Pediatric Bone, Second Edition: Biology & Diseases

PEDIATRIC BONE BIOLOGY & DISEASES SECOND EDITION Edited by FRANCIS H. GLORIEUX Genetics Unit, Shriners Hospital, McGil...

Author: Francis H. Glorieux | John M Pettifor | Harald Juppner

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PEDIATRIC BONE BIOLOGY & DISEASES SECOND EDITION Edited by

FRANCIS H. GLORIEUX Genetics Unit, Shriners Hospital, McGill University, Montreal, Quebec, Canada

JOHN M. PETTIFOR Mineral Metabolism Research Unit. Department of Pediatrics, Chris Hani Baragwanath Hospital, Soweto, Johannesburg, South Africa

HARALD JU¨PPNER Endocrine Unit, Department of Medical and Pediatrics, Massachusetts General Hospital, and Harvard Medical School, Boston, Massachusetts, USA

AMSTERDAM • BOSTON • HEIDELBERG • LONDON NEW YORK • OXFORD • PARIS • SAN DIEGO SAN FRANCISCO • SINGAPORE • SYDNEY • TOKYO Academic Press is an imprint of Elsevier

Academic Press is an imprint of Elsevier 32 Jamestown Road, London NW1 7BY, UK 225 Wyman Street, Waltham, MA 02451, USA 525 B Street, Suite 1800, San Diego, CA 92101-4495, USA First Edition 2003 Second Edition 2012 Copyright Ó 2012, 2003 Elsevier Inc. All rights reserved. No part of this publication may be reproduced, stored in a retrieval system or transmitted in any form or by any means electronic, mechanical, photocopying, recording or otherwise without the prior written permission of the publisher Permissions may be sought directly from Elsevier’s Science & Technology Rights Department in Oxford, UK: phone (+44) (0) 1865 843830; fax (+44) (0) 1865 853333; email: [email protected]. Alternatively, visit the Science and Technology Books website at www.elsevierdirect.com/rights for further information Notice No responsibility is assumed by the publisher for any injury and/or damage to persons or property as a matter of products liability, negligence or otherwise, or from any use or operation of any methods, products, instructions or ideas contained in the material herein. Because of rapid advances in the medical sciences, in particular, independent verification of diagnoses and drug dosages should be made Medicine is an ever-changing field. Standard safety precautions must be followed, but as new research and clinical experience broaden our knowledge, changes in treatment and drug therapy may become necessary or appropriate. Readers are advised to check the most current product information provided by the manufacturer of each drug to be administered to verify the recommended dose, the method and duration of administrations, and contraindications. It is the responsibility of the treating physician, relying on experience and knowledge of the patient, to determine dosages and the best treatment for each individual patient. Neither the publisher nor the authors assume any liability for any injury and/or damage to persons or property arising from this publication British Library Cataloguing-in-Publication Data A catalogue record for this book is available from the British Library Library of Congress Cataloging-in-Publication Data A catalog record for this book is available from the Library of Congress ISBN : 978-0-12-382040-2 For information on all Academic Press publications visit our website at www.elsevierdirect.com Typeset by TNQ Books and Journals Pvt. Ltd. Printed and bound in United States of America 12 13 14 15 10 9 8 7 6 5 4 3 2 1

Preface to the Second Edition

The first edition of this book, published in 2003, was sufficiently well received that we find it timely to publish a second edition 8 years later. A great deal of new knowledge has enriched the field of pediatric bone health and diseases during this period. We have kept in this second edition the same overall organization set in the first one. There have been changes in the list of contributors with a few declining and new ones embarking with enthusiasm. The first part of the book deals with basic information on the major systems controlling the development, structure and homeostasis of bone tissue, with one chapter added on dental development and maturation. The second part describes the tools and techniques validated for the evaluation of bone development and its abnormalities. To follow suggestions received, a new chapter on radiographic imaging completes this section. The third part deals with specific disorders or groups of disorders with emphasis where appropriate on basic aberrations and their clinical expression. As was requested by several readers, we have tried to provide more information on management of the various diseases. We hope that the new edition will be well received, as pediatricians and researchers working in the field of pediatric bone diseases see the need for such a textbook. The realm of pediatric bone diseases continues to rapidly expand and we hope that the book will be useful

to those interested in it, to the scientists at the bench to understand the clinical expression of the biological pathways and functions they study in the laboratory, and to the clinicians who wish to connect their observations with complex biological mechanisms and to the underlying genetic defects as they evaluate their patients’ response to established or new therapies. We wish to express our gratitude to Mara Conner and Megan Wickline at Elsevier/Academic Press for their understanding, patience and support. We also extend our appreciation to all the contributors to this large effort. In the preface of the first edition we expressed the hope that the content of the book would give substance to the recognition that “pediatric osteology” had come of age as a medical specialty, which has frequently provide substantial new insights into biology by the fact that an increasing number of these pediatric diseases has now been defined at the molecular level and thus may have significant implications also for the understanding of adult bone diseases. Although those who have invested their professional life into it come from a spectrum of different backgrounds, they now identify themselves more clearly with this emerging field. May the book be of help in that effort, and also entice younger colleagues to join in an exciting journey.

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List of Contributors

Judith E. Adams MBBS, FRCR, FRCP, FBIR Consultant Radiologist, Manchester Royal Infirmary & Professor of Diagnostic Radiology, University of Manchester, UK

John Damilakis Associate Professor, University of Crete, Faculty of Medicine, Department of Medical Physics, Iraklion, Crete, Greece

Laura K. Bachrach MD Professor of Pediatrics, Department of Pediatrics, Stanford University School of Medicine, Palo Alto, CA, USA

Marie B. Demay MD Endocrine Unit, Massachusetts General Hospital, Harvard Medical School, Boston, MA, USA

Murat Bastepe MD, PhD Assistant Professor of Medicine, Endocrine Unit, Department of Medicine, Massachusetts General Hospital and Harvard Medical School, Boston, MA, USA

Michael J. Econs MD Professor of Medicine, Indiana University School of Medicine, Indianapolis, IN, USA Serge Ferrari MD Division of Bone Diseases, University Hospitals and Medical Faculty of Geneva, Switzerland

Clemens Bergwitz MD Assistant Professor of Medicine, Massachusetts General Hospital, Department of Medicine, Division Endocrinology, Massachusetts General Hospital, Boston, MA, USA

Mary Fewtrell Reader in Childhood Nutrition, Honorary Consultant Paediatrician, Childhood Nutrition Research Centre, UCL Institute of Child Health, London, UK

Maria Luisa Bianchi MD Bone Metabolism Unit, Istituto Auxologico Italiano IRCCS, Milan, Italy

Thomas J. Gardella PhD Associate Professor of Medicine, Massachusetts General Hospital, Department of Medicine, Division of Endocrinology, Massachusetts General Hospital, Endocrine Unit, Boston, MA, USA

Paolo Bianco MD Professor of Pathology, Sapienza University of Rome, Director, Anatomic Pathology, Umberto I University Hospital, Rome, Italy

Francis H. Glorieux OC, MD, PhD Professor of Surgery, Pediatrics and Human Genetics, McGill University, Adjunct Professor of Pediatrics, University of Montreál, Director of Research, Shriners Hospital for Children, Montreal, Canada

Nick Bishop MD Head, Academic Unit of Child Health, Professor of Paediatric Bone Disease, University of Sheffield, Sheffield Children’s Hospital, Sheffield, UK Lynda F. Bonewald PhD Vice Chancellor for Research Interim, Curator’s Professor, Lee M and William Lefkowitz Professor, Director, Bone Biology Research Program Director, UMKC Center of Excellence in Mineralized Tissues University of Missouri at Kansas City School of Dentistry, Department of Oral Biology, Kansas City, MO, USA

Michel Goldberg DDS Professeur e´merite, UMR-S 747INSERM Universite´ Paris Descartes Nick Harvey MD Lecturer and Honorary Consultant in Rheumatology, The MRC Lifecourse Epidemiology Unit, University of Southampton, Southampton General Hospital, Southampton, UK

Jean-Philippe Bonjour MD Division of Bone Diseases, University Hospitals and Medical Faculty of Geneva, Switzerland

Priv.Dozent Dr Wolfgang Ho¨gler Consultant Paediatric Endocrinologist, Birmingham Children’s Hospital, Honorary Senior Lecturer, University of Birmingham, UK

Noe¨l Cameron Professor of Human Biology, Centre for Global Health and Human Development, Loughborough University, Loughborough, Leicestershire, UK

Ingrid A. Holm MD, MPH Division of Genetics, Program in Genomics, and the Manton Center for Orphan Disease Research, Children’s Hospital Boston and Harvard Medical School, Boston, MA, USA

Thomas O. Carpenter MD Departments of Pediatrics (Endocrinology) and Orthopaedics and Rehabilitation, Yale University School of Medicine, New Haven, CT, USA

Katharina Ja¨hn PhD University of Missouri-Kansas City, School of Dentistry, Kansas City, MO, USA

Thierry Chevalley MD Division of Bone Diseases, University Hospitals and Medical Faculty of Geneva, Switzerland

Harald Ju¨ppner MD Professor of Pediatrics, Endocrine Unit and Pediatric Nephrology Unit, Department of Medicine and Pediatrics, Massachusetts General Hospital and Harvard Medical School, Boston, MA, USA

William G. Cole MD, PhD Chief of Pediatric Surgery, University of Alberta, Canada

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xii

LIST OF CONTRIBUTORS

Frederick S. Kaplan MD Departments of Orthopedic Surgery, Medicine, and the Center for Research in FOP and Related Disorders, University of Pennsylvania School of Medicine, Philadelphia, PA, USA Christopher S. Kovacs MD, FRCPC, FACP Professor of Medicine (Endocrinology and Metabolism), Obstetrics & Gynecology, and BioMedical Sciences, Memorial University of Newfoundland, Attending and Consultant Specialist (Endocrinology and Metabolism), Health Sciences Centre and Eastern Health Corporation, St. John’s, Newfoundland, Canada Henry M. Kronenberg MD, PhD Endocrine Unit, Massachusetts General Hospital and Harvard Medical School, Boston, MA, USA Mary B. Leonard MD, MSCE Associate Professor of Pediatrics & Epidemiology, The Children’s Hospital of Philadelphia, Department of Pediatrics, Department of Biostatistics and Epidemiology, University of Pennsylvania School of Medicine, Philadelphia, PA, USA Fanxin Long PhD Associate Professor of Medicine, and Associate Professor of Developmental Biology, Division of Endocrinology, Metabolism and Lipid Research, Department of Medicine, Washington University in St Louis, MO, USA Christa Maes PhD Laboratory of Experimental Medicine and Endocrinology (LEGENDO), Katholieke Universiteit Leuven, Belgium

John M. Pettifor MBBCh, PhD Mineral Metabolism Research Unit, Department of Paediatrics, University of the Witwatersrand and Chris Hani Baragwanath, South Africa Anthony A. Portale MD Professor of Pediatrics, University of California San Francisco, San Francisco, CA, USA John T. Potts MD Endocrine Unit, Department of Medicine, The Massachusetts General Hospital and Harvard Medical School, Boston, MA, USA Ann Prentice PhD Director, MRC Human Nutrition Research, Elsie Widdowson Laboratory, Fulbourn Road, Cambridge, UK Frank Rauch MD Associate Professor, Genetics Unit, Shriners Hospital for Children, Montreal, Canada Jean-Marc Retrouvey Division of Orthodontics, McGill University, Montreal, Quebec, Canada David L. Rimoin MD, PhD Medical Genetics Birth Defects Center, Cedars-Sinai Health System and Department of Pediatrics and Medicine, UCLA School of Medicine, Los Angeles, CA, USA Rene Rizzoli MD Division of Bone Diseases, University Hospitals and Medical Faculty of Geneva, Switzerland David Rowe Department of Genetics and Developmental Biology, University of Connecticut Health Center, Farmington, CT, USA Isidro B. Salusky MD Professor of Pediatrics, Division of Pediatric Nephrology, Director, Pediatric Dialysis Program, Director, General Clinical Research Center, David Geffen School of Medicine at UCLA, Los Angeles, CA, USA

Daniella Magen MD Division of Pediatric Nephrology, Meyer Children’s Hospital, Rambam Health Care Campus and Laboratory of Molecular Medicine, Rappaport Faculty of Medicine and Research Institute Technion e Israel Institute of Technology, Haifa, Israel

Ste´phane Schwartz DDS Assistant Director, Dental Clinic Montreal Children’s Hospital MUHC (McGill University Health Center) Associate Professor, McGill University

David D. Martin MD Paediatric Endocrinology and Diabetology, University Children’s Hospital, Tu¨bingen University, Tu¨bingen, Germany

Nick Shaw MD Consultant Paediatric Endocrinologist, Birmingham Children’s Hospital, Honorary Senior Lecturer, University of Birmingham, UK

Marc D. McKee PhD Professor, Faculty of Dentistry, Department of Anatomy & Cell Biology and Faculty of Medicine, McGill University, Montreal, Quebec, Canada

Eileen M. Shore PhD Departments of Orthopedic Surgery, Genetics, and the Center for Research in FOP and Related Disorders, University of Pennsylvania School of Medicine, Philadelphia, PA, USA

Walter L. Miller MD Distinguished Professor of Pediatrics, University of California San Francisco, San Francisco, CA, USA Geert Mortier MD Department of Medical Genetics, University of Antwerp, Antwerp, Belgium Zulf Mughal Consultant in Paediatric Bone Disorders & Honorary Senior Lecturer in Child Health, Royal Manchester Children’s Hospital, Central Manchester University Hospitals NHS Foundation Trust, Manchester, UK Amaka C. Offiah HEFCE Clinical Senior Lecturer, Academic Unit of Child Health, Sheffield Children’s NHS Foundation Trust, Sheffield, UK Bram Perdu MD Department of Medical University of Antwerp, Antwerp, Belgium

Genetics,

Farzana Perwad MD Assistant Professor of Pediatrics, University of California San Francisco, San Francisco, CA, USA

Rene´ St-Arnaud PhD Genetics Unit, Shriners Hospital for Children, Montreál (Que´bec) and Departments of Surgery and Human Genetics, McGill University, Montreál (Que´bec), Canada Andrea Superti-Furga Professor, University of Lausanne, Division of Pediatrics, Centre Hospitalier Universitaire Vaudois, Lausanne, Switzerland Rajesh V. Thakker MD, ScD, FRCP, FRCPath, FMedSci Professor of Medicine, Academic Endocrine Unit, Nuffield Department of Medicine, Oxford Centre for Diabetes, Endocrinology and Metabolism (OCDEM), Churchill Hospital, Headington, Oxford, UK Sheila Unger MD, FRCP University of Lausanne, Service of Medical Genetics, Centre Hospitalier Universitaire Vaudois, Lausanne, Switzerland Filip Vanhoenacker MD Department of University Hospital of Antwerp, Belgium

Radiology,

LIST OF CONTRIBUTORS

Wim Van Hul MD Department of Medical Genetics, University and University Hospital Antwerp, Belgium Katherine Wesseling Perry MD David Geffen School of Medicine at University of California, Los Angeles, CA, USA Michael P. Whyte MD Medical-Scientific Director, Center for Metabolic Bone Disease and Molecular Research, Shriners Hospital for Children, Professor of Medicine, Pediatrics, and Genetics, Washington University School of Medicine, St. Louis, MO, USA

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Shlomo Wientroub MD Department of Pediatric Orthopedic Surgery, Dana Children’s Hospital, Tel Aviv Medical School, Tel Aviv, Israel Israel Zelikovic MD Division of Pediatric Nephrology, Meyer Children’s Hospital, Rambam Health Care Campus, and Laboratory of Developmental Nephrology, Department of Physiology and Biophysics, Rappaport Faculty of Medicine and Research Institute, Technion e Israel Institute of Technology, Haifa, Israel

C H A P T E R

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Bone Cell Biology: Osteoclasts, Osteoblasts, Osteocytes K. Ja¨hn, L.F. Bonewald University of Missouri-Kansas City, School of Dentistry, 650 E 25th Street, Kansas City, MO 64108, USA

INTRODUCTION TO BONE

RANKL, ephrins, and sclerostin as described in this chapter. Moreover, the extracellular matrix (ECM) on which osteoblasts and osteoclasts attach and in which osteocytes reside, influences the function of these cells. Although previously viewed as mainly a support structure for bone cells, it is now clear that the bone ECM controls and directs bone cell function. This chapter will provide an overview of how bone cells coordinate their actions to generate, maintain and remove bone mass.

Depending upon field of study, bone is described very differently. From a material, biochemical, or physical point of view, bone is described as a composite material that is made up of an organic component consisting of collagenous and non-collagenous proteins and a mineral phase consisting of calcium and phosphate [1]. Often bone is viewed as a dead, hard material with only a structural purpose, to hold the body in place, to protect internal organs, and to serve as attachment sites for skeletal muscles to achieve movement. Yet bone is far from being either dead or static. From a biological point of view, bone is a complex living tissue in which this composite material of organic and mineral components is created and maintained by at least three major cell types, namely osteoblasts, osteocytes and osteoclasts (see schematic bone remodeling unit in Figure 1.1) [2]. In the first and second decades of human life, bone is constantly growing. Towards the end of the second and into the third decades, dynamic remodeling takes place to maintain the skeleton with the potential for increasing bone mass. After the third decade, bone mass starts its inevitable decline. Considering this simplified time course of human bone development, it is obvious that bone is dynamically controlled and remodeled. As a dynamic connective tissue, bone is constantly responding to external forces such as loading of the skeleton and to internal and external signals such as cytokines, growth factors and hormones. Several hormones have been shown to play important roles in the skeleton such as the estrogens and androgens, parathyroid hormone (PTH) and 1,25-dihydroxyvitamin vitamin D3 (1,25,D3). In addition to these external signals, bone cells are in constant communication with each other and cells of the immune and hematopoietic systems, through factors such as osteoprotogerin,

Pediatric Bone, Second Edition DOI: 10.1016/B978-0-12-382040-2.10001-2

OSTEOCLASTS The cells that resorb the mineralized ECM of bone are called osteoclasts. They arise from hematopoietic progenitors that also give rise to macrophages. The osteoclast precursor cells are recruited to the bone surface where they fuse to form multinucleated cells [3] (see schematic image in Figure 1.2). The co-stimulating molecule RANKL (RANK ligand), which is expressed by osteoblasts, stromal cells and osteocytes, is the most commonly known cellular activation pathway of osteoclast-precursors and the osteoclastogenic cascade of transcription factors [3,4]. Osteoclasts are very rare in bone, only two to three cells per mm3 bone can be found [5]. The active form of these multinucleated giant cells is present in specialized cavities on the bone surface, known as Howship’s lacunae. Osteoclasts show a high level of polarization when attached to the bone surface. The most characteristic feature in this state is the ruffled border, which consists of finger-like cytoplasmic projections [6] and is turned towards the bone surface. The osteoclast seals the cavity around its ruffled border and then secretes protons and a variety of proteolytic enzymes, i.e. collagenases, gelatinases, into the cavity, to carry out the organic breakdown of the bone ECM [7].

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Copyright Ó 2012 Elsevier Inc. All rights reserved.

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1. BONE CELL BIOLOGY: OSTEOCLASTS, OSTEOBLASTS, OSTEOCYTES

FIGURE 1.1

The bone remodeling unit. This drawing of a bone remodeling unit depicts an active osteoclast sitting in its Howship’s lacunae releasing protons and enzymes into a sealed zone to degrade and digest the mineralized bone matrix. Cuboidal-shaped osteoblasts are positioned behind the osteoclasts secreting type I collagen and other non-collagenous proteins to form the osteoid e the newly deposited, unmineralized bone matrix. Several differentiation stages of the osteoblast phenotype are illustrated. The spindle-shaped bone lining cells are covering all inactive bone surfaces, as well as the bone remodeling unit to create a micromilieu where theoretically cross-talk takes place between osteoclasts and osteoblasts. Transition stages from the osteoblast to the osteocyte are shown including the early osteoid osteocyte to the mature embedded osteocyte. The role of the osteocyte to create a communication network between practically all cells on the bone surface is demonstrated by the extensive cell processes connecting osteocytes, osteoblasts, bone lining cells and osteoclasts.

The sole function of the osteoclast is to resorb bone. The mature osteoclast is described histologically as a multinucleated, tartrate resistant, acid phosphatase positive cell. However, macrophage polykaryons can have these same characteristics; so the “gold standard” for identifying an osteoclast is the formation of resorption lacunae or “pits” on a mineralized surface (see histological image in Figure 1.2). Other characteristics of the osteoclast include the expression of calcitonin receptors, enzymes such as cathepsin K and matrix metalloprotein-9 (MMP-9) that play a role in matrix degradation, and the vacuolar proton pump for the transport of protons to the resorption lacunae. For the osteoclast to resorb, it must form a “sealing zone” around the periphery of its attached area to concentrate not only proteases but also protons into this limited area. Underneath the cell, within the ruffled border, the pH is reduced to approximately 2e3 which enhances the degradation of mineralized matrix [8]. As osteoclast precursors are derived from hematopoietic precursors, the same stem cells that become granulocytes and monocytes/macrophages, cell lines such as RAW 267.4 and MOPC-5, are available that represent osteoclast precursors for experimental studies. These

cells can form TRAP positive multinucleated cells that resorb bone and form resorption lacunae on dentin [9]. It has been well known for the last 10e15 years in the bone field that osteoclast precursors require supporting cells for osteoclast formation. The importance of factors produced by supporting cells such as macrophage colony stimulating factor (M-CSF) to induce proliferation of osteoclast precursors has been validated [10]. Critical factors and cell surface molecules involved in osteoclast formation have only recently been elucidated with the discovery of RANK ligand (RANKL) and osteoprotogerin (OPG) [11,12]. The osteoclast precursor expresses a receptor known as RANK (receptor activator of NFkB) that signals through the NF-kB pathway. The binding of the cell membrane bound ligand, RANKL, activates RANK receptor. However, the soluble factor, OPG, acting as a “decoy” receptor can bind to RANKL, preventing osteoclast formation. The expression of RANKL on the surface of supporting cells occurs when these cells are exposed to bone resorbing cytokines, hormones, and factors such as interleukins 1, 6, 11, PTH, PTH-related protein (PTHrp), oncostatin M, leukemia inhibitory factor, prostaglandin E2, or 1,25 D3 [13]. These factors upregulate RANKL to a level

PEDIATRIC BONE

THE OSTEOBLAST CELL LINEAGE

FIGURE 1.2 Osteoclast differentiation and activity. The activation of osteoclast precursors to fuse and form active osteoclasts is shown in the schematic image (upper panel). Arising from a hematopoietic precursor that is activated by M-CSF and RANKL, the pre-osteoclast is formed and fuses with others to produce multinucleated TRAP positive cells, which finally form a sealing zone to attach to the bone matrix to form the bone resorption cavity underneath a ruffled border. Active, TRAP positive stained osteoclasts (red) can be seen on the histological image taken from the distal region of a mouse femur. Methyl green was used as counterstaining to label all cell nuclei. (See color plate section.)

capable of overcoming the effects of circulating OPG, thereby resulting in osteoclast formation. Efforts to generate osteoclasts without supporting cells have only recently been accomplished in vitro by using an artificial, soluble form of RANKL [14]. A new therapeutic called Denosumab which is an anti-RANKL antibody has been approved for the treatment of osteoporosis [15,16].

THE OSTEOBLAST CELL LINEAGE Osteoblasts are known as the cells that form bone, characterized by their unique ability to secrete a type I collagen-rich ECM that eventually mineralizes.

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Osteoblasts were first named by Gegenbaur in 1864 [17], and since then their function as the major proteinexpressing cells in bone has been widely studied. All of the cells and stages of differentiation with the osteoblast lineage are often quickly summarized as “the osteoblast”. However, at least four to five major maturation stages of the osteoblast cell lineage are commonly accepted, the immature osteoblast referred to as preosteoblasts, mature osteoblasts, osteoid ostecytes, early osteocyte, and mature osteocytes as terminally differentiated osteoblasts embedded in the bone matrix [18]. Franz-Odendaal et al. in 2006 postulated a total of eight different maturation stages for the transition of osteoblasts to osteocytes based on morphological features as well as expression profiles of the cells [19]. The recent classification of osteoblast-lineage clearly distinguishes between active surface osteoblasts that secrete the ECM and several maturation stages of osteoid osteoblasts and young osteocytes to mature and deeply embedded osteocytes. Lining cells on the bone surface are thought also to be terminally differentiated osteoblasts on the bone surface, but the lineage of these cells has not been completely validated (a short summary of osteoblast maturation stages and markers can be found in Figure 1.3). Osteoblasts arise from multipotent progenitor cells of mesenchyme origin, mesenchymal stem cells. Friedenstein and co-workers were the first to characterize these clonogenic cells by their adherence to tissue culture plastic surfaces, and discovered that this isolated population is able to differentiate into a variety of mature cell types ranging from osteoblasts, to chondrocytes or adipocytes, under the appropriate conditions [20,21]. The niche for osteoprogenitors in bone is the periosteum, the endosteum, and the marrow stroma [22,23]. With bone trauma, fracture, or other conditions requiring bone repair, mesenchymal progenitor cells serve as a stem cell reservoir that are recruited to the injury site. However, unlike the hematopoietic stem cell, the identification of mesenchymal stem cells has not been as thoroughly characterized. One can identify single hematopoietic stem cells, but this has not been accomplished for mesenchymal stem cells. At present, a population of cells with a series and panel of markers has been identified with the capacity to differentiate into osteoblasts [24]. Chemotactic molecules such as osteopontin [25] and various members of the transforming growth factor-b family [26,27], which are stored in great amounts in the bone ECM, when released attract progenitor cells to the injured site or to a previously resorbed, remodeling site. These progenitors differentiate into matrix-producing osteoblasts. A “master control gene” necessary for progenitor cells to differentiate into osteoblasts is the transcription factor Runx2 (Runt-related transcription factor-2; also

PEDIATRIC BONE

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FIGURE 1.3 The osteoblast lineage. The upper panel represents the differentiation of a pre-osteoblast into an osteocyte. While the preosteoblast descended from the mesenchymal progenitor cell is mainly but not entirely expressing type I collagen, the mature osteoblast also expresses a variety of non-collagenous proteins for ECM synthesis, including bone sialoprotein and osteocalcin, a bone specific marker. During the differentiation into mature osteocytes, several proteins are upregulated such as E11/gp38, a marker for embedding and early osteocytes and sclerostin a marker for late, mature osteocytes. The histological image from the distal region of a mouse femur (stained with hematoxylin and eosin) shows an active cuboidal osteoblast on the bone surface (purple arrow), an early osteocyte (blue arrow) and a mature, stellar-shaped osteocyte deeply embedded in the bone matrix (green arrow). (See color plate section.)

named Cbfa1, Osf2, Pebp2a1, or Aml3) [28,29]. Runx2 is of crucial importance during osteoblast differentiation. This was dramatically demonstrated by deleting or “knocking-out” the gene in mice. Newborn mice possessed a cartilagenous, but non-mineralized skeleton and died upon birth due to inability to breathe. The importance of this “master control gene” was highlighted in 1997, when a total of five papers were

published in Cell describing the role of Runx2 during osteoblast differentiation and the dramatic effects of its targeted deletion in mice models leading to the complete absence of bone formation [28,30e33]. Runx2 was the first true osteoblast-specific transcription factor identified through its binding site within the promotor of the osteocalcin gene [34]. Runx2 is known to control osteoblast differentiation by subsequently activating the expression of osteoblast phenotype-specific genes such as type I collagen and osteocalcin [34,35]. While Runx2 is the earliest and most specific marker of osteogenesis identified to date [36], several homeodomain transcription factors, such as Msx2, Dlx5, Bapx1, and Hoxa-2 have been suggested to regulate Runx2 expression. A second crucial transcription factor required for osteoblast differentiation during development that acts downstream of Runx2 is osterix (Osx) [35]. Mice lacking Osx have a similar phenotype to those lacking Runx2. Nakashima et al. demonstrated that osterix-null mice do not show signs of intramembranous or endochondral bone formation with the absence of bone markers such as osteocalcin, osteopontin, or osteonectin [35]. The active, mature osteoblasts exhibit a cuboidal or polygonal cell shape. As these cells are responsible for the secretion of the bone matrix or osteoid, osteoblasts contain abundant endoplasmatic reticula and enlarged Golgi apparatus [37]. The major protein secreted by osteoblasts is type I collagen, a fibrillar extracellular matrix protein that determines the tensile strength of bone. Type I collagen is not exclusively found within bone, it is in fact one of the most abundant proteins in vertebrates and is present in skin, tendons and ligaments. Each type I collagen molecule is composed of three polypeptide chains (one a2 and two a1 chains) that are organized as a right-handed triple helix [38e40]. Osteoblasts further express a variety of so-called non-collagenous proteins to form the unmineralized matrix in the osteoid seam [41,42]. The specific function of most of these proteins is still not completely understood. Osteocalcin (OCN) is one of the most prominent non-collagenous proteins that is bone-specific in expression. This small 49 amino acid protein contains, in most species, uncommon post-transcriptional modification, gammacarboxylation, of three glutamic acids (Gla). These calcium-binding residues lead to the formation of “Gla”-helices which bind to hydroxyapatite (the inorganic component of the ECM of bone) [38]. OCN is expressed during the late stage of osteoblast differentiation in vitro [43]. Moreover, in embryonic bone, it is upregulated with the early onset of hydroxyapatite crystal formation [40]. Therefore, OCN has been suggested to be one of the main regulators of bone turnover, and mineralization [44]. Recent findings also imply a role for this calcium-binding protein in its undercarboxylated form in the hormonal regulation of glucose

PEDIATRIC BONE

THE OSTEOCYTE

and fat metabolism [45]. This has opened a whole new area of investigation and supports the novel hypothesis that bone can function as an endocrine organ. Other cells derived from the osteoblastic-lineage are bone lining cells. This origin of lining cells has been very controversial. Bone lining cells are thought to be resting osteoblasts, pre-osteoblasts or post-osteoblasts. We know that these cells cover almost all surfaces in adult bone to build a connective tissue barrier. More recently, bone lining cells were acknowledged as active cells that participate in bone resorption and formation. Moreover, these cells appear to be involved in homeostatic processes as they are in communication with the osteocyte network and are an element of the strain sensing network [37]. Bone lining cells may also be one of the major sources for active osteoblasts and could be seen as a reservoir for pre-osteoblasts if subjected to the right stimulus. This theory was proposed by Dobnig et al., where intermittent treatment of adult rats with PTH led to a reactivation of bone lining cells to bone forming osteoblasts that led to increased bone formation [46].

