Applied Biosystems). For in-house sequencing: ABI PRISM 3100 Genetic Analyzer (Applied Biosystems) or an alternative system.

14 Target Genes by ChIP Display

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Methods

CD begins after ChIP has been completed with the cells and antibodies of interest. To identify Runx2 targets in osteoblasts, we usually ChIP half the chromatin collected from two confluent 100-mm plates of MC3T3-E1 cells with 5 µg of Runx2-specific antibodies (Santa Cruz, cat. no. 10758X). The other half is immunoprecipitated with 5 µg of non-specific IgG to control for non-specific IP. Each of the specific and non-specific ChIPs is done in duplicate or triplicate to control for experimental variation. Thus, we typically initiate a CD experiment for a given experimental condition with two or three ChIP samples and two or three mock-ChIP samples. Each of the samples is cleaned with an Enzymatic Reaction Cleanup Kit (Qiagen) and eluted in 50 µL H2O, which is sufficient for about 100 separate CD reactions. Each CD reaction is then displayed in one lane of an acrylamide gel, and will typically yield up to two targets. Although this chapter does not describe the ChIP procedure, two critical points must be emphasized: (1) No salmon sperm DNA should be used for ChIP intended for CD. Instead, substitute with bacterial tRNA (see Note 3); and (2) high-quality ChIPs are essential for the success of CD. Before a CD protocol is initiated, one must ensure that the starting ChIP material is enriched for at least one, and preferably more, known targets. We recommend the use of real-time PCR to determine the enrichment levels, with both internal (non-targets) and external (IgG) controls (see Note 4). If the enrichment levels are satisfactory (∼10-fold or better), then this material may be used for CD.

3.1

Digestion and Linker Ligation

1. Resuspend each of the two long linker oligos (“A” and “T”) and the short linker oligo in dH2O at a 100 µM concentration. 2. Mix 15 µL of the short linker oligo with 15 µL of the long “A” linker oligo. Add 10 µL of 10× React2 buffer and 60 µL of dH2O. Likewise, mix 15 µL of the short linker oligo with 15 µL of the long “T” linker oligo and add 10 µL of 10× React2 buffer and 60 µL of dH2O. Mix each well. 3. Anneal oligos by placing samples in a beaker of boiling water for 1 minute. Turn off heat and allow water to return to room temperature without removing samples. After cooling, annealed linkers may be stored at −20 °C. 4. Mix 10 µL of each ChIP sample with 2 µL of 10× React1 buffer, 1 µL of SAP, and 7 µL of H2O. Incubate at 37 °C for 30 minutes. Heat denature for 15 minutes at 65 °C (see Note 5). 5. Add 1 µL (10 units) of AvaII to each sample. Incubate at 37 °C for 30 minutes (see Note 6). 6. Add 2 µL each of both annealed linkers, 3 µL of 10× T4 ligase buffer and 1 µL T4 DNA ligase (400 U). Incubate overnight at 16 °C.

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7. Add 0.5 µL of AvaII and 0.5 µL of T4 Ligase to each sample. Incubate 1 hour at room temperature (see Note 7). Purify the samples on QuickSpin columns using Enzymatic Reaction Clean-Up Buffer. Elute with 50 µL dH2O.

3.2

PCR Amplification

1. Make a PCR master mix containing 1× PCR Buffer, 1.5 mM MgCl2, 0.2 mM each dNTP, 0.025 U/µL Taq DNA Polymerase (see Note 8), and 0.5 µM of one or two of the eight possible primers (see Note 9). 2. Add 18 µL of the PCR master mix to 2 µL of each replicate of each columnpurified ChIP sample (see Notes 10 and 11). 3. Mix by pipetting up and down, and then place tubes in a thermocycler at room temperature (see Note 12). 4. Amplify the three samples representing each IP using two or three annealing temperatures between 68–70 °C. Specifically, amplify each set of three samples using the following program: (1) 72 °C for 30 seconds; (2) 95 °C for 5 minutes; (3) 95 °C for 1 minute; (4) 68 °C [or 69 or 70 °C] for 1 minute; (5) 72 °C for 1 minute; (6) go to 3, 45 times; (7) 72 °C for 5 minutes; (8) End (see Figs. 14.2A and 14.3, and Note 13).

a

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Fig. 14.2 Target display and identification. a. DNA samples from three independent ChIPs and three mock ChIPs were. DNA samples from three independent ChIPs and three mock ChIPs were subjected to LM-PCR with one of the 36 possible primer combinations (Fig. 14.1G). Each PCR was performed with an annealing temperature of either 68 or 69 °C as indicated. The products were then resolved side-by-side on an 8% polyacrylamide gel. Bands indicated with white arrowheads are not putative targets because they are comparably present in both the ChIP and the mock lanes. Bands representing a putative target are indicated with black arrowheads. The bands labeled i and ii were excised from the gel, purified and re-amplified. b. The re-amplified products from panel A were treated with the restriction enzymes HaeIII (Ha), HinfI (Hi), and MspI (Ms). The digestion products were resolved on a 4% agarose gel. This example demonstrates that targets can be identified even in the presence of contaminating bands, seen in the lanes containing undigested products (U). The bands of interest, identified based on their size (compare with panel A), are indicated with arrows. A sub-fragment likely to have arisen from the band of interest is indicated in each of the HinfI lanes with an arrowhead. Indeed, after sequencing these AvaII-HinfI sub-fragments and after mapping of the “hit” to the mouse genome, it was shown to reside between two AvaII sites separated by 473 bp, consistent with the size of the band of interest that was excised from the acrylamide gel (panel A, and see Section 3.4.11.). M, Molecular weight marker.

14 Target Genes by ChIP Display a M

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Fig. 14.3 Effect of dilution and annealing temperature on CD. a. Three dilutions (1:1, 1:4, 1:16) of Runx2 ChIP (+) or mock ChIP (−) were amplified with one of the 36 possible primer combinations (Fig. 14.1G) at an annealing temperature of 68, 69, or 70 °C, and the products were resolved on an 8% non-denaturing polyacrylamide gel. Arrowheads indicate a putative target that was reproducible at all dilutions, and with both 70 and 69 °C, but not 68 °C, as the annealing temperature. b. Another example, using a different primer set. Arrowheads indicate a putative target that appeared only with annealing temperatures of 69 and 70 °C, and only without dilution. Although this candidate is inferior to the one in panel A, it can be considered a putative target, but might very well prove to be a false-positive during the validation steps (Sections 3.4.11. and 3.4.12.). M, Molecular weight marker.

3.3

PAGE and Target Isolation

We recommend that the CD procedure be completed with one or two of the 36 primer combinations (represented by the 36 boxes in Fig. 14.1G) before moving on to additional families. Only very experienced investigators should consider performing all of the LM-PCR reactions with all possible primer combinations (see Fig. 14.1G) prior to PAGE and target identification. 1. Cast a 1× TBE, 8% native polyacrylamide gel for resolving nucleic acids. We use gels 14 cm long and 17 cm wide. 2. After polymerization, load the wells with 1× sample loading buffer and pre-run in 1× TBE for 30 minutes at 150 V.

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3. Add 5 µL of 5× sample loading buffer to each sample after amplification and mix well. 4. Flush the wells of the gel thoroughly with a syringe. Using gel loading pipette tips, carefully apply 10 µL of each sample to the bottom of each well. We typically load samples immunoprecipitated with specific antibodies on one-half of the gel, whereas samples immunoprecipitated with non-specific antibody are loaded on the other half (see Fig. 14.2A). We load replicates next to each other and place molecular weight markers in the middle and on each side of the gel. 5. Allow the gel to run at 150–200 V, until the xylene cyanol dye is 3 cm from the bottom of the gel. 6. Remove the gel and incubate in 1× TBE supplemented with 100 ng/mL ethidium bromide. Visualize bands using a UV transilluminator. Capture and print the image. 7. Note any co-migrating bands that are reproducible in the ChIPs with the specific antibody. If these do not co-migrate with bands in the control lanes, then they are considered candidate targets (see Figs. 14.2A and 14.3, and Note 14). 8. Cut the bands that represent good candidate targets out of the gel using a fresh blade for each band. Be careful to avoid neighboring bands and lanes. 9. Place each excised gel piece in a Spin-X column, and freeze at −80 °C. Spin the columns at maximum speed in a microcentrifuge for 15 minutes at room temperature.

3.4

Target Identification

In principle, targets can be identified by cloning into a vector followed by sequencing. However, we prefer to identify targets without cloning, by directly sequencing sub-fragments obtained through secondary digestion as described in the following section (see Note 15). 1. Re-amplify the eluted DNA using the same primers and conditions as in Section 3.2., in a total reaction volume of 50 µL. 2. Split each amplified DNA sample into five 10 µL aliquots. Digest 2–4 of these aliquots with different restriction enzymes, each in a final volume of 20 µL (see Note 15). 3. After digestion, add 5 µL of 5× sample loading buffer to each sample, and mix thoroughly. Load 10 µL of each sample on a 4% agarose gel containing 100 ng/mL ethidium bromide. Load the undigested sample in lane #1, followed by the digested samples (see example in Fig. 14.2B). Repeat this for each group of samples, separating the groups with 1-kb DNA ladder. Allow the gel to run at 100 volts for 2–3 hours. 4. Place the agarose gel on a UV transilluminator, and note the pattern of bands (sub-fragments) resulting from the restriction digestion for each candidate target. Look for identical patterns of sub-fragments, which appear to have originated from a common target. Excise several such sub-fragments from the gel for sequencing (see Note 16).


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5. Purify the DNA on MinElute columns (Qiagen) according to the manufacturer’s protocol. Sequencing of the eluted DNA can be performed either in house or using a service lab. Steps 6–9 describe the sequencing procedure used in our lab. 6. Mix 10 µL of each eluted DNA sample with 2 µL 5× BigDye Sequencing Buffer, 4 µL Ready Reaction Premix, and 5 pmol of the primers used in Step 3.2.1. Add dH2O up to 20 µL. Place samples in a thermocycler and run the following program: (1) 95 °C for 4 minutes; (2) 93 °C for 30 seconds; (3) 50 °C for 15 seconds; (4) 60 °C for 4 minutes; (5) Go to 2, 30 times; and (6) End. 7. Remove samples from the thermocycler and clean DNA with AutoSeq G-50 Microcentrifuge Columns. 8. Place samples in SpeedVac and dry under low heat. Resuspend in 10 µL Hi-Di Formamide. 9. Transfer samples to a 96-well plate. Heat to 100 °C for 5 minutes and cool immediately on ice for 2 minutes. Load plate into ABI PRISM 3100 Genetic Analyzer and begin run. 10. Map the resulting sequences within the genome of interest, using BLAST or a similar search engine. 11. Search the regions surrounding the “hit” for AvaII recognition sequences. The distance (in bps) between the two sites on either side of the sequenced region should closely match the size of the fragment that was excised from the polyacrylamide gel. Additionally, the identities of the nucleotides at the ends of each fragment should match the primer pair used to generate the fragment in Section 3.2. Finally, the fragment should contain restriction enzyme recognition sequences at positions that are compatible with the digestion patterns that were seen on the agarose gel (step 3.4.4). 12. Validate each “hit” of interest using conventional ChIP assay with locus-specific primers (see Note 17). Annotated genes near confirmed targets are potentially regulated by the transcription factor of interest.