THE OSTEOCYTE The mature differentiated stage of the osteoblast lineage becomes apparent when osteoblasts are surrounded by the ECM of bone. Osteoblasts then undergo morphological changes; they develop long slender-like cell processes, lose many of their cytoplasmic organelles and develop into stellar-shaped cells [18]. This very distinctive osteoblast maturation stage is then called an osteocyte. Osteocytes possess a very unique location in bone, being trapped within lacunae similar to small “caves” inside the bone matrix, where these cells form a connective network by sending their dendritic processes through small “tunnels”, called canaliculi, that connect and span throughout the whole bone volume. It is unclear why, considering that osteocytes are also the most abundant cell type found in bone [47], their specific role within bone biology was overlooked for decades. This is thought to be due to the difficulty of isolating and removing mature osteocytes from the mineralized matrix. The proposed main function of osteocytes was to sense distribution and amount of mechanical strain that is applied to the bone [48,49]. A key factor that led to the discovery of this particular function of osteocytes is their location within the load-bearing ECM of bone [50]. Moreover, the importance of the integrity of the osteocytic network is highlighted in aging or microdamaged bone or with certain treatments, i.e. glucocorticoids, where the loss of viable osteocytes causes bone

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tissue to become a truly inactive, dead tissue [51,52]. The transfer of the strain information between osteocytes is realized through their long slender-like processes e the most dramatic morphological feature of osteocytes. Whereas numerous osteoblast cell lines have been generated for experimental studies, only one osteocyte cell line has been generated to date, the MLO-Y4 cell that possesses dendritic processes and early markers for osteocytes. This cell line has been and is being used extensively to elucidate osteocyte function [53e59]. Gap junctions at the end of these processes enable the osteocytes to communicate. Gap junctions are specialized cellecell contact points which are formed by members of the connexin protein family [60,61]. Six of these transmembrane proteins are joined across the extracellular gap between two adjacent cells to form a channel structure allowing the passage of small metabolites, ions and intracellular signaling molecules below a size of 1 kDa. But what causes this form of intracellular signaling due to mechanical stimulation? In theory, there are at least three different forms of mechanical strain on bone cells e hydrostatic pressure, direct cell strain and fluid flow-induced shear stress. The most commonly accepted theory is load-induced fluid flow [48,49,60]. Interstitial fluid is squeezed through the porous ECM and through the lacunarecanaliculi system in response to bone deformations by physiological loading and the shear stresses then act directly on the outer cellular structures of the osteocytes [62]. The most prominent molecules that have been involved in mechanotransduction in bone are nitric oxide [63], adenosine triphosphate (ATP) [64,65], prostaglandin (PGE2) [49,66] and calcium [67]. Osteocytes have recently come into the spotlight as they appear to be multifunctional [68]. One function is to contribute to the regulation of mineral homeostasis. It has been shown that osteocytes can regulate calcium availability by removing and replacing their mineralized matrix under normal, healthy conditions such as lactation [68,69]. In addition to the regulation of calcium, osteocytes have a major role in the regulation of phosphate homeostasis. Osteocytes regulate both biomineralization and phosphate through molecules such as phosphate regulating factor with homologies to endopeptidase on chromosome X (Phex), dentin matrix protein-1 (Dmp1), and fibroblast growth factor (FGF23), all expressed by osteocytes [70e73]. Both Dmp1 and Phex appear to downregulate the expression of FGF-23, which allows more efficient reabsorption of phosphate by the kidney thereby maintaining sufficient circulating phosphate to maintain normal bone mineral content. Dmp1 null mice have a similar phenotype to hyp mice carrying a Phex mutation, that of osteomalacia and rickets due to elevated FGF-23 levels in osteocytes

PEDIATRIC BONE

6


[73,74]. While autosomal dominant hypophosphatemic rickets in patients is due to mutations in Phex [75], autosomal recessive hypophosphatemic rickets is due to mutations in Dmp1 [74] and both show elevation of FGF-23 in osteocytes. Based on these observations, we proposed that the osteocyte lacunocanalicular network can function as an endocrine system, targeting distant organs such as kidney [74]. Another function of osteocytes is to act as orchestrators of both bone formation and bone resorption. Osteocytes produce a factor, sclerostin, which inhibits osteoblastic bone formation [76,77]. This molecule is now the target of therapeutics as an antibody to sclerostin has been shown to enhance bone mass and promote bone healing [78e81]. Conversely, osteocytes can support osteoclast formation, especially in their dying or apoptotic state [82,83]. Osteocytes appear to express both RANKL and OPG [84]. What is not clear is how factors produced by osteocytes can reach the bone surface, but Dallas and co-workers have shown by dynamic imaging that osteocytes can extend their dendritic processes into marrow and vascular spaces suggesting a mean for the delivery of factors [85,86].

CONCLUSION Not only are bone cells in constant communication with each other, but the ECM can also influence the function of bone cells. Bone is a reservoir of factors ready to be released during resorption that can modify the bone coupling process or provide circulating growth factors. A number of transcription factors as described above have been identified that are specific for bone induction and development. Clearly, these growth factors and transcription factors are regulated by a number of circulating hormones such as parathyroid hormone, estrogen, and 1,25(OH)2D3. As outlined in this chapter, another layer of complexity is added due to the fact that bone structure is also regulated by mechanical load or lack of mechanical loading. Understanding the normal physiology of bone and its diseases has led to the generation of therapeutics important in the prevention and treatment of bone disease and acceleration and initiation of bone repair. Studies are also underway to identify means to treat or reverse abnormal bone development based on studies of bone cell origin, function, and their regulation.

References [1] Cowin SC. Bone Mechanics Handbook. New York: CRC Press; 2001. [2] Bilezikian JP, Lawrence GR, Rodan GA. Principles of Bone Biology. Academic Press; 2002.

[3] Takahashi N, Udagawa N, Takami M, Suda T. Cells of bone: osteoclast generation. In: Bilezikian JP, Raisz LG, Rodan GA, editors. Principles of Bone Biology. Academic Press; 2002. p. 109e26. [4] Matsuo K, Irie N. Osteoclast-osteoblast communication. Arch Biochem Biophys 2008;473:201e9. [5] Meunier PJ, Coindre JM, Edouard CM, Arlot ME. Bone histomorphometry in Paget’s disease. Quantitative and dynamic analysis of pagetic and nonpagetic bone tissue. Arthritis Rheum 1980;23:1095e103. [6] Roodman GD. Advances in bone biology: the osteoclast. Endocr Rev 1996;17:308e32. [7] Hill PA. Bone remodelling. Br J Orthod 1998;25:101e7. [8] Roodman GD. Advances in bone biology: The osteoclast. Endocr Rev 1996;17:308e32. [9] Chen W, Li YP. Generation of mouse osteoclastogenic cell lines immortalized with SV40 large T antigen. J Bone Miner Res 1998;13:1112e23. [10] Kodama H, Nose M, Niida S, Yamasaki A. Essential role of macrophage colony-stimulating factor in the osteoclast differentiation supported by stromal cells. J Exp Med 1991;173:1291e4. [11] Yoshizawa T, Handa Y, Uematsu Y, et al. Mice lacking the vitamin D receptor exhibit impaired bone formation, uterine hypoplasia and growth retardation after weaning. Nat Genet 1997;16:391e6. [12] Simonet WS, Lacey DL, Dunstan CR, et al. Osteoprotegerin: a novel secreted protein involved in the regulation of bone density. Cell 1997;89:309e19. [13] Aubin JE, Bonnelye E. Osteoprotegerin and its ligand: A new paradigm for regulation of osteoclastogenesis and bone resorption. Medscape Womens Health 2000;5:5. [14] Quinn JM, Elliott J, Gillespie MT, Martin TJ. A combination of osteoclast differentiation factor and macrophage-colony stimulating factor is sufficient for both human and mouse osteoclast formation in vitro. Endocrinology 1998;139:4424e7. [15] Bekker PJHD, Rasmussen AS, Murphy R, et al. A single-dose placebo-controlled study of AMG 162, a fully human monoclonal antibody to RANKL, in postmenopausal women. J Bone Miner Res 2004;19:1059e66. [16] Rizzoli RYU, Kirkpatrick P. Denosumab. Nat Rev Drug Discov 2010;9:591e2. ¨ ber die Bildung des Knochengewebes. Natur[17] Gegenbaur C. U wissenschaften 1864. [18] Aubin JE, Liu F. The osteoblast lineage. In: Bilezikian JP, Raisz LG, Rodan GA, editors. Principles in Bone Biology. Academic Press; 1996. p. 51e67. [19] Franz-Odendaal TA, Hall BK, Witten PE. Buried alive: how osteoblasts become osteocytes. Dev Dyn 2006;235:176e90. [20] Friedenstein AY, Lalykina KS. Lymphoid cell populations are competent systems for induced osteogenesis. Calcif Tiss Res 1970;(Suppl):105e6. [21] Friedenstein AJ, Chailakhyan RK, Gerasimov UV. Bone marrow osteogenic stem cells: in vitro cultivation and transplantation in diffusion chambers. Cell Tiss Kinet 1987;20:263e72. [22] Burger EH, Boonekamp PM, Nijweide PJ. Osteoblast and osteoclast precursors in primary cultures of calvarial bone cells. Anat Rec 1986;214:32e40. [23] Aubin JE, Triffitt JT. Mesenchymal stem cells and osteoblast differentiation. In: Bilezikian JP, Raisz LG, Rodan GA, editors. Principles of Bone Biology. Academic Press; 2002. p. 59e82. [24] Dominici M, Le Blanc K, Mueller I, et al. Minimal criteria for defining multipotent mesenchymal stromal cells. The International Society for Cellular Therapy position statement. Cytotherapy 2006;8:315e7.

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[46] Dobnig H, Turner RT. Evidence that intermittent treatment with parathyroid hormone increases bone formation in adult rats by activation of bone lining cells. Endocrinology 1995;136:3632e8. [47] Parfitt AM. The cellular basis of bone turnover and bone loss: a rebuttal of the osteocytic resorption-bone flow theory. Clin Orthop Relat Res 1977;127:236e47. [48] Knothe Tate ML. Whither flows the fluid in bone? An osteocyte’s perspective. J Biomech 2003;36:1409e24. [49] Klein-Nulend J, van der PA, Semeins CM, et al. Sensitivity of osteocytes to biomechanical stress in vitro. FASEB J 1995;9:441e5. [50] Lanyon LE. Osteocytes, strain detection, bone modeling and remodeling. Calcif Tissue Int 1993;53(Suppl.1):S102e6. [51] Dunstan CR, Somers NM, Evans RA. Osteocyte death and hip fracture. Calcif Tissue Int 1993;53(Suppl.1):S113e6. [52] Frost HM. In vivo osteocyte death. J Bone Joint Surg Am 1960;42-A:138e43. [53] Kato Y, Windle JJ, Koop BA, Mundy GR, Bonewald LF. Establishment of an osteocyte-like cell line, MLO-Y4. J Bone Miner Res 1997;12:2014e23. [54] Bonewald L. Establishment and characterization of an osteocyte-like cell line, MLO-Y4. J Bone Miner Metab 1999;17:61e5. [55] Cheng BZ, Luo J, Sprague E, Bonewald LF, Jiang JX. Expression of functional gap junctions and regulation by fluid flow in osteocyte-like MLO-Y4 cells. J Bone Miner Res 2001;16:249e59. [56] Plotkin LI, Bellido T. Glucocorticoids induce osteocyte apoptosis by blocking focal adhesion kinase-mediated survival. Evidence for inside-out signaling leading to anoikis. J Biol Chem 2007;282:24120e30. [57] Kogianni GMV, Noble BS. Apoptotic bodies convey activity capable of initiating osteoclastogenesis and localized bone destruction. J Bone Miner Res 2008;23:915e27. [58] Bakker ADSV, Krishnan R, Bacabac RG, et al. Tumor necrosis factor alpha and interleukin-1beta modulate calcium and nitric oxide signaling in mechanically stimulated osteocytes. Arthritis Rheum 2009;60:3336e45. [59] Guo DKA, Guthrie J, Veno PA, Harris SE, Bonewald LF. Identification of osteocyte-selective proteins. Proteomics 2010;10:3688e98. [60] Cherian PP, Cheng B, Gu S, Sprague E, Bonewald LF, Jiang JX. Effects of mechanical strain on the function of Gap junctions in osteocytes are mediated through the prostaglandin EP2 receptor. J Biol Chem 2003;278:43146e56. [61] Jiang JX, Siller-Jackson AJ, Burra S. Roles of gap junctions and hemichannels in bone cell functions and in signal transmission of mechanical stress. Front Biosci 2007;12:1450e62. [62] Weinbaum S, Cowin SC, Zeng Y. A model for the excitation of osteocytes by mechanical loading-induced bone fluid shear stresses. J Biomech 1994;27:339e60. [63] Vatsa A, Mizuno D, Smit TH, Schmidt CF, MacKintosh FC, Klein-Nulend J. Bioimaging of intracellular NO production in single bone cells after mechanical stimulation. J Bone Miner Res 2006;21:1722e8. [64] Jørgensen NRGS, Civitelli R, Steinberg TH. ATP- and gap junction-dependent intercellular calcium signaling in osteoblastic cells. J Cell Biol 1997;139:497e506. [65] Genetos DCKC, Zhang Y, Yellowley CE, Donahue HJ. Oscillating fluid flow activation of gap junction hemichannels induces ATP release from MLO-Y4 osteocytes. J Cell Physiol 2007;212:207e14. [66] Robinson JA, Chatterjee-Kishore M, Yaworsky PJ, et al. Wnt/ beta-catenin signaling is a normal physiological response to mechanical loading in bone. J Biol Chem 2006;281:31720e8.

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[67] Hung CT, Pollack SR, Reilly TM, Brighton CT. Real-time calcium response of cultured bone cells to fluid flow. Clin Orthop Relat Res 1995:256e69. [68] Bonewald LF. The amazing osteocyte. J Bone Miner Res 2011;26:229e38. [69] Qing H, Bonewald LF. Osteocyte remodeling of the perilacunar and pericanalicular matrix. Int J Oral Sci 2009;1:59e65. [70] Bonewald LF. Osteocytes as dynamic, multifunctional cells. Ann NY Acad Sci 2007;1116:281e90. [71] Thompson DL, Sabbagh Y, Tenenhouse HS, et al. Ontogeny of Phex/PHEX protein expression in mouse embryo and subcellular localization in osteoblasts. J Bone Miner Res 2002;17: 311e20. [72] Nampei A, Hashimoto J, Hayashida K, et al. Matrix extracellular phosphoglycoprotein (MEPE) is highly expressed in osteocytes in human bone. J Bone Miner Metab 2004;22: 176e84. [73] Liu S, Zhou J, Tang W, Jiang X, Rowe DW, Quarles LD. Pathogenic role of Fgf23 in Hyp mice. Am J Physiol 2006;291:E38e49. [74] Feng JQ, Ward LM, Liu S, et al. Loss of DMP1 causes rickets and osteomalacia and identifies a role for osteocytes in mineral metabolism. Nat Genet 2006;38:1310e5. [75] Autosomal dominant hypophosphataemic rickets is associated with mutations in FGF23. Nat Genet 2000;26:345e8. [76] Winkler DGSM, Geoghegan JC, Yu C, et al. Osteocyte control of bone formation via sclerostin, a novel BMP antagonist. EMBO J 2003;22:6267e76. [77] Beighton P. Sclerosteosis. J Med Genet 1988;25:200e3. [78] Li X OM, Warmington KS, Morony S, et al. Sclerostin antibody treatment increases bone formation, bone mass, and bone strength in a rat model of postmenopausal osteoporosis. J Bone Miner Res 2009;24:578e88.

[79] Li X WK, Niu QT, Asuncion FJ, et al. Inhibition of sclerostin by monoclonal antibody increases bone formation, bone mass and bone strength in aged male rats. J Bone Miner Res 2010 Jul [Epub ahead of print]. [80] Tian XJW, Li X, Paszty C, Ke HZ. Sclerostin antibody increases bone mass by stimulating bone formation and inhibiting bone resorption in a hindlimb-immobilization rat model. Bone 2010 Sep; [Epub ahead of print]. [81] Agholme FLX, Isaksson H, Ke HZ, Aspenberg P. Sclerostin antibody treatment enhances metaphyseal bone healing in rats. J Bone Miner Res 2010;25:2412e8. [82] Noble BS, Peet N, Stevens HY, et al. Mechanical loading: biphasic osteocyte survival and targeting of osteoclasts for bone destruction in rat cortical bone. Am J Physiol Cell Physiol 2003;284; C934eC43. [83] Tatsumi SIK, Amizuka N, Li M, et al. Targeted ablation of osteocytes induces osteoporosis with defective mechanotransduction. Cell Metab 2007;5:464e75. [84] Kramer IHC, Keller H, Pegurri M, et al. Osteocyte Wnt/betacatenin signaling is required for normal bone homeostasis. Mol Cell Biol 2010;30:3071e85. [85] Kamel SAVP, Jiang J, Bonewald LF, Dallas SL. Comprehensive imaging of osteocytes and their dendrites in situ using widefield fluorescence, confocal microscopy and time lapse imaging. 31st Annual Meeting of the American Society for Bone and Mineral Research (abstract) 2009 Sep. [86] Veno PALY, Kamel SA, Feng JQ, Bonewald LF, Dallas SL. A membrane-targeted GFP selectively expressed in osteocytes reveals cell and membrane dynamics in living osteocytes. 32nd Annual Meeting of the American Society for Bone and Mineral Research (abstract) 2010 October 16.

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Bone Matrix and Mineralization Marc D. McKee 1, William G. Cole 2 1

Faculty of Dentistry, and Department of Anatomy and Cell Biology, McGill University, Montreal, Quebec, Canada 2 Divisions of Orthopaedics and Pediatric Surgery, The Stollery Children’s Hospital, and University of Alberta, Edmonton, Alberta, Canada

STRUCTURAL HIERARCHY IN BONE

intimate intercalation of nanocrystals of mineral with the organic scaffolding extracellular matrix [1]. In fact, it is precisely this nanoscale deposition of trillions of inorganic, tiny plate-like crystallites within a softer organic matrix that provides for the inherent flexibility of bones that allows them to be deformed during mechanical challenge, but then return to their original organization characteristic of any given skeletal element. Beyond the macroscale, bone tissue is largely organized as a mosaic of different connected geometric subunits. These are generally referred to as cylindrical osteons (Haversian systems), half-cylindrical hemiosteons, or generally as irregularly disposed intervening bone tissue having a variety of forms related to the forces placed upon it. These subunits are constructed together either loosely as cancellous (spongy) trabecular bone, or densely as compact, generally cortical bone. Collectively, these osseous subunits are “welded” together at their adhesive interfaces by so-called cement lines (really cement planes) [4]. The organic and inorganic components of cement lines likely ensure molecular bonding across this adhesive interface, safeguarding tissue cohesion during bone remodeling as defective, fatigued (microfractured) bone is resorbed, and new bone is deposited to fill the resorption site leaving an intervening cement line at this temporospatially distinct boundary [5]. The cell teams responsible for these alternating cycles of bone resorption followed by formation are called bone multicellular/metabolic units (BMUs) and consist of osteoblasts, osteocytes and osteoclasts that leave behind nascent bone packets bonded to the residual bone [6]. Collectively, this cellular activity e together with the abundant mineralized matrix e creates the necessary physiologic and functional properties of bone, and allow for its carefully orchestrated removal and replacement otherwise know as bone turnover. Figure 2.1 presents a montage of light and transmission

Remarkable in its structural complexity and function, bone tissue serves multiple purposes depending on its anatomic location and on its surrounding and attached tissues. Indeed, hierarchical organization of bone extending from the macroscale to the nanoscale provides for a variety of functions and biomechanical properties that result entirely from its basic structural elements and how they are assembled and interact with one another [1]. Bone tissue is a hard, tough and durable material, able to withstand substantial loads and impacts, and able to respond dynamically e through the actions of cells e to a variety of mechanical challenges. In doing so, the skeleton ultimately repairs and replaces itself throughout the lifespan. Further to this, bone tissue also serves as an important vital ion reservoir, harboring abundant quantities of key mineral ions such as calcium and phosphate which can be mobilized upon demand and released for systemic use by cells throughout the body, and for something no less important than providing nutrient mineral ions to breastfeeding infants during lactation [2]. Bone’s hardness and toughness come from the fact that it is a composite material, consisting primarily of a protein-rich, interconnected organic matrix network whose macromolecular assemblies are interwoven in a way that provides a scaffold for mineral deposition [1]. Mineralization of this extracellular matrix by calcium and phosphate complexation to form crystalline salts that harden bones (and teeth) is not haphazard, but in fact occurs in a way that is almost unimaginably controlled at the molecular level e indeed, like most cellular processes. The inorganic phase (mostly crystalline, as a substituted form of hydroxyapatite [3]) that forms in the skeleton provides exactly the required hardness, while at the same time ensuring requisite toughness by


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10

2. BONE MATRIX AND MINERALIZATION

(A)

(C)

(B)

(E)

(D)

FIGURE 2.1 Light (A) and transmission electron (BeE) micrographs showing osteoblast-lineage bone cells (osteoblasts, Ob; osteocytes, Oc), unmineralized bone extracellular matrix (Osteoid), and the mineralized matrix (MM) of bone. (A) Cuboidal osteoblasts align at forming bone surfaces, with some becoming incorporated into the bone as osteocytes. (B, C) Osteoblasts secrete a collagen fibril-rich layer of extracellular matrix known as the osteoid which in its deeper regions show patches of mineralized matrix. Mineralization becomes more confluent at the mineralization front where it permeates throughout the fibrillar and interfibrillar matrix compartments. (D, E) Some osteoblasts become trapped within the extracellular matrix eventually to become embedded within mineralized bone as osteocytes. Osteocyte cell processes extend into the mineralized bone matrix within small channels termed canaliculi (arrows).

electron micrographs illustrating the cells, matrix and mineral of bone, and their relationships at the ultrastructural level. The subsequent sections of this chapter provide details on the composition of the extracellular matrix of bone, with a section at the end providing a discussion of matrix mineralization, and how proteins in the matrix are thought to regulate this process.

ORGANIC MATRIX OF BONE This section focuses on the organic matrix which constitutes approximately 20e30% by weight of bone [7]. The organic matrix contains approximately 90% collagen (by weight) and 10% non-collagenous proteins, proteoglycans, and lipids; although on a molar basis,

the latter category roughly equals collagen. Plasma proteins are also present in the organic matrix. Many plasma proteins permeate bone deriving from its rich vasculature, but have little or no affinity for bone, whereas others such as a2-HS glycoprotein (fetuin A) have a higher affinity for the matrix and/or mineral of bone [7]. A description of the primary structure, synthesis, and assembly of the major macromolecules of bone follows. Major bone collagens are described first, followed by the small, leucine-rich, interstitial proteoglycans (SLRPs), and then by non-collagenous proteins of bone. Some macromolecules are not discussed, including the microfibrillar proteins, such as type VI collagen and the fibrillins, as well as the lipoproteins. Details concerning the latter macromolecules as well as other macromolecules of the extracellular matrix and further details about the

PEDIATRIC BONE

ORGANIC MATRIX OF BONE

described macromolecules can be found online at the National Center for Biotechnology Information (http://www.ncbi.nlm.nih.gov/), National Library of Medicine, National Institutes of Health.

Type I Collagen Type I collagen e the most abundant protein of bone e is also present in periosteum, perichondrium, ligaments, tendons, annulus fibrosus, menisci, dermis, sclera, dentin, fascia, and the adventitial layers of viscera [8,9]. It is one of the fibrillar collagens. The characteristic feature of the fibrillar collagens is that they contain a long, continuous triple helix that assembles into highly organized collagen fibrils [10]. The fibrils have high tensile strength, which is critical for the function of bone and other tissues. Each molecule of type I collagen is composed of two a1(I) chains and one a2(I) chain [10]. A small number of molecules contain three a1(I) chains. The importance of type I collagen in normal development is highlighted by the phenotypes that result from the homozygous or heterozygous loss of the a1(I) chains. Homozygous loss in the Mov 13 mouse, in which the a1(I) gene is inactivated by insertional mutagenesis, results in a prenatal lethal phenotype because of the lack of type I collagen in the tissues [11]. Heterozygous loss of one a1(I) allele, either in the Mov 13 mouse or in humans, yields the osteogenesis imperfecta type IA phenotype [12]. The a1(I) and a2(I) chains are encoded by the COL1A1 and COL1A2 genes, respectively. The COL1A1 gene is located on chromosome 17q21.3-q22 and the COL1A2 gene on chromosome 7q21.3-q22 [13,14]. Both genes have a similar structure, but because exons 33 and 34 are fused in COL1A1, the COL1A1 gene contains 51 exons, whereas the COL1A2 gene contains 52 exons [15,16]. The differences of 18 kb for COL1A1 and 38 kb for COL1A2 are attributable to differences in the sizes of their introns. For the pre-pro-a2(I) chain, the signal peptide is encoded by part of exon 1; the N-propeptide is encoded by part of exon 1, exons 2e5, and part of the junctional exon 6; the N-telopeptide is encoded by part of exon 6; the triple helix is encoded by part of exon 6, exons 7e48, and part of the junctional exon 49; the Ctelopeptide is encoded by part of exon 49; the C-propeptide is encoded by part of exon 49 and exons 50e52 [10,15]. The same arrangement exists for the pre-proa1(I) chain, except that the exon numbering is reduced by one beyond exons 33 and 34, which are fused in COL1A1 [10]. The exons encoding the main triple helix of the two chains have similar sizes [14]. Each exon encodes complete GlyeXeY triplets, where X and Y are often proline so that they commence with a codon for Gly

11

and end with a codon for Y. All exons encoding the triple-helical domain are 45, 54, 99, 108, or 162, base pairs (bps). It is likely that 54 bp was the ancestral exon size. The 108- and 162-bp exons arise from loss of introns. The 45- and 99-bp exons result from recombinations between two 54-bp exons [17]. The pre-pro-a1(I) chain contains 1464 amino acid residues [10,15]. It contains a signal peptide of 22 residues, an N-propeptide of 139 residues, an N-non-helical telopeptide of 17 residues, a main triple helix of 1014 residues, a C-non-helical telopeptide of 26 residues, and a C-propeptide of 246 residues. The mature tissue form of the a1(I) chain contains 1057 residues, including the main triple helix and the N- and C-telopeptides. The N-propeptide begins with a globular domain of 86 residues, including a von Willebrand type C repeat and 10 cysteine residues. It is followed by a triple-helical domain of 48 amino acids and a short globular domain. The procollagen N-proteinase cleavage site is at the Pro161eGln162 bond, numbered from the start of the signal peptide. Allysine cross-linking sites are located at residues 170 and 1208. There are two hydroxylysines that can be glycosylated at residues 265 and 1108. The mammalian collagenase cleavage site is at the Gly953e Ile954 bond. Proline residue 1164 may be 3-hydroxylated. There are two potential RGD (Arg-Gly-Asp) cell attachment sites at residues 745e747 and 1093e1095. Procollagen C-proteinase cleaves at Ala1218eAsp1219. The globular C-propeptide contains four cysteine residues at positions 1259, 1265, 1282, and 1291 that are involved in interchain disulfide bonding [18]. It also contains cysteine residues that are involved in intrachain disulfide bonding between residues 1299 and 1462 and between residues 1370 and 1415. There is also a putative Asn1365 site for attachment of an N-linked oligosaccharide. The pre-pro-a2(I) chain is shorter than the prepro-a1(I) chain, although the main triple-helical domains are the same size [16,19]. The pre-pro-a2(I) chain has a signal peptide of 22 amino acid residues, an N-propeptide of 57 residues, an N-non-helical telopeptide of 11 residues, a main helical domain of 1014 residues, a C-terminal non-helical telopeptide of 15 residues, and a globular C-propeptide of 247 residues. The very short globular domain of the N-propeptide contains only two cysteine residues. It is followed by a triple-helical domain of 42 residues and a second short globular domain. Procollagen N-proteinase cleaves at the Asn79eGln80 bond. Lysine residues 177 and 1023 are potential sites for hydroxylation and glycosylation. There are potential cell attachment sites at RGD sequences at positions 777e779, 822e824, and 1005e1007. The procollagen C-proteinase cleavage site is at the Ala1119eAsp1120 bond. Three sites for interchain disulfide bonds are at residues 1163, 1186, and 1195.

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Intrachain disulfide bonds may form between cysteine residues at positions 1203 and 1364 and between cysteine residues at positions 1272 and 1317. There is a potential N-linked oligosaccharide attachment site at Asn1267. The structures of the C-propeptide of human type III procollagen and the C-telopeptides of type I collagen have been described [20,21]. The structures of the Cpropeptides of types I and III procollagen are likely homologous. In contrast to the elongated structure of the C-terminal triple-helical region and the Ctelopeptide, the C-propeptide has a low-resolution structure composed of three large lobes and one small lobe [21]. This structure is readily interpretable in terms of the subunit composition and known positions of inter- and intrachain disulfide bonds. Among the eight cysteines found in the C-propeptide domains of type III procollagen, cysteines 1e4 are involved in interchain disulfide bonding, whereas cysteines 5e8 form intrachain disulfide bonds [22]. Consequently, it is likely that the three large lobes correspond to the intrachain disulfide-bonded region of each of the three polypeptide chains, whereas the small lobe corresponds to the junction region containing the interchain disulfide bonds and linking to the rest of the procollagen chain. Such an arrangement would place the chain-recognition region at the core of the structure, well positioned to determine the specificity of chainechain interactions [21]. The C-telopeptide of type I collagen had a hairpin conformation, with the C terminus folded back onto the triple helix [20]. After being transcribed, the pre-mRNA for the prepro-a1(I) and pre-pro-a2(I) chains undergoes exon splicing, capping, and the addition of a polyA tail, which gives rise to mature mRNAs. The mRNAs are translated in polysomes bound to the rough endoplasmic reticulum [23]. Formation of the triple-helical heterotrimeric type I procollagen molecule requires the synthesis of the constituent pro-a1(I) and pro-a2(I) chains from their distinct mRNAs. Transfer to the Golgi apparatus and movement into the secretory pathway require completion of both post-translational modifications and triple helix formation. The [pro-a1(I)]2[proa2(I)] heterotrimer is the major product, whereas stable [pro-a1(I)]3 homotrimer is formed in only small amounts. A [pro-a1(I)][pro-a2(I)]2 heterotrimer or a [pro-a2(I)]3 homotrimer have not been detected in cell cultures or tissues. Efficient type I procollagen heterotrimeric assembly appears to require that the elongating nascent pro-a1(I) and pro-a2(I) chains be inserted into the same compartments of the rough endoplasmic reticulum and that a mechanism for chain selection be operational within the compartment. Coordinated expression of the COL1A1 and COL1A2 genes is one way of ensuring

that the appropriate 2:1 ratios of the protein chains are available for the assembly of heterotrimers [24e26]. However, the ratios of the pro-a1(I):pro-a2(I) mRNAs vary widely, even though the resultant heterotrimers maintain the protein chain ratio of 2:1 [27]. Consequently, chain selection probably plays an important role in molecular assembly [28]. In support of the latter proposal, distinct chain recognition sequences of 15 amino acid residues have been identified within the Cpropeptides of the type I procollagen chains [29]. The interactions between the C-propeptides that lead to registration of nascent pro-a1(I) chains occur while the chains are still associated with polysomes [30e32]. Nascent collagen chains are translated on complex polyribosomal aggregates associated with more than one strand of mRNA. Consequently, it is likely that the organization of the mRNA translocons is a critical factor in molecular assembly [23]. Elongation pauses are also a prominent feature of synthesis of pro-a1(I) and proa2(I) chains [33]. There also appears to be an interaction between the translation complexes for the synthesis of the two chain types [28]. These interactions may serve two purposes: to bring the translation complexes to the same regions of the endoplasmic reticular membrane to ensure the co-localization of the nascent chains for interaction and to regulate or coordinate the rates of synthesis of the two pro-a chains. In many systems producing type I collagen, the pro-a1(I):pro-a2(I) ratio is greater than 2:1 [34]. In such situations, it is likely that all of the pro-a2(I) mRNA is engaged in heterotrimer synthesis, whereas the excess pro-a1(I) chains may either produce [pro-a1(I)]3 trimer or be degraded [35]. The signal peptides are removed as the nascent chains enter the rough endoplasmic reticulum. Both procollagen chains undergo hydroxylation and glycosylation of specific residues as a prerequisite for the formation of a stable triple-helical heterotrimer. Many of these enzymatic modifications occur as co-translational events. Approximately 100 proline residues in the Y position of GlyeXeY repeats undergo 4-hydroxylation by prolyl 4-hydroxylase. A single proline residue in the X position undergoes 3-hydroxylation by prolyl 3hydroxylase. A variable number of lysine residues, also in the Y position, may undergo lysyl hydroxylation by lysyl hydroxylase. Hydroxylation of proline residues to hydroxyproline is critical for the formation of a stable triple helix. The hydroxylases require the procollagen chains to be in a nascent state and various co-factors need to be present, such as ferrous ions, molecular oxygen, a-ketoglutarate, and ascorbate [36]. Prolyly 4-hydroxylase, also called procollagen proline 2 oxoglutarate 4-dioxygenase, is a tetramer consisting of two a and two b subunits with a molecular weight of 240 kDa. The b subunit is also known as protein

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disulfide isomerase. The gene for the b subunit, P4HB, is located on chromosome 17q25 [37]. The gene is expressed ubiquitously. There are two a subunits whose genes are named P4HA1 and P4HA2 [38]. The P4HA1 gene is located on chromosome 10q21.3-23.1 [39], and the P4HA2 gene is located on chromosome 5q31 [38]. The mRNAs encoded by P4HA1 and P4HA2 are expressed ubiquitously, although the ratios of the mRNAs vary between tissues. There are two a (I) subunit mRNA isoforms that result from mutually exclusive alternative splicing of exons 9 or 10 of P4HA1 [40]. Tetramers containing either two a (I)/two b subunits or two a1(II)/two b subunits have similar enzyme activities [38]. Co-expression of recombinant P4HA1, P4HA2, and P4HB in insect cells did not show any tetramers that contained both P4HA1 and P4HA2 protein chains [41]. Proline 986 of the triple helical region of pro a1(I) chain is the single proline residue that undergoes 3hydroxylation. Cartilage-associated protein (CRTAP), encoded by CRTAP, forms a complex with prolyl-3hydroxylase-1 (P3H1), encoded by LEPRE1 and the prolyl cis-trans isomerase cyclophilin B (CyPB), encoded by PPIB that is required for type I collagen 3-prolyl hydroxylation [42]. There appear to be at least four lysyl hydroxylases. The enzyme is also called procollagen-lysine 2 oxoglutarate 5-dioxygenease. Lysyl hydroxylase 1, an 85-kDa membrane-bound homodimeric protein, is localized to the cisternae of the rough endoplasmic reticulum [43]. It hydroxylates specific lysine residues in XeLyseGly sequences. Its gene, PLOD1, is located on chromosome 1p36.3-p36.2 [43]. It is expressed in the skeleton and in many other tissues. Lysyl hydroxylase 2 is encoded by PLOD2, which is located on chromosome 3q23-q24 [44]. It also forms homodimers. It is highly expressed in non-skeletal tissues as well as in bone [45]. Lysyl hydroxylase 2 modifies lysyl residues in the telopeptides of type I collagen involved in intermolecular cross-link formation. Lysyl hydroxylase 3 is encoded by PLOD3, which is located on chromosome 7q36 [46,47]. It is also expressed mostly in non-skeletal tissues. There is also a putative telopeptide lysyl hydroxylase that is specific for bone. The gene called either BRKS or TLH1 was predicted from linkage studies of patients with Bruck syndrome type I to be located on chromosome 17p12 [48]. Recently, FKBP10, which encodes FKBP65, which has prolyl cis-trans isomerase activity, was identified as a possible cause of some cases of Bruck syndrome type I e the gene is located on chromosome 17q21.2 [49]. When lysyl residues become hydroxylated, they may serve as a substrate for a glycosyltransferase and for a galactosyltransferase, which add glucose and galactose, respectively, to the hydroxyl group. These modifications also require the chains to be in the nascent

13

state. Increased levels of glycosylation tend to reduce the size of the collagen fibrils presumably caused by interference with packing of the molecules. A mannose-rich oligosaccharide may also attach to an asparagine residue in the C-propeptide. Chaperones play an important role in the assembly of procollagens in the rough endoplasmic reticulum. Protein disulfide isomerase, which has both enzymatic and chaperone functions, interacts transiently with procollagen chains early in the procollagen assembly pathway [50,51]. Release of prolyl 4-hydroxylase, including its b-subunit protein disulfide isomerase, from the triple-helical domain coincides with the assembly of thermally stable triple-helical molecules. However, if triple helix formation is prevented, prolyl 4-hydroxylase remains associated with the triple-helical domain, suggesting a role for the enzyme in preventing aggregation of this domain. Protein disulfide isomerase is also able to associate independently with the C-propeptide of monomeric procollagen chains prior to trimer formation, indicating a role for this isomerease in coordinating the assembly of heterotrimeric molecules [50]. It is also able to catalyze disulfide bond formation within and between the C-propeptides. HSP47, also called collagen-binding protein 2, is a heat shock protein that resides in the endoplasmic reticulum [52]. It interacts transiently with procollagen during its folding, assembly, and transport from the endoplasmic reticulum of mammalian cells. It has been suggested to carry out a diverse range of functions, such as acting as a molecular chaperone facilitating the folding and assembly of procollagen molecules, retaining unfolded molecules within the endoplasmic reticulum, and assisting the transport of correctly folded molecules from the endoplamic reticulum to the Golgi apparatus. The association of HSP47 with procollagen coincides with the formation of a collagen triple helix. HSP47 may form a chaperone complex with FK506binding protein 10 (FKBP10) that interacts with procollagen [49]. The importance of HSP47 in normal type I collagen biosynthesis is highlighted by the phenotype of mice lacking the heat shock protein [52]. The homozygous mice did not live longer than 11.5 days, and their tissues were deficient in collagen fibrils. The findings indicated that type I collagen was unable to form a rigid triple-helical structure without the assistance of HSP47 and that HSP47 was essential for normal development. HSP47 is encoded by SERPINH1 which is located on chromosome 11q13.5 [53]. The C-propeptides undergo registration and stabilization by the formation of interchain and intrachain disulfide bonds in the rough endoplasmic reticulum. The formation of the disulfide bonds is catalyzed by the enzyme protein disulfide isomerase, which is also the b-subunit of prolyl 4-hydroxylase. This process

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is critical for the correct alignment of the main triplehelical domain because the helix winds up from the C terminus. Triple helix formation is initiated in the rough endoplasmic reticulum immediately after the synthesis of the pro-a chains and after the formation of the interchain disulfide bonds within the C-propeptide [54,55]. It is likely that formation of the triple helix is a posttranslational event because the production of triplehelical molecules requires approximately 8 or 9 minutes after completion of the synthesis of the pro-a chains. The helix propagates from a single C-terminal nucleation site toward the N terminus and is interrupted by the random occurrence of peptide bonds in the cis configuration. The C-propeptide and C-telopeptide do not appear to play a role in nucleation of triple helix formation [56]. However, a minimum of two hydroxyproline-containing GlyeXeY triplets at the C-terminal end of the triple helix are required for nucleation to occur [56]. Direct nuclear magnetic resonance measurements of chick calvarial collagen showed that approximately 16% of the XePro and 8% of XeHyp bonds were cis in the unfolded collagen [57]. Many studies have shown that the rate-limiting step in the zipper-like propagation of the helix is the process of cis-trans isomerization [54,55,57,58]. Peptidyl prolyl cis-trans isomerase catalyzes the cis-trans isomerization of XePro peptide bonds in collagen [59]. FKBP65, encoded by FKBP10, also has prolyl cis-trans isomerase activity [60]. Protein disulfide isomerase is also present in the rough endoplasmic reticulum. However, protein disulfide isomerase does not appear to act as a cisetrans isomerase [59]. Full hydroxylation of proline residues in the Y position of GlyeXeY triplets also enhances the rate of propagation of the triple helix from the site of nucleation to the N terminus [61]. Biophysical studies using model collagen peptides have also shown that the folding of GlyeXeY peptides is best described as an all-or-none, third-order reaction [62e64]. The formation of the triple helix occurs in the rough endoplasmic reticulum. The type I procollagen molecules move to the Golgi apparatus, where oligosaccharides may be added to a C-propeptide asparagine residue. The molecules are secreted from the cell, during which (or soon after) the N- and C-propeptides are rapidly cleaved. The N-propeptide is specifically cleaved by procollagen 1 N-endoproteinase. The enzyme is encoded by the ADAMTS2 gene, which is an abbreviation for a disintegrin-like and metalloproteinase with thrombospondin type I motif [65]. The enzyme exists in a long and a short form as a result of alternative splicing [66]. The C-propeptide is cleaved by procollagen C-endoproteinase, which is the same as bone morphogenetic protein-1 (BMP-1) [67]. The gene is located on chromosome 8q21 [68]. Procollagen C-endoproteinase is a secreted, neutral zinc metalloproteinase. The Drosophila equivalent gene

is called tolloid (TLD). There are two isoforms of the human enzyme as a result of alternative splicing [69]. The long form appears to be an inactive proenzyme that can be activated by removal of the pro-domain. There are four mammalian BMP-1/TLD-like proteases [70]. One of them, tolloid-like-1 (TLL1), is also an astracin-like metalloproteinase. Its gene, TLL1, is located on chromosome 4q32-q33 [70]. The activity of procollagen C-endoproteinase is enhanced by procollagen C-endopeptidase enhancer, which is a glycoprotein that binds to the C-propeptide and enhances the activity of the Cproteinase enzyme [71]. Its gene, PCOLCE, is located on chromosome 7q21.3-q22 approximately 6 Mb from the COL1A2 gene that encodes pro-a2(I) chains of type I procollagen [72]. A second procollagen C-endopeptidase enhancer has been isolated. Its gene, PCOLCE2, is located on chromosome 3q21-q24 [73]. A number of functions have been proposed for the released a1(I) N-propeptide, including prevention of premature intracellular molecular association, facilitation of transcellular transport and secretion, regulation of extracellular fibrillogenesis, and feedback regulation of procollagen synthesis [74]. However, there is little evidence to support these proposals. For example, deletion of exon 2 of COL1A1 in mice, which deleted the 65-amino acid cysteine-rich globular domain of the N-propeptide of pro-a1(I) chains, did not produce any demonstrable anomalies in type I collagen biosynthesis, collagen cross-linking, or collagen fibrillogenesis [74]. Following removal of the N- and C-propeptides, the type I collagen molecules can self-assemble, cross-link, undergo further growth, and pack into thick collagen fibers. The nucleation steps that initiate the formation of collagen fibrils may commence within crypts on cell surfaces [75,76]. Various biophysical studies as well as rotary shadowing electron microscopy have shown that type I collagen monomers are rod-like structures with a length of approximately 300 nm and a diameter of approximately 1.4 nm. The overall helical symmetry is left-handed, with 10 residues in three turns and a pitch of approximately 3 nm. The three helical chains are further coiled about a central axis to form a right-handed helix with a repeat distance of approximately 10 nm [77,78]. The high content of glycine and its occurrence in every third residue of the triple-helical domain give rise to a polymer of tripeptide units with the formula (eGlyeXeYe)n. Glycine is the smallest amino acid; as such, it is the only amino acid that can pack tightly at the center of the triple-stranded collagen fibril monomers. The side chains of amino acids in the remaining eXe and eYe positions protrude from the chain, and this arrangement allows a variety of amino acid residues to be accommodated in the molecule. The high imino acid content, particularly the high 4-hydroxyproline

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content, has a stabilizing effect on the triple-helical structure. In type I collagen, the triple-helical configuration occurs throughout 95% of the rod-like monomer. The N- and C-telopeptides do not contain glycine residues at every third position. The long triple-helical domain not only provides the molecule with the stability required for its biomechanical functions but also makes it resistant to enzymatic cleavage apart from specific peptide bonds that can be cleaved by mammalian collagenases. The collagen monomers are able to undergo spontaneous self-assembly into fibrils. The fibrils are crossstriated as a result of the assembly of molecules in a parallel array but with a stagger of approximately one-quarter of their length. The periodicity of the cross-striated fibril is a result of each monomer having five highly charged regions at approximately 67-nm intervals. The repeat period e called a D period e is approximately 67 nm in length and contains 234 amino acids. The overall length of the collagen fibril monomer is 4.4 D units, which also corresponds to 300 nm. Because of the non-integral length of the monomers, overlapping by D divides the fibril into overlap zones that include the N- and C-termini of the molecules and gap zones that do not. The quarter-stagger arrangement of the collagen molecules provides the appropriate substrate conformation for the action of lysyl oxidase. The enzyme, which is a copper-dependent amine oxidase, requires molecular oxygen for activity. It acts on specific lysine and hydroxylysine residues to produce the corresponding aldehydes that are required for the formation of covalent collagen cross-linkages. The enzyme, which is also called protein-lysine 6-oxidase, is encoded by LOX, which is located on chromosome 5q23.3-q31.2 [79]. Alternative splicing produces three mRNA isoforms [79]. There are also four lysyl oxidase-like loci. LOXL1 is located on chromosome 15q22, LOXL2 on chromosome 8p21-p21.2, LOXL3 on chromosome 2p13.3, and LOXL4 on chromosome 10q24 [80,81]. The lysine aldehyde pathway occurs primarily in adult dermis, cornea, and sclera, whereas the hydroxylysine aldehyde pathway predominates in bone, ligaments, tendons, and embyronic dermis. The first step in both pathways is the oxidative deamination of the 3-amino group in telopeptide lysine and hydroxylysine residues to form their corresponding aldehydes, called allysine and hydroxyallysine, respectively. In the lysine aldehyde pathway, two allysines may condense to form the aldol condensation product, which forms intramolecular bonds. Aldimine cross-links are formed when allysine in the telopeptides reacts with lysine or hydroxylysine residues in adjacent helices to provide covalent intermolecular cross-linkages. In the hydroxylsine aldehyde pathway, hydroxyallysine can condense with

15

a hydroxylysine residue to form a reducible cross-link that can undergo an Amadori rearrangement to form hydroxylysino-5-oxo-norleucine. The hydroxylysinederived aldimine cross-links can also occur as galactosyl or glucosylgalactosyl derivatives. In the hydroxylysine aldehyde pathway, the major mature cross-link is based on trivalent 3-hydroxypyrididinium residues [82]. It includes hydroxylysylpyridinoline, derived from three hydroxylysine residues, and lysylpyridinoline, derived from two hydroxylysine residues and one lysine residue. These two cross-links are naturally fluorescent and can be assayed directly in tissue hydrosylates as well as in blood and urine. Insights into the importance of posttranslational modifications and preferred intermolecular binding partners for telopeptide and helical cross-linking domains in regulating cross-link type and placement are being obtained using ion-trap mass spectrometry and peptide-specific antibodies [83]. The formation of collagen fibrils has been studied extensively. Of particular interest are in vitro studies that use intact procollagen as well as procollagen lacking either the N-propeptide (pC) or C-propeptide (pN) with fibril formation initiated by the addition of specific Nand C-proteases [84,85]. This approach leads to the formation of fibril-like structures with the characteristic collagen D-periodic banding pattern but with a distinctive, bipolar needle-like morphology that is different from that of fibrils isolated from native tissue [76,85]. These fibril structures show a single polarity reversal where the orientation of the collagen is reversed with amino termini at both ends of the fibrils [85]. Newly formed fibrils with characteristic D periodicity have been isolated from various embyronic tissues [86]. These fibrils also frequently show a single polarity reversal [87]. Fibrils may increase in size by the fusion of small (1e10 mm) segments, and the lateral association of long or short fibrils may lead to thicker fibrils [86,88]. The growth of type I collagen fibrils appears to be partly regulated by other collagen molecules that are included within the heterotypic fibers [89]. For example, other collagens in the heterotypic type I collagen fibrils of dermis include types III, V, XII, and possibly XIV collagen [90]. In bone, the other collagens are type V and type V/XI hybrid molecules [91]. It is likely in these various types of heterotypic type I collagen fibrils that the N-propeptides that remain attached to the collagens, other than type I collagen, play a role in regulating the growth of the fibrils [90]. A number of other molecules, including the small leucine-rich proteoglycans such as decorin, fibromodulin, lumican, as well as hyaluronan, also appear to regulate the growth of fibrils [92]. The molecular packing of collagen fibrils has been determined mainly in tissues such as tendon. X-ray diffraction studies indicate the presence of three-

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dimensional crystallinity admixed with liquid-like lateral order [93,94]. The lateral unit cell, which contains five molecules in cross-section, gives rise to row lines with a maximum spacing of 3.8 nm. Electron microscopy of a transverse section of tendon fibrils reveals a similar periodicity (z4 nm) orientated radially with respect to the fibril center. A feature of the model is that molecules are tilted obliquely in a plane oriented 30 to the fibril surface [94], which results in the helicoidal organization of collagen fibrils. An additional feature of the model is that the fibril surface is coated in molecular ends, which has important consequences for fibril growth. For example, the persistence of the N-propeptides of types III, V, or XI collagen might prevent their incorporation into the center of the fibril, thereby forcing all N termini to the surface of the fibril, with prevention of further accretion and limiting fibril diameter [93]. An alternative molecular packing model is the fivestranded Smith microfibril [94]. The microfibril, with a diameter of approximately 4 nm, is the minimum filamentous structure that possesses an axial D repeat. Although their existence is still debated, evidence indicates that they do exist [93]. Three-dimensional image reconstructions of 25-nm diameter collagen fibrils show evidence of a 4-nm repeat in transverse section, which might correspond to ordered arrays of microfibrils, particularly at the level of gapeoverlap junctions. Bone and other connective tissues have distinctive collagen fiber sizes and distinctive suprafibrillar architectures, as seen by polarized light microscopy [95]. In woven bone, the collagen fibrils are generally randomly distributed. Lamellar bone contains collagen fibrils that are arranged in parallel layers or sheets running in different directions. Bone osteons have a lamellar structure in which the lamellae are arranged in concentric cylinders.

Type V Collagen Type V collagen was first identified in human placenta and dermis, but later studies showed that it is widely expressed in type I collagen-containing tissues including bone [96,97]. The type V collagen molecules exist as heterotrimers a1(V)2a2(V) a1(V) a2(V) a3(V) and as homotrimers a1(V)3 [98,99]. An apparently distinct a4(V) chain is synthesized by Schwann cells [100]. Type V collagen chains also form heterotypic molecules with type XI collagen chains. For example, the highly homologous a1(V) and a1(XI) chains may yield an a1(V) a1(XI) a2(V) trimer in bone and cartilage [91]. The following description of type V collagen is limited to the a1(V) and a2(V) chains because the a3(V) and a4(V) genes are not expressed in bone, although the a3(V) gene is expressed in ligament attachments to bone [101].