4

Notes

1. Target genes can be identified in comprehensive gene expression studies (e.g., using microarrays) based on their response to over-expression or activation (e. g., by ligand) of transcription factors. However, there are at least four problems with expression-based approaches. First, it is generally difficult to tell whether a responsive gene is a direct or indirect target of the transcription factor of interest. Second, in experiments involving over-expression of transcription factors, the response of some genes may be forced by exaggerated concentrations of the protein, resulting in physiologically insignificant results. Third, such studies do not provide information on the location of the cis-acting regulatory elements. Last, expression studies are unable to disclose genes, to which the transcription factor of interest binds without functional consequences under the experimental

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conditions employed. Such genes may become responsive to the transcription factor under different conditions, for example, in the presence of a specific extracellular stimulus and/or a particular co-activator (either in the same or a different cell type). CD is a “sampling” method and will only disclose a fraction of all the targets. High throughput methods for transcription factor target discovery include ChIPChip (5–7), SABE (8), STAGE (9), ChIP-PET (10), GMAT (11), SACO (12), and DamID (13). These methods are also used to map histone modifications, a task for which CD is not well suited. However, the high throughput methods share requirements, which render them prohibitively expensive for many labs. First, the amount of starting material needed is relatively large. Second, complex statistical analysis is needed to discern true targets from the large number of non-specifically immunoprecipitated fragments. Third, complicated equipment is required and the price is extremely high. This is exacerbated by the possibility of poor reproducibility, which usually necessitates many repetitions, especially in the beginning of a project with a new transcription factor. In addition, unlike some of the high throughput methods listed in the preceding, CD can lead to novel targets before a complete screen is finished. Of the 36 reaction types that constitute a full screen (see Fig. 14.1G), one can start with one or a subset of reactions and continue only if more targets are desirable. Furthermore, CD provides the opportunity to compare occupancy between different cells, and pursue only those with interesting phenotypes. For example, one can display side-byside targets occupied in two different cell types, or under different physiological conditions, and then pursue the identification of only selected targets. Any ChIP protocol can be adapted for CD. However, the typical high background of non-specifically immunoprecipitated fragments, which can be tolerated in conventional ChIP assays of candidate targets, is the Achilles’ heel of methods intended to identify unknown targets. Although CD is relatively forgiving in this sense, it will not distinguish between targets and contaminating DNA. Therefore, extra care should be taken to prevent contamination. Related to this, the beads used for immunoprecipitation should be pre-adsorbed with bacterial tRNA instead of salmon sperm DNA. tRNA will not be ligated or amplified during the following steps. Alternatively, the pre-adsorption step can be skipped altogether. Enrichment for known targets is a measure of the success of a ChIP experiment. Using real-time PCR with locus-specific primers, one compares the concentrations of known targets to those of non-targets after ChIP. To more rigorously measure the enrichment factor, the known-target-to-non-target ratio in the ChIP is compared to the respective ratio in samples immunoprecipitated with nonspecific antibodies, which should in theory be equal to 1 for single-copy genes. In our lab, we achieve a ~10-fold enrichment for Runx2 targets in MC3T3-E1 cells. Prior to AvaII treatment, we dephosphorylate the DNA with SAP to prevent ligation of linkers to the ends produced by sonication. We include this step even though we are not sure that sonication produces ends that are compatible with


6.

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8.

9.

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ligation. SAP was chosen because it can be efficiently heat denatured to prevent dephosphorylation of fragments generated later for LM-PCR by AvaII digestion. A key feature of CD is the restriction digestion, a step that standardizes the size of a diverse group of DNA fragments obtained from a given target after sonication and ChIP. The ability of CD to disclose a given target depends on the presence of two recognition sites for the restriction enzyme in the vicinity of that target. We use the 5-bp cutter AvaII because the genomic distribution of its cognate sites can be expected to result in many sonication fragments (which themselves are on the order of 300–900 bp) containing two AvaII sites. Sites for 6-bp cutters occur too infrequently, so it is unlikely that two sites would be present in one sonication fragment. Of course even for 5-cutters such as AvaII, many sonication fragments will not be long enough and targets will be missed. Digestion with 4-bp cutters, however, would result in a large number of fragments and would unnecessarily increase the complexity of the DNA fragment pool. Of several possible 5-bp cutters, we chose AvaII because it is active in salt-free React1 buffer, eliminating the need for desalting prior to ligation. Conveniently, AvaII is an inexpensive and stable enzyme. Noteworthy, digestion by AvaII can be blocked by CpG methylation. Although this enzyme property may limit target discovery, it can be considered advantageous because methylated regions may be associated with less accessible genes. Be that as it may, among the initial Runx2 targets identified by CD, a fair number were associated with CpG islands. The second dose of AvaII is added to digest concatemers of AvaII fragments that might have been generated during linker ligation. Re-digestion with AvaII recycles these concatemers, whereas the presence of ligase facilitates linker ligation. Because the linkers do not restore the AvaII site (see Fig. 14.1F), their ligation to digested DNA fragments is irreversible even in the presence of AvaII. We use Taq DNA polymerase because it lacks proofreading activity. This prevents “editing” of the nested primers, which may anneal to the template even when the nucleotides at positions +3 or +6 (see Fig. 14.1F) are not complementary. Such mismatches would be perfect substrates for a 3′–5′ proofreader, which would then use the edited primer to amplify the fragment even when it should not be amplified in that specific reaction. By amplifying targets in individual families (see Fig. 14.1G), we increase the signal-to-noise ratio for one group of targets at a time. Because all fragments originating from a given target have the same nucleotides at the ends, they remain in the same family and the signal is not eroded. In contrast, the background fragments are scattered into many families (see Fig. 14.1E). This feature of CD also provides investigators with the choice of performing either a full or a partial screen, i.e., using as many primer combinations (families) as they wish. In our search for Runx2 target genes in MC3T3-E1 cells, we limited the initial CD screen to eight families, those that are amplified by a single primer each (1), and later continued with other families (2).

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10. In addition to biological replicates, we strongly encourage the use of technical replicates. There is a huge number of non-specifically immunoprecipitated fragments in each ChIP sample that are all competing for a limited number of primers in the amplification reaction. Usually, a subset of these fragments wins out early on in the competition, often for random reasons. The so-called “founder effect” and the complexity of the template DNA result in a different pattern of amplification from run-to-run of the same sample (see Fig. 14.2A). Although fragments that represent true targets are present at a higher concentration and should be amplified reproducibly, technical replicates help one to distinguish between true targets and spuriously amplified fragments (see Fig. 14.2A). 11. As an alternative to using direct replicates, it may be worthwhile to perform the PCR on different dilutions of the ChIP material. Seeing a particular band with all dilutions enhances one’s confidence that it truly represents a target (see Fig. 14.3A). Other bands, which are visible with lower dilutions, but not with higher dilutions (see example in Fig. 14.3B) can be either dismissed or pursued contingent upon validation (step 3.4.12). 12. Do not pre-heat the thermocycler, because this might compromise amplification efficiency. The linkers used for CD are unphosphorylated oligonucleotide duplexes; thus, only the short oligo of the duplex is ligated to the AvaII fragment. The long oligo is kept in place through non-covalent interactions and will be lost upon heating. To allow the polymerase to fill-in the lost sequence before the strands are completely separated, we start at room temperature and hold the reaction for 30 seconds at 72 °C. Hot-start polymerases complexed with antibodies should not be used in this PCR as they become active only after the reaction is heated to 95 °C. 13. Due to the large amount of non-specifically precipitated fragments in a ChIP sample, non-specific amplification is a technical challenge that must be overcome by CD. To this end, we use short (14-bp) primers and high annealing temperatures. Unfortunately, it is possible for the annealing temperature selected to be too stringent for any amplification to occur. Because it is difficult to predict what annealing temperature will yield the right balance between too stringent and too permissive, we recommend that replicates be performed at slightly different annealing temperatures. This can be accomplished easily using a thermocycler with gradient capabilities. Two or three temperatures between 68–70 °C usually provide good results (see Figs. 14.2 and 14.3). 14. In theory, targets should appear as co-migrating bands in all of the ChIP lanes but none of the mock ChIP lanes. In practice, such perfect scenarios are infrequent (see Fig. 14.2A), and investigators must exercise their own good judgment in selecting bands for further analysis. In any case, each putative target must be validated using conventional ChIP assay with locus-specific primers (step 3.4.12). 15. There are at least three reasons for the secondary amplification and digest. First, it is useful as a screen to confirm that co-migrating bands from different lanes actually contain the same sequence. Second, it helps to eliminate contamination from overlapping and neighboring bands present on the polyacrylamide gel. Restriction digestion results in a unique pattern of sub-fragments arising from


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the target; sub-fragments that participate in such a pattern will likely lead to identification of true targets (see Fig. 14.2B). Third, the secondary digestion facilitates the sequencing step. Because the primers used for sequencing are the same as those used for amplification, they would anneal to both ends of an intact fragment, producing two simultaneous and thus uninterpretable sequences. Digestion of the fragments is intended to create a sequence-able template with only one end to which a primer can anneal. Because most of the fragments isolated after PAGE are up to 500-bp in length, it is best to start the secondary digestion with enzymes that have a short recognition sequence, usually 4-bp cutters. Restriction enzymes that work well in our hands for the secondary digestion are HaeIII, MspI, RsaI, and HinfI, all of which are active in the PCR buffer. If these enzymes do not yield sequence-able sub-fragments, digestion with a panel of additional 4-bp cutters, or even 5- or 6-bp cutters alone or in combination, is advisable. Again, to avoid a desalting step, it is helpful to select enzymes that are active in the PCR buffer (a.k.a. primer extension mix). 16. Bands excised from the agarose gel must have one intact end containing the PCR primer in order to be sequenced. Because some bands may have lost both original ends during the secondary digest, we usually excise more than one band from the agarose gel, with preference toward those that are only slightly smaller than the undigested fragment. It is also advisable to excise several replicate sub-fragments because even sequence-able sub-fragments can become corrupted during gel extraction or the sequencing step itself. Having replicates increases the odds of obtaining a sequence. 17. For validation by conventional ChIP, there are at least two important considerations. First, CD does not disclose the exact sequence to which transcription factors bind. The discrete binding site may lie anywhere within the AvaII fragment, or even just outside the fragment. Thus, when designing primers for validation, it may be useful to target more than one sequence within, or immediately surrounding, the AvaII fragment. Cognate binding motifs found in the region can provide guidance for primer design. Second, validation of a target disclosed by CD is most likely to occur when using the same ChIP material that was used as input for the CD experiment. This is due to the fact that transcription factor occupancy at some loci may be sensitive to minor alterations in culture conditions. For this reason, it may be wise not to use all of the original ChIP material for CD, so that the leftover can be employed for validation. In addition, we recommend the validation of each target in independent ChIPs, which would demonstrate that occupancy by the transcription factor of interest occurs regardless of minor alterations to the culture conditions. Acknowledgments ChIP Display was developed with NIH grant R21 AR49297 and further employed in projects supported by the NIH (RO1 DK071122 and RO1 CA109147) and the Department of Defense (PC040322). The work was conducted in a facility constructed with support from Research Facilities Improvement Program grant no. C06 (RR10600-01, CA62528-01, RR14514-01) from the NIH/NCRR. BF holds the J. Harold and Edna L. LaBriola Chair in Genetic Orthopaedic Research at the University of Southern California.

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References 1. Barski, A. and Frenkel, B. (2004) ChIP Display: novel method for identification of genomic targets of transcription factors. Nucleic Acids Res, 32, e104. 2. Pregizer, S., Barski, A., Gersbach, C.A., Garcia, A.J. and Frenkel, B. (2007) Identification of novel Runx2 targets in osteoblasts: Cell type-specific BMP-dependent regulation of Tram2. J Cell Biochem, 102, 1458–1471. 3. Jariwala, U., Prescott, J., Jia, L., Barski, A., Pregizer, S., Cogan, J.P., Arasheben, A., Tilley, W.D., Scher, H.I., Gerald, W.L. et al. (2007) Identification of novel androgen receptor target genes in prostate cancer. Mol Cancer, 6, 39. 4. Prescott, J., Jariwala, U., Jia, L., Cogan, J.P., Barski, A., Pregizer, S., Shen, H.C., Arasheben, A., Neilson, J.J., Frenkel, B. et al. (2007) Androgen receptor-mediated repression of novel target genes. Prostate, 67, 1371–1383. 5. Cawley, S., Bekiranov, S., Ng, H.H., Kapranov, P., Sekinger, E.A., Kampa, D., Piccolboni, A., Sementchenko, V., Cheng, J., Williams, A.J. et al. (2004) Unbiased mapping of transcription factor binding sites along human chromosomes 21 and 22 points to widespread regulation of noncoding RNAs. Cell, 116, 499–509. 6. Horak, C.E. and Snyder, M. (2002) ChIP-chip: a genomic approach for identifying transcription factor binding sites. Methods Enzymol, 350, 469–483. 7. Ren, B., Robert, F., Wyrick, J.J., Aparicio, O., Jennings, E.G., Simon, I., Zeitlinger, J., Schreiber, J., Hannett, N., Kanin, E. et al. (2000) Genome-wide location and function of DNA binding proteins. Science, 290, 2306–2309. 8. Chen, J. and Sadowski, I. (2005) Identification of the mismatch repair genes PMS2 and MLH1 as p53 target genes by using serial analysis of binding elements. Proc Natl Acad Sci U S A, 102, 4813–4818. 9. Kim, J., Bhinge, A.A., Morgan, X.C. and Iyer, V.R. (2005) Mapping DNA-protein interactions in large genomes by sequence tag analysis of genomic enrichment. Nat Methods, 2, 47–53. 10. Wei, C.L., Wu, Q., Vega, V.B., Chiu, K.P., Ng, P., Zhang, T., Shahab, A., Yong, H.C., Fu, Y., Weng, Z. et al. (2006) A global map of p53 transcription-factor binding sites in the human genome. Cell, 124, 207–219. 11. Roh, T.Y., Ngau, W.C., Cui, K., Landsman, D. and Zhao, K. (2004) High-resolution genomewide mapping of histone modifications. Nat Biotechnol, 22, 1013–1016. 12. Impey, S., McCorkle, S.R., Cha-Molstad, H., Dwyer, J.M., Yochum, G.S., Boss, J.M., McWeeney, S., Dunn, J.J., Mandel, G. and Goodman, R.H. (2004) Defining the CREB regulon: a genome-wide analysis of transcription factor regulatory regions. Cell, 119, 1041–1054. 13. van Steensel, B. and Henikoff, S. (2000) Identification of in vivo DNA targets of chromatin proteins using tethered dam methyltransferase. Nat Biotechnol, 18, 424–428.