The gene for the a1(V) chain, COL5A1, is located on chromosome 9q34.2-q34.3 [102]. The gene has 66 exons, more than the number of exons in type I and II collagens. Exon 1 encodes the signal peptide of 36 amino acid residues and one base of the N-propeptide. Exons 2e14 encode the remainder of the N-propeptide, with exon 14 being a junctional exon that encodes the end of the N-propeptide and the beginning of the triple-helical domain. The N-propeptide contains 505 amino acid residues and the N-telopeptide contains 17 residues. The pro-a1(V) N-propeptide is similar in size and domain structure to the N-propeptides of the proa1(XI) and pro-a2(XI) chains [103e105]. The Npropeptides of the latter three chains all contain a very large globular domain, immediately downstream of the signal peptide, that is much larger than, and has no apparent homology to, the cysteine-rich globular domains of type IeIII collagens. The globular domains of the pro-a1(V), pro-a1(XI), and pro-a2(XI) chains are bisected by a cluster of two cysteines into a basic Nterminal subdomain and a C-terminal subdomain rich in acidic residues and tyrosines [103e106]. The latter region contains 27 tyrosine residues and 73% of the total number of tyrosine residues in the pre-pro-a1(V) chain. Approximately 40% of the N-propeptide tyrosine residues are sulfated [107]. The N-terminal subdomain is analogous to the thrombospondin 1 motif and is identical to the proline- and arginine-rich peptide (PARP) motif [108]. The derived three-dimensional structure of PARP suggests a conserved nine b-stranded structure [109,110]. The PARP motif is also present in the N-propeptides of the pro-a1(XI) and pro-a2(XI) chains but not in the pro-a2(V) chains. The globular domain of the N-propeptide of the pro-a1(V) chain is followed by an interrupted triple-helical domain of 25 GlyeXeY repeats and then another short non-collagenous region prior to the main triple-helical domain. Pro-a1(V), proa1(XI), and pro-a2(XI) sequences share similarities in all subdomains of their N-propeptides, with the exception of the acidic globular variable region [104]. There is no evidence of alternative splicing in the Npropeptide of pro-a1(V) chains, although it occurs in the pro-a1(XI) and pro-a2(XI) N-propeptides [103]. There are two hypothetical N-proteinase cleavage sites (Ala-Gln) at positions 541e542 and 546e547. The main triple-helical domain of a1(V) chains is similar to those of type IeIII collagens in that each non-junctional exon begins with a complete codon for glycine and ends with a complete codon for a residue in the Y position of GlyeXeY triplets [102,111]. The codons are commonly 45 or 54 bps. The main triplehelical domain is encoded by exons 14e62 [102]. These exons encode a main triple-helical domain of 1014 amino acid residues. Lysine residues at positions 642 and 1482, which are important for intermolecular

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cross-linkages, are preserved. The helical domain lacks the Gly-Ile/Leu mammalian cleavage site at position 775e776 of type IeIII collagens [112]. The absence of this sequence explains the lack of cleavage of the a1(V) chains by mammalian collagenase. Carboxyl terminal of the main triple helix is the C-telopeptide of 33 amino acid residues and the Cpropeptide of 233 residues. They are encoded by exons 62e66. The putative C-proteinase cleavage site is located at position 1605e1606 (AlaeAsp). The C-propeptide has a high homology with the same region of pro-a1(XI) chains. Seven cysteine residues and their vicinities are conserved. The pro-a chains that can form homotrimers or both homo- and heterotrimers of pro-a chains have eight cysteine residues, but the pro-a chains that form heterotrimers only have seven cysteine residues [17]. The C-propeptide of pro-a1(V) chains has eight cysteine residues, whereas the C-propeptide of the pro-a1(XI) chains has seven cysteine residues [113]. The pro-a1(V) chain contains numerous sites for potential attachment of N-linked oligosaccharides. It has a heparin binding site at residues 897e929 [114]. Heparin binding sites are also present in a1(XI) and a2(XI) chains but not a2(V) chains [115]. Binding of extracellular chondroitin sulfate E to type V collagen may facilitate cell binding and matrix assembly [116]. There are RGD cell attachment sites at positions 645e647 and 663e665. The gene for the a2(V) chain, COL5A2, resides on chromosome 2q14-q32 [117,118]. The gene, which spans approximately 67 kb, is located in a tail-to-tail orientation with the COL3A1 gene. The intergenic distance is approximately 22 kb. The two genes contain 51 exons and share almost identical structures. COL5A2 encodes a pre-pro-a2(V) chain of 1496 amino acid residues [119,120], and it includes a signal peptide of 26 residues. The pro-a2(V) N-propeptide is encoded by four complete exons and partially by the junctional exon. The N-propeptide of 167 residues includes an aminoterminal globular subdomain of 82 residues, in which there is a von Willibrand type C-like repeat. This subdomain contains a central cluster of 10 cysteine residues flanked on both sides by short, hydrophilic sequences. It is similar to the equivalent subdomain of the Npropeptide of pro-a1(I) chains but dissimilar to the equivalent subdomain of the pro-a1(V) chain [111,121]. The N-propeptide also includes an interrupted helical subdomain of 78 residues and a non-helical subdomain of seven residues. The N-proteinase cleavage site is at the AsneGln bond at position 193e194 of the full-length chain. The N-telopeptide contains 15 residues, including the lysine cross-linking site at residue 175. The main triple-helical domain contains 1017 residues and is followed by a C-telopeptide of 26 residues and a Cpropeptide of 246 residues. The C-proteinase cleavage

17

site is located at the GlyeAsp bond at position 1250e1251. There are seven RGD sequences that are potential cell binding sites. There are several sites for the attachment of N-linked oligosaccharides. Also, there are three cysteine residues, at positions 1293, 1299, and 1325, for intermolecular cross-linking and sites for intrachain disulfide cross-links at positions 1333e1494 and 1402e1447. In contrast to type I procollagen processing, in which the N- and C-propeptides are rapidly cleaved following secretion, type V procollagen molecules retain their Npropeptides. A variety of type V collagen molecules retaining all or parts of the N-propeptides have been extracted from tissues and from tissue culture medium [109,122,123]. Rotary shadowing confirmed the retention of the N-propeptides on some of the type V collagen molecules extracted from tissues [124]. Nonetheless, within pro-a1(V)2pro-a2(V) heterotrimers, some proa1(V) N-propeptides and pro-a2(V) C-propeptides are enzymatically removed by bone morphogenetic protein-1-like enzymes. Pro-a1(V) C-propeptides are processed by furin-like proprotein convertases in vivo [125]. When type V collagen epitopes are unmasked in tissues, this collagen is found to co-localize with type I collagen [126]. The exact spatial relationship between these collagens in the type I collagen fibrils is unclear. However, co-polymeric assembly is likely because cross-linkages between types I and V collagens have been isolated from bone [127]. It is also likely that type V collagen regulates the formation of the type I collagen fibrils [128]. Type V collagen molecules are capable of forming homotypic fibrils in vitro with or without an apparent 67-nm cross-striation pattern [129]. Increasing the quantity of type V collagen relative to type I collagen decreased the final fibril diameter [130]. It has been proposed that the retained N-propeptides of type V collagen molecules protrude from the surface of the type I collagen fibrils, where they regulate the growth of the fibrils [89,130]. Confirmation of the importance of the amino-terminal extension of the a2(V) chain was provided by the phenotype of mice in which the COL5A2 gene was engineered to lack exon 6, which normally encodes the N-telopeptide of a2(V) chains. The mice showed abnormal type I collagen fibrillogenesis in the dermis [131]. Although type V collagen, with the exception of its N-propeptides, is buried within the type I collagen fibrils, its main triple-helical domain is known to bind to thrombospondin, heparin, heparan sulfate, decorin, and biglycan [114,132e135]. Type V collagen contains seven RGD sequences on the a2(V) and two on the a1(V) chains that may enable type V collagen to attach to various cell types. These interactions may involve the a1b1 and a2b1 integrins [136].

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Small, Leucine-rich Interstitial Proteoglycans The small, leucine-rich interstitial proteoglycans (SLRPs) are a large family of extracellular matrix glycoproteins/proteoglycans that share a leucine-rich repeating structural motif [137]. Many of the family members bind to various collagens and to growth factors such as TGF-b1 [138]. The gene family can be divided into subfamilies based on similarities in amino acid sequences and gene organization. The type I subfamily includes decorin and biglycan, which contains an N-terminal domain substituted with chondroitin or dermatan sulfate chains. These proteoglycans show 57% protein sequence identity and are encoded by genes composed of eight exons with exon/intron junctions in conserved positions [139,140]. Fibromodulin and lumican constitute the type II subfamily and exhibit 48% protein sequence identity. Their genes are composed of three exons with conserved exon/intron junctions [141,142]. Other members of the latter family include the proline- and arginine-rich and leucine-rich repeat protein (PRELP) and osteomodulin. The class III SLRP expressed in cartilage is epiphycan, also called PG-Lb [143]. Chondroadherin represents its own subfamily, with a different gene organization and a different amino acid composition [144]. The leucine-rich repeating extracellular glycoproteins/proteoglycans have core proteins of 32e42 kDa [145]. The protein chains can be divided into N-terminal, central, and C-terminal domains. The N-terminal domains are least conserved, but in all members of the family they contain four Cys residues, which form intrachain disulfide bonds. The glycosaminoglycan chains in decorin and biglycan are O-glycosidically linked to Ser residues in the N-terminal region, providing polyanionic properties to the proteoglycans. In contrast, the N-terminal domains of fibromodulin and lumican carry clusters of negatively charged Tyr sulfate residues. The different leucine-rich repeat proteoglycans and glycoproteins have similar C-terminal domains, which contain approximately 50 amino acid residues. This domain contains two Cys residues involved in an intrachain disulfide bond, leading to the formation of a 34- to 41-residue loop. The common central domain constitutes approximately 60e80% of the total protein. In most of the members of the family, it contains 10 or 11 repeats of a 20- to 25-residue-long leucine-rich motif, with Asn and Leu residues in conserved positions. Each leucine-rich repeat contains the motif LXXLXLXXNXL, with each motif being separated by nine to 18 amino acids. Up to 30 adjacent leucine-rich repeats have been described in some leucine-rich repeat proteins. The leucine residues in the motif may be replaced by alanine, valine, isoleucine, phenylalanine, tyrosine, or methionine. The asparagine

residue at position 9 may be replaced by cysteine or threonine. There are consensus site Asn residues in the central repeat domain for substitution with carbohydrates. For example, the latter Asn sites are partially substituted with keratan sulfate in fibromodulin and lumican. The three-dimensional structure of one member of the family, ribonuclease inhibitor, showed that the leucinerich repeats form a horseshoe-shaped coil of parallel, alternating a-helices and b-sheets stabilized by interchain hydrogen bonds [146]. Results of structural studies of decorin and biglycan are in accordance with the latter x-ray crystallographic findings [147]. Decorin, fibromodulin, and lumican bind to fibrillar collagens in vitro, leading to delayed fibril formation and the formation of thinner fibrils [148]. These changes are likely attributable to binding of the leucine-rich repeat glycoproteins/proteoglycans to the surface of the axially growing fibril, which inhibits the incorporation of additional triple-helical collagen monomers [85]. Binding of the leucine-rich repeat proteoglycan to collagen alters the surface properties of the fibrils and may affect the interactions between individual collagen fibrils as well as between the fibrils and the matrix. Competitive binding and displacement of proteoglycans may regulate the growth of collagen fibrils during skeletal development [149]. Decorin and fibromodulin bind to distinct and apparently separate sites in the gap region of the D period of the collagen fibril in vivo [150,151]. Decorin has been shown to bind to a small region of the C-terminus of the triple-helical domain of a1(I) chains, close to one of the intermolecular crosslinking sites [152]. This region corresponds to the c1 band of the collagen fibril D period. Proteoglycan core proteins of decorin, biglycan, and fibromodulin, prepared as fusion proteins, each bound TGF-b1 [153]. There was negligible binding to several other growth factors. Intact decorin, biglycan, and fibromodulin, isolated from bovine tissues, competed with the fusion proteins for TGF-b1 binding. Affinity measurements suggest a two-site binding model with KD values ranging from 1 to 20 nM for the high-affinity binding site and from 50 to 200 nM for the low-affinity binding site. Stoichiometry indicated that the highaffinity binding site was present in one of 10 proteoglycan core molecules and that each molecule contained a low-affinity binding site. Tissue-derived biglycan and decorin were less effective competitors for TGF-b binding than fibromodulin or the non-glycosylated fusion proteins. Removal of the chondroitin/dermatan sulfate chains of decorin and biglycan (fibromodulin is a keratan sulfate proteoglycan) increased the activities of decorin and biglycan, suggesting that the glycosaminoglycan chains may hinder the interaction of the core proteins with TGF-b. The fusion proteins competed for the binding of radiolabeled TGF-b to Mv1Lu cells and

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endothelial cells. Affinity labeling showed that the binding of TGF-b to betaglycan and the type-I receptors in Mv1Lu cells and to endoglin in endothelial cells was reduced, but the binding to the type II receptors was unaffected. Latent recombinant TGF-b1 precursor bound slightly to fibromodulin and not at all to decorin and biglycan. These findings indicate that the three decorin-type proteoglycans each bind TGF-b isoforms and that slight differences exist in their binding properties. They, and possibly other members of this family of proteoglycans, may regulate TGF-b activities by sequestering TGF-b in the extracellular matrix. Representative members of the subfamilies of small, leucine-rich interstitial proteoglycans are described next. Decorin and Biglycan The decorin gene, DCN, and the biglycan gene, BGN, are located on chromosomes 12ql3.2 and Xq28, respectively [154]. Both genes have a similar eight exon gene structure. The decorin gene is approximately 45 kbs, whereas the biglycan gene is approximately 8 kbs. Exon 1 of decorin, which encodes the 50 untranslated region, exists as exon 1a and exon 1b, which undergo alternative splicing [140]. The decorin transcript can also undergo additional alternative splicing to yield five isoforms, AeE. Isoform A, which contains the full amino acid coding sequence, is the product of transcript variants Al from exon 1a and A2 from exon 2b. Isoform B lacks the exons 3 and 4 sequence, isoform C lacks the exons 3e5 sequence, isoform D lacks the exons 4e7 sequence, and isoform E lacks the exons 3e7 sequence. There are several polyadenylation sites, of which two account for the 1.6- and 1.9-kb mRNAs isolated from various connective tissues and cultured cells [140]. The biglycan mRNA sizes, also related to the use of different polyadenylation sites, are 2.1 and 2.6 kbs [139]. The BGN gene is subject to X inactivation because there is no homologous gene on the Y chromosome. However, the pseudoautosomal expression of BGN probably results from the regulation of the BGN gene by a gene or genes that escape X inactivation [155]. Preprodecorin contains 359 amino acid residues [156]. It includes a signal peptide of 16 residues, an Npropeptide of 14 residues, and a mature chain of 329 residues. Intrachain disulfide bonds are present between Cys54 and Cys67 as well as between Cys313 and Cys346. Twelve leucine-rich repeats are present in the central domain between residues 73 and 359. The O-linked chondroitin sulfate or dermatan sulfate attachment site is at residue 34. Residues 211, 262, and 303 provide potential N-linked oligosaccharide attachment sites. The core protein of 359 amino acid residues has a predicted molecular weight of 39 kDa, whereas the protein extracted from tissues has a molecular weight of

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approximately 130 kDa, of which the chondroitin sulfate chain contributes approximately 40 kDa [157]. Preprobiglycan contains 368 amino acid residues [139]. It contains a signal peptide of 16 residues, a propeptide of 21 residues, and a mature protein of 331 residues. A region of proteoglycan N-terminal homology is present from residues 57 to 81 of the full-length protein. Ten leucine-rich repeats are located between residues 91 and 315. Residues 316e368 contain a domain with proteoglycan C-terminal homology. Residues 42, 47, 180, and 198 are potential chondroitin sulfate or dermatan sulfate attachment sites. Biglycan contains two substituted sites, whereas decorin contains only one. Asn residues 270 and 311 are possible sites for N-linked oligosaccharides. The secreted core protein has a predicted molecular weight of 38 kDa, whereas the intact tissue protein has a molecular weight of approximately 270 kDa, with the two chondroitin sulfate chains contributing 40 kDa [158]. BMP-1 cleaves probiglycan at a single site, removing the propeptide and producing a biglycan molecule with an N-terminus identical to that of the mature form found in tissues [159]. The BMP-1-related proteases, mammalian Tolloid and mammalian Tolloidlike 1 (mTLL-1), have low levels of probiglycan-cleaving activity. Wild-type mouse embryo fibroblasts produce only fully processed biglycan, whereas the fibroblasts derived from embryos homozygous null for the Bmp1 gene, which encodes both BMP-1 and mammalian Tolloid, produce predominantly unprocessed probiglycan, and fibroblasts homozygous null for both the Bmp1 gene and the mTLL-1 gene produce only unprocessed probiglycan. Consequently, all detectable probiglycanprocessing activity in the mouse embyronic fibroblasts is accounted for by the products of these two genes. The importance of decorin and biglycan in the formation of the connective tissues is demonstrated by the anomalies observed in mice that fail to express these genes. In decorin-deficient mice, the mice have fragile skin attributable to coarse and irregular collagen fibrils, confirming the importance of decorin in normal collagen fibrillogenesis [160]. Biglycan-deficient mice show reduced longitudinal growth and decreased bone mass, indicating an important role for biglycan in bone health [161]. Mice lacking both decorin and biglycan have more profound osteopenia compared to mice deficient in only one of these SLRPs [162]. Fibromodulin and Related Small Proteoglycans Representative members of the fibromodulin subfamily also include lumican, PRELP, and osteomodulin, also known as osteoadherin. They are found in most connective tissues but in greatest abundance in cartilage, tendon, and ligament [163]. The following descriptions will be restricted to fibromodulin and osteomodulin which are both present in bone.

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The fibromodulin gene, FMOD, is approximately 8.5 kbs and is located on chromosome 1q32 [164]. The gene contains three exons with conserved exon/intron junctions. Exon 1 encodes the 50 untranslated region, exon 2 contains 983 bps and encodes the major part of the translated region, and exon 3 encodes the last 50 nucleotides of the translated region as well as the 30 untranslated region. The fibromodulin mRNA is approximately 3 kbs. The highest expression of the gene is in hyaline cartilages, tendon, and ligament, whereas lower levels of expression occur in most other connective tissues [163]. Fibromodulin is a 59-kDa protein [163]. The fulllength protein contains 376 amino acid residues, including a signal peptide of 18 amino acid residues and a mature protein of 358 amino acid residues. The mature protein is composed of a central region containing leucine-rich repeats with up to four keratan sulfate chains flanked by disulfide-bonded terminal domains. The amino-terminal domain contains 10 tryosine residues that are partially sulfated [141]. The keratan sulfate chains are attached by N-glycosidic linkages from Nacetylglucosamine to asparagine residues in the central domain of the molecule [165]. In contrast to other SLRPs, fibromodulin does not contain glycosaminoglycan chains in the amino-terminal domain and does not contain chondroitin/dermatan sulfate in any domain. Mice lacking fibromodulin showed altered tendon structure because of the high proportion of thin, irregular collagen fibrils [145]. These relatively mild changes confirmed the important role of fibromodulin in regulating collagen fibrillogenesis in vivo. The gene for osteomodulin (also known as osteoadherin), OMD, is located on chromosome 9q22.31. The full-length human protein contains 421 amino acid residues. The primary structure of bovine osteomodulin was obtained by nucleotide sequencing of a cDNA clone from a primary bovine osteoblast expression library [166,167]. The entire translated primary sequence corresponds to a 49,116-Da protein with a calculated isoelectric point for the mature protein of 5.2. The dominating feature is a central region consisting of 11 leucinerich repeats ranging in length from 20 to 30 residues. The full, primary sequence contains four putative sites for tyrosine sulfation, three of which are at the N-terminal end of the molecule. There are six potential sites for Nlinked glycosylation, some of which are substituted with keratan sulfate chains. Osteomodulin shows 42% sequence identity to bovine keratocan and 38% identity to bovine fibromodulin, lumican, and human PRELP. Unique to osteomodulin is the presence of a large and very acidic C-terminal domain. The distribution of cysteine residues resembles that of other leucine-rich repeat proteins except for two centrally located cysteines. Northern blot analysis of RNA samples from various

bovine tissues showed a 4.5-kilobase pair mRNA for osteomodulin to be expressed in bone only. Osteomodulin mRNA was detected by in situ hybridization in mature osteoblasts located superficially on trabecular bone. Osteomodulin binds to hydroxyapatite and cells. Cell binding is mediated by integrin avb3 [167].

Thrombospondins The five members of the thrombospondin family are designated thrombospondin-1 to thrombospondin-5 [168]. Thrombospondin-5 is also known as cartilage oligomeric matrix protein (COMP). The thrombospondins are multimeric, multidomain glycoproteins that function at cell surfaces and in the extracellular matrix. The latter functions are also referred to as matricellular functions. The thrombospondin-1 gene, THBS1, is located on chromosome 15ql5 [169]. The full-length protein monomer contains 1170 amino acids, including a signal peptide of 31 amino acids. The thrombospondin-2 gene, THBS2, is located on chromosome 6q27 [170]. The full-length protein contains 1172 amino acids, including a signal peptide of 18 amino acid residues. The thrombospondin-3 gene, THBS3, on chromosome 1q21 encodes a protein monomer of 956 amino acids, including a signal peptide of 36 amino acid residues [171]. The thrombospondin-4 gene, THBS4, on chromosome 5ql3 encodes a full-length protein monomer of 961 amino acid residues, including a signal peptide of 21 residues [172]. Finally, the thrombospondin-5 gene, COMP, located on chromosome 19pl3.1 encodes a fulllength monomer of 757 amino acids, including a signal peptide of 20 amino acid residues [173]. Each thrombospondin is expressed in multiple tissues, particularly during fetal life. Similarly, most tissues express multiple members of the thrombospondin family [174]. Consequently, most of the thrombospondins are expressed in bone and cartilage during embryogenesis [175]. COMP is the most abundant form in postnatal growth plates [176]. Thrombospondin-2 and COMP are also present in postnatal bone and bone marrow stromal cells [176]. The thrombospondins are divided into subfamily A and subfamily B [174]. Thrombospondin-1 and thrombospondin-2 monomers, the members of subfamily A, share a complex modular structure and can assemble into disulfide-linked trimers. The modules from the amino terminus include the N-terminal domain, the oligomerization sequence, the procollagen homology region, three type 1, three type 2, and seven type 3 repeating units, and a globular C-terminal domain. Thrombospondins-3e5 are in subfamily B. They have unique N-terminal regions and lack the procollagen homology domain and type 1 repeats, but they contain

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four copies of the type 2 repeat and are assembled as pentamers. The globular amino- and carboxyl-terminal domains are connected to the remainder of the monomers by thin, flexible stalk regions [177]. The conformations of the stalk regions and the termini are determined, at least in part, by the calcium ion concentration [178]. The oligomerization motifs in all the thrombospondins are likely to be in a coiled-coil conformation, giving rise in subfamily A to trimers and in subfamily B to pentamers [179]. The procollagen homology region of thrombospondin-1 also folds into a stable, compact, and disulfide-bonded monomer. Isolated type 3 repeats of COMP produce a 14.2-nm rod-like structure [180]. The C-terminal domain and the adjoining type 3 repeats may fold together, in the presence of calcium, to form a C-terminal globular structure [180]. Thrombospondin-1 and thrombospondin-2 can form homotrimers and heterotrimers [181]. Thrombospondin trimers and pentamers are stabilized by the formation of interchain disulfide bonds between their corresponding oligomerization domains. Pentamerization of COMP involves the formation of interchain disulfide bonds between cysteine residues 68 and 71 [182]. Intrachain disulfide bonds are also present in the type 3 repeats. There are also specific sites for the addition of N- and O-linked sugars and for the C-mannosylation of tryptophan [183]. In the extracellular matrix, the thrombospondins bind to other macromolecules as well as to cell surface receptors. Many of the interactions involve specific domains within the monomers. Some of the interactions are sequence specific, but many are also dependent on the ionic environment and the conformation of the monomers. Each thrombospondin-1 monomer can bind approximately 35 calcium ions, whereas subfamily B thrombospondins, such as COMP, are expected to bind almost double that number of calcium ions [184]. Major changes in conformation of the type 3 repeats and the molecule follow the binding of calcium ions. Binding sites are also present for heparin and heparan sulfate proteoglycan, decorin, various integrins, as well as a wide range of proteases, cytokines, and growth factors [183]. The binding sites and the consequences of the interactions have been most thoroughly studied for thrombospondin-1 and thrombospondin-2. For example, the activities of thrombin, plasmin, neutrophil cathepsin G, elastase, urokinase plasminogen activator, plasminogen activator inhibitor, and MMP-2 are modified following binding to the latter thrombospondins [185]. The small latent TGF-b1 complex with the latencyassociated peptide binds to the WSIIWSPW motif in the second type 1 repeat of thrombospondin-1 [186]. An intermolecular activation effect of the KRFK motif

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in the first type 1 repeat of thrombospondin-1 releases mature, active TGF-b1. Thrombospondin-2, which lacks the KRFK motif, can bind the latent TGF-b1 complex but cannot activate it [186]. Thrombospondin-4 and COMP bind to type I and II collagens [187]. COMP also binds type IX collagen and MMP-19 and -20, whereas thrombospondin-4 also binds to laminin, fibronectin, and matrilin-2 [187]. Cell surface interactions with thrombospondin-1 have been studied in detail [174]. The amino-terminal domain, type 1 repeats, type 3 repeats, and the Cterminal domain all interact with the cell surface. Each region appears to act via different cell surface receptors. The interactions are calcium-dependent and appear to be important for the maintenance of cell adhesion, spreading, migration, and shape. Cell attachment activity also appears to be important for thrombospondins-2 and -4 and COMP [188]. The C-terminal domains of thrombospondin-4 and COMP bind types I, II, and IX collagens in a zinc iondependent manner [187,189]. The binding of the latter thrombospondins is to the amino and carboxyl propeptides and two triple-helical sites of these collagens. The pentameric structure of thrombospondin-4 and COMP favors the use of multiple sites of interaction, which are likely to stabilize the extracellular matrix. Further insights into the functions of thrombospondin (TSP)-1, -2, -3 and -5 have been obtained from murine gene knockout studies. Mice lacking thrombospondin-1 show decreased embryonic viability, early onset pneumonia, increased circulating monocytes, and reduced TGF-b1 activation in the inflamed lungs and pancreas [190]. TSP1-deficient mice have no reported appendicular skeletal defects but have craniofacial dysmorphism, that is more pronounced in compound TSP1/2-deficient mice, spinal lordosis and altered rates of wound healing and tissue remodeling in various injury models. Mice lacking thrombospondin-2 have increased thickness and density of the longbone cortices, attributed to increased bone formation [191]. The TSP2-deficient mice also show resistance to ovariectomy-induced bone loss, differential surface responsiveness to mechanical loading, increased bone formation and reduced chondrogenesis in response to fracture, and altered bone mass and geometry associated with aging [168]. The TSP2-deficient mice have fragile skin, lax tendons, abnormal fibrillar collagen organization, increased vascular density, prolonged bleeding time, and accelerated skin wound healing [191]. The changes in the skin and tendons indicate that thrombospondin-2 normally plays an important role in collagen fibrillogenesis. Abnormal fibroblast interactions with the extracellular matrix and increased production of active MMP-2 were also identified in cell cultures from TSP2-deficient mice.