Chapter 15

Application of the Laser Capture Microdissection Technique for Molecular Definition of Skeletal Cell Differentiation In Vivo Dafna Benayahu, Rina Socher, and Irena Shur

Abstract Laser capture microdissection (LCM) method allows selection of individual or clustered cells from intact tissues. This technology enables one to pick cells from tissues that are difficult to study individually, sort the anatomical complexity of these tissues, and make the cells available for molecular analyses. Following the cells’ extraction, the nucleic acids and proteins can be isolated and used for multiple applications that provide an opportunity to uncover the molecular control of cellular fate in the natural microenvironment. Utilization of LCM for the molecular analysis of cells from skeletal tissues will enable one to study differential patterns of gene expression in the native intact skeletal tissue with reliable interpretation of function for known genes as well as to discover novel genes. Variability between samples may be caused either by differences in the tissue samples (different areas isolated from the same section) or some variances in sample handling. LCM is a multi-task technology that combines histology, microscopy work, and dedicated molecular biology. The LCM application will provide results that will pave the way toward high throughput profiling of tissue-specific gene expression using Gene Chip arrays. Detailed description of in vivo molecular pathways will make it possible to elaborate on control systems to apply for the repair of genetic or metabolic diseases of skeletal tissues. Keywords Laser capture microdissection, skeletal tissues, gene expression, RNA isolation, chromatin immunoprecipitation.

1

Introduction

Skeletal tissues develop from mesenchymal stem cells that differentiate to chondrogenic, osteogenic, hematopoietic-supporting and skeletal muscle cells (1–5). Skeletal tissues are active throughout life, in skeletal growth and remodeling, and are regulated by multiple factors, including systemic hormones and local regulatory factors (5–8). A cell’s differentiation is coordinated by activation or repression of transcription factors implicated in regulation of proliferation, and of functional From: Methods in Molecular Biology, Vol. 455: Osteoporosis: Methods and Protocols Edited by Jennifer J. Westendorf © Humana Press, Totowa, NJ

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genes for certain cytokines, their receptors, and ECM proteins that serve a crucial role in skeletal differentiation. A decrease in osteoblast activity and an increase in bone resorption by osteoclasts enhance bone remodeling, which results in osteoporosis. Studying the alterations in cell differentiation will teach us more about the connection between stem cell differentiation and skeletal pathology which occurs in different metabolic disorders including osteoporosis. Disfunction of skeletal cells leads to osteopenia and osteoporosis, diseases that are responsible for millions of fractures annually, mostly involving the lumbar vertebrae, hip, and wrist. Studying the changes in bone physiology in animal models will shed light on the type of changes that may contribute to develop modalities to attenuated bone loss. The laser capture micro-dissection (LCM) method allows the selection of individual or clustered cells from intact tissues. Subsequent RNA/DNA and/or protein extraction enables one to interpret the molecular mechanisms that function in vivo in the particular cells and tissues. LCM provides the opportunity to uncover the molecular mechanisms controlling cellular fate in the natural microenvironment. This technique is particularly useful for selecting cells from skeletal tissues that are difficult to study individually because of their anatomical complexity (10, 11). The LCM technique is particularly attractive for in vivo characterization of mesenchymal cells because it allows the definition of molecular mechanisms that drive differentiation of distinct cells in skeletal tissues. Single cell analysis sheds light on alterations in regulatory processes, which occur in osteoporosis, or changes of physiological status, such as aging. The use of LCM overcomes the problem of tissue heterogeneity and allows for the selection of defined cells while minimizing contamination from neighboring unrelated tissues. It is now realistic to extract small numbers of cells, isolate RNA, DNA and protein, and perform various molecular analyses, including chromatin immunoprecipitation (ChIP), to compare different types of cells from various tissues. Such analysis provides reliable interpretation of gene function in specific tissues. An analysis of tissue-specific gene expression in vivo enables one to determine the specific tissue control systems required for their differentiation. In addition, these methods enable discovery of tissue-specific genes and provide detailed descriptions of in vivo molecular pathways. The presented method can be used to obtain data that could be applied to prevent or delay the onset of age-related skeletal disabilities and diseases. Acquired knowledge about the intrinsic control of osteoblast transcription factors and gene expression in vivo will serve to develop novel diagnostic and therapeutic tools for various pathologies of the bone.

2

Materials

For molecular analyses, use gloves throughout the entire procedure along with RNase-free instruments and reagents.

15 The Laser Capture Microdissection Technique

2.1

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Preparation of Samples for Laser Capture Microdissection

1. Frozen samples are embedded in Tissue-Tek embedding medium (OCT, Torrance, CA, United States). 2. One-millimeter-thick sterile glass slides, or membrane covered slides (PALM, Germany). 3. Phosphate buffer solution (PBS) containing 40 units of ribonuclease inhibitor. 4. 70% ethanol for fixation.

2.2

Histological Staining

1. Mayer’s hematoxylin (H&E) solution. 2. 70%, 96%, 100% ethanol solutions.

2.3

Laser Pressure Catapulting of the Samples

1. Catapult Buffer: 0.5 µM EDTA pH 8.0, 20 µM Tris-HCl, pH 8.0. 2. Proteinase K: 20 mg/mL in 100 µL of double distilled water (DDW) that is DEPC-treated (see Note 1). 3. Sterile mineral oil.

2.4 1. 2. 3. 4. 5. 6. 7. 8. 9. 10. 11. 12. 13. 14.

RNA Analysis RNAeasy total kit (Qiagen, Germany). Trizol (Sigma) for extraction in a phenol precipitation-based method. PALM RNA-extraction kit (PALM). RNeasyMini (Qiagen). Absolutely RNA nanoprep kit (Stratagene, La Jolla, USA). High Pure RNA tissue kit (Roche, Switzerland). Purescript RNA isolation kit (Gentra, Minneapolis, USA). ArrayPure Nano-scale RNA purification kit (Epicentre Biotechnologies, Madison, USA). SuperScript III reverse transcriptase (Invitrogen, Carlsbad, USA). iScript Select cDNA synthesis kit (Bio-Rad). Reverse-i MAX Blend kit (ABgene, United Kingdom). PCR mix (Sigma, USA). Tris Borate EDTA (TBE) buffer. SeaKem LE Agarose (Cambrex, New Jersey, USA).

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Chromatin Immunoprecipitation (ChIP) for LCM Samples

1. SDS Lysis Buffer: 1% SDS, 10 mM EDTA, 50 mM Tris-HCl, pH 8.1. 2. ChIP Dilution Buffer: 0.01% SDS, 1.1% Triton X-100, 1.2 mM EDTA, 16.7 mM Tris-HCl, pH 8.1, 167 mM NaCl. 3. Low Salt Wash Buffer: 0.1% SDS, 1% Triton X-100, 2 mM EDTA, 20 mM TrisHCl, pH 8.1, 150 mM NaCl. 4. High Salt Wash Buffer: 0.1% SDS, 1% Triton X-100, 2 mM EDTA, 20 mM TrisHCl, pH 8.1, 500 mM NaCl. 5. LiCl Wash Buffer: 0.25 M LiCl, 1% IGEPAL-CA630, 1% deoxycholic acid (sodium salt), 1 mM EDTA, 10 mM Tris-HCl, pH 8.1. 6. TE Buffer: 10 mM Tris-HCl, 1 mM EDTA, pH 8.0. 7. Elution Buffer: 1% SDS, 0.1 M NaHCO3.

3 3.1

Methods Preparation of Slides

1. Freeze isolated tissues in liquid nitrogen. 2. Embed frozen tissue in optimal cutting temperature (OCT) Tissue-Tek embedding medium. 3. Cut 6-µm thickness sections at −20 °C using a cryostat (e.g., Jung Frigocut 2800N microtome, Leica). 4. Mount cryo-tissue sections on the regular 1-mm thick sterile glass slides, or membrane covered slides. 5. Sections may be fixed or deep frozen at −80 °C for the longer storage (see Note 2).

3.2

Fixation

1. Air-dry slides for 20 seconds. 2. For RNA isolation it is recommended to pipet the RNase inhibitor solution on the slide, then dip the mounted sections for 90 seconds into 70% ethanol. The dip in RNase-free water (see Note 3). 3. Allow the fixed sections to dry at the room temperature for 60 minutes.

3.3

Histological Staining

Sections may be used unstained if areas of interest are recognized based on morphology, or can be stained for better visualization (see Note 4). Mayer’s hematoxylin/ eosin (H&E) is a routine histological staining, which results with nuclei stained in blue and cytoplasm is pink/red (Fig. 15.1).


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Fig. 15.1 Morphology of stained (a) and unstained (b,c) sections. Sections stained with H&E are used for the visualization of periosteum (P) and bone (B). The isolated areas are marked by arrows.

1. Immerse the sections for 3 minutes in Mayer’s hematoxylin solution followed by 10 minutes of rinsing in water. 2. Immerse the sections for 3 minutes in eosin, quickly dip in water. 3. Following staining, slides are dehydrated in increasing ethanol solutions of 70%, 96%, and 100%. 4. Air dry for 60 minutes.

3.4

Specimen Isolation and Collection

3.4.1

Laser Cutting (Microdissection) of the Samples

1. Search for an area of interest on the section. 2. Pipette 5 µL of ethanol onto the depiction area to improve the visualization of the cells. 3. Mark the cells or cell area with the software tool. The ethanol will evaporate rapidly. Next you will catapult the marked cells or cell areas.

3.4.2

Laser Pressure Catapulting of the Samples

Use the cap with liquid that will allow the catapulted samples to stick to the cap. You will add autoclaved mineral oil (PCR oil) or catapult buffer in the inner ring of the cap. Aqueous solutions dry out after a while. The catapulted cells or cell areas stick onto the wet inner surface of the cap and do not fall down after the catapulting procedure (see Note 5). Alternatively use silicon caps that are clear adhesive caps without liquid. There is no danger of RNase activity in the absence of water, nor evaporation and crystal formation during extended sample harvesting. On the other hand, low energy is needed for catapulting due to short target distance. 1. Pipet 3–6 µL of catapult buffer into the middle of the cap (an optional method is described in the following). 2. Put the cap into the cap holder. 3. Perform laser microdissection and laser pressure catapulting of selected cells or cell areas into the cap (see Note 6).

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4. Remove the cap from the cap holder and put it onto a 0.5-mL microfuge tube containing lysis buffer or pipette lysis buffer into the tube with the attached cap. 5. Mix and perform the lysis of the cells by inversion. 6. Centrifuge the sample at 16,000 g for 3 minutes. 7. If not continue with isolation procedure, store the samples in lysis buffer at −80 °C. 3.4.3

Looking Into the Cap to See the Catapulted Samples

The efficiency of catapulting is observed by looking into the collection device (e.g., cap) with the 5×, 10×, 40×, and 63× objectives. By using the software function “go to checkpoint” the lid is moved out of the light path and the cap can be lowered further toward the objectives. Normally most catapulted areas/cells can be found within the small inner ring of the caps. 3.4.4

Getting the Collected Cells from the Cap into the Tip of the Tube

After microdissection, the fluid from the cap is spun down in a bench centrifuge (3 minutes, 16,000 g) and samples can be stored for later use. For RNA extraction, add an appropriate lysis buffer to the tube, close the tube, and mix by inversion. Then spin down the lysate as described in the preceding. (For future RNA isolation/ analysis, the tube is placed on ice or stored at −80 °C.)