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In the TSP3-deficient mice, skeletal development was initially normal but, by 9 weeks of age, there was accelerated ossification of the secondary ossification centers of the femoral heads [192]. The TSP3-null mice also had an increased femoral cortical bone periosteal circumference and parameters of mechanical strength. The phenotype of TSP4-deficient mice has not been reported to date. However, the C-terminal domain of TSP4 binds to collagens I, II, III and V, as well as laminin, fibronectin and matrilin-2 [174]. TSP5-deficient mice have a mild skeletal phenotype with growth plate disorganization and mild exercise-induced articular cartilage flattening [193]. These skeletal changes are worse in compound TSP1/5 and TSP3/5-deficient mice [193]. Missense mutations of THBS5, particularly those that affect calcium binding and protein folding, are associated with the pseudoachondroplasia and multiple epiphyseal dysplasia phenotypes in mice and humans [194].

Osteonectin Osteonectin, initially isolated from demineralized bone matrix, was named according to its ability to bind to calcium, hydroxyapatite, and collagen, and to nucleate hydroxyapatite deposition [195,196]. The protein has also been called secreted protein, acidic and rich in cysteine (SPARC), basement membrane 40, or “culture shock” protein [197]. Osteonectin is an antiadhesive protein because it inhibits cell spreading, induces rounding of cells, and disassembles focal adhesions [198]. Other activities of osteonectin include calcium-dependent binding to collagens and thrombospondin, binding to platelet-derived growth factor-AB and -BB, and regulation of cell proliferation and MMP expression [199]. The gene SPARC or ON, which contains 10 exons, is located on chromosome 5q31.3-q32 [200]. The gene is expressed at high levels in tissues undergoing morphogenesis, remodeling, and wound repair. It is also made by cells of osteoblastic lineage and the hypertrophic chondrocytes of the growth plate [201]. The SPARC gene is also expressed in several postnatal non-skeletal tissues, including salivary and renal tubular epithelium [202]. Osteonectin is the most abundant non-collagenous protein in mineralized bone matrix in some species [196]. The SPARC gene encodes a full-length protein of 303 amino acids, including the signal peptide of 17 amino acid residues. The core molecular weight is 33 kDa [203]. The protein extracted from bone has an apparent molecular weight of 43 kDa attributable to post-translational modifications such as glycosylation. The mature human protein consists of 286 amino acid residues divided into three domains [204]. An aminoterminal acidic segment (residues 1e52) binds five to eight calcium ions with low affinity and mediates

interactions with hydroxyapatite. A follistatin-like domain (residues 53e137) contains five disulfides and an N-linked oligosaccharide at Asn [99]. Finally, an ahelical domain contains two EF hand, high-affinity, extracellular calcium binding sites (EC domain, residues 138e286). Crystal structure analysis showed that the follistatin and extracellular calcium domains interact through a small interface that involves the EF hand pair of the extracellular domain [205]. The elongated follistatin domain is structurally related to serine protease inhibitors of the Kazal family. Residues implicated in cell binding, inhibition of cell spreading, and disassembly of focal adhesions cluster on one side of osteonectin opposite the binding epitope for collagens and the Nlinked oligosaccharide. Crystal structure analysis also has shown that the collagen-binding epitope in the helix aA is partially masked by helix aC [205]. Deletion of helix aC produced a 10-fold increase in collagen affinity, similar to that seen after proteolytic cleavage of this helix [206]. Five residues were crucial for collagen binding: R149 and N156 in helix aA and L242, M245, and E246 in a loop region connecting the two EF hands of osteonectin. These residues were spatially close and formed ˚ , which matches the diameter of a flat ring of 15 A a triple-helical collagen domain. Nearly identical binding characteristics were displayed by type I and IV collagens. The absence of Sparc in transgenic mice gives rise to aberrations in the structure and composition of the extracellular matrix that result in cataracts, severe osteopenia, and impaired wound healing [207e209]. Sparcnull mice have decreased trabecular bone volume and fail to demonstrate an increase in bone mineral density in response to a bone-anabolic parathyroid hormone treatment regimen [209,210]. By 6 months of age, the mice develop severe eye pathology [207]. Sparc-null mice have greater deposits of subcutaneous fat and larger epididymal fat pads in comparison with wildtype mice. The dermis and fat pads of the SPARCdeficient mice contain less collagen I and the fibers are of smaller diameter than in controls [211e213].

Osteocalcin Osteocalcin is a small protein that accounts for approximately 10% of the non-collagenous protein of bone [214]. It is also called bone g-carboxyglutamate or bone Gla protein. It is one of several Gla proteins found in the skeleton. Other Gla proteins, such as protein S, are made elsewhere and are deposited in the bone matrix from the circulation [215]. The Gla proteins have in common the presence of glutamic acid residues that have been g-carboxylated by a specific g-carboxylase that requires vitamin K as a co-factor [216]. These

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[319] Roschger P, Paschalis EP, Fratzl P, Klaushofer K. Bone mineralization density distribution in health and disease. Bone 2008;42:456e66. [320] Bresler D, Bruder J, Mohnike K, Fraser WD, Rowe PS. Serum MEPE-ASARM-peptides are elevated in X-linked rickets (HYP): implications for phosphaturia and rickets. J Endocrinol 2004;183:R1e9. [321] Addison WN, Masica DL, Gray JJ, McKee MD. Phosphorylation-dependent inhibition of mineralization by osteopontin

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ASARM peptides is regulated by PHEX cleavage. J Bone Miner Res 2010;25:695e705. [322] Pampena DA, Robertson KA, Litvinova O, Lajoie G, Goldberg HA, Hunter GK. Inhibition of hydroxyapatite formation by osteopontin phosphopeptides. Biochem J 2004;378: 1083e7. [323] Addison WN, McKee MD. ASARM mineralization hypothesis: a bridge to progress. J Bone Miner Res. 2010;25: 1191e2.

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C H A P T E R

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Prenatal Bone Development Fanxin Long Department of Medicine, Department of Developmental Biology, Washington University School of Medicine, Washington University in St Louis, Missouri, USA

Intramembranous Ossification

Proper formation of the skeleton requires precise control of both skeletal patterning and skeletal cell differentiation. Skeletal patterning usually refers to the specification of position, number, and shape of the skeletal elements, parameters believed to be determined before the appearance of skeletal cell types. This very important aspect was dealt with extensively in the previous edition of this chapter by St.-Jacques and Helms, but will not be discussed here. Instead, this update will focus on the more recent findings, especially those from mouse genetic studies, regarding the molecules that control the formation of chondrocytes and osteoblasts. Before this discussion, however, the chapter will briefly reiterate the different modes and the time line of bone formation in the human embryo, largely based on the previous edition of the chapter by St.-Jacques and Helms.

Intramembranous ossification begins toward the end of the second month of gestation in humans. The process has been most extensively studied in the developing cranium, where it is often preceded by a cellular proliferation at specific sites in the mesenchyme, and becomes histologically evident when a cluster of pale-staining stellate cells aggregate and take on a rounded basophilic appearance [3]. The aggregated mesenchymal cells gradually differentiate into mature secretory osteoblasts that actively produce the collagen I-rich extracellular matrix that is characteristic of bone. This differentiation process occurs in a small number of sites within the territory of each intramembranous bone, and these sites are called ossification centers. It is important to note that, although the initial condensation occurs in an avascular milieu, the differentiation of osteoblasts and the onset of mineralization are intimately related to blood vessel invasion, first in the surrounding mesenchyme and ultimately in the bone rudiment [4]. The inital bone tissue (spicule) is irregularly shaped and completely surrounded by the osteoblasts that secreted it. Some of the osteoblasts soon become encased in the matrix and become known as osteocytes. Each osteocyte is enclosed in its own lacuna but extends projections through small channels called canaliculi to maintain contact with neighboring osteocytes and intercellular fluids outside of the ossification center. Less differentiated osteoprogenitor cells present at the periphery proliferate and eventually differentiate into new osteoblasts, which continue to add bone to form a well-defined longer structure called a trabecula (beam). Multiple trabeculae soon connect and form a scaffolding characteristic of cancellous (or spongy) bone [3]. The first bone formed in the embryo is of an immature type, displaying relatively high cellularity and an almost random orientation of collagen fibers [5]. At approximately the time of birth,

MODES OF BONE FORMATION Most skeletal elements of modern vertebrates can be traced to the endoskeleton (skeleton formed inside the body) found in primitive vertebrates [1]. Vestiges of the exoskeleton (skeleton formed outside the body), however, are present in the skull and pectoral girdle. Although such exoskeletal derivatives have become thoroughly integrated with elements of the endoskeleton, they still form differently from the endoskeleton during embryogenesis. These bones, termed dermal because of their historical association with the skin, form by direct differentiation of mesenchymal cells into osteoblasts through intramembranous ossification. On the other hand, bones derived from the primitive endoskeleton develop first into cartilage templates but are subsequently replaced by bone through endochondral ossification [2].


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3. PRENATAL BONE DEVELOPMENT

this woven bone is gradually replaced by the more mature lamellar bone characterized by successive layers of uniformly oriented collagen fibers.

Endochondral Ossification This process also initiates with the aggregation of undifferentiated mesenchymal cells to form condensations (Fig. 3.1). However, unlike in intramembranous ossification, cellular proliferation does not appear to play an important part in this process, as these condensations are the result of an increase in cell packing mediated by changes in the extracellular matrix and cellecell adhesion molecules [6]. By their positions, shapes, and sizes, these prechondrogenic condensations prefigure the different skeletal elements [7]. As in intramembranous ossification, the initial condensation forms in an avascular environment, but here it remains avascular. In the core of these condensations, cells differentiate into chondrocytes that secrete a cartilage matrix characterized by the presence of types II, IX, and XI collagen and specific proteoglycans such as aggrecan. At the periphery of the condensation, cells surrounding the cartilage core flatten and form a thin membrane of stacked cells called the perichondrium, which insulates the cartilage from the surrounding mesenchyme [8]. Perichondrial cells retain chondrogenic potential and probably contribute to the radial expansion of the cartilage by appositional growth. Initially, the chondrocytes and perichondrial cells proliferate rapidly, and this proliferation together with the deposition of new matrix drives the growth of the elements. At a certain stage specific for each element, chondrocytes in the center E10.5

E12.5

E14.5

E15.5 – 18.5

FIGURE 3.1 Endochondral bone formation. Time line is based on mouse embryonic days, and the events are depicted for the tibia. The timing of events may differ in other skeletal elements. Cartilage initially develops from mesenchymal condensation within an avascular environment. Vascularization occurs following the hypertrophy of cartilage. (Courtesy of Dr Matthew Hilton.)

undergo progressive maturation. They acquire a flattened appearance and become organized in columns along the longitudinal axis of the developing skeletal element. Columns are separated by relatively thick lateral partitions of extracellular matrix, the longitudinal septa, whereas chondrocytes within a column are separated by thin transverse septa. Further maturation of these cells leads to hypertrophy characterized by cell enlargement, cessation of proliferation, and secretion of a distinct extracellular matrix rich in type X collagen that becomes progressively calcified [9]. These changes are accompanied by vascular invasion of the hypertrophic cartilage, and by differentiation of the inner perichondrium cells into osteoblasts, which secrete a layer of primary bone to form the bone collar [8]. At this stage, the thin layer of tissue covering the newly formed bone becomes known as the periosteum and continues to supply osteoblasts that produce the bone matrix of the diaphysis. Changes in the composition and properties of the cartilage matrix in the hypertrophic zone, including calcification of the longitudinal septa and the release of angiogenic factors, trigger its invasion by capillaries [10]. This results in the death of the terminally differentiated hypertrophic chondrocytes and degradation of the uncalcified transverse cartilage septa by invading “chondroclasts”, a cell type of ill-defined origin associated with the invading capillary sprouts. The invading blood vessels are believed to carry in osteoprogenitors that eventually differentiate into osteoblasts. Remnants of the calcified longitudinal septa act as templates on which the osteoblasts secrete bone matrix and establish the primary ossification center. Osteoclasts, which are bone-resorbing cells of hematopoietic origin, also appear within the ossifcation center, contributing to the formation of the marrow cavity. The synchronous maturation and columnar organization of the chondrocytes result in a characteristic histological structure known as the embryonic growth plate in which zones of proliferation, maturation, hypertrophy, and bone formation can be identified, linearly progressing from the articular ends (epiphysis) towards the midshaft (diaphysis) of the element. Continued proliferation of the less mature chondrocytes at the epiphysis, followed by their hypertrophy and their eventual replacement by trabecular bone near the diaphysis results in a distal displacement of the growth plate and longitudinal growth of the skeletal element. In a typical long bone, this growth is accompanied by resorption of the older trabeculae by osteoclasts, thus leaving significant trabecular bone only at the region immediately adjacent to the growth plate (metaphysis). Continued deposition of cortical bone by the periosteum (subperiosteal bone) leads to radial growth. Eventually, a secondary ossification center forms at the center of the epiphysis distal to the growth plate, from which

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MOLECULAR REGULATION OF SKELETAL DEVELOPMENT

ossification proceeds radially in all directions. Formation of the secondary ossification center does not modify the general organization of the growth plate, which remains active until cessation of growth after puberty. In the long bones of humans, the growth plate eventually disappears with the only cartilage remaining at the articular surface. As in intramembranous ossification, the first bone tissue produced in the trabeculae and the diaphysis of the long bones is of a woven type. Remodeling of this immature bone following its resorption by osteoclasts leads progressively to replacement by mature lamellar bone.

TIMING AND SEQUENCE OF BONE FORMATION IN HUMANS All skeletal elements do not form simultaneously in the embryo. Generally, chondrification (cartilage formation) proceeds in a rostralecaudal (head-to-tail) fashion in the axial skeleton and in a proximaledistal (shoulderto-fingertip) fashion in the limb skeleton. This is a direct consequence of the developmental origin of the different elements from the progressively forming somites and the outwardly elongating limb buds, respectively. The chondrification sequence is rapid, and early in the seventh gestational week the cartilage primordia of all the elements of the axial and appendicular skeleton are present [11]. In the cephalic region, cartilage formation proceeds in a caudal-to-rostral direction. It starts at the beginning of week 7 and continues well into week 8 [12]. Thus, the appearance of the cartilage primordia of the skull base occurs after that of most axial and appendicular elements, but it precedes ossification of the membranous bones of the skull vault. Almost all primary ossification centers appear between weeks 7 and 12 of embryonic life in humans. In contrast, the secondary ossification centers such as those in the epiphyses appear over a long period, from late fetal life until puberty. The clavicle is the first bone in the body to ossify, with the mandible and maxilla following almost immediately. Generally, the facial and calvarial centers appear before the basicranial centers, followed by the hyoid centers [13]. More than 100 centers of ossification appear during human skull formation, but extensive fusion takes place between many of them, reducing the number to 45 by the time of birth [14]. In the axial skeleton, the costal centers appear first, followed by the primary vertebral centers and the sternal centers. The second to 11th rib centers appear at approximately the same time. The centers in the first and 12th ribs appear later in that order. The vertebrae ossify from three primary centers, one for the body and one for each neural arch. In the spine, as in the

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thoracic cage, the sequence is not strictly cephalocaudal because ossification of the vertebral body starts first in the lower thoracic and upper lumbar regions and propagates in both directions from this area. Some delay is observed before the first cervical and the last two sacral centers appear. Ossification centers in the neural arches appear in a cephalocaudal sequence except in the atlas and axis, in which ossification is slightly delayed [13]. The sternum arises as a pair of cartilaginous bands that fuse along the midline as the ventral body wall develops. This fused cartilage precursor subsequently subdivides into craniocaudal elements, most of which will fuse again and will ossify after the fifth month to form the body of the sternum [15]. The primary ossification centers of the pectoral girdle appear before those of the pelvis in the following order: clavicle, scapula, ilium, ischium, and pubis. The centers in the humerus, femur, and then radius, ulna, and tibia appear at approximately the same time. The fibula differentiates slightly later. The bones of the hand invariably appear before their counterparts in the foot. These differ in that centers in the distal phalanges of the hand appear before those of the metacarpals, whereas the distal phalanges centers of the foot appear after the metatarsal centers [13]. The proximal and middle phalanges ossify last. Of the tarsal and carpal bones, only the talus and calcaneous generally begin to ossify before birth, with the cuboid sometimes and the lateral cuneiform rarely ossifying before birth [3].

MOLECULAR REGULATION OF SKELETAL DEVELOPMENT EpithelialeMesenchymal Interaction The requirement for an early tissue interaction to induce mesenchymal condensation is best demonstrated by tissue dissociationerecombination experiments with craniofacial tissues. Embryonic mandibular mesenchyme can be separated from the mandibular epithelium and cultured in vitro. Mesenchyme isolated before migration or at an early stage after migration of mesenchymal cells to the final site of bone formation fails to form bone in culture. However, if early mesenchyme is combined in vitro with mandibular epithelium or if mesenchyme is isolated at a late stage, then osteogenesis takes place, indicating a requirement for epithelialemesenchymal contact [16]. The signal from the epithelium appears to be permissive in nature. First, mandibular mesenchyme forms bone in culture when recombined with epithelia from other regions of the embryo as well, provided that they are of the correct age. Furthermore, the shape of the bone

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produced is independent of the source of epithelium but is dictated by the nature of the mesenchyme [16]. Finally, an epithelium such as the mandibular is incapable of inducing bone formation in a mesenchyme that is normally non-osteogenic. Such interactions have been studied in detail mostly in craniofacial structures and have been shown to be essential to induce formation of scierai ossicles and scierai cartilage, mandible and Meckel’s cartilage, maxilla, palatine bone, otic vesicle, and calvarial bones, and also clavicle and limb cartilage [17,18]. The molecular nature of the epithelial signal(s) is unclear but probably involves in many cases the bone morphogenic proteins (BMPs), which form a subfamily of secreted proteins among the TGF-b superfamily. Members of this group are characterized by a conserved C-terminal domain with several cysteine residues and undergo cellular processing by cleavage of an inactivating N-terminal domain. They act as dimers to bind a type II transmembrane receptor and induce recruitment and phosphorylation of the cytoplasmic tail of one of the type I receptors. Phosphorylation of the type I receptor initiates signal transduction through various pathways, including the SMAD family of intracellular signaling molecules. At least 20 different BMPs and the related growth and differentiation factors (GDFs) have been identified in vertebrates [19]. Members of this family of growth factors were first identified as proteins present in demineralized bone matrix, which has the remarkable capacity to induce ectopic bone formation in subcutaneous implants [20,21]. Purified recombinant BMPs share this capacity and are able to potentiate chondrocyte and osteoblast differentiation of cells in vitro [22]. Consequently, BMPs have long been considered in vivo “bone inducers”. However, BMPs induce ectopic bone by recapitulating the events occurring during endochondral bone formation [23], and therefore may not act by inducing ossification (production of bone matrix) per se. Instead, the bone-forming ability of BMPs probably reflects their role in the induction of mesenchymal condensations [24]. Several BMPs known to have cartilage/boneinducing potential accumulate in the extracellular material (ECM), including epithelial basal lamina [25,26]. In the case of the mandible, an epithelialemesenchymal interaction involving BMP signaling has been documented. BMPs-2, -4, and -7 are found in the distal mandibular epithelium whereas the homeobox-containing transcription factor Msx1 is present in the mandibular mesenchyme. Msx1 function in the mesenchyme was demonstrated by Msx1 knockout mice, which exhibited mandibular defects [27]. Expression of Msx1 requires mandibular epithelium [28] and can be induced by ectopic BMPs [29], thus implicating these molecules

in the early epithelialemesenchymal required for mandible formation.

interaction

Condensation Skeletal condensations have been studied most intensely in the developing limb in vivo and by means of micromass cultures of limb mesenchyme in vitro. The condensation stage is a very transient phase, rapidly followed by differentiation of the aggregated cells. At the histological level, it is characterized by a significant increase in cell-packing density in specific areas of the mesenchyme. Cells of the precartilaginous condensations can be visualized by their affinity for the lectin peanut agglutinin (PNA) and their transient upregulation of a number of markers, including versican, tenascin, syndecan, N-CAM, N-cadherin, thrombospondin-4, type I collagen, and heparan and chondroitin sulfate proteoglycans [30]. Cell proliferation plays a role in condensation of the calvarial, scierai, and mandibular primordia but not in the precartilaginous condensations of the limb, where cell movements appear to be more important [7]. Regardless of the mechanism, the high cell density generated in the condensations leads to increased cellecell contact as well as the formation of gap junctions facilitating intercellular communication [31,32]. These appear to be essential for differentiation to take place, and the extent of cellular condensation correlates with the level of subsequent chondrogenesis [33]. Two cell adhesion molecules implicated in the condensation process are N-cadherin and N-CAM. Ncadherin belongs to a group of calcium-dependent transmembrane glycoproteins that mediate cellecell adhesion by homotypic interactions through their extracellular domains. The protein cytoplasmic domain interacts with the actin cytoskeleton via the catenin molecules. N-CAM is a member of the large immunoglobulin superfamily of membrane glycoproteins and also mediates cellecell adhesion via homotypic interactions but in a calcium-independent manner. These molecules are expressed at high levels in condensing mesenchyme but disappear in differentiating cartilage and can be detected later only in the perichondrium. Much evidence shows that perturbing the functions of N-cadherin and N-CAM causes reduction or alteration of chondrogenesis both in vitro and in vivo [33]. Conversely, overexpression of N-cadherin and N-CAM in micromass cultures stimulates chondrogenesis [34,35]. Cellecell adhesion in the digit condensations was also shown to depend on Epheephrins interactions. The Eph receptors belong to the family of receptor tyrosine kinases. They are characterized by a unique cysteine-rich motif in their extracellular domain, followed by two fibronectin type III motifs. They interact with a family of at least eight ephrin ligands associated

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with the cell membranes via a transmembrane domain or a GPI anchor [36]. EphrineEph signaling has been implicated in the sorting and adhesion of mesenchymal cells in both mouse and chicken [37e39]. Interestingly, in Hoxa13 mutant mice, expression of EphA7 was reduced in mesenchymal condensations correlating with altered adhesiveness, poorly resolved condensations, and defective chondrogenesis [38], whereas misexpression of Hoxd11 or Hoxa13 in the developing chick limb affected condensation size and cell adhesiveness [40,41]. Thus, Hox transcription factors may control skeletal patterning via the regulation of ephrineEph signaling, at the condensation stage. Interactions between cells and the ECM also significantly affect condensation. Prior to condensation, mesenchymal cells secrete an ECM rich in hyaluronan (HA) that facilitates cell movement but prevents close cellecell interactions. As condensation begins, a transient increase in hyaluronidase activity leads to controlled hydrolysis of HA and reduced intercellular space, thus favoring cellecell interactions [42,43]. Condensation also coincides with upregulation of a number of ECM proteins, including type I collagen, fibronectin, and various proteoglycans [33]. Syndecan-3 is an integral membrane protein that is also a heparan sulfate proteoglycan that interacts with ECM components and heparin-binding growth factors [44]. It is transiently expressed at high levels during formation of the precartilage condensations and downregulated (except in the perichondrium) after chondrocyte differentiation. Antibodies against syndecan-3 impair the formation of precartilaginous condensations in micromass cultures of chick limb mesenchyme [45]. Fibronectin is a dimeric proteoglycan present in the ECM of many tissues and plays an important role in cell migration and differentiation. Fibronectin interaction with the cellular integrin receptors can activate signaling through the focal adhesion kinase and the integrin-linked kinase pathways. Prior to condensation, fibronection is distributed throughout the intercellular space of the mesenchyme, but it accumulates in the condensations and reaches its maximal level of expression just prior to overt chondrogenesis [46]. Interaction between extracellular fibronectin and heparin-like molecules of the mesenchymal cell surface is crucial for the formation of precartilage condensations [47,48]. Antibodies specific to a certain isoform of fibronectin also perturbed chondrogenesis [49]. Expression of the aforementioned cellecell and cellematrix adhesion molecules is modulated by signaling factors. For instance, treatment of pluripotential cells or limb bud mesenchyme with TGF-b stimulated expression of fibronectin, N-CAM, N-cadherin, and tenascin [50,51]. Exposure of limb bud mesenchymal cells to certain BMPs also correlated with an increase in N-cadherin and N-CAM expression and

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stimulates chondrogenesis [33]. Functionally, BMP/ GDF gain-of-function experiments in chick [52,53] and mouse embryos [54,55] led to hyperplasia and, in some cases, fusion of limb cartilage elements. Conversely, broad repression of BMP signaling inhibited cartilage development [54,56]. The important role of BMP signaling in chondrogenesis has been further demonstrated by genetic experiments in the mouse. Earlier gene deletion studies of BMPs in the mouse have been complicated by either early lethality or functional redundancy among the family members. For instance, null mutations in Bmp-2 or -4, as well as in the BMP receptor type 1A gene (Bmp-r1A), lead to early embryonic lethality [57e59], whereas mice null for Bmp-3 or Bmp-3b (also known as Gdf10) display no obvious embryonic phenotype in the skeleton [60,61]. However, by using the Cre-loxP technology, simultaneous deletion of Bmp2 and Bmp4 in the prechondrogenic limb mesenchyme with Prx1-Cre led to the failure of the formation of certain chondrogenic condensations [62]. These results indicate that a threshold of Bmp signaling is necessary for chondrogenesis. The fact that all chondrogenic condensations were not equally affected by the loss of Bmp2 and Bmp4 suggests that the less affected skeletal element may require a different set of Bmp family members. As in the mouse, mutations affecting BMP signaling in humans also lead to defects in skeletal elements. For instance, mutations in GDF5 have been found to cause brachypodism (bp) in the mouse, and HuntereThompson acromesomelic dysplasia as well as Grebe syndrome in humans [63]. Moreover, mutations in the gene encoding the BMP antagonist Noggin (NOG) also cause two autosomal dominant disorders e proximal symphalangism (SYM1) and multiple synostose syndrome (SYNS1), both characterized by multiple joint fusions [64]. Although the precise function of BMP signaling during mesenchymal condensation remains to be fully elucidated, studies have suggested that BMP/GDF signaling controls recruitment of cells into the early condensations [52,55]. Consistent with this view, genetic deletion of noggin, an extracellular antagonist of BMP proteins, resulted in enlarged cartilage elements and the lack of joints in the mouse [65]. Analyses of chondrogenesis in micromass cultures of limb bud mesenchyme established that BMP signaling is indeed required for formation of prechondrogenic condensations but also for differentiation of mesenchymal cells into chondrocytes [66,67]. More recently, by using an imaging system that dynamically visualizes limb mesenchymal cells undergoing successive phases of cartilage formation in vitro, Barna and Niswander found that BMP signaling was required for the “compaction” event that coalesces the small cellular aggregates into a round cluster of tightly associated cells with a distinct outer boundary,

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a prerequisite for chondrogenic differentiation [68]. Importantly, the compaction event appeared to be independent of Sox9 function, as Sox9-null cells exhibited no defect up to this point, although they subsequently became segregated from the chondrogenic mesenchymal condensations and adopted a “fibroblastoid” morphology distinct from the round morphology typical of the cells that remained within the condensation. Thus, BMP signaling appears to play a critical role in the formation of mesenchymal condensation before Sox9 becomes important. The molecular nature for this early requirement of BMP however, is not known at present. Interactions of the mesenchymal cells with other cell types also affect condensation. For instance, in the developing limb, there is an inverse relationship between condensations and blood vessel distribution. Vessels are initially present throughout the limb mesenchyme but undergo local regression from sites at which precartilaginous condensations will form shortly thereafter. It was shown that this vascular regression is essential for the condensation step and subsequent chondrogenesis in the developing limb of chick embryos, although the molecular mechanism was unclear. Interestingly, a more recent study showed that the chondrogenic primordia played a role in patterning the limb vasculature through the expression of vascular endothelial growth factor (VEGF) [69]. Thus, early chondrogenic events may trigger the vascular regression, which may in turn reinforce chondrogenesis.

Chondrocyte Differentiation The overt differentiation of condensed mesenchymal cells into chondrocytes is characterized by a shift in production of ECM and adhesion molecules. Whereas N-CAM, N-cadherin, and type I collagen are downregulated, collagen types II, IX, and XI, as well as the large proteoglycan aggrecan, are upregulated. Sox9, an HMG domain DNA-binding protein, is a key transcription factor regulating chondrocyte differentiation. Mutations in this gene cause the rare and severe dwarfism campomelic dysplasia in humans [70]. During embryogenesis, Sox9 is expressed in all prechondrogenic condensations, where it precedes the expression of the Col2a1 gene, which encodes the major cartilage matrix protein type II collagen. Because Sox9þ/ males were sterile, the function of Sox9 in chondrogenesis was initially studied by analyzing the fate of Sox9/ ES cells in chimeric mouse embryos. It was shown that the mutant cells were excluded from prechondrogenic condensations and failed to express any chondrocytespecific markers [71]. Interestingly, a more recent study, by applying live imaging techniques to limb mesenchymal condensations in vitro, demonstrated that the Sox9/ cells participated in the initial mesenchymal

condensations normally, but later segregated from the prechondrogenic clusters [68]. Tissue-specific knockout of Sox9 by the Cre-loxP technology confirmed that Sox9 is indispensable for chondrogenesis [72]. Sox9 likely performs multiple functions during chondrocyte development. Live imaging of mesenchymal condensation in vitro with Sox9/ cells indicated that Sox9 was required not for the initial mesenchymal condensation, but rather performed a subsequent role in maintaining the round morphology typical of the prechondrogenic cells [68]. Because Col2a1, one of the best known target genes of Sox9, was not yet activated in a majority of the prechondrogenic cells, the function of Sox9 at this stage was likely to be independent of type II collagen. Thus, in addition to activating chondrocyte-specific genes, Sox9 appears to play an earlier role that remains to be elucidated. Two other members of the Sox family, L-Sox5 and Sox6, interact with the same sequences and together with Sox9 activate transcription of the Col2a1 and aggrecan genes. L-Sox5 and Sox6 have partly redundant functions and single mutants displayed limited skeletal abnormalities, but the Sox5/Sox6 double mutants showed severe generalized chondrodysplasia characterized by poor differentiation of chondrocytes [73]. Thus, L-Sox5 and Sox6 are potent enhancers of chondrocyte differentiation. In addition to its role in mesenchymal condensation as discussed above, BMP signaling also stimulates chondrocyte differentiation after the condensation stage. BMPs transduce signals by binding to complexes of type I and II serine/threonine kinase receptors. In the most extensively studied mechanism, known as the “canonical” pathway, ligand binding induces phosphorylation of the receptors which, in turn, phosphorylates and activates receptor Smads (R-Smads) 1, 5 and 8 [74]. The activated R-Smads then complex with Smad4 to enter the nucleus, eventually regulating gene expression. Simultaneous deletion of the type I receptors Bmpr1a and Bmpr1b in the chondrogenic lineage with Col2-Cre resulted in a severe form of general chondrodysplasia where a majority of the cartilage promodia that prefigure the endochondral bones were absent [75]. Moreover, simultaneous removal of Smad1 and Smad5 in chondrogenic cells, regardless the status of Smad8, essentially abolished cartilage formation [76]. These findings therefore establish that canonical BMP signaling through R-Smads is indispensable for the formation of cartilage following the activation of the Col2a1 gene. Interestingly, deletion of Smad4, the binding partner for Smad1, 5 or 8, with a similar Col2Cre line, did not severely affect cartilage formation [77]. The discrepancy between the R-Smad- and Smad4-deficient mice raises the possibility that BMPSmad1/5/8 signaling may control cartilage development in a Smad4-independent manner. Indeed, BMPs

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were shown to induce R-Smad nuclear localization in Smad4-null colon cancer cells [78], but whether this occurs during cartilage development is currently unknown. In contrast to BMP, some members of the Wnt family inhibit chondrocyte differentiation. The Wnt genes encode a large family (19 members in mouse and humans) of cysteine-rich secreted glycoproteins sharing homology with the Drosophila signaling factor wingless. Wnt proteins control morphogenesis and tissue patterning in a wide variety of organs [79]. Depending on the cell context, Wnt ligands interact with several types of membrane receptors including those of the Frizzled (Fz) family and the low-density lipoprotein receptor related proteins Lrp5 and 6 to activate a variety of intracellular signaling cascades. Among the most extensively studied is the b-catenin-dependent, canonical pathway. In this mechanism, binding of Wnt ligands stabilizes bcatenin, which enters the nucleus to interact with the LEF/TCF family of transcription factors, resulting in transcriptional activation of downstream target genes. Other b-catenin-independent pathways include those mediated by PKC or the Rho family of small GTPases, although the latter was recently shown also to participate in b-catenin signaling [80]. Studies in chick embryonic limbs or limb bud micromass cultures showed that ectopic expression of Wnt1 or Wnt7a inhibited chondrocyte differentiation [81,82]. Consistent with the view that the antichondrogenic role of Wnt proteins is mediated through b-catenin, overexpression of a stabilized form of b-catenin in embryonic limb mesenchyme resulted in a near complete loss of all limb cartilage elements in the mouse, due to an early differentiation defect reflected by the loss of Sox9 expression in the limb mesenchyme [83]. In addition, canonical Wnt signaling also plays further inhibitory roles at subsequent stages after Col2a1 expression is activated, as conditional overexpression of the stabilized b-catenin by Col2-Cre led to severe achondrodysplasia [84]. Similar to canonical Wnt signaling, Notch signaling also suppresses chondrogenesis. Notch signaling mediates communication between neighboring cells to control cell fate decisions in all metazoans [85,86]. The mammalian genome encodes four Notch receptors (Notch1e4) and at least five ligands (Jagged1, 2 and Delta-like 1, 3, 4). In the canonical Notch pathway, binding of the ligands to the Notch receptors present on the neighboring cell surface triggers two successive intramembrane proteolytic cleavages of the receptors mediated by the g-secretase complex and resulting in the release of the Notch intracellular domain (NICD) [87e89]. Upon its release from the plasma membrane, NICD translocates to the nucleus where it interacts with a transcription factor of the CSL family (RBPJk/ CBF-1 in mammals) to activate transcription of target

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genes [90]. Expression studies revealed that multiple ligands and receptors are expressed in the prechondrogenic mesenchyme. Abolition of Notch signaling in the embryonic limb mesenchyme by the removal of either the g-secretase activity or the transcriptional effector RBPjk resulted in an acceleration of chondrocyte differentiation [91]. Conversely, forced expression of the constitutively active NICD in the limb mesenchyme completely abolished chondrogenesis. Importantly, the antichondrogenic effect of NICD was dependent on RBPjk, as simultaneous removal of the latter fully rescued the chondrogenic defect caused by NICD overexpression [91]. In addition to the direct suppressive role in chondrogenesis, Notch signaling is also known to control axial skeletal patterning through the regulation of somitogenesis. This was evidenced by the findings that Delta-like 3 (Dll3) [92], presenilin 1 (PS1) [93,94], a catalytic subunit of the g-secretase complex, or lunatic fringe, a glycosyltransferase that modifies Notch proteins [95] exhibited defects in the axial skeleton. Consistent with the mouse studies, human mutations in Dll3 [96] were found to cause spondylocostal dysostosis. Retinoid signaling via the retinoic acid receptor a (RARa) also acts to inhibit early chondrocyte differentiation [67,97]. In these studies, transgenic mice were generated to express a constitutively active RARa in the limb mesenchyme. Cells expressing the transgene did participate in mesenchymal condensations in vivo and in vitro but failed to differentiate into chondroblasts and maintained instead a prechondrogenic phenotype. In contrast, the addition of an RARa-selective antagonist to cultures of these cells was sufficient to stimulate chondroblast differentiation and cartilage formation, even when BMP signaling was repressed. Furthermore, inhibition of RAR activity correlated with PKA activation, increased Sox9 expression, and enhanced Sox9 transcriptional activity [67]. In the developing mandibular process, Sox9 and the homeobox transcription factor Msx2 are induced by BMP-4, and the relative amount of these two factors appears to determine where chondrogenesis takes place [98]. These two factors are also expressed in a subpopulation of cranial neural crests cells that will populate the mandibular area. In this population, Msx2 represses chondrogenic differentiation until cell migration is completed within the mandibular primordium [99]. It is not known whether the antagonistic interaction between Sox9 and Msx2 is direct or mediated by other factors.

Chondrocyte Proliferation and Maturation PTHrP (parathyroid hormone-related peptide) is a paracrine factor that shares homology with PTH (parathyroid hormone) in its N-terminal domain and binds

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a common receptor, the PTH/PTHrP receptor 1 (PTHR1). In developing long bones in rodents, Pthrp is expressed at highest levels by cells of the periarticular perichondrium, and at lower levels by the proliferating chondrocytes near the articular surface [100]. The Pth-r1 gene is expressed at low levels by proliferating chondrocytes and at high levels by the maturing chondrocytes adjacent to the Ihh-secreting cells, as well as osteoblastic cells in the perichondrium [101,102]. Targeting inactivation of the gene encoding either PTHrP or its receptor in the mouse resulted in neonatal lethality due to severe skeletal dysplasia in which premature maturation of chondrocytes led to short-limb dwarfism and excessive bone formation at birth [103]. Moreover, Pthr1/ chondrocytes present in the growth plate of chimeric mouse embryos developed from both Pthr1/eand wild-type ES cells underwent hypertrophy prematurely in a cell autonomous manner [104]. These studies identified a primary function for PTH signaling in the growth plate, which is to suppress the rate of chondrocyte hypertrophy. PTHrP acts by activating cAMP-dependent signaling pathways, in part by increasing PKA-dependent phosphorylation and transcriptional activity of Sox9 [105,106]. In keeping with the negative role of PTHrP in hypertrophy, misexpression of PTHrP in chondrocytes through a transgene markedly delayed chondrocyte maturation and bone formation, resulting in a completely cartilaginous endochondral skeleton at birth [107]. In humans, inactivating mutations in PTH-R1 cause Blomstrand chondrodysplasia characterized by advanced skeletal maturation with shortness of long bones and increased bone density [108e110]. Conversely, gain-of-function mutations in PTH-R1 cause Jansen metaphyseal chondrodysplasia [111,112], which was recapitulated in mice overexpressing such a mutant receptor [113]. Yet another activating mutation in PTH-R1 was identified in some enchondromatosis, a condition characterized by the presence of benign cartilage tumors adjoining the growth plate [114]. However, another research group could not confirm these findings [115], while others identified additional novel PTH-R1 mutations [116]. The expression of Pthrp in the periarticular region is strictly dependent on Indian hedgehog (Ihh), a member of the Hh family that includes additionally Sonic hedgehog (Shh) and Desert hedgehog (Dhh) in mammals. The Hedgehog (Hh) family of proteins plays fundamental roles in animal development conserved from flies to humans [117e119]. Smoothened (Smo), a seven-pass transmembrane protein is indispensable for transducing the Hh signal. Hh signaling ultimately controls the processing and subcellular localization of the Gli transcription factors (Gli1e3) that regulate expression of downstream target genes. In the developing cartilage, Ihh is primarily expressed by

prehypertrophic chondrocytes (chondrocytes immediately prior to hypertrophy) and early hypertrophic chondrocytes; Ihh signals to both immature chondrocytes and the overlying perichondrial cells [101,102]. Definitive evidence for the physiological function of Ihh signaling came from mouse genetic knockout studies. Ihh/ embryos exhibited multiple defects in the developing cartilage of the endochondral skeleton, these including a 50% reduction in chondrocyte proliferation and premature hypertrophy of chondrocytes [101, 120]. Subsequent genetic studies of Smo, revealed that whereas direct Ihh input was required for proper proliferation of chondrocytes [120,121], it did not appear to be critical for the regulation of chondrocyte hypertrophy, which instead depended primarily on PTHrP whose expression was induced by Ihh [101,120,122]. Later experiments however, did reveal that direct Ihh signaling does play a role within the proliferating population, in controlling the formation of the columnar from the round chondrocytes [123,124]. The control of PTHrP by Ihh appears to be through direct Hh signaling in the target cells, as localized removal of Smo led to a corresponding loss of PTHrP expression within the periarticular region [123]. Finally, the role of Ihh in chondrocyte proliferation and PTHrP expression is mainly mediated through the derepression of Gli3 repressor function, as simultaneous deleleton of both Gli3 and Ihh restored normal proliferation and hypertrophy [125,126]. Overall, these studies have led to a model in which Ihh and PTHrP jointly regulate chondrocyte proliferation and maturation. The insulin-like growth factors 1 and 2 (IGF-1 and IGF-2) are general growth-promoting factors that also directly regulate chondrocytes. These factors and their receptor (IGF-1R) are closely related to insulin and the insulin receptor (IR), respectively. IR and IGF-1R are tyrosine kinase receptors, activated as heterotetrameric complexes, which act via the docking protein insulin receptor substrate 1 (IRS-1) to initiate signal transduction by the mitogen-activated protein kinase or the phosphatidyl-inositol 3 kinase pathways [127]. IGF-2 (and to a lesser degree IGF-1) also interacts with a second receptor, the mannose-6-phosphate/IGF-2 receptor (IGF-2R). This is a monomeric receptor devoid of a signaling domain. It essentially acts as a scavenger to prevent accumulation of toxic levels of IGF-2 (and possibly IGF-1). These receptors are ubiquitously expressed, indicating the wide spectrum of cell types that insulin/IGF signaling acts on. IGF-1 and -2 are both locally produced by chondrocytes, in addition to the circulating IGF-1 produced in the liver that acts as an endocrine factor. Gene inactivation studies in the mouse have confirmed the essential role of the IGF signaling pathway in overall growth of the mouse embryo development, primarily through the control of cell proliferation [128e130]. Genetic experiments with

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single and compound mutant embryos in IGF and Ihh signaling indicated that the two signaling pathways appear to stimulate chondrocyte proliferation in parallel mechanisms [131]. Fibroblast growth factor (FGF) signaling is also recognized as an essential component of the control of chondrocyte proliferation [132]. FGFs are polypeptide growth factors with diverse biological functions [133]. The human and mouse genomes contain 22 FGF and four FGFR genes. Most FGFs function by binding to and activating cell surface tyrosine kinase FGF receptors (FGFR). FGFs also bind to heparin or heparan sulfate proteoglycans that facilitate the FGF-binding and activation of FGFRs. Signaling via FGFRs is propagated through recruitment and phosphorylation of a variety of signaling proteins, that is triggered by the autophosphorylation of the activated receptors and tyrosinephosphorylation of the closely linked docking proteins [134]. As a result, multiple signaling modules including MAPK, PI3K, STAT1 and PKC are activated. Activating mutations in the transmembrane domain of the receptor FGFR-3 cause a spectrum of chondrodysplasia including hypochondroplasia, achondroplasia and thanatophoric dysplasia, depending on the degree of activation of the receptor [132,135,136]. Similarly, mouse models harboring the activating mutations in FGFR3 phenocopied many aspects of these diseases and revealed that reduced chondrocyte proliferation was the main cellular defect [137,138]. Conversely, FGFR3 deletion in the mouse led to increased proliferation and postnatal skeletal overgrowth [139,140]. Thus, FGFR-3 signaling normally suppresses chondrocyte proliferation in the growth plate. FGFR-3 signaling in chondrocytes is likely to be activated by FGF-18, as FGF-18/ embryos exhibited a similar overproliferation phenotype in chondrocytes [141,142]. The intracellular signaling may involve the signal transducer and activator of transcription (STAT) family of transcription factors [143,144]. Others have shown that FGFs activated the MAPK pathway to upregulate Sox9 expression and activity during chondrogenesis [145].

Osteoblast Differentiation Regardless of their origin from either the intramembranous or the endochondral process, osteoblast differentiation requires the same key transcription factors. One such factor is Runt domain factor 2/core binding factor a1 (Runx2/Cbfa1). Runx2 is a transcription factor isolated by virtue of its binding to cis-regulatory elements of the gene encoding osteocalcin, an osteoblast-specific protein. Runx2 is first expressed in cells of the mesenchymal condensations believed to contain common osteochondroprogenitor cells. With development, its expression progressively increases in

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preosteoblasts and then in mature osteoblasts but is downregulated in chondrocytes except for the hypertrophic chondrocytes where Runx2 is expressed. Runx2/ Cbfa1 is indispensable for osteoblast differentiation, as demonstrated by knockout studies in the mouse. The skeleton of Runx2/ mice, which die at birth, is completely devoid of osteoblasts due to a differentiation arrest [146,147]. Runx2 is also required for osteoblast function beyond differentiation [148]. However, ectopic expression of Runx2 was not sufficient to induce osteoblast differentiation in the limb mesenchyme [149], and overexpression in osteoblasts caused osteopenia due to a blockage in osteoblast maturation [150]. Consistent with the fundamental importance of Runx2 in osteoblast differentiation and function, haploinsufficiency of Runx2 in humans causes cleidocranial dysplasia (CCD), an autosomal dominant disorder characterized primarily by a delay in closure of cranial sutures, reduced or absent clavicle, and dental anomalies [151]. Osterix (Osx, Sp7) is another transcription factor essential for osteoblast differentiation. Osx was discovered as upregulated in C2C12 cells by BMP during osteoblast differentiation [152]. Genetic deletion of Osx led to a complete lack of osteoblasts in the mouse embryo. The relatively normal Runx2 expression in the Osx/ embryos indicates that Osx functions in osteoblast differentiation downstream of Runx2. Furthermore, in the long bones of these mutant embryos, some cells normally destined to become osteoblasts adopted the chondrocyte phenotype, indicating that the Runx2-positive cells are bipotential and can differentiate into chondrocytes as opposed to osteoblasts in the absence of Osx. ATF4, a member of the basic leucine zipper family of transcription factors, plays an important role at a later stage than Runx2 and Osx during osteoblast differentiation [153]. Atf4/ embryos exhibited a severe deficit in the formation of mature osteoblasts even though Runx2 and Osx were expressed. In addition to its role in directly regulating the transcription of the mature osteoblast marker osteocalcin, Atf4 also controls the osteoblast phenotype by stimulating amino acid import to ensure a high level of protein translation of type I collagen, the main constituent of the bone matrix. Besides transcription factors, a number of developmental signals have been identified to control osteoblast differentiation in the embryo [154] (Fig. 3.2). Deletion of Ihh in the mice caused profound defects not only in chondrocytes, as discussed earlier, but also in osteoblast development in the endochondral skeleton [101,120]. In particular, loss of Ihh arrested osteoblast development at a primitive stage prior to expression of the earliest known markers including Cola1(I), AP and Runx2, and before the activation of canonical Wnt signaling in the lineage [155]. Thus, Ihh functions genetically upstream of canonical Wnt signaling during osteoblast

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Fgf, Bmp Hh

MSC

Wnt, Fgf

Runx2+

Wnt, Bmp

Osx+

OB

Notch

FIGURE 3.2 Multiple developmental signals control osteoblast

differentiation. Y: stimulation; t: inhibition. MSC: mesenchymal stem cells; OB: mature osteoblasts. (Figure modified from [154].)

development. Through genetic deletion of Smo to cellautonomously remove Hh responsiveness, it was shown that direct Ihh input was required in the perichondrial osteoprogenitors for development of the osteoblast lineage [120,121]. The control of Ihh over osteoblast differentiation requires both derepression of Gli3 repressor and activation of the Gli2 activator [156,157]. The importance of Gli2 activity is consistent with the finding that Gli2-null embryos (lethal at birth) showed impaired osteoblast formation [158]. The importance of canonical Wnt signaling in osteoblast differentiation during embryogenesis was suggested by genetic deletion of b-catenin from early osteogenic progenitors in the mouse [83,155,159,160]. These studies demonstrated that b-catenin is required for the progression both from Runx2- to Osterix-positive stage [155], and from Osx-positive cells to mature osteoblasts [160]. However, b-catenin appears to be dispensable in mature osteoblasts, as its deletion by Col1-Cre or OC-Cre did not affect osteoblasts per se, but instead decreased the expression of Opg by osteoblasts, and therefore indirectly enhanced osteoclast formation [161,162]. Additionally, non-canonical Wnt signaling mechanisms have also been shown to stimulate osteoblast differentiation [163,164]. Mouse genetic studies have also demonstrated the importance of BMP signaling in bone formation. A critical threshold level of BMP2/4 signaling was shown to be required for trabecular bone formation in the long bones, and specifically that the BMP signal controlled the progression from Runx2- to Osterix-positive cells [62]. Contrary to BMP2 and 4, BMP3 is a negative regulator of bone mass, as BMP3-null mice contained twice as much trabecular bone volume than the control littermates at 5e6 weeks of age [60]. It is possible that BMP3 inhibits osteoblastogenesis by counteracting BMP2, as suggested by in vitro studies [60]. Besides its role in osteoblast differentiation, BMP signaling was also shown to regulate the function of mature osteoblasts. Most notably, genetic deletion of BMPR1A using Og2-Cre (osteocalcin2-Cre) decreased osteoblast function

without an obvious effect on osteoblast numbers [165]. Similarly, overexpression of noggin, a secreted inhibitor for BMPs, under the osteocalcin promoter caused a reduction in osteoblast function and a lower bone mass in postnatal mice [166]. Mouse genetic studies have also revealed important roles for FGF signaling in the osteoblast lineage. Mice lacking the mesenchymal splice form of FGFR-2 exhibited a defect in osteoblast differentiation due to a partial loss of Runx2 [167]. Conversely, mice harboring a gain-of-function mutation in FGFR-2 showed an increase in osteoblast numbers associated with an increase in Runx2 expression and osteoblast differention [168]. Likewise, a gain-of-function mutation of FGFR-1 stimulated Runx2 expression and enhanced osteoblast differentiation in the calvaria, although the status of the long bones was not reported [169]. Tissue-specific deletion of FGFR-1 revealed that FGFR-1 signaling in osteoprogenitors promotes osteoblast differentiation without affecting Runx2 expression, whereas FGFR-1 signaling in mature osteoblasts inhibits the cells’ mineralization activity [170]. In contrast, mice lacking FGFR-3 showed a decrease in bone mineral density due to defects in mineralization, even though the number of osteoblasts was increased [171]. Finally, FGF-18-null embryos exhibited defects in the formation of mature osteoblasts despite normal Runx2 expression [141,142]. Thus, Fgf signaling likely promotes osteoblast differentiation, as well as proliferation and mineralization of mature osteoblasts. In contrast to the signals discussed above, Notch normally inhibits osteoblast differentiation. Genetic removal of either the g-secretase activity, or Notch recptors in the embryonic limb mesenchyme caused excessive bone formation that began in the embryo and culminated in adolescent mice [156]. The phenotype was due to increased osteoblast differentiation from the osteogenic precursors. Consistent with the negative role of Notch in osteoblast differentiation, forced-expression of NICD in osteoblastic precursors reduced osteoblast numbers and caused osteopenia [172]. Interestingly, forced-expression of NICD at a later stage of the osteoblast lineage led to sclerosis owing to excessive proliferation of the immature osteoblasts, highlighting stagespecific functions of constitutive Notch activation in the osteoblast lineage [173,174]. In humans, Notch1 haploinsufficiency caused ectopic osteoblast differentiation and calcification in the aortic valves [175,176].

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Postnatal Bone Growth: Growth Plate Biology, Bone Formation, and Remodeling Christa Maes 1, Henry M. Kronenberg 2 1

Assistant Professor, Katholieke Universiteit Leuven, Laboratory of Experimental Medicine and Endocrinology, Gasthuisberg O&N I, Leuven, Belgium 2 Professor of Medicine, Harvard Medical School; Chief, Endocrine Unit, Massachusetts General Hospital, Boston, Massachusetts, USA

ENDOCHONDRAL AND INTRAMEMBRANOUS BONE FORMATION

The precise arrangement of the individual anatomic elements of the skeleton involves actions and crosstalk of several morphogens including fibroblast growth factors (FGFs), sonic hedgehog (Shh), bone morphogenetic proteins (BMPs) and Wnts during fetal development [1]. These mechanisms underlying the early condensation, segmentation, differentiation and patterning events are reviewed in Chapter 3. The initial cartilage mold forms when mesenchymal cells condense and then differentiate into chondrocytes, under the direction of transcription factors including Sox-9 [2]. The chondrocytes synthesize a characteristic extracellular matrix (ECM) rich in collagens II, IX, and XI and aggrecan, among other matrix molecules. As such, a cartilaginous model or anlage is established that prefigures the future bone. The cartilage mold enlarges both through proliferation along the entire length of the mold and through matrix production. In the center of the mold, chondrocytes stop proliferating, enlarge (hypertrophy), and change the constituents of the matrix to include, for example, type X collagen. The matrix is subsequently mineralized, specialized osteoclasts called chondroclasts invade the region of hypertrophic chondrocytes along with blood vessels and preosteoblasts, and the hypertrophic chondrocytes undergo apoptosis. The invading cells come from the connective tissues surrounding the cartilage mold called the perichondrium. Perichondrial cells carry out a distinct genetic program that partly regulates the cartilage mold via the secretion of signaling molecules that regulate chondrocyte proliferation and differentiation, such as BMPs and parathyroid hormone-related peptide (PTHrP). In turn, the perichondrium is regulated by the cartilage. Perichondrial cells adjacent to prehypertrophic/

Intramembranous ossification is responsible primarily for the formation of most of the craniofacial elements, as will be discussed below. In contrast, all the long bones of the axial skeleton (vertebrae and ribs) and the appendicular skeleton (limbs) develop and grow through endochondral ossification (Fig. 4.1). This process encompasses the deposition of true bone matrix on top of scaffolding cartilaginous anlagen, and is controlled by a series of distinct cell types. The dramatic, asymmetric lengthening of the appendicular long bones is driven to a large extent by the proliferation, differentiation and matrix deposition of chondrocytes that form and make up cartilage, and of osteoblasts that build mineralized bone. Osteoclasts, giant multinuclear cells that break down (“resorb”) bone, contribute to the further modeling and remodeling of the deposited bone. In this lifelong process, packets of bone are constantly being removed and replaced to ensure an adequate shape, position, density and functionality of the mineralized bone framework. Bone growth is absolutely dependent on angiogenesis, putting vascular endothelial cells in the picture as another crucial cell type in bone. Over the last decades, intensive research has focused on the cellular and molecular control of bone development, growth and remodeling. In vitro cell systems have been extremely instructive, but insights derived from genetically altered mice and lessons from human inherited disorders have advanced the field tremendously, as reviewed in this chapter.