3.5

RNA Isolation

Working with RNA is more demanding than working with DNA, because of the chemical instability of the RNA and the ubiquitous presence of RNases. In a case that LCM-obtained cells are going to be used for the RNA extraction the following steps should be taken: 1. Designate a special area for RNA work only. 2. Clean benches with 100% ethanol or a special cleaning solution (e.g., RNase Zap). 3. Always wear gloves. After putting on gloves, do not touch surfaces or equipment to avoid reintroduction of RNase to decontaminated material. 4. Use sterile, disposable plastic ware. 5. Use filtered pipetter tips. 6. Glassware should be baked at 180 °C for 4 hours. (RNases can maintain activity even after prolonged boiling or autoclaving!) 7. Use reagents that are RNase-free. 8. All solutions should be made with DEPC-(diethylpyrocarbonate) treated H2O. 9. For best results use either fresh samples or samples that have been quickly frozen in liquid nitrogen or at −80 °C. (This procedure minimizes degradation of RNA by limiting the activity of endogenous RNases.) All required reagents should be kept on ice.


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10. To ensure RNase-free slides perform sterilization by autoclaving. Sections that are mounted on slides maybe treated to reduce the chance of contamination with exogenous RNases with RNase-ZAP followed by two separate washings in DEPCtreated water and drying at 37 °C up to 55 °C for 30 minutes up to 2 hours. 11. Store RNA by aliquoting in ethanol or RNA buffer at −80 °C and maintain the RNA at this temperature for long-term storage. Store prepared slides at −80 °C too. 12. RNA is not stable at elevated temperatures; therefore, avoid high temperatures (> 65 °C) because they affect the integrity of RNA. 13. RNA extraction is performed by using kits for the small-scale RNA purification. Kits are available from various companies (see Materials). 14. Two options are available to quantify RNA at a small scale: (1) Use a nanodrop to measure absorbance at A260 at the nanometer range. The assay can measure as low as 1 ng/µL and it is minimally affected by contamination likely to be found in small nucleic acid preps. (2) LabChips for analyzing RNA by the pico or nano Lab Chip Kit (Agilent) for quantitation on Agilent 2100 bioanalyzer (Agilent Technologies, Palo Alto, CA). The resulting electropherograms determine both integrity and concentration, of the RNA sample.

3.6

Reverse Transcription and RT-PCR

RNA reverse transcription (RT) to make cDNA includes the first strand cDNA synthesis with subsequent amplification by PCR using gene specific primers. It could be performed using several reverse transcriptases (e.g., Sensiscript III Reverse Transcriptase; iScript RNase H+ MMLV reverse transcriptase for sensitive detection using 1 pg to 1 µg of input total RNA; Reverse-iT MAX Blend, which includes a unique combination of RTases. The AMV component allows reverse transcription to be carried out at up to 57 °C when secondary structure is problematic, and the MMuLV provides maximum yields. RNase inhibitor is included in the blend to prevent degradation from RNAse A.) Both random hexamers and anchored oligo-dT can be used to prime the RT reactions. Random primers generate the most diverse pool of cDNA, whereas oligo-dT primers anneal only to mRNA poly-A tails and thus will be biased toward the 3′ end.

3.7

PCR Analysis

PCR analyses are performed based on protocols available at manufacturer web sites for the PCR kits. Two major technologies are used: (1) standard semi-quantitative PCR with the end-point product detection visualized on agarose gels for detection of PCR amplification is based on size determination of the product, or (2) real-time detection kinetic PCR is based on the detection and quantitation of a fluorescent reporter, which increases in direct proportion to the amount of PCR product in a reaction. By recording the amount of fluorescence emission at each cycle, it is possible

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to monitor the PCR reaction during the exponential phase, in which the first significant increase in the amount of PCR product correlates to the initial amount of target template. Specificity of amplification is assessed with a melting study. The presence of specific amplification products is confirmed by a single peak on the melting curve, which indicates an absence of non-specific amplification and primer–dimer formation.

3.8

Chromatin Immunoprecipitation from Tissues Samples Retrieved by LCM

1. Cryosections are fixed in 1% formaldehyde for 10 minutes. 2. Wash in PBS and air-dry for 30 minutes at room temperature. 3. Laser cut (microdissection) and laser pressure catapult (LPC) selected cells or cell areas. 4. Catapulting 2,500–3,000 cells is necessary for each sample. 5. Cups are inserted into the autoclaved 0.5-mL tubes and centrifugation is performed at 16,000 g for 3 minutes. 6. After the centrifugation the samples are processed for ChIP. 7. Resuspend cell pellet in 100 µL of SDS lysis buffer with protease inhibitors and incubate for 10 minutes on ice. 8. Sonicate for 3 minutes at max frequency using cup horn with ultrasonic water bath (e.g., MIXL-2020 Sonicator, MiSonix, Inc.) (see Note 7). 9. Centrifuge sample for 10 minutes at 16,000 g at 4 °C, and transfer the supernatant to a new 1.5 mL-microcentrifuge tube. Discard the pellet. 10. Dilute the sonicated cell supernatant 10-fold in ChIP Dilution Buffer with protease inhibitors (see Notes 8 and 9). 11. Add the antibody of interest for immunoprecipitation (the amount will vary per antibody) to the 1 mL of supernatant fraction and incubate overnight at 4 °C with rotation. 12. Add 30 µL of protein-A agarose/salmon sperm DNA (50% slurry). Incubate for 1 hour at 4 °C with rotation to collect the antibody/protein complex. 13. Pellet agarose by gentle centrifugation at 100–170 g at 4 °C for 1 minute. Carefully remove the supernatant that contains unbound non-specific DNA (see Note 10). 14. Wash the protein A agarose/antibody/protein complex for 5 minutes on a rotating platform with 1 mL of each of the buffers: (1) low salt wash buffer, (2) high salt wash buffer, (3) LiCl wash buffer, and (4) TE buffer. 15. Elute the protein–DNA complex from the antibody by adding 250 µL freshly prepared elution buffer to the pelleted protein A agarose–antibody–protein complex. Mix by vortex and incubate at room temperature for 15 minutes with rotation.


16. 17. 18. 19. 20.

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Spin down agarose, and carefully transfer the supernatant fraction to another tube and repeat elution. Combine eluates, the total volume is now 500 µL. Add 20 µL of 5 M NaCl to the combined eluates (500 µL) and reverse proteinDNA cross-links by heating at 65 °C for 4 hours (see Notes 11 and 12). Add 10 µL of 0.5 M EDTA, 20 µL of 1 M Tris-HCl, pH 6.5, and 2 µL of 10 mg/ mL proteinase K to the combined eluates and incubate for 1 hour at 45 °C. Recover DNA by phenol/chloroform extraction and ethanol precipitation (see Notes 13 and 14). Wash pellets with 70% ethanol and air dry. Resuspend pellets in TE buffer for PCR (Fig. 15.2 and Tables 15.1 and 15.2).

Fig. 15.2 ChIP analysis was performed with the antibody that specifically recognizes chromatin remodeling protein CReMM/CHD9 (12-13). The unbound fraction was used for the Re-ChIP with anti-polymerase-II (Pol-II) antibody. We analyzed the amplification of Cbfa1, Biglycan, and PPARgamma promoters using specific primers listed in Table 15.1. The PCR products were visualized in an agarose gel and the results are summarized in Table 15.2. In: Input fraction; B: Bone, P: Periost, 1-ChIP with anti-CReMM/CHD9 antibody; 2- reChIP with anti-Pol-II antibody.

Table 15.1 PCR primers and expected size for PCR product Gene Forward primer Reverse primer

Expected PCR product

Cbfa1

AAAGCCACAGTG GTAGGCAG

TGAAGCATTCAC ACAATCCAA

142 bp

Biglycan

CCATATCCTGC TTCTCCCA

CCAGGGACCAA GTGTAAGGA

223 bp

PPARγ

ATGGTTATTCCA TTTGGGGG

CTTAAGGCCTTT GCCCTTTT

203 bp

Table 15.2 Results of chip and re-chip of periost and endochondral bone Re-ChIP with Pol-II ChIP with DB P B P B −

+

+

+

Cbfa1

−

+

−

+

Biglcan

−

−

−

+

PPARγ

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Notes

1. Addition of proteinase K is an optional step. The solution should be freshly prepared. 2. The best results are obtained when freshly prepared cryo-sections are used. 3. If OCT or another tissue freezing medium was used, it is important to get rid of the medium on the slide before fixation; otherwise, it will interfere with laser efficiency. Removing of the medium is achieved by gentle washing the slide for about 1 minute in water. 4. H&E-staining of cryo-sections is also suitable for RNA preparation. Slides should be treated carefully with solutions prepared with RNase–free water. The whole procedure is shortened from standard 10 to 3 minutes to minimize the activity of RNases. Alternatively, if RNA extraction is performed from the unstained slides, every ninth slide in the serial section could be stained with H&E to provide better visual reference. 5. For single cells or very small areas, spinning down from oil may be difficult; therefore, aqueous solutions should be preferred. 6. When using membrane mounted samples the dissected membrane acts as a backbone. The selected area/cell can be catapulted with a single laser shot from a remaining “bridge” at the border. Morphological integrity is completely preserved with this procedure. Important: Laser settings have to be adjusted to cells or tissue in concern. 7. After this step samples can be frozen at −80 °C. 8. A portion of the diluted cell supernatant (1%) should be kept. This is the input sample of cross-linked histone-DNA complexes that will be reversed by adding 1 µL of 5 M NaCl and heating at 65 °C for 4 hours. 9. For a negative control, perform a no-antibody immunoprecipitation by incubating the supernatant fraction with 30 µL of protein A agarose/salmon sperm DNA (50% slurry). 10. Supernatant (unbound fraction) can be further used for the subsequent ChIP analysis of the protein–DNA complexes that can be precipitated from the unbound fraction, by adding an antibody of interest to the unbound fraction and repeating the incubation step overnight, i.e., Re-chip. Then add protein A agarose/ salmon sperm DNA for 1 hour and pellet agarose by gentle centrifugation at 100–170 g at 4 °C for 1 minute. Carefully remove the supernatant that contains unbound non-specific DNA. 11. At this step the sample can be stored at −20 °C and the procedure continued at a later date. 12. Include the input material and perform the protein–DNA cross-link reversal by adding 1 µL of 5 M NaCl per 20-µL sample and heating to 65 °C for 4 hours. 13. The addition of an inert carrier, such as 20 µg glycogen or yeast tRNA, provides for better visualization of the DNA pellet. 14. Alternatively, DNA purification can be performed by using commercially available kits for the genomic DNA purification.


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Acknowledgments This study was supported by funding from Ramot at Tel Aviv University and by the Chief Scientist of Ministry of Commerce and consortium “Bereshit” and the CellProm program from the EEC 6th framework to D.B.

References 1. Friedenstein, A. J., Piatetzky, S., II, Petrakova, K. V. (1966) Osteogenesis in transplants of bone marrow cells. J Embryol Exp Morphol 16, 381–390. 2. Benayahu, D. (2000) The hematopoietic microenvironment: the osteogenic compartment of bone marrow: cell biology and clinical application. Hematology 4, 427–435. 3. Jiang, Y., Jahagirdar, B. N., Reinhardt, R. L., et al. (2002) Pluripotency of mesenchymal stem cells derived from adult marrow. Nature 418, 41–49. 4. Pittenger, M. F., Mackay, A. M., Beck, S. C., et al. (1999) Multilineage potential of adult human mesenchymal stem cells. Science 284, 143–147. 5. Benayahu D, Akavia UD, Shur I. (2007). Differentiation of bone marrow stroma-derived mesenchymal cells. Current Medical Chemistry. 14 (2):173–9. 6. Syed, F., Khosla, S. (2005) Mechanisms of sex steroid effects on bone. Biochem Biophys Res Commun 328, 688–696. 7. Benayahu, D., Shur, I., Ben-Eliyahu, S. (2000) Hormonal changes affect the bone and bone marrow cells in a rat model. J Cell Biochem 79, 407–415. 8. Compston, J. (2001) Sex steroids and bone. Physiol Rev 81, 419–447. 9. Locklin, R., Williamson, M., Beresford, J., et al. (1995) In vitro effects of growth factors and dexamethasone on rat marrow stromal cells. Clin Orthop 313, 27–35. 10. Benayahu, D., Akavia, U. D., Socher, R., Shur, I. (2005) Gene expression in skeletal tissues: application of laser capture microdissection. J Microsc 220(Pt 1), 1–8. 11. Benayahu D, Socher R, Shur I. 2007. Laser Capture Microdissection of Bone Cells. Methods in Molecular Biology/Molecular Medicine Osteoporosis Editor: Jennifer Westendorf published by Humana Press, USA. 12. Shur, I., Benayahu, D. (2005) Characterization and functional analysis of CReMM, a novel chromodomain helicase DNA-binding protein. J Mol Biol 352, 646–655. 13. Shur, I., Socher, R, Benayahu, D. (2006) In vivo association of CReMM/CHD9 with promoters in osteogenic cells. J Cell Physiol 207, 374–378. 14. Shur, I., Salomon, R., Benayahu, D. (2006) Dynamic interactions of chromatin-related mesenchymal modulator, a chromodomain helicase-DNA-binding protein, with promoters in osteoprogenitors. Stem Cells 24, 1288–1293. 15. Marom, R., Shur, I., Hager, G. L., Benayahu, D. (2006) Expression and regulation of CReMM, a chromodomain helicase-DNA-binding (CHD), in marrow stroma derived osteoprogenitors. J Cell Physiol 207(3), 628–35.

Chapter 16

Quantitative Trait Loci Mapping Dong-Hai Xiong, Jian-Feng Liu, Yan-Fang Guo, Yan Guo, Tie-Lin Yang, Hui Jiang, Yuan Chen, Fang Yang, Robert R Recker, and Hong-Wen Deng

Abstract This chapter presents current methods for mapping quantitative trait loci (QTLs) in natural populations especially in humans. We discussed the experimental designs for QTL mapping, traditional methods adopted such as linkage mapping approaches and methods for linkage disequilibrium (LD) mapping. Multiple traits and interaction analysis are also outlined. The application of modern genomic approaches, which mainly exploit the microarray technology, into QTL mapping was detailed. The latter are very recent protocols and are less developed than linkage and association methods at present. The main focus of this chapter is technical issues although statistical issues are also covered to certain extent. Finally, we summarize the limitations of the current QTL approaches and discuss the solutions to certain problems. Keywords QTL, linkage, association, linkage disequilibrium, microarrays, eQTL, whole genome association.

1 1.1

Introduction Experimental Design for QTL Mapping

The classical quantitative genetics theory presumes that quantitative traits are controlled by an infinite number of genes, each with an infinitesimal effect, and are influenced by environmental factors. However, in addition to the polygenes with infinitesimal effects, there are other loci that may have major effects on quantitative traits (1) Such genes are called quantitative trait loci (QTL). A quantitative trait locus is defined as a region of the genome (i.e., segment of chromosome) that harbors one or more genes affecting a quantitative trait (2). The QTL concept is also used for trait variables with discrete distribution, for which continuous underlying processes can be assumed. These traits are often referred to as threshold traits. QTLs are believed to play an important role in the genetic mechanism underlying quantitative traits or complex discrete traits. In the past several decades, QTL

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mapping has been greatly developed and widely used in various fields, such as medical genetics, livestock production, plant breeding and population genetics of model organisms. Particularly, searching for QTLs by means of genome-wide linkage and/or association scans has become a major method to identify the genes underlying complex traits/diseases in humans. The aim of QTL mapping is to estimate positions and effects of the unobserved QTL through the observed quantitative traits and markers, and finally to estimate the variance components and the heritability of the quantitative traits. The designs of such mapping experiment involve choosing the appropriate crossing types, the parental strains, and phenotyping and genotyping strategies. Sound experimental designs help maximize the statistical power of QTL mapping. According to the distinct characteristics of natural populations and inbred lines, different experimental design strategies need to be taken for mapping QTLs underlying complex traits of interest. To detect the effect of QTLs in natural populations, it is necessary to generate linkage disequilibrium. In inbred lines, we need to employ the reasonable crossing designs to identify the effects of the chromosomal regions linked to the genetic markers.

1.2

Methods for QTL Mapping in Humans: Linkage Analysis

Related individuals are likely to share genetic loci controlling the variation of quantitative traits (i.e., blood pressure, body mass index, bone mass density, etc.). If a marker locus is linked to a QTL, the sharing pattern would be similar. Model-free linkage methods are commonly used in QTL mapping because there is no need to assume the underlying inheritance model of the traits of interest. A popular modelfree linkage method is to exploit the inverse relationship between the difference of trait values within sib-pairs and their marker identity-by-descent (IBD) scores (3).

1.3

Methods for QTL Fine Mapping in Humans: Association Mapping

1.3.1

What Is the Association?

Genetic linkage studies based on pedigree data have limited resolution because of the relatively small number of meiosis events within the general pedigrees. Association mapping (also called linkage disequilibrium mapping) can be used to further narrow down the linkage region. The classical definition of linkage disequilibrium (LD) refers to the non-random association of alleles at tightly linked loci. Consider two markers that were on the same chromosome. We designate the corresponding alleles and frequencies as A/a (pa.pa), B/b (pb, pb). Four haplotypes consisting of these two markers are possible: AB, Ab, aB, and ab. If the frequencies of alleles A, a, B, and b in the

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population were all 0.5 and these two loci were in linkage equilibrium, then we would expect the frequencies of each of the four haplotypes in the population to be 0.25. However, the significant deviation of the haplotype frequencies from 0.25 suggested the LD between the two loci. 1.3.2

Measures of LD

One measure of LD is D, calculated as (4): D = p(AB) × p(ab) – p(Ab) × p(aB) where p(AB) is the frequency of the AB haplotype in the population, and likewise for the other haplotypes. In practice, it is the standardized D—namely D′—that is widely used: D' = D/Dmax where Dmax is the maximum value of |p(AB) – p(A) × p(B)|. D′ varies between 0 (no LD) and ±1 (complete association) and is less dependent on the allele frequencies than D. As a rule of thumb, D′>0.33 is often taken as the threshold level of LD above which associations are considered as apparent. Hill et al. (5) proposed another statistic, r2, which is: r2 =

D2 PA Pa PB Pb

where pA is the frequency of the A allele in the population, and likewise for the other alleles in the population. The r2 parameter can also be calculated for two loci with multiple (more than two) alleles as follows, which is usually necessary for microsatellite markers: n m

r2 = ∑ ∑ i

j

Dij2 Pi Pj

where locus 1 has n alleles, locus 2 has m alleles, pi is the frequency of allele i of locus one, pj is the frequency of allele j of locus two, and Dij = pij – pi × pj, with pij being the frequency of haplotype ij.

1.3.3

Tagging SNP and Haplotype

Originally, haplotype was defined as “a set of alleles of closely linked loci on a chromosome that tend to be inherited together.” In the single nucleotide polymorphism (SNP) era, the haplotype concept is based on SNPs and closely related to the LD concept. There is remarkable genomic variability in LD across the human genome

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with a series of high LD regions separated by short discrete segments of very low LD. Those high LD regions usually exhibit limited haplotype diversity, so that a small number of haplotypes account for most of the chromosomes in such genomic regions (haplotype blocks) in a population. Thus far, there are a number of ways to define haplotype blocks (6, 7). For example, Gabriel et al. (7) adopted the standardized gametic disequilibrium coefficient D′ to identify the high LD regions. Within these regions, allelic dependence made it unnecessary to study the redundant SNPs. Only the representative SNPs such as tag SNPs need to be genotyped and analyzed to make the association studies much more cost-effective while maintaining the statistical power at the same time. In association studies, both SNP markers and haplotypes constructed by individual SNPs can be used. Whether a specific method is more effective depends on whether the true causal allele is in higher LD and has more closely matched allele frequency with the marker set that method utilizes.

1.4 Multiple Traits and Interaction Analysis Genetic multivariate analyses in which the correlations between the phenotypes are explicitly modeled can provide greater statistical power to identify quantitative trait loci (QTLs) whose effects are too small to be detected by univariate analysis of the individual traits (8). The multivariate framework also serves to formally integrate results from separate univariate analysis. Joint genetic linkage analysis of multiple correlated traits has been shown to improve the power to detect, localize, and estimate the effect of possible QTLs or disease susceptibility loci (9, 10). Pleiotropic effects or close-linkage loci that each influence different traits can also be identified by joint genetic analysis (11). Epistasis, defined as the interaction between different genes, has become a hot topic in complex disease genetics in recent years. For complex traits, such as diabetes, asthma, and osteoporosis, the search for susceptibility loci has, to date, been less successful than for simple mendelian disorders. Part of this complexity of mapping complex disease genes can be attributed to epistasis or gene–gene interactions (12). It is hoped that the identification of the mode of gene interaction will facilitate understanding of the pathological mechanisms involved in complex diseases.

1.5

Microarrays and QTLs

1.5.1

Expression Quantitative Trait Loci Analyses

In the year of 2001, Jansen (13) firstly proposed the concept of “genetical genomics,” aimed at merging the genetic map and genomics to unravel the function and interrelation of gene products and gene action from expression profiling, such as gene expression, protein, enzymatic, and metabolite levels. The experiments in

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yeast, plant, mouse, and human exemplified the merit of this kind of combination (14, 15). For example, through analyzing the co-segregation of eQTL with clinical QTL, Yaguchi (16) identified four candidate genes from 106 genes in a QTL spanning 10 Mb. Schadt (15) found two independent genomic regions for fat-pad-mass trait according to a hint from two separate gene expression clusters. 1.5.2

Whole Genome Association

With the near completion of the International HapMap Project and the rapid improvements in microarray technology, there comes the era of whole genome association (WGA), in which a dense set of SNPs across the genome is genotyped to survey the most common genetic variation for a role in complex diseases or traits. Because no assumptions are made about the genomic location of the causal variants, this approach could exploit the strengths of association studies without having to guess the identity of the causal genes. Therefore, the WGA approach represents an unbiased yet fairly comprehensive option that can be attempted even in the absence of convincing evidence regarding the function or location of the causal genes (17). To date it has been used to find causal genes and/or polymorphisms underlying complex diseases or traits such as rheumatoid arthritis (RA), lung cancer, alcoholism, and obesity (18–23). It can be imagined that the WGA approach will also be applied to dissect the causative genetic variants influencing other complex diseases in the near future.

2 2.1

Materials Linkage Study

The necessary material for mapping QTLs is a linkage map of polymorphic marker loci that adequately covers the whole genome. There are several criteria for the marker loci: (1) highly polymorphic, so that individuals are likely to carry different alleles at each locus; (2) abundant to cover the genome comprehensively; (3) neutral, both with respect to the quantitative trait of interest and to reproductive fitness; and (4) co-dominant, that all possible genotypes at a marker locus can be identified phenotypically. The final criterion is less stringent, as dominant/recessive markers can also be used in some designs. The most frequently used marker for QTL mapping is microsatellite marker.

2.2

Association Study

In association studies, single nucleotide polymorphisms (SNPs) are the most widely used markers. SNPs can be classified as noncoding or coding SNPs. Non-coding SNPs are generally located in introns, non-transcribed or untranslated regions, and

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the intergenic regions. Coding SNPs may cause non-synonymous or synonymous mutations, depending on whether there is any associated amino acid change. Various systems have been developed to meet the challenge of genotyping a large number of SNPs for disease association studies (24). The features of SNP genotyping methods are: (1) endonuclease cleavage (e.g., restriction site analysis and invader assay), (2) primer extension (e.g., SNPstream, multiplex mini-sequencing, pyrosequencing, MassArray, fluorescence polarization, SnaPShot, etc.), (3) allele-specific PCR (e.g., FRET primers, AlphaScreen, intercalating dye, etc.), (4) oligonucleotide ligation (e.g., OLA (oligonucleotide ligation assay), microarray ligation, ligase chain reaction, padlock probes, etc.), and (5) hybridization methods (e.g., Genechip microarrays, TaqMan, Molecular Beacons, etc.). For genome-wide association studies requiring hundreds of thousands of SNP markers, highly multiplexed and high-density microarray–based SNP genotyping systems (e.g., GeneChip 500 K, BeadArray, SNPstream, etc.) have been rapidly developed, offering the potential of genome-wide SNP mapping of genes involved in complex diseases in the near future (25).

2.3

eQTL Mapping

Generally, there are two types of microarray technologies: in situ synthesized oligonucleotide microarray (developed by companies such as Affymetrix, Agilent, and Febit AG) and spotted array based on the physical deposition of biomolecules (typically oligonucleotides or cDNAs) in glass or nylon membrane matrices. In Affymetrix, three categories of expression analysis arrays (for genes, transcripts, and genomes, respectively) are available at http://www.affymetrix.com/products/ arrays/index.affx. 2.3.1

Instruments

1. Agilent 2100 Bioanalyzer (Agilent, Palo Alto, CA). 2. Hybridization Oven 640 (Affymetrix, Santa Clara, CA). 3. High-solution Laser scanner: GeneChip Scanner 3000 (Affymetrix), Axon 4000a scanner (Axon Instruments). 4. GeneChip Fluidics Station 450 (Affymetrix). 5. GeneAmp PCR System 9700 and 7900HT Fast Real-Time PCR System (Applied Biosystems, Foster, CA). 2.3.2

Total RNA/Poly-A mRNA Isolation from Mammalian Cells

1. RNeasy Mini Kit (Qiagen, Valencia, CA) for total RNA isolation, or Oligotex Direct mRNA kit (Qiagen) for isolating mRNA directly from mammalian cells, or Oligotex mRNA kit (Qiagen) for separating mRNA from total RNA.