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4. POSTNATAL BONE GROWTH: GROWTH PLATE BIOLOGY, BONE FORMATION, AND REMODELING

FIGURE 4.1 Stepwise schematic diagram of long bone development through endochondral ossification. Around embryonic day 12 (E12) in mice, mesenchymal progenitor cells condense and differentiate into chondrocytes to form the cartilage anlagen that prefigures future long bones. Chondrocytes in the center become hypertrophic, while cells in the surrounding perichondrium differentiate into osteoblasts forming a bone collar, the provisional cortical bone. The hypertrophic cartilage core is subsequently invaded by blood vessels, eroded and replaced by bone and marrow (“primary ossification center”, POC). In the metaphysis, hypertrophic cartilage of the growth cartilage is continually replaced with trabecular bone, a process that relies on metaphyseal vascularization and mediates longitudinal bone growth. Around postnatal day 5 (P5), epiphyseal vessels invade the avascular cartilage and initiate a secondary center of ossification (SOC). Discrete layers of residual chondrocytes form growth plates between the epiphyseal and metaphyseal bone centers to support further postnatal longitudinal bone growth. Ultimately, (in humans) the growth plates close and growth stops. A detailed view of the perinatal bone structure (boxed area) is provided in Figure 4.2.

hypertrophic chondrocytes respond to Indian hedgehog (Ihh) and other signals from the chondrocytes to become pre-osteoblasts [3]. Some of these pre-osteoblasts differentiate into mature osteoblasts that lay down and mineralize a collagen I-containing matrix, the “bone collar”, around the cartilage mold. This bone collar forms the initiation site of the cortical bone, the dense outer envelope of compact, lamellar bone that provides the long bone with most of its strength and rigidity (see Fig. 4.1). The cortical bone grows by its remodeling by endosteal osteoclastic bone resorption and periosteal new bone formation, as pre-osteoblastic cells proliferate and differentiate. In humans, the cortical bone further enlarges by the formation of Haversian systems around central blood vessels. Other pre-osteoblasts formed in the perichondrium invade the developing bone along with the blood vessels and chondroclasts that accumulated in the region just prior to the cartilage invasion and erosion, and become true osteoblasts inside the new cavity [4]. These osteoblasts lay down bone matrix dominated by collagen type I on top of the calcified matrix left behind by the hypertrophic chondrocytes. This is the primary spongiosa, which is subsequently remodeled by further waves of osteoclasts and osteoblasts to form the secondary spongiosa, the forerunner of mature

trabecular bone. The marrow space is enlarged by continual osteoclast activity, and hematopoietic precursors migrate from the fetal liver to establish marrowbased hematopoiesis. Near the ends of the bone, the same cellular sequence of chondrocyte hypertrophy, vascular invasion, and bone formation subsequently occurs at so-called secondary ossification centers. The remaining discrete layers of residual growth cartilage between the expanding bone in the primary and secondary ossification centers at the opposing ends of the long bone are then called the growth plates [5]. The chondrocytes of the growth plates provide the prime engine for further postnatal longitudinal bone growth. This process is typified by precise temporal and spatial regulation of chondrocyte proliferation and differentiation. In fetal life, all non-hypertrophic chondrocytes vigorously proliferate. As a formal growth plate forms, however, the chondrocytes near the top of the growth plate slow down their rate of proliferation dramatically and are thought of as “reserve” or “resting” cells that may serve as stem cells for all other chondrocytes. Under controls that are poorly understood, resting cells periodically become proliferating chondrocytes that undergo several rounds of proliferation. These proliferating cells flatten out, with their short axis parallel to the long axis of the long bone, and form

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CELLULAR AND MOLECULAR CONTROL OF SKELETAL GROWTH

columns of stacked proliferating chondrocytes, with each column representing descendants of a distinct clone of cells. The lengthening of the bone derives primarily from the large number of proliferating chondrocytes and the matrix secreted by all of the chondrocytes, and from the substantial enlargement that each chondrocyte undergoes when it hypertrophies. Because the chondrocytes and their matrix ultimately disappear, however, normal osteoblast function in the primary spongiosa and in the area adjacent to the growth plate is also essential to allow the expansion of the bone length generated by chondrocyte activity to be translated into permanent lengthening. This fact is illustrated, for example, by the short bones of mice in which the essential transcription factor Runx2 was inactivated in osteoblasts [6,7]. Another absolute requirement for endochondral bone growth to proceed is the progressive neovascularization of the region where cartilage becomes replaced by primary spongiosa. Early studies revealed that blocking of the bone’s blood supply in vivo resulted in reduced longitudinal growth [8,9]. Molecularly, this process strongly depends on vascular endothelial growth factor (VEGF) signaling, as shown first by experiments inhibiting VEGF action in juvenile mice by administration of a soluble VEGF receptor chimeric protein (sFlt-1) [10]. These mice showed impaired vascular invasion of the growth plate and, concomitantly, trabecular bone formation and bone growth were reduced while the hypertrophic cartilage zone became enlarged. Thus, endochondral bone growth involves rigorous coupling of maturation and activity of chondrocytes, osteoclasts and osteoblasts, and vascular invasion. Ultimately, at least in humans, the growth plates completely disappear or “close” during puberty, in a process that actively requires the action of estradiol acting on estrogen receptor a (ERa) in both boys and girls (see below), and longitudinal bone growth stops. Remodeling of existing bone, replacing the primary spongiosa by lamellar bone in the secondary spongiosa and renewing the cortical bone, takes place throughout adult life, ensuring optimal mechanical properties of the skeleton and contributing to mineral ion homeostasis. This continual bone turnover is accomplished through the balanced action of osteoclasts and osteoblasts (see below) and results in a dynamic organization of honeycomb plate-like structures or trabeculae in the interior of the bone that are surrounded by blood vessels and bone marrow and housed within the cortical bone. Different from the endochondral ossification process outlined above, a small number of bones, primarily the flat bones of the skull, form without a cartilage mold by a mechanism termed intramembranous bone formation [11]. The development of these bones also starts with mesenchymal progenitor cells forming

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condensations at the sites where the bones will be formed [12]. Yet, in the mesenchymal condensations of intramembranous bones, cells do not differentiate into chondrocytes but instead differentiate directly into pre-osteoblasts and then osteoblasts that deposit “osteoid” or bone matrix rich in type I collagen. As the osteoblasts mature further, the bone matrix becomes progressively mineralized. The growth of these bones occurs by waves of proliferation and differentiation of pre-osteoblasts at the perimeter of the growing bones. As the perimeters of adjacent bones approach each other, a specialized structure, the suture, forms. At the suture, proliferating mesenchymal cells differentiate into osteoblasts. Normal development of the cranium requires that the skull expands in close coordination with the growth of the underlying brain; sutures normally do not close completely until brain expansion ceases [11,13e15]. Bone growth is regulated by a large variety of transcriptional, paracrine and endocrine signaling systems. In this chapter, we focus on a few such systems, chosen because of their established importance in normal human physiology and disease. The cellular and molecular control of skeletal development and growth is first considered, followed by a discussion of some of the prime endocrine controlled regulatory systems of bone growth and remodeling.

CELLULAR AND MOLECULAR CONTROL OF SKELETAL GROWTH Spontaneous mutations in humans and mice and experimental manipulation of genes, either deletions or overexpression (causing loss- or gain-of-function) have identified many growth and transcription factors and signaling cascades involved in skeletal development and growth by regulating the coordinated differentiation and/or actions of the various cell types of bone, namely chondrocytes, vascular endothelial cells, osteoblasts and osteoclasts.

Growth Plate Biology: Chondrocyte Proliferation and Differentiation In endochondral bones, the initial conversion of progenitor cells in the mesenchymal condensations into chondrocytes is determined by the expression of the key transcription factor Sox9, whereas the combined action of Sox9, 5 and 6 is required to direct the subsequent differentiation of chondrocytes throughout all phases of the chondrocyte lineage [16,17]. These phases encompass the round upper or “reserve” chondrocytes, the flattened, columnar proliferating chondrocytes, the

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FIGURE 4.2 Schematic view of the cellular structure of developing long bones. The epiphysis is composed of chondrocytes, organized in layers of proliferation (round periarticular and flat columnar proliferating cells), progressive differentiation towards the metaphysis (prehypertrophic and hypertrophic chondrocytes), and cell death (apoptosis). The avascular cartilage is supplied by the epiphyseal blood vessel network that overlays its surface. In the metaphysis, blood vessels invading the terminal hypertrophic chondrocytes, osteoclasts resorbing the cartilage, and osteoblasts building bone on the cartilage remnants, all act coordinately to replace the cartilage anlagen with bone and marrow.

pre-hypertrophic, and the hypertrophic chondrocytes, characteristically organized as stratified layers in the growth cartilage (Fig. 4.2). The progression of the chondrocytes through these stages is the driving force of actual bone lengthening and is tightly controlled by a myriad of local signaling molecules. Some of the best-characterized ones are the pathway governed by PTHrP and Ihh, and the FGFs; their importance is reflected by the fact that mutations in their signaling receptors are causal to a number of severe human dwarfing conditions. Mutations in the PTH/PTHrP receptor cause Jansen and Blomstrand chondrodysplasia [18,19], and constitutive activation of FGF receptor (FGFR)3 leads to human achondroplasia and thanatophoric dysplasia [20e23]. We will therefore briefly review the basic mechanisms by which these signaling

systems control the pace of proliferation and differentiation of growth chondrocytes. The PTHrP/Ihh pathway provides a major regulatory system in growth chondrocytes. Ihh, a member of the conserved family of hedgehog proteins, is produced by the pre-hypertrophic and early hypertrophic chondrocytes of the growth cartilage. By as yet not fully clarified mechanisms involving signaling through the receptor Patched (Ptc), Ihh acts directly on cells expressing PTHrP to stimulate the expression of PTHrP in chondrocytes located near the periarticular ends of the bone. PTHrP in turn signals back to its receptor (the PTH/ PTHrP receptor that also responds to PTH in osteoblasts and kidney) that is expressed at low levels by proliferating chondrocytes and strongly by pre-hypertrophic cells. The result of this PTHrP signaling is that the further chondrocyte differentiation to hypertrophy is slowed down and chondrocytes are kept in the proliferative state, thereby maintaining the adequate length of columns. By preventing premature hypertrophic differentiation, the generation of cells that can produce Ihh is consequently slowed down and, therefore, the production of PTHrP is lowered. Thus, PTHrP and Ihh regulate each other via a negative feedback signaling pathway that controls the pace of chondrocyte differentiation in the growth plate [24e26]. In addition, mutant mouse models revealed that Ihh regulates endochondral bone growth through PTHrP-independent mechanisms as well. These include a direct role of Ihh signaling in the regulation of chondrocyte proliferation and accelerated conversion of round to columnar cells, and Ihhinduced direct actions on the perichondrium controlling bone collar formation [27e30]. Perichondrial cells are driven into the osteoblast lineage under the influence of Ihh stimulating the expression of Runx2, an essential transcription factor in osteoblast development (see below). Conversely, Runx2 also regulates Ihh expression and hypertrophic differentiation [24,30,31]. Also beyond development, Ihh continues to be essential in regulating and maintaining the growth plate and sustaining postnatal bone growth, as shown by inducible mutagenesis in mice [32]. The FGF pathway involves multiple ligands and receptors that regulate cell proliferation, differentiation, and/or apoptosis in several processes, including skeletal patterning and bone growth. FGFs are involved in skeletogenesis at virtually every step from the initial outgrowth of the limb to postnatal lengthening of the long bones and formation of the intramembranous bones of the skull [33]. A total of 22 different FGFs have been identified in humans and mice. Expression of FGF-7, -8, -17, and -18 occurs in the perichondrium surrounding the growth plate, whereas FGF-2 is expressed in the periosteum of bone. FGF-2 and -9 are both expressed by

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chondrocytes, whereas FGF-2 is also expressed by osteoblasts. In the growing skull, FGF-2, -4, and -9 are expressed in the mesenchyme of the suture, whereas FGF-18 and -20 are expressed by differentiating osteoblasts [33]. Four genes encode receptors for the FGF family (called FGF receptors 1 to 4 [FGFR1e4]), although the actual number of receptors is larger because of splice variants of these genes. Each encodes a transmembrane protein with tyrosine kinase activity that is triggered by ligand binding at the plasma membrane. The extracellular domain contains three immunoglobulin domains that contribute to ligand-binding specificity and affinity. In the endochondral bones, FGFR3 is expressed by proliferating chondrocytes, FGFR1 is expressed by prehypertrophic and hypertrophic chondrocytes, and FGFR2 is expressed by perichondrial cells. In the sutures of the growing intramembranous bones of the skull, FGFR1 is found in the mesenchyme, whereas both FGFR1 and FGFR2 are expressed in the differentiating osteoblasts of sutures. FGFR3 is expressed late at the osteoblastic front of sutures [33]. A series of inherited human diseases and mutations in mice have demonstrated the importance of individual components of this complicated network of ligands and receptors in bone development. First, activating mutations in FGFR3 result in chondrodysplasias and dwarfism. Genetic analysis of the role of FGF signaling in the growth plate has, however, been complicated by the multiple roles of FGF signaling during early stages of development. Secondly, mutations in FGFR1 or 2 cause craniosynostosis, which results in skeletal dysplasias characterized by premature fusion of one or more cranial sutures, such as occur in Apert, Crouzon, and other syndromes [34]. The molecular mechanisms that contribute to these FGFR gain-of-functions mutations are complex and include constitutive (ligand-independent) or ligand-dependent FGFR activity [34]. The current insights in the FGF/FGFR signaling functions in these two settings, endochondral and intramembranous ossification, are briefly discussed here and further (see below), respectively. Humans with constitutively active FGFR3, due to point mutations such as a glycine-to-arginine mutation at residue 380 in the receptor’s transmembrane domain, have a form of dominantly inherited short-limbed dwarfism called achondroplasia. Both milder and more severe chondrodysplasias are caused by other activating mutations of FGFR3 [23]. Corresponding transgenic animal models have helped clarify the mechanisms whereby activation of FGFR3 leads to dwarfism. Activation of FGFR3 in humans and mice or overexpression of FGF-2 in transgenic mice lead to a decrease in chondrocyte proliferation in the growth plate and increased apoptosis. Conversely, mice missing the

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FGFR3 gene, normally expressed in proliferative chondrocyte columns, have elongated columns and longer bones than normal [33,35,36]. These findings were surprising since activation of FGF receptor tyrosine kinases typically leads to a positive effect on cellular proliferation in most tissues. Substantial evidence, however, suggests that FGFR3 activation in chondrocytes leads to activation of STAT1, which activates the cell cycle inhibitor p21Waf1/Cip1 and thereby decreases chondrocyte proliferation; the decrease in chondrocyte proliferation caused by FGF signaling in vivo and in vitro is blocked in chondrocytes from STAT1/ mice [37e39]. In addition, in transgenic mice, activation of FGFR3 leads to a decrease in expression of Ihh and its receptor and downstream target Ptc, whereas in mice lacking FGFR3, Ihh and Ptc expression are upregulated [40e42]. Since Ihh stimulates chondrocyte proliferation, this provides a further, indirect mechanism supporting the decrease in chondrocyte proliferation after activation of FGFR3. Besides modulating proliferation, FGFR3 signaling in vivo also appears to affect chondrocyte differentiation, as mouse models of achondroplasia and thanatophoric dysplasia show delayed hypertrophic differentiation of chondrocytes [42e44]. During chondrocyte differentiation, the FGF and BMP pathways appear generally to antagonize each other [45]. Moreover, BMP-4 expression is altered in FGFR3 mutant mouse models, and both FGF signaling and BMP signaling regulate Ihh production. Thus, each of these pathways has multiple mechanisms for communicating with each other as they regulate chondrocyte proliferation and differentiation. Due to the plethora of FGFs expressed in the skeleton and the many actions of FGFs early in development, it remains difficult to identify the specific FGF ligands that normally activate FGFR3 in the growth plate. FGF18 [46e48] and FGF-9 [49] appear to be the most important FGF ligands in regulating chondrogenesis identified so far. These ligands activate FGFR3 expressed on proliferating chondrocytes. In this context, the observation that mice missing FGF-18 have a growth plate phenotype that closely resembles that of the FGFR3 knockout mouse is interesting, since it suggests that FGF-18, synthesized in the perichondrium surrounding the growth plate, is the primary activator of FGFR3 in chondrocytes [46e48]. This phenotype thus highlights the importance of interactions between cells in the perichondrium and chondrocytes in the growth plate [3]. Besides the abovementioned growth factors, chondrogenesis is also affected by several other growth factor and hormones, including BMPs, growth hormone (GH) and insulin-like growth factors (IGFs), thyroid hormone, estrogen, androgen, 1a,25-dihydroxyvitamin D3 (1,25(OH)2D3), glucocorticoids, as discussed below and/or in other chapters.

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Blood Vessel Invasion of Cartilage and Bone Vascularization A crucial aspect of bone growth is the vascularization of the expanding bone center. This is particularly evident during endochondral ossification: the cartilage intermediate of the long bones represents a unique mesenchymal tissue in that it is itself avascular, while fully differentiated hypertrophic cartilage undergoes angiogenic invasion, a process that is associated with the decay of the terminal cartilage and its replacement by bone. The replacement of the initially avascular cartilage template by highly vascularized bone and marrow tissue occurs through three consecutive vascularization events (see Fig. 4.1). First, the initial vascular invasion of the cartilage anlagen during embryonic development (sometimes called quiescent angiogenesis) involves endothelial cells invading from the perichondrial tissues and organizing into immature blood vessels in the primary center of ossification. Second, capillary invasion at the metaphyseal border of the growth cartilage mediates rapid bone lengthening. Third, vascularization of the cartilage ends initiates the formation of secondary ossification centers (see Fig. 4.1). Vascularization is an absolute requirement for endochondral bone development and growth to proceed, as a physical blockage of blood vessel invasion into cartilaginous fetal skeletal explants completely halted their development [50], and blocking of the bone’s blood supply in vivo resulted in reduced longitudinal growth [8,9]. In fact, as a rule, any type of bone formation occurs in close spatial and temporal association with vascularization of the ossified tissue, a concept termed angiogeniceosteogenic coupling [51,52]. The reasons that the vascular system is crucial for bone growth, homeostasis and repair obviously include its intrinsic function to supply oxygen, nutrients and growth factors/hormones to the bone cells as required for their specified activities. On top of that, the blood vessels are thought also to serve to bring in (precursors of) osteoclasts that will degrade the cartilage or bone extracellular matrix, to remove end products of the resorption processes, and to bring in progenitors of osteoblasts that will deposit bone [4,53,54]. Here, we will briefly review the basic molecular players currently known to govern the survival of the avascular cartilage, its invasion by blood vessels and the vascularization of bone. Endochondral cartilaginous condensations typically develop for a prolonged period of time as an avascular tissue. Immature chondrocytes are resistant to vascular invasion due to the production of angiogenic inhibitors [55,56]. As the fetal growth plates expand in the absence of blood vessels during development, chondrocytes in the center of the cartilage consequently face the challenge of hypoxia [57,58]. The main mediator of the

cellular responses to hypoxia is the transcription factor hypoxia-inducible factor (HIF)-1. HIF-1 is a heterodimer of two proteins, HIF-1a and HIF-1b; HIF-1b is constitutively expressed, whereas HIF-1a is the hypoxia-responsive component of the complex [59]. Particularly the stability of the HIF-1a protein is hypoxia sensitive, through oxygen-dependent hydroxylation of specific residues within its amino acid sequence. In non-hypoxic conditions, a family of HIF prolyl 4-hydroxylases is responsible for the hydroxylation of two proline residues (P402 and P564) in the oxygen dependent degradation domain (ODDD) of HIF-1a. The E3 ubiquitin ligase Von Hippel-Lindau (VHL) binds to the hydroxylated HIF-1a, and targets it to the proteasome for degradation [60,61]. In hypoxic conditions, this hydroxylation of HIF1a does not occur, and HIF-1a is stabilized and able to translocate to the nucleus, heterodimerize with the b-subunit, and initiate its transcriptional program along with other co-factors. As a result, a variety of pathways involved in the cellular adaptation to hypoxia are activated, including key regulators of glucose utilization and cell metabolism (stimulating anaerobic glycolysis), angiogenesis, and erythropoiesis [62]. In the last years, studies using genetically modified mice have shed light on how the genetic program controlled by hypoxia modulates endochondral bone development, by establishing the essential and nonredundant role of HIF-1a in chondrogenesis in vivo [57,58,63,64]. Conditional deletion of HIF-1a in limb bud mesenchyme or in chondrocytes, achieved by the use of the Cre-loxP strategy, caused massive cell death of the inner chondrocytes in the developing growth plate. This cell death region corresponds to the central hypoxic region area of the fetal growth plate [58,65]. Thus, HIF-1a is required for survival of the hypoxic chondrocytes located in the center of the growth cartilage, furthest from blood vessels. The precise mechanisms by which HIF-1a prevents chondrocyte death have yet to be fully explained. Likely, the transcriptional activity of HIF-1a is, on the one hand, required for turning on vital oxygen-sparing metabolic pathways that allow chondrocytes to survive and differentiate in their hypoxic environment. Activation of anaerobic glycolysis in mammalian hypoxic cells is HIF-1a-dependent [66]. During skeletogenesis, phosphoglycerate-kinase1 (PGK), a key enzyme of anaerobic glycolysis, was found to be strikingly stronger expressed in developing cartilage than in the surrounding tissues; HIF-1a-deficient chondrocytes, however, failed to upregulate PGK expression [58]. On the other hand, HIF-1a may additionally provide an indirect survival effect by inducing the expression of its downstream target VEGF, a principal regulator of blood vessel formation [67]. Deletion of VEGF from cartilage causes a cell death phenotype in the center of the fetal growth plate that closely mimics

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what is observed in HIF-1a deficient growth plates [65,68]. Moreover, hypoxia increases VEGF accumulation in chondrocytes in vitro in an HIF-1a-dependent manner [69,70]. Thus, HIF-1a-mediated upregulation of VEGF and induction of angiogenesis in the perichondrial tissues surrounding the avascular cartilage is likely one of the mechanisms by which chondrocytes survive hypoxia. In addition to the low level of expression of VEGF noted in the central, hypoxic immature chondrocytes of the growth cartilage [65,68], VEGF is at all times expressed abundantly by late hypertrophic chondrocytes, where it is critical for blood vessel invasion and replacement of cartilage by bone [10,71e73]. Inhibition of VEGF action in juvenile mice by administration of a soluble VEGF receptor chimeric protein (sFlt-1) impaired vascular invasion of the growth plate; concomitantly, trabecular bone formation and bone growth were reduced and the hypertrophic cartilage zone became enlarged likely due to reduced osteoclast-mediated resorption [10]. Further mouse genetic studies univocally established that VEGF is an essential physiological mediator of all three key vascularization stages of endochondral bone development (see above and Fig. 4.1). These mouse models include the Cre/LoxP-mediated conditional inactivation of the VEGF gene in chondrocytes, and genetically engineered mice expressing only one of the three major VEGF isoforms [65,68,72e75]. Altogether, these models have exposed multiple essential roles of VEGF and its isoforms in endochondral ossification, not only as a key inducer of vascularization but also as a direct modulator of bone development by affecting the various cell types involved. Perichondrial cells, osteoblasts and osteoclasts are all well documented to express several of the VEGF receptors and to respond to VEGF signaling by enhanced recruitment, differentiation, activity and/or survival (reviewed in [71,76,77]. The current model is that VEGF is produced at high levels by hypertrophic chondrocytes and is partly sequestered in the cartilage matrix upon its secretion. Trapped VEGF can be released from the matrix by proteases such as matrix metalloproteinase (MMP) 9 secreted by osteoclasts/chondroclasts during cartilage resorption. VEGF can then bind to its receptors on endothelial cells and stimulate the guided attraction of blood vessels to invade the terminal cartilage [78]. Osteoblasts and osteoclasts, or precursors thereof, associated with the newly vascularized bone region may at the same time be affected by the VEGF signaling in their recruitment, proliferation, differentiation and function. These pleiotrophic actions of VEGF on the various cells in the bone environment may contribute to the tight coordination of vascularization, ossification, and matrix resorption that is characteristically seen in endochondral bone development and growth [71,77].

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Since the loss-of-function models pointed out that VEGF is a positive regulator of bone development, skeletal growth and fracture repair [10,72,73,79,80], VEGF appears as an interesting target for stimulating fracture healing [81]. While it is being currently tested in preclinical models as a potential bone regeneration therapy, data using mice overexpressing VEGF temporally in the skeleton suggest that increasing the tightly coupled processes of angiogenesis and osteogenesis might be associated with potential side effects including disturbances in hematopoiesis [82]. In general, the actions of VEGF are highly dose dependent, and its physiological levels must be under very strict control mechanisms. Although the regulation of VEGF expression in the skeleton is still largely unresolved and may vary depending on the developmental stage and specific location and cell type, several mechanisms have been implicated to date. Hypoxia is a crucial trigger of VEGF expression in chondrocytes, osteoblasts, and possibly osteoclasts, through mechanisms that at least in part involve HIF [69,70,83e85]. During early bone development, Runx2 may be an important inducer of VEGF expression and blood vessel invasion into cartilage [86]. In addition, several hormones (including PTH, GH, 1,25(OH)2D3) and locally produced growth factors (e.g. FGFs, transforming growth factor-b [TGFb], BMPs, IGFs, plateletderived growth factor [PDGF]) have been demonstrated to be involved, at least in vitro, in the regulation of VEGF expression (see [76]).