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QIAshredder for simple and rapid homogenization of cell lysates (Qiagen). RNase inhibitor, 20 U/µL (Ambion, Austin, TX). RNase-free DNase set (Qiagen). DEPC-treated water (Ambion). RNase zap (Ambion).

2.3.3

Quantification of RNA/cRNA

1. RNA 6000 ladder (Ambion). 2. RNA 6000 Nano kit (Agilent, Palo Alto, CA). 2.3.4

cDNA Synthesis

1. Poly-A RNA control kit (Affymetrix). 2. Sample cleanup module (Affymetrix). 3. One-cycle cDNA synthesis kit for 1–15 µg of total RNA or 0.2–2 µg of mRNA, or two-cycle cDNA synthesis kit for 10–100 ng of total RNA (Affymetrix, Santa Clara, CA). 4. Cy-labeled dNTPs for spotted array (GE Healthcare). 5. Biotin-labeled cRNA synthesis. 6. GeneChip expression 3′-amplification reagent for IVT labeling (Affymetrix). 7. Sample cleanup module (Affymetrix). 2.3.5

Hybridization, Wash, and Stain

1. Hybridization control kit (Affymetrix). 2. GeneChip hybridization, wash, and stain kit (Affymetrix or home made). 3. Wash buffer A and B (Affymetrix or home made). 2.3.6 1. 2. 3. 4.

RT-PCR

TaqMan gold RT-PCR Kit (Applied Biosystems, Foster, CA). TaqMan gene expression assays (Applied Biosystems). Housekeeping gene control (Applied Biosystems). AmpliTaq Gold PCR Master Mix (Applied Biosystems).

2.4


In whole-genome genotyping, currently there are two most representative genotyping platforms: Illumina Sentrix HumanHap550 Genotyping Beadchip, and Affymetrix

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GeneChip500 k. Both allow genotyping of >500,000 SNPs across the human genome. According to the HapMap Project, the characterization of 300,000–500,000 tag SNPs is sufficient to provide good genomic coverage of all possible common polymorphisms in various ethnic groups. The Sentrix HumanHap550 Genotyping BeadChip (Illumine, San Diego, CA) contains over 555 K SNP loci with high-density tagSNP content that provides comprehensive genomic coverage across multiple populations. The GeneChip Human Mapping 500 K Array Set (Affymetrix) also enables highly powered whole-genome association studies. It is comprised of a set of two arrays that enable genotyping of >500,000 SNPs. One array uses the Nsp I restriction enzyme (~262,000 SNPs), whereas the second uses Sty I (~238,000 SNPs). Total genomic DNA with the concentration of 50 ng/µL as determined by PicoGreen is required for this 500 K array set. We will elaborate the materials and major steps of WGA analysis using GeneChip human mapping 500 K array set.

2.4.1

Instruments for Genomic DNA Purification and Quantification

1. Sorvall RT7 plus refrigerated centrifuge (Sorvall, Guelph, ON). 2. VWR Multi-Tube Vortexer (VWR, West Chester, PA). 3. SpectraMax Plus384 High-Throughput Microplate Spectrophotometer (Molecular Devices, Palo Alto, CA).

2.4.2

Instruments and Software for WGA Using Affymetrix GeneChip500 k Platform

1. Dual 96-Well GeneAmp PCR System 9700 (Applied Biosystems). 2. SpectraMax Plus384 High-Throughput Microplate Spectrophotometer (Molecular Devices). 3. Maxi-Mix* II Mixer (VWR). 4. QIAvac Multiwell Unit (Qiagen). 5. GeneChip Hybridization Oven 640 (Affymetrix). 6. GeneChip Operating Software (GCOS) (Affymetrix). 7. GeneChip Fluidics Station 450 (Affymetrix). 8. GeneChip Scanner 3000 7 G (Affymetrix). 9. GeneChip Genotyping Analysis Software (GTYPE) (Affymetrix).

2.4.3 DNA Purification and Quantification 1. Puregene DNA purification kit (Gentra) containing: a. RBC lysis solution. Store at room temperature. b. Cell lysis solution. Store at room temperature.

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c. Protein precipitation solution. Store at room temperature. d. RNase A solution. Store at room temperature. e. DNA hydration solution. Store at room temperature. 2. Quant-iT PicoGreen dsDNA Assay Kit (Invitrogen) containing: a. Quant-iT PicoGreen dsDNA reagent is dissolved at dimethylsulfoxide (DMSO) and stored at −20 °C. b. 20× TE: 200 mM Tris-HCl, 20 mM EDTA, PH7.5. Store at room temperature. c. Lambda DNA standard with the concentration of 100 µg/mL. Store at 2–6 °C.

2.4.4

WGA Using Affymetrix GeneChip500 k Platform

1. Affymetrix GeneChip Mapping 250 K Nsp and/or Sty Assay Kit (Affymetrix, Santa Clara, CA), containing: a. b. c. d. e. f. g. h. i.

Adaptor Nsp or Sty, 50 µM. DNA labeling reagent, 30 mM. Fragmentation reagent. Oligo control reagent, 0100. PCR primer 002, 100 µM. Terminal deoxynucleotidyl transferase. 5× terminal deoxynucleotidyl transferase buffer. Reference genomic DNA 103 (use as a positive control). 10× fragmentation buffer.

2. Sty I (10,000 U/mL) and NE buffer 3 (New England Biolabs). 3. Nsp I (10,000 U/mL) and NE buffer 2 (New England Biolabs). 4. BSA (bovine serum albumin). 5. T4 DNA ligase. 6. T4 DNA ligase buffer. 7. G-C melt (5 M) (Clontech). 8. Molecular grade biology water. 9. dNTP (2.5 mM each). 10. Clontech TITANIUM Taq DNA polymerase (50×) (Clontech): a. 50× Clontech TITANIUM Taq DNA polymerase. b. 10× Clontech TITANIUM Taq PCR buffer. 11. 2% TBE Gel:BMA reliant precast (2% SeaKem Gold) (Cambrex, East Rutherford, NJ). 12. Gel loading solution (Sigma). 13. All purpose Hi-Lo DNA Marker (Bionexus). 14. 5 M TMACL (Tetramethyl Ammonium Chloride) (Sigma). 15. 10% Tween-20, diluted to 3% in molecular biology grade water. 16. MES hydrate SigmaUltra (Sigma). 17. MES sodium salt.

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18. 19. 20. 21. 22. 23. 24. 25. 26.

DMSO. 0.5 M EDTA. Denhardt’s solution (Sigma). HSDNA (herring sperm DNA). Human Cot-1 DNA. 20× SSPE. SAPE (streptavidin, R-phycoerythrin conjugate) (Invitrogen). Biotinylated anti-streptavidin antibody (Vector Laboratories). Distilled water.

3

Methods

3.1

Experimental Design for QTL Mapping

3.1.1

Experimental Design for Model Organism Population

QTL mapping results in model organisms such as inbred mouse lines can be applied to the mapping of QTL or disease susceptibility loci in humans through comparative genomics approaches (26). Two experimental designs are often used for QTL mapping in inbred lines, i.e., back cross design and F2 design. Both of these designs usually start with a cross between two inbred lines from the F1 population. An alternative to the cross between inbred lines is to study natural outbred populations such as the human populations.

3.1.1.1

Back Cross

Starting with two completely inbred parental lines, P1 and P2, the back cross (BC) design examines marker/trait associations in the progeny formed by back crossing the F1 to one of the parental lines. The BC design is depicted (Fig. 16.1). Two parental lines have different marker and QTL genotypes. They are mated to form an F1. There is a general assumption with BC design that the two parental lines are homozygous for alternate alleles of both loci, so all F1 individuals have the same heterozygous genotypes in both marker locus and QTL. The F1 is then mated to one of the parental lines. The BC progeny can be classified into two groups according to the recombination status (Fig. 16.1). There are only two marker genotype groups for the BC design, i.e., individuals with marker genotype M 2M2 and those with M1M 2 (Fig. 16.1). The difference between the means of these two progeny marker genotype groups can be calculated to represent genotypic effect. The significance of such genotypic effect can be tested by traditional methods such as analysis of variance (ANOVA),

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Fig. 16.1 The back cross design, r denotes the recombination rate between mark locus and QTL.

or t-test. Under the null hypothesis that there is no QTL linked to the marker, the ratio of the marker mean-squares to the residual mean-squares follows a central F-distribution when we perform ANOVA. The significant deviation of such statistic from the central F-distribution is the evidence that a QTL is linked to the marker. The t statistic can be derived as the ratio of the difference between the means of two genotype groups to the standard error of this difference. Such t-statistic will have a central t-distribution under the null hypothesis. Both of the preceding two tests can be easily conducted by most statistical packages, such as SAS, SPSS, etc. The contrast between marker group means in a back cross gives an estimate of the additive QTL effect (a), which is unbiased only in absence of dominance (d = 0), and recessive or partly recessive QTL may not be detected. This problem can be overcome by backcrossing to both parental lines (1). Furthermore, the back cross design is usually less powerful than the F2 design because it only detects differences between heterozygotes and one of the homozygotes when performing one back cross between F1 and the parental line.

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F2 Design

F2 design is depicted in Fig. 16.2. Firstly as in BC design, two homozygous inbred parents are mated to form a heterozygous F1. Then the F1 individuals are mated with each other to generate F2 progeny that include three non-recombinants and seven recombinants. The F2 design detects marker/trait associations in the progeny from an intercross mating of the F1 individuals, as three genotypes are generated at each marker locus, the estimation of the degree of dominance associated with the detected QTL is possible in the F2 design.

Fig. 16.2 The F2 design.

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Similarly, significance of a segregating QTL can be detected by ANOVA, including all three marker genotypes. Additionally, comparisons between different marker genotypes can be tested by t-test. The main difference of interest is that between the means of the two distinct homozygotes. Note that, although it is possible to test for significance of additive and dominance effects, the test statistic is always the function of recombination rate, additive effect and/or dominance effect, i.e., QTL effect and recombination rate are always confounded. This concern is also present in BC design. Although F2 and back cross are the most widely used designs, other line crosses can offer further advantages. For example, the F1 can be used to create recombinant inbred lines (RIL) and doubled-haploid lines (DHL), which allow marker–trait associations to be scored in a completely homozygous background and across multiple environments (27).

3.1.2

Experiment Design for Human Population

Designs using inbred lines are frequently used in laboratory animals (e.g., mice) and plants. However, in human populations, inbred lines are unavailable. Instead, experimental designs should be based on the analysis of families within existing populations. Two main types of experimental designs are commonly used for QTL mapping studies in human populations, including: (1) sib-pair design for analysis of many small nuclear full-sib families based on a single generation; and (2) general pedigree design for analysis of large families with complex pedigree structures based on multiple generations.

3.1.2.1

Sib-pair

Sib-pair design in human outbred population was first proposed by Haseman and Elston in 1972 (28). This method is most suitable for human populations, which include many small full-sib families. The main idea of this method is illustrated in Fig. 16.3. We consider the simplest case to introduce this design, i.e., assuming each parent has different alleles in both the marker locus and the linked QTL locus denoted by M1, M2, M3, M4, and Q1, Q2, Q3, Q4, respectively. For simplicity, only three kinds of nonrecombinant sib-pairs are shown in Fig. 16.3. In sib-pair 1, both individuals’ marker alleles are identical by descent (IBD). In sib-pair 2, only M 1 alleles carried by two individuals are paternal IBD. In sib-pair 3, none of the marker alleles are IBD. According to the assumption, sibpairs with more marker loci IBD should also have higher probability of IBD in QTL and hence are expected to be more similar than sib-pairs sharing no IBD marker alleles.

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Fig. 16.3 The full sib-pair design.

Because QTL genotypes cannot be directly observed, in theory, we can detect the linked QTL compare means of different marker genotypes as BC or F2 design. However, these comparisons are not available because the differences between different marker genotypes maybe show negative or positive in different families. In order to resolve this issue, Haseman and Elston (28) proposed a regression model based on squared differences. They defined: x1 j = m + g1 j + e1 j x2 j = m + g2 j + e2 j where x1j and x2j are phenotypes for sibs 1 and 2 of family j, m is the general mean, gij and eij are the QTL and residual effects respectively for sib i.

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Then let Yj = (x1j – X2j)2 = (g1j + e1j – g2j – e2j)2. If the two sibs share IBD of both QTL alleles, then g1j = g2j and so Yj = (e1j – e2j)2, which will be less than Yj for other sibs sharing no allele IBD or sharing only one allele IBD of QTL. According to this relationship between Yj’s, presence of a linked QTL can be detected by the following regression: Y j = a + bp j + e j where pj is the proportion of alleles IBD at the marker of interest between sib-pair, a is the y-intercept and b is the regression coefficient which will have a negative value. Even with incomplete linkage between the marker and the QTL, b will still be negative. Under null hypothesis, b should not be significantly different from zero. Through the squared sib-pair trait difference regressed on the proportion of marker alleles, the sib-pairs are estimated to share identity by descent: A significantly negative regression coefficient suggests linkage between the marker and a QTL. We need to note that, in this regression model, when both parents have complete marker informative, pj can be inferred unequivocally. However, it is impossible for all families in which both parents are fully informative in all marker loci. In the case that parents cannot provide complete marker information, pj can be estimate as fj2 + 0.5fj1, where fj2 and fj1 are the probabilities that sib-pair j has one or two marker alleles IBD, respectively.

3.1.2.2

General Pedigree

In general pedigreed design, mixed model based on variance components estimation is the popular method. Under this model, QTL mapping by variance components in general pedigrees has been developed independently by both the human genetics (29) and animal breeding (30) communities. Consider the “polygenic + QTL” model used to estimate additive genetic variance and QTL variance in a population with general pedigree. Using matrix algebra form, the animal model can be written: y = Xβ + Za + Zq + e where y is a vector of phenotypes of all pedigreed individuals, β is a vector of fixed effects, X is a design matrix relating the appropriate fixed effects to each individual, a is a vector of random effects (polygenic additive effects), q is a vector of additive QTL effects, Z is an incidence matrix relating the appropriate random effects to each individual, and e is a vector of residual errors. We depict the general pedigree design in Fig. 16.4. For any pair of individuals in the pedigree, the genetic covariance between them is a function of 2Θij and Rij. where Qij is the coefficient of co-ancestry, the probability that an allele randomly drawn from individual i is IBD with an allele randomly drawn from individual j.

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Fig. 16.4 QTL mapping in general pedigrees. The pedigree contains 10 individuals denoted by number 1, 2, …, 10. All of individuals are typed at a marker locus with four alleles A, B, C, and D. 2Θij (twice the coefficient of coancestry) and Rij (the IBD coefficient at the marker) between each pair of individuals is shown.

Note that the coefficient of co-ancestry is obtained from the pedigree structure of the individuals concerned instead of other information, such as marker genotypes. Rij is the proportion of alleles that two individuals i and j actually share IBD at a chromosomal location. Two alleles are IBD if we can trace them back to a single copy in a common ancestor. To obtain an REML solution of this model, marker data are used to infer Rij. Using all markers on a chromosome to estimate Rij at each location is known as multipoint mapping. It should be pointed out that Rij is an estimate rather than a probability, which varies at each test location.

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Through this model, we can obtain estimates of the additive genetic variance, the variance attributable to a QTL at the location considered and the likelihood of of the REML solution (l1). A likelihood ratio test statistic (LRT) can be used to perform chi-square test as follows: LRT = – 2ln(l0 –l1) Under the null hypothesis of no QTL at the test location the test statistic follows a χ2 distribution (29, 30). Under the different null hypothesis of no QTL anywhere on the chromosome, the test statistic appears to approximate to a χ2 distribution with 1 degree of freedom (30). Using general pedigree design, mapping QTL utilizes more pedigree information than sib-pair designs, and hence appears more powerful to detect QTL (30, 31).

3.2

QTL Mapping in Humans: Linkage Analysis

3.2.1

Regression Analysis

Regression tries to predict a variable (Y) in terms of another variable (X) via a functional relationship Y = f(X). The functional relationship f could be linear (Y = a + bX), quadratic (Y = a + bX + cX2), logarithmic (Y = a log X), or more complex relationships. The simplest one is linear regression. The statistical problem of linear regression is to determine the “best” values of a and b that minimize the deviations from the linear relationship. This is done by “the least squares errors” method. Since some of the errors are positive and some are negative, minimizing the sum of errors (Σei) is not optimal. It is better to minimize the sum of squared errors (Σei2), that is (Σ(Yi – a – bXi)2), to minimize with respect to a, b. The best values: b0 = Cov (X,Y ) / Var (X ), a0 = Mean(Y ) – b0 Mean(X ) Haseman and Elston (1972) developed a regression approach for detecting linkage based on the squared difference in quantitative trait values of sib-pairs (Y) and their ∧ estimated marker IBD scores (p m ). The basis of the regression is the equation: ∧

∧

E (Y⎥ p m ) = a + b p m ,

(see Note 1)

where there is no dominance in the trait and b = −2p(1 − p)a2 (1 − 2q)2; p being the allele frequency of A1, a the conditional expectation of the trait given genotype A1A1, and q the recombination fraction between the QTL and the marker locus. Function of q, a test for no linkage (i.e., q = 0.5) is equivalent to testing b = 0 in Equation (1). The test can be performed via the usual t statistic based on the least squares regression estimate of b.

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Based on the traditional Haseman-Elston method, many extensions have been made so that it can be used for linkage analysis in more scenarios. First, Amos and Elston extended the above regression procedure to other relative pairs (32). Then, Fulker and Cardon applied the Haseman-Elston regression equation to interval mapping (33). Tiwari and Elston extended the traditional Haseman-Elston procedure to the case of two unlinked QTLs which might interact epistatically (34). Recently, Elston et al. (3) suggested that the mean-corrected cross-product of the sib-pair trait values carry more linkage information than the squared sib-pair trait difference.

3.2.2

Variance Components Analysis

Another popular statistical approach for QTL mapping is to dissect the genetic variation within the quantitative trait. Although the methods assume specific probability distributions for trait values, the advantage of using these methods is that larger sibships or entire pedigrees can be simultaneously analyzed (35). The general variance components model is given by: Y=m+g+G+e where m is the overall mean of the quantitative trait, g is a random effect due to a major gene with additive variance sa2 and dominance variance sd2, G is a random polygenic effect with variance sG2 and e is the non-shared environmental effect (or random error) with variance se2. The trait values of individuals in a pedigree are usually assumed to be distributed as multivariate normal with dispersion matrix V, where the variance of the trait value of each individual is sa2 + sg2 + sG2 + se2 and the covariance between the trait values of two individuals is given by fσa2 + ∆sd2 + fsG2, where f is the coefficient of relationship between the two individuals and ∆ is the probability that the two individuals share both their alleles IBD at the major locus. Conditioning on IBD score (p) at a marker locus, the preceding covariance is given by: f (q , p )s a2 + g(q , ∆ )s d2 + fs G2 , where q is the recombination fraction between the QTL and the marker locus. The log-likelihood of the data is given by: c−

1 1 log | V | − ∑ (Y − m1)′V −1 (Y − m1), ∑ 2 P 2 P

where c is a constant, Y and m1 are, respectively, the vector of trait values and that of the means within a pedigree and the summation is over independent pedigrees. The variance components methods use the maximum likelihood method to estimate the parameters. The test for linkage is equivalent to testing sa2 = 0 versus sa2 > 0.

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The usual likelihood ratio test statistic is asymptotically distributed as a 1– : 1– mixture 2 2 of a χ12 variable and a point mass at zero. Based on the basic framework, Almasy and Blangero expanded the variance components model to calculate multipoint IBD probability using pedigrees of arbitrary size. They also extended the model to incorporate multiple QTLs. In addition, the model can also incorporate other environmental co-variates, gene by environment interaction terms, epistatic effects and so on.

3.3

QTL Fine Mapping in Humans: Association Mapping

3.3.1

Population-Based Association Mapping

Genetic association studies aim to correlate differences in between-group disease frequencies (or trait values for continuous variables) with differences in allele frequencies at candidate loci. The association study adopting a random sample chosen from a population is usually called population-based association study. The populationbase design is often more powerful and feasible to be conducted than the family-based design. However, this approach is prone to the confounding effect of population stratification that refers to differences in allele frequencies between cases and controls due to systematic differences in ancestry rather than association of genes with disease (see Note 2). Some solutions, such as genomic control (GC) and structured association (SA) have been developed and widely used to correct for the stratification confounder (36–38).

3.3.2

Transmission-Disequilibrium Test

The most popular method for family-based design is the transmission disequilibrium test (TDT) (39). TDT is carried out with data on transmission of marker alleles from heterozygous parents for that marker to affected offspring (case). In TDT, cases’ genotypes are compared to those of their parents to explore whether a specific allele, or marker, at a locus of interest appears to be transmitted in excess of what is expected on the basis of mendelian inheritance. Such apparent excess transmission indicates that cases are being selected for that allele, thereby providing evidence that this allele is a risk factor for disease. The test is unaffected by population stratification. The general procedure to conduct such study is as follows: 1. Affected probands are ascertained. 2. The probands and their parents are typed for the marker. 3. Those parents who are heterozygous for marker M1 are selected. They may or may not be affected. Let a be the number of times a heterozygous parent transmits M1 to the affected offspring, and b be the number of times the other allele is transmitted.

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The TDT statistic is (a − b)2/(a + b). It follows a χ2 distribution with 1 degree of freedom, provided the numbers are reasonably large. Whatever association test is used, the problem of multiple testing must be addressed. A full Bonferroni correction (dividing the nominal p value by N, the total number of individual tests, to obtain an experiment-wide threshold) is overconservative for large values of N. Thus other methods, such as false discovery rate (FDR) (40) and permutation test (41), were proposed to solve this problem. FDR method controls the expected proportion of incorrectly rejected type I errors in a list of rejected hypotheses. It is a less conservative comparison procedure with optimal power at a cost of increasing the likelihood of obtaining type I errors. The permutation test uses a Monte Carlo procedure to provide a global assessment of significance. It is more powerful than the Bonferroni correction procedure when alleles at linked markers are associated.

3.4

Multiple Traits and Interaction Analysis

3.4.1

Principal Component Analysis

It has been suggested that evaluating multiple phenotypes jointly is more powerful than using of each interrelated phenotypes separately (42). One of the most commonly used methods is principal component analysis (PCA), which can be performed by classic statistical package SPSS or SAS. PCA uses the information from the entire correlation coefficient matrix of some phenotypes to produce a smaller number of hypothetical factors (components) that help explain correlations among the original variables. PCA includes two stages: (1) factor extraction using PCA, and (2) rotation of the principal components using varimax option. Factors are defined by extracting the eigenvalues (which represent the variance explained by each of the principal components) >1.0. The results of PCA generate a new uncorrelated combination of the original phenotypes with a reduction of the number of variables. The principal component scores (the correlation coefficients between the variables and the factors) are extracted as “new” phenotypes representing the original phenotypes for other statistical genetic analysis.

3.4.2

Multivariate Linkage Analysis

In univariate linkage analyses, a variance component approach is widely used (29, 35), which is based on specifying expected phenotypic covariances between relatives as a function of the IBD relationships at one or more genetic loci. Briefly, the covariance matrix for a pedigree is given by: Ω = Πσ 2m + 2Φσ 2g + Iσ 2e

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where smij2 is the additive genetic variance caused by the major locus, Π is a matrix whose elements (πmij) provide the probability that individuals i and j share IBD at a QTL linked to a genetic marker locus; P is a function of the estimated IBD matrix for the genetic marker itself and a matrix of the correlations between the proportions of genes IBD at the marker and at the QTL. sg2 is the genetic variance due to residual additive genetic factors, F is the kinship matrix for the entire pedigree, Se2 is the variance due to individual-specific environmental effects, and I is an identity matrix. By assuming underlying multivariate normality of the trait within pedigrees, the likelihood of any pedigree can be easily written and numerical procedures can be used to estimate the variance component parameters. The basic variance component model can extend to a multivariate framework (see Note 3). In the multivariate linkage model, the phenotype covariance is further decomposed to include the genetic correlation between traits caused by additive genetic effects and the shared effects of the QTLs, such that the covariation between two individuals for two traits is given by: Ω=

Ω 11Ω 12 Ω 12 Ω22

where Ω is a covariance matrix of 2×2 covariance matrices, where the elements are defined by: Ωab = Πmsmasmb + 2Φrgs gasgb + I reseaseb where a and b are 1 or 2, and rg and re are the additive genetic and environmental correlations between the two traits, respectively. The genetic correlation estimates the proportion of genes shared in common between the two traits. If a = b, then rg is 1, and the covariance of a pair of relatives simplifies to Equation 1. In this model, the marker-specific genetic correlation (rm) is assumed to be 1. This approach can be conducted in program SOLAR (43).