Ossification: Osteoblast Differentiation and Activity Osteoblasts are highly specialized cells able to deposit and mineralize large amounts of matrix rich in type I collagen. Like chondrocytes, osteoblasts are cells of mesenchymal origin; they are thought to descend from mesenchymal stem cells that are pluripotent and have the capacity to differentiate into a variety of cell types, including adipocytes, chondrocytes and osteoblasts [87e89]. The prime transcription factors required to directing the cells into the osteoblastic lineage are b-catenin, Runx2 (previously termed Cbfa1), and Osterix, as discussed below [90,91]. Once committed, pre-osteoblasts further differentiate into mature osteoblasts secreting type I collagen and other bone matrix components, together called osteoid. Subsequently, osteoblasts direct the mineralization of the osteoid in a process that requires active alkaline phosphatase (ALP), expressed on the osteoblast membrane. These mature osteoblasts express characteristic genes including the gene encoding osteocalcin. Ultimately, the cells die through apoptosis, are converted to bone lining cells or become embedded within the bone matrix as osteocytes. Osteocytes are the most abundant

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cell type in postnatal bone, playing roles in the sensing of mechanical forces, the regulation of bone formation and resorption, and the control of phosphate homeostasis [92,93] (see further). Here, we will discuss a number of crucial transcription factors and extracellular signaling molecules that have been shown to control various aspects of osteoblast biology, including Wnt/b-catenin signaling, Runx2 and Osx, and actions elicited by BMPs, hedgehogs (Hh), and FGFs. At least some of these are currently known to play a role in inherited human bone diseases. Wnt/b-catenin Signaling Wnts (wingless-type MMTV integration site) are a large family of secreted growth factors (19 different members in the mouse and human genomes) that play essential roles in multiple developmental processes. Wnts are also required for adult tissue maintenance, and perturbations in Wnt signaling can lead to tumor formation and other diseases [94,95]. In the skeletal system, mutations in Wnt signaling components lead to skeletal malformations and diseases such as osteoporosis-pseudoglioma and osteoarthritis (see below). Wnt pathway modulation, particularly neutralizing inhibitors of Wnt signaling, has emerged as a promising strategy to improve bone mass. These drugs are exciting breakthroughs but are not without risks; the challenges include tissue-specific targeting and consequently, long-term safety [96e98]. Wnts can transduce their signals through several different downstream signaling pathways. The best understood pathway is the canonical or Wnt/b-catenin pathway. Central to this pathway is the regulation of the protein stability of b-catenin, which acts as a transcription factor in the canonical Wnt pathway and is also involved in cell adhesion by binding to membranous cadherins. In the absence of Wnts, cytoplasmic b-catenin is constitutively degraded through its phosphorylation by glycogen synthase kinase 3-b (GSK3-b) that targets it to the ubiquitineproteasome pathway. Upon signaling by Wnts, b-catenin is stabilized and translocates to the nucleus, where it interacts with T-cell factor (TCF)/lymphoid enhancer factor (LEF) family transcription factors to regulate the expression of its target genes. In addition, other pathways affecting GSK3-b (e.g. phosphatidyl inositol 3 [PI3]-kinase pathways) can also modulate b-catenin transcriptional activity [94,95,99,100]. The importance of b-catenin in skeletal biology was proven by conditional b-catenin loss- and gain-offunction mouse models elucidating its role as a crucial transcription factor in (i) determining osteoblast lineage commitment of early osteo-chondroprogenitors [101e104] and (ii) coupling osteoblast to osteoclast activity [105]. The first set of studies provided evidence

that, in mesenchymal progenitor cells in the perichondrium and calvarium where cells are destined to become osteoblasts, Wnt signaling is high, leading to high levels of b-catenin inducing the expression of genes that mediate osteoblast differentiation (such as Runx2) while inhibiting transcription of genes required for chondrocyte differentiation (such as Sox9). In the absence of b-catenin, these cells become chondrocytes instead of osteoblasts, as revealed via genetic modifications in mice. Conversely, in the inner region of the condensations, Wnt signaling must be low, as these cells become chondrocytes (see [100]). Secondly, inactivation of bcatenin in differentiated osteoblasts [105] or osteocytes [106] revealed that its transcriptional activity is important for stimulating osteoblastic production of osteoprotegerin (OPG or TNFRSF11B, see below), an inhibitor of osteoclast formation. As such, b-catenin plays a role in the coupling between osteoblast and osteoclast activity, the central principle of bone remodeling and maintenance of bone mass (see below). Upon Wnt stimulation, the ligands bind to two synergistically acting families of Wnt (co)-receptors, the Frizzled (Fzd) receptor family members (10 known) and low-density lipoprotein receptor-related proteins (LRP5 or LRP6). This leads to the recruitment of axin to LRP5/6 in the plasma membrane. The sequestering of axin at the plasma membrane on its turn likely leads to the disassembly of the b-catenin destruction complex, thus mediating the stabilization and transcriptional activity of b-catenin. Activation of the pathway is normally constrained by the expression of secreted Wnt inhibitors such as Dickkopfs (Dkks), sclerostin, and secreted frizzled-related proteins (Sfrps). Members of the Dkk family and sclerostin bind to LRP5/6, and thereby block the ability of Wnt ligands to interact with these co-receptors. Proteins of the Sfrp family can directly bind to Wnt ligands, offering another way to inhibit activation of the pathway [94,95,98,107]. Mutations in the Wnt receptor complexes and Wnt antagonists have been clinically associated with changes in bone mineral density and fractures. Loss-of-function mutations in the LRP5 co-receptor cause the autosomal recessive disorder osteoporosis-pseudoglioma syndrome (OPPG), which is characterized by low bone mass, ocular defects, and a predisposition to fractures [108]. These findings were recapitulated in germline LRP5 knockout mice, which developed a low bone mass phenotype similar to patients with OPPG due to decreased osteoblast proliferation [109]. Conversely, gain-of-function mutations in the same group that render LRP5 with reduced affinities for the secreted antagonists Dkk1 and sclerostin result in high bone mass phenotypes [110e113]. There is debate as to whether these actions of LRP5 are direct or indirect actions on bone; argumentation to the latter is the

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finding that activation of LRP5 in the duodenum, rather than in osteoblasts, increased osteoblast proliferation by suppressing serotonin secretion; the lower blood levels of gut-derived serotonin lead to an increase in bone mass through less serotonin binding to its osteoblast receptor, 5-hydroxytryptamine (Htr1b), and activation of the cAMP responsive element binding protein (CREB) transcription factor [114]. Mutations in SOST have also been identified in patients, namely those diagnosed with van Buchem disease and sclerosteosis, diseases associated with high bone mass [115,116]. Altogether, these findings over the last decade highlighted the important role of canonical Wnt signaling in regulating human bone mass. Extensive research is therefore being conducted in this area based on the prospect that it can possibly lead to pharmacological intervention useful in the management of osteoporosis. Among the candidate therapeutics are small molecule inhibitors of GSK3b, neutralizing antibodies to Dkk1, secreted Frizzled-related protein 1, and sclerostin. Since osteocytes are the major producer of sclerostin, inhibition of this Wnt antagonist appears as a particularly promising strategy to prevent bone loss without causing adverse side effects, such as related to the tumorigenicity and toxicity to other tissues of activated Wnt signaling [96e98]. Runx2 and Osterix: Essential Transcription Factors in Osteoblastogenesis Runx2 (runt related transcriptional factor 2, previously known as core-binding factor a1 [Cbfa1], Osf2, or AML3) and Sp7/Osterix determine the osteoblast lineage from mesenchymal stem cells along with canonical Wnt signaling. In the process of osteoblast differentiation, these factors and canonical Wnt signaling molecules inhibit mesenchymal cells from differentiating into chondrocytes and adipocytes. Runx2, a transcription factor of the ancient runt family, is the earliest known marker essential for the production of the osteoblast lineage. Functional studies have demonstrated that it is a central regulator of osteoblast differentiation and function, and absolutely essential for the induction of osteoblasts and the formation of endochondral and intramembranous bone. The dominant role of Runx2 is illustrated by the fact that Runx2-deficient mice completely lack osteoblasts and do not form bone at all e instead a completely cartilaginous skeleton develops without any true bone matrix [7,117]. In heterozygous (haploinsufficient) Runx2 mutant mice, the defect in osteoblast differentiation is limited to intramembranous bones [117]. The resulting phenotype in these mice closely resembles the cleidocranial dysplasia (CCD) syndrome in humans, a dominantly inherited developmental disorder of bone, and RUNX2 is mutated in most CCD patients [118,119]. Consistent with its function as an early transcriptional regulator of osteoblast

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differentiation, Runx2 is also an early molecular marker of the osteoblast lineage, being highly expressed in perichondrial mesenchyme and in all osteoblasts. Pre-hypertrophic chondrocytes also express Runx2, and Runx2 plays important roles in cartilage biology as well. Runx2 can be sufficient to induce differentiation of mesenchymal cells into osteoblasts: its ectopic expression in non-osteoblastic cells leads to the expression of osteoblast-specific genes in vitro and in vivo [6]. Runx2 binds to an osteoblast-specific cis-acting DNA element present in the promoter of most genes expressed in osteoblasts; as such, Runx2 mediates osteoblast differentiation by inducing ALP activity, and regulating the expression of a variety of bone matrix protein genes, including the Col1a1, osteopontin, osteonectin, bone sialoprotein, and osteocalcin genes and Runx2 itself [120,121]. The regulation of different stages of osteoblast differentiation by Runx2 is, however, very complex; Runx2 transcriptional regulation involves interactions with a myriad of transcriptional activators and repressors and other co-regulatory proteins that are under continued investigation. The current model is that Runx2 triggers the expression of major bone matrix protein genes and the acquisition of an osteoblastic phenotype at an early stage of osteoblast differentiation, while inhibiting the late osteoblast maturation stages and the transition into osteocytes. As such, Runx2 may play an important role in maintaining a supply of immature osteoblasts. Runx2 must thus be suppressed for immature osteoblasts to become fully mature osteoblasts, which form mature bone with regularly and densely packed collagen fibrils and high mineralization [120,121]. Furthermore, Runx2 regulates the expression of RANKL and OPG in osteoblasts, thus affecting osteoclast differentiation (see below). Runx2 is also required for the expression of Osterix (encoded by the Osx or Sp7 gene), an SP family transcription factor with three zinc finger motifs. Osterix is expressed in osteoblast progenitors, osteoblasts, and at a lower level also in pre-hypertrophic chondrocytes [4,122]. Like Runx2-deficient mice, mice lacking Osterix showed complete lack of osteoblasts and absence of both intramembranous and endochondral bone formation [122]. Thus, Osterix is a third transcription factor that is essential for osteoblast differentiation. Since Runx2 is expressed in the mesenchymal cells of Osx-null mice but Osterix is not expressed in Runx2-null mice, it can be concluded that Osterix acts downstream of Runx2 [122]. Furthermore, the Osx gene contains a consensus Runx2-binding site in its promoter region, suggesting that Osterix might be a direct target of Runx2 [123]. The transcriptional activity of Osterix involves its interaction with NFATc1 cooperatively forming a complex that binds to DNA and induces the expression of Col1a1 [124]. The subtleties of how Osterix regulates

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osteoblast differentiation and function, and which osteoblastic genes are directly regulated by Osterix are as yet to be elucidated. Expression of genes characteristic of mature osteoblasts (such as those encoding bone sialoprotein, osteopontin, osteonectin, osteocalcin) was absent in cells surrounding chondrocytes in Osx-null mice, and instead these cells express genes characteristic of chondrocytes (Sox9, Sox5, Col2a1) [122]. Osterix has also been reported to inhibit chondrogenesis in vitro [125,126]. Thus, Osterix may be important for directing precursor cells away from the chondrocyte lineage and toward the osteoblast lineage. Besides b-catenin, Runx2, and Osterix, various other (non-bone-specific) transcription factors are also involved in the genetic control of osteoblast differentiation and function, albeit not to a similar critical extent as their inactivation does not completely abrogate bone formation. These include ATF4, Msx1 and Msx2, Dlx5, Dlx6 and Dlx3, Twist, activator protein-1 (AP1) and its related molecules (Fos/Jun), and Schnurri-2 and -3 (for reviews and references therein, see [91,120,127,128]). Secreted Local Signaling Molecules Involved in Osteoblastogenesis A myriad of morphogens and signaling molecules control the activity of the transcription factors just described. These molecules include Wnts, as discussed before, as well as locally produced BMPs and TGF-b, Hedgehogs (particularly Ihh), FGFs, Notch ligands, IGFs, PDGF, and systemic factors like PTH, GH, prostaglandins, estrogens, androgens, 1,25(OH)2D3 and glucocorticoids. The roles of the signaling pathways involving BMPs, Ihh, and FGFs in osteoblastogenesis and bone formation are briefly reviewed here; the endocrine control of bone growth and remodeling is provided further (see below and other chapters). The family of growth factors that has received a tremendous amount of attention is that of the BMPs. These are members of the transforming growth factorb superfamily of growth factors that can, when applied locally, induce de novo bone formation; some BMPs, including BMP-2 and BMP-7, are therefore clinically used in orthopedics [129,130]. BMPs transduce signals through serine/threonine kinase receptors, composed of type I and II subtypes, activating intracellular Smads that relay the BMP signal to target genes in the nucleus [131, 132]. The analysis of their functions in vivo has relied mostly on gene deletion experiments. Although several BMPs affect skeletal patterning and joint formation, it has proved difficult using this approach to elucidate how they affect osteoblast differentiation. BMPs are crucial for regulating development in almost all the principal organs and tissues [131,133]. In the morphogenesis of the skeleton, BMPs have been

implicated in early limb patterning, mediating the correct formation, size or shape of the mesenchymal condensations, as well as converting the condensing mesenchymal cells into chondrocytes [134e136]. Their functions in osteogenesis during development, postnatal bone formation and remodeling, and fracture repair have therefore been hard to document in vivo because, in some cases, the inactivation of specific BMPs led to severe early defects and lethality (e.g. BMP-2, -4), precluding investigation of the later stages. In other cases, removal of individual family members (such as BMP-7) displayed no defects in skeletogenesis, presumably due to functional redundancy between the various BMP ligands and receptors [137]; around 20 BMP family members have been identified to date, and perichondrial cells, osteoblasts, and chondrocytes express multiple BMPs, BMP receptors, and BMP antagonists. Recent and ongoing studies therefore employ conditional (site-specific) and/or combined (multiple targets) mutagenesis strategies. As such, the double knockout of BMP-2 and BMP-4 in the limb completely disrupted osteoblast differentiation, demonstrating the crucial roles of these two BMPs in osteoblast differentiation [138]. Effects of BMP signaling in later stages of the osteoblast differentiation are suggested by studies using BMP antagonists. Targeting noggin overexpression to differentiated osteoblasts by the osteocalcin promoter results in osteopenia by 8 months of age [139]. Likewise, overexpression of gremlin, another BMP antagonist, in differentiated osteoblasts results in reduced bone mineral density and fractures [140]. On the other hand, a recent report using an osteoblast-targeted deletion of BMP receptor signaling indicated that BMP signaling in osteoblasts physiologically induces bone resorption by enhancing osteoclastogenesis via the RANKL-OPG pathway, thereby reducing bone mass [141]. Overall, signaling by BMPs plays an important role in a variety of cell types in bone such as osteoblasts, chondrocytes, and osteoclasts, and the precise mechanisms underlying their actions during bone growth have not yet been fully elucidated due to the considerable complexity and involvement of a myriad of interacting pathways. Nevertheless, particularly BMP-2 has been shown to play a critical role in osteogenic differentiation: it promotes the commitment of pluripotent mesenchymal cells to the osteoblast lineage [142,143] and has been demonstrated to induce the expression of both Runx2 and Osx during osteoblastogenesis [144e147]. The activity of BMP-2 as a potent inducer of bone formation is being applied to repair bone defects in humans [129,130,148]. Moreover, a human genetic study indicated that polymorphisms of BMP-2 gene expression are linked to a high risk for osteoporosis [149].

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The Hedgehog family member Ihh is required for endochondral but not for intramembranous bone formation, by controlling osteoblast differentiation in the perichondrium of long bones. This statement is based on the finding that Ihh-deficient mice have disorganized growth plates, as expected (see above), but also completely lack osteoblasts in bone formed through endochondral ossification; however, these do appear to be present in the skull, the mandibles, and the clavicles e bones that form by intramembranous ossification [30,150]. In these mice, Runx2 is expressed in chondrocytes but not in perichondrial cells. As well, nuclear b-catenin is absent in the perichondrial cells of Ihh-deficient mice [103]. Thus, Ihh is required for inducing the initial activity of Runx2 and b-catenin in perichondrial cells and triggering them to become endochondral osteoblasts, thereby coupling chondrocyte maturation to osteoblast differentiation during endochondral ossification. Chondrocyte-derived Ihh remains crucial for sustaining trabecular bone also in the postnatal skeleton [32]. Skeletal abnormalities have been described in mutant mice lacking some of the intracellular mediators of Ihh signaling termed Gli proteins (three related transcription factors Gli1, Gli2 and Gli3); Gli2 mediates Ihh-induced osteoblast differentiation in mesenchymal cell lines by associating with Runx2 and stimulating its expression and function, as well as inducing BMP-2 expression [151,152]. FGF signaling has also been implicated in the proliferation of immature osteoblasts and the anabolic function of mature osteoblasts in vivo [46,48,153e155]. Whether and how osteoblast differentiation per se is affected by FGFs is currently elusive, but evidence indicates that FGF signaling induces BMP-2 expression and stimulates the expression and transcriptional activities of Runx2 [156e158]. Conversely, Runx2 has been demonstrated to form a complex with Lef1 or TCF that binds to the promoter region of the gene encoding FGF18, inducing its expression [159]. These findings underscore once more how the various pathways that are essential for endochondral and intramembranous bone development physically and functionally converge. Moreover, the ultimate outcome relies on the complex integration of both stimulatory and inhibitory signaling. In any case, FGFs and their receptors have important roles in the growth of intramembranous bones and the bone adjacent to growth plates in endochondral bone growth. In the cranial vault, inappropriately rapid differentiation of osteoblasts at the sutures, when combined with normal or increased cell proliferation, can lead to craniosynostosis, the premature fusion of sutures. This is a serious condition because it does not allow the normal growth of the brain and neural structures. As noted earlier, FGFs and their receptors are expressed in sutures. Activating mutations of FGFR1 or 2 in humans

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can cause craniosynostosis [34]. The ligands most crucial for activating FGF receptors in sutures and osteoblasts of the limb are uncertain. However, genetic evidence in mice established that both FGF-2 and FGF-18 are important [46e48,158]. Thus, multiple FGFs and FGF receptors are needed for normal osteoblast production and differentiation.

Bone (Re)Modeling: Osteoclastogenesis, Bone Resorption, and the Coupling Principle The Role of Osteoclasts in Bone Modeling and Remodeling Osteoclasts are giant multinucleated cells that have the unique capacity to efficiently degrade calcified tissues. During development and growth of the endochondral bones, the main tissue to be resorbed is the calcified cartilage matrix produced by hypertrophic chondrocytes. Coinciding in time and place with the vascular invasion of the terminally differentiated chondrocytes, the calcified cartilage is co-invaded by osteoclasts, or a postulated related cell type termed “chondroclasts” [160]. Osteoclasts are not strictly essential for the excavation of the marrow space and its colonization by blood vessels and bone-forming osteoblast lineage cells to replace the hypertrophic chondrocytes during endochondral bone development [4,161]. However, the degradation, digestion and removal of the septa of calcified cartilage does require osteoclasts, as mice lacking osteoclasts, such as in the case of receptor activator of nuclear factor-kB (RANK)-deficiency (see below), display poorly remodeled osteocartilaginous structures instead of normal trabecular bone [4,161]. Moreover, such mice present with disorganized growth plates and impaired longitudinal bone growth, leading to a small body size and short limbs [161]. Not surprisingly, antiresorptive drugs such as bisphosphonates can interfere with bone modeling (shaping) during growth [162,163]. Bone, like many other organs in the body, goes through a tremendous amount of destruction and growth during childhood. Bisphosphonates potently decrease the activity of osteoclasts, which can lead to the accumulation of growth plate residues within trabecular bone tissue. Calcified cartilage has a high mineral density and therefore contributes to increase densitometric results, but is less resistant to fractures than is normal bone. Low remodeling activity might also delay bone healing after injury. This is of clinical interest, for instance in the treatment of juvenile osteoporosis during growth and of osteogenesis imperfecta (OI). OI is a genetic disease characterized by increased bone fragility and low bone mass due to the synthesis and secretion of defective type I collagen

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molecules by the osteoblasts [164]. In the majority of OI patients, the disease is inherited in an autosomal dominant manner and can be linked to mutations in one of the two genes encoding collagen type I a chains (COL1A1 and COL1A2). This can be viewed as a near complete arrest of the normal osteoblast function in the most severe forms. Human OI patients exhibit characteristic short stature, which has been attributed to increased incidence of bone fractures, defective bone matrix production, and poor mineralization [165], hence illustrating the critical role played by the osteoblasts in the control of longitudinal growth. Disrupted growth plate structure was observed in severe OI patients and in various mutant mouse strains modeling the disease [165,166]. Pamidronate, a nitrogen-containing bisphosphonate that interferes with osteoclast activity, is now used worldwide to treat children and adolescents with moderate to severe forms of OI [167,168]. The hypothesis initially underlying the use of an anti-osteoclast medication in an osteoblast disorder such as OI was that a decrease in the activity of the bone resorbing system might compensate for the inadequate functioning of the bone-forming cells. This treatment has been reported to increase lumbar spine bone mineral density and metacarpal cortical width, to decrease fracture rate, and to improve mobility. These results are very encouraging, but the fact that bisphosphonates so effectively inhibit osteoclastic action raises concern that this form of treatment could further compromise longitudinal bone growth in OI children; therefore, treatment with bisphosphonates during growth is mainly reserved for patients with significant clinical problems [164,167,168]. Throughout further postnatal life, osteoclasts primarily act to degrade mineralized bone during the constant process of bone remodeling. Indeed, bone is continuously renewed via a process of self-destruction followed by a regeneration process. Its purposes are to repair damaged bone, remove old bone, and facilitate skeletal responses to changes in loading requirements and physiologic needs in ensuring mineral homeostasis. This dynamic nature of the skeleton is achieved by the coordinated actions of osteoclasts, osteoblasts, osteocytes within the bone matrix and osteoblast-derived lining cells that cover the surface of bone. In short, remodeling initiates with signals that stimulate osteoclast formation, followed by osteoclast-mediated bone resorption, a reversal period, and then a long period of bone matrix formation and mineralization mediated by osteoblasts [169,170] (Fig. 4.3). Packets of the existing trabecular and cortical mineralized bone tissue that are renewed during remodeling are called bone remodeling units (BRUs) or bone multicellular units (BMUs) [169]. The bone in an activated BRU is first removed by osteoclastic bone resorption in a process that takes a few weeks (see Fig. 4.3). During

FIGURE 4.3 Three-phase model of bone remodeling. The skeleton is a metabolically active organ that undergoes continuous remodeling throughout life. Bone remodeling involves the removal of mineralized bone by osteoclasts followed by the formation and subsequent mineralization of bone matrix by osteoblasts. Initiation starts with recruitment of hematopoietic precursors and their differentiation to osteoclasts, induced by osteoblast lineage cells that express osteoclastogenic ligands such as RANKL. Osteoclasts become multinucleated and resorb bone. Transition is marked by switching from bone resorption to formation via coupling factors, possibly including diffusible factors (e.g. hormones), membrane-bound molecules (e.g. ephrins), and factors embedded in bone matrix that become released upon osteoclastic bone resorption and can stimulate osteoblast recruitment, differentiation and/or activity. During the termination phase, the resorbed lacuna is refilled through bone formation by osteoblasts that later flatten to form a layer of lining cells on the bone surface or become osteocytes connected by canaliculi within the bone.

the time lag that occurs between the end of resorption and the beginning of formation, called the reversal phase, osteoblastic cells differentiate on the bone surface and deposit the organized matrix that then becomes mineralized. Hence, the lost bone is replaced by osteoblastic bone formation, lasting 3e4 months for one packet. This discrepancy in the kinetics of bone resorption and formation partly explains how increased resorption, even when accompanied by coupled increased formation, can cause bone loss, for example, in estrogen deficiency or hyperparathyroidism [169]. Excessive osteoclast activity can be the basis of several pathological conditions including osteoporosis, a common low bone mass disorder typically prevalent in postmenopausal women, but also periodontal disease, rheumatoid arthritis, multiple myeloma, Paget’s disease and metastatic cancer. On the other hand, impaired osteoclast differentiation and/or function leads to osteopetrosis, a rare human disease characterized by increased bone mass and obliteration of the bone marrow cavity. Many efforts have been made to dissect the regulatory pathways controlling osteoclast differentiation and function; the most important of those are reviewed here.

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Osteoclastogenesis Osteoclasts derive from hematopoietic precursor cells of the myelomonocytic lineage and share a common precursor with macrophages. The early differentiation of the bipotential macrophage/osteoclast precursor cells is regulated by PU.1, a versatile hematopoietic cellspecific transcriptional regulator of the ETS-family. Particularly, the commitment to macrophages/osteoclasts is dependent on high level/activity of PU.1; consequently, PU.1 null mice lack both macrophages and osteoclasts, and are osteopetrotic [171,172]. This is a cell autonomous defect that can be corrected by bone marrow transplantation [172]. PU.1 regulates the lineage fate decision of the early progenitors by directly controlling expression of the c-Fms gene that is a key determinant of differentiation into the macrophage/osteoclast lineage (see below) [173]. PU.1 also regulates the transcription of another key osteoclastogenesis control gene, receptor activator of nuclear factor-kB (RANK) (see below) in myeloid progenitors [174]. Although mature macrophages and osteoclasts have some cell surface markers in common, the latter express high levels of tartrate-resistant acid phosphatase (TRAP), cathepsin K, vitronectin, calcitonin receptor and avb3 integrin, that typify the osteoclast lineage (see below). PU.1 is the earliest marker of osteoclast differentiation; yet, the signals that induce its expression and that of other cell-specific transcription factors involved in osteoclastogenesis are as yet unresolved. Among the other transcription factors that control osteoclast differentiation and function is NF-kB, which appears to play a role early during osteoclast differentiation. NF-kB is formed as a dimer composed of various combinations of proteins: p50, p52, p65, c-Rel, RelA and RelB [175]. Mice deficient in both p50 and p52 harbor an osteopetrosis phenotype due to an arrest of osteoclast differentiation; the presence of a large number of macrophages in these mice places NF-kB downstream of PU.1 in the genetic pathway controlling osteoclast differentiation [176,177]. NF-kB increases the expression of another transcription factor, c-Fos, which is the cellular homolog of the v-Fos oncogene and a major component of the activator protein (AP)-1 transcriptional complex [178]. The deletion of c-Fos in mice caused an early arrest of osteoclast differentiation leading to osteopetrosis, while again macrophage numbers were enhanced [179,180]. c-Fos interacts with the master transcription factor for osteoclastogenesis, nuclear factor of activated T cells c1 (NFATc1), to induce osteoclast-specific genes such as those encoding TRAP and calcitonin receptor (see below) [181,182]. After the precursor cells have committed to the osteoclast lineage, they become subjected to a complex multistep process culminating in the generation of

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mature, multinuclear activated osteoclasts; these steps include proliferation, maturation and fusion of differentiating precursor cells, and finally activation of resorption. Several secreted molecules and signaling proteins control the process of osteoclast differentiation and function. The requirement for secreted molecules to control osteoclastogenesis was first documented with the genetic elucidation of a classical mouse mutation called op/op [183]. Mice homozygous for this recessive mutation lack osteoclasts and macrophages. The osteopetrotic phenotype of op/op mice is not cured by bone marrow transplantation, indicating that it is non-cell-autonomous defect [116,183]. The gene mutated in op/op mice encodes the growth factor macrophage colony-stimulating factor (M-CSF) [183]. The second critical cytokine essential for osteoclastogenesis is receptor activator of nuclear factor-kB ligand (RANKL) (also known previously as osteoprotegerin ligand [OPGL], osteoclast differentiation factor [ODF], or tumor necrosis factor [TNF]-related activationinduced cytokine [TRANCE]). As M-CSF, this signaling molecule is also strongly expressed by bone marrow stromal cells (i.e. osteoblast progenitors) and osteoblasts; M-CSF is produced in both a soluble and a membrane-bound form, while RANKL is made exclusively as a membrane protein by osteoblasts. The process of osteoclastogenesis requires direct cell-to-cell interaction between stromal/osteoblastic cells and osteoclast precursors presumably because of these key membrane-bound ligands. Both PTH and 1,25(OH)2vitaminD3 as well as several other osteotropic factors stimulating resorption increase the expression of RANKL on stromal/osteoblastic cells. Furthermore, not only osteoblasts but also chondrocytes and T cells synthesize and secrete RANKL and are able to support osteoclastogenesis. This may be important physiologically during osteoclast differentiation and invasion at the hypertrophic cartilage during endochondral bone development and growth, and definitely plays a major role pathologically. For instance, production of RANKL by T cells has been implicated as an activator of osteoclastic resorption in inflammatory-mediated bone and cartilage destruction such as seen in several autoimmune disorders including rheumatoid arthritis, likely working synergistically with TNF-a (for reviews, see [181,182,184e186]. M-CSF and RANKL affect several steps of the osteoclastogenesis cascade by binding to their respective receptors, c-Fms and RANK, that are expressed by osteoclast progenitors and osteoclasts at all stages of differentiation. M-CSF signaling is essential early on in the lineage for proliferation, differentiation and survival of the osteoclast/macrophage precursors. RANKL is an essential inducer of multiple aspects of osteoclastogenesis, including osteoclast differentiation, fusion, activation of

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mature osteoclasts to resorb mineralized bone, and cell survival [181,182,184e186]. Through these actions, RANKL potently stimulates bone resorption. RANKL acts via binding to its signal transducing receptor RANK, a member of the TNF