3.4.3

Epistasis and Gene–Environment Interaction Analysis

Methods for the detection of epistasis vary according to whether one is performing linkage or association analysis, and dealing with a quantitative or a qualitative trait (see Note 4). Here, we focus on quantitative traits, as most surrogate phenotypes for osteoporosis are quantitative. For genetic linkage analysis, one popular method is the variance component model. Test for epistasis uses an extension of the variance component model (43), which can be written as: Ω = Πm1σ 2 m1 + Π m 2 σ 2 m 2 + Π m1 m 2 σ 2 m1 m 2 +2Φσ 2g + Ισ e2

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2 2 where Sm1 and Sm2 are the additive genetic variance due to the observed marker m1 and m2, Pm1 and Pm2 are the matrixes whose elements provide the probability that any two individuals i and j share IBD for the observed genetic marker m1 and m2, 2 is epistatic component, and Pm1m2 is the element by element matrix multiplicaSm1m2 tion of Pm1 and Pm2 serves as the coefficient for the epistatic component. This model can be implemented in program SOLAR (43) by comparing it with models that do not contain these components using maximum-likelihood methods. Another popular method of linkage analysis is the Haseman-Elston method (28) and its extensions (3), which is based on a regression model. The squared difference or trait similarity between the members of a pair of relatives in a regression equation is a function of the underlying IBD sharing probabilities. To estimate epistasis between loci, it is required to include IBD sharing probabilities at different loci as predictors in the regression framework. In the quantitative genetics literature, the general epistatic model is a standard multiple linear (as opposed to logistic) regression equation (44):

y = m + a1x1 + d1z1 + a2x2 + d2z2 + iaax1x2 + iad x1z2 + idaz1x2 + idd z1z2 where y is a quantitative phenotype of interest, xi and zi are dummy variables related to the underlying genotype at locus i. The coefficients µ, a1, d1, a2, d2 represent genetic parameters to be estimated corresponding to the mean effect and additive and dominance effects at loci 1 and 2, whereas iaa, iad, ida, and idd correspond to epistatic interaction effects. An advantage of this formulation is that, provided the parameter estimates are almost independent, we can test the epistasis between specific alleles at the two loci rather than merely testing the overall existence of any interaction between two loci.

3.5

Using Microarrays in QTL Mapping

3.5.1

eQTL Mapping: Measuring Gene Expression

1. Extract mRNA from the samples (see Note 5). For oligonucleotide microarrays, mRNA should be converted to cDNA first, which in turn will be used to generate biotin-labeled cRNA. But for spotted microarrays, the labeled “target” could be cRNA or cDNA. 2. RNA extraction and quantification: RNA should be extracted using various kits according to different sample types (Qiagen). Usually the quantification of total RNA amount is done on Agilent 2100 Bioanalyzer (see Note 6). 3. Convert mRNA to cDNA using reverse transcriptase and a poly-T primer. (For the spotted array, the cDNA is synthesized with Cy-labeled dNTPs.) 4. Amplify resulting cDNA using T7 RNA polymerase in the presence of biotinUTP and biotin-CTP. Each cDNA will yield 50–100 copies of biotin-labeled cRNA.

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5. Quantify the cRNA and determine the cRNA yield. 6. Fragment an appropriate amount of cRNA at 94 °C using fragmentation buffer to produce cRNA fragments of length 35–200 nucleotides. 7. Hybridize to chip using hybridization oven. 8. Wash away non-hybridized material and stain hybridized biotin-labeled cRNA with streptavidin-phycoerythrin and wash again. 9. Scan the chip in a high-resolution laser scanner. For oligo-nucleotide microarrays, absolute gene expression level is acquired. For spotted arrays, to which tested sample and the control was co-hybridized, expression values are based on the competitive hybridization of the two samples. 10. Validate microarray gene expression level with RT-PCR by reverse transcribing cDNA from RNA with reverse transcriptase (Applied Biosystems) and performing real-time quantitative PCR in a 50-µL reaction volume with ideal housekeeping gene control (Applied Biosystems).

3.5.2


Similar to traditional candidate gene association approaches, both population-based and family-based designs are commonly used in whole genome association studies to locate genes and/or polymorphisms associated with complex diseases (see Note 7). Taking account for the high cost of whole genome association study and that many polymorphisms within a small genomic region may be in strong linkage disequilibrium (LD) with each other, the two-stage approach has been adopted, in which a subset of the data is used to select a proportion of promising SNPs from a larger set of all known polymorphic sites within a candidate gene or a candidate region, and only these promising SNPs are typed in additional subjects (45). Therefore, it is quite important to choose the sub-sample for promising SNP selection. Too large a sample obviously defeats the purpose of reducing costs by limiting the number of SNPs to be genotyped in additional subjects, whereas too small a sample could lead to unstable estimates of the parameters of a model that can be applied to the analysis of the data of the additional subjects. Nevertheless, as the cost of high-volume genotyping plummets and haplotype tagging information from the International HapMap project rapidly accumulates in public databases, such two-stage designs may soon become unnecessary.

3.5.2.1

Isolation Total Genomic DNA from Whole Blood

1. Add 5 mL of whole blood (or bone marrow) to a 50-mL centrifuge tube containing 15 mL of RBC lysis solution. Invert to mix and incubate 5 minutes at room temperature; invert again at least once during the incubation. 2. Centrifuge for 5 minutes at 2,000 g. Remove supernatant leaving behind the visible white cell pellet and about 200–400 µL of the residual liquid. Vortex the tube vigorously to resuspend the cells in the residual liquid.

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3. Add 5 mL of cell lysis solution to the resuspended cells and pipette up and down to lyse the cells. 4. Add 1.67 mL of protein precipitation solution to the cell lysate. 5. Vortex vigorously at high speed for 20 seconds to mix the protein precipitation solution uniformly with the cell lysate. Centrifuge at 2,000 g for 5 minutes. 6. Pour the supernatant containing the DNA (leaving behind the precipitated protein pellet) into a 15- or 50-mL tube containing 5 mL of 100% isopropanol (2-propanol). Mix the sample by inverting gently 50 times. 7. Centrifuge at 2,000 g for 3 minutes; the DNA will be visible as a small white pellet. 8. Pour off supernatant and drain tube briefly on clean absorbent paper. Add 5 mL of 70% ethanol and invert tube several times to wash the DNA pellet. Centrifuge at 2,000 g for 1 minute. Carefully pour off the ethanol. 9. Invert and drain the tube on clean absorbent paper and allow sample to air dry for 10–15 minutes. 10. Add 500 µL of DNA hydration solution. 11. Rehydrate DNA by incubating at 65 °C for 1 hour and overnight at room temperature. If possible, tap tube periodically to aid in dispersing the DNA. 12. For storage, sample may be centrifuged briefly and then transferred to a 1.5-mL tube. Store DNA at 4 °C. For long-term storage, store at −20 °C or −80 °C.

3.5.2.2

Quantitation of Genome dsDNA

1. Prepare a 1× TE working solution by diluting the concentrated buffer 20-fold with sterile, distilled, DNase-free water. 2. Prepare an aqueous working solution of the Quant-iT PicoGreen reagent by making a 1:200 dilution of the concentrated anhydrous dimethylsulfoxide (DMSO) solution in 1× TE. 3. Prepare a 2 µg/mL stock solution of bacteriophage lambda dsDNA in 1× TE. Determine the DNA concentration on the basis of absorbance at 260 nm (A260) in a cuvette with a 1-cm pathlength; an A260 of 0.04 corresponds to 2 µg/mL dsDNA solution. 4. Dilute the 2 µg/mL DNA stock solution into plastic test tubes as shown in the table that follows. Add 1.0 mL of the aqueous working solution of the Quant-iT PicoGreen Reagent to each plastic test tube. Mix well and incubate for 2–5 minutes at room temperature, protected from light. Vol (µL) DNA stock

Vol (µL) 1× TE

Vol (µL) PicoGreen working solution

Final DNA conc. in PicoGreen assay

1,000 100 10 1 0

0 900 990 999 1,000

1,000 1,000 1,000 1,000 1,000

1 µg/mL 100 ng/mL 10 ng/mL 1 ng/mL Blank

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5. Measure sample fluorescence using the fluorescence microplate reader (excitation ~480 nm, emission ~520 nm). 6. Subtract the fluorescence value of the reagent blank from that of each of the samples. Use corrected data to generate a standard curve of fluorescence versus DNA concentration. 7. Add 1.0 mL of the aqueous working solution of the Quant-iT PicoGreen Reagent to each sample. Incubate 2–5 minutes at room temperature, protected from light. 8. Measure fluorescence of sample using instrument parameters that correspond to those used when generating standard curve (see step 5). 9. Subtract fluorescence value of reagent blank from that of each sample. Determine DNA concentration of the sample from standard curve.

3.5.2.3

Whole Genome Association Analysis

1. Dilute the DNA sample to 50 ng/µL using reduced EDTA:TE buffer and aliquot 5 µL of each sample per well to a 96-well plate. 2. Aliquot 14.75 µL of the Digestion Master Mix (1 µL of Nsp I or Sty I restriction enzyme, 2 µL of corresponding NE buffer, 2 µL of BSA, and 9.75 µL of H2O) to digest the genomic DNA. Put the plate on a thermal cycler and run the digest program (37 °C for 120 minutes, 65 °C for 20 minutes, and 4 °C hold). 3. Aliquot 5.25 µL of the ligation master mix (0.75 µL of adaptor Nsp I or adaptor Sty I, 2.5 µL of T4 DNA ligase buffer, 2 µL of T4 DNA ligase) into each digested DNA sample to ligate the digested samples. Put the plate on a thermal cycler and run the digest program (16 °C for 180 minutes, 70 °C for 20 minutes, and 4 °C hold). 4. Dilute each DNA ligation reaction by adding 75 µL of molecular biology-grade H2O per 25 µL ligated DNA. 5. Transfer 10 µL of diluted ligated DNA to the corresponding well of three reaction plates. Aliquot 90 µL of PCR Master Mix (39.5 µL of H2O, 10 µL of titanium Taq PCR buffer, 20 µL of GC-Melt, 14 µL of dNTP, 4.5 µL of PCR primer 002, and 2 µL of TITANIUM Taq DNA polymerase) to three reaction plates and perform PCR program using GeneAmp PCR system 9700 (one cycle of 94 °C for 3 minutes, 30 cycles of 94 °C for 30 seconds, 60 °C for 45 seconds, and 68 °C for 15 seconds, one cycle of 68 °C for 7 minutes, and 4 °C hold). 6. Run 3 µL of each PCR product mixed with 3 µL of 2× gel loading dye on 2% TBE gel at 120 V for 1 hour. Check the average size of ligated DNA between 200 and 1,100 bp. 7. Add 8 µL of 0.1 M EDTA to each PCR reaction. Consolidate three PCR reactions for each sample into one well of the clean-up plate. Apply a vacuum and maintain at ~600 mbar until the wells are completely dry by QIAvac multiwell unit. Wash the PCR products by adding 50 µL of molecular biology grade water and dry the wells completely.

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8. Add 45 µL of RB buffer to each well. Cover the plate with PCR plate cover film and moderately shake the clean-up plate on a plate shaker for 10 minutes at room temperature. Transfer the purified PCR product to a fresh plate. 9. Use spectrophotometric analysis to determine the purified PCR product yield. Add 2 µL of the purified PCR product to 198 µL molecular biology grade water and mix well. Read the absorbance at 260 nm. And 1 absorbance unit at 260 nm equals 50 µg/mL for double-stranded PCR product. 10. Transfer 90 µg of each of the purified DNA samples to a new plate. Bring the total volume of each well up to 45 µL by adding the appropriate volume of RB buffer. 11. Add 5 µL of 10× fragmentation buffer to each sample. Dilute the stock of fragmentation reagent to 0.05 U/µL, and add 5 µL of diluted fragmentation reagent to the fragmentation plate. Run the fragment program (37 °C for 35 minutes, 94 °C for 15 minutes, and 4 °C hold). 12. Dilute 4 µL of fragmented PCR product with 4 µL of gel loading dye and run on 4% TBE gel at 120 V for 30 minutes to 1 hour. Check the average size of fragment DNA

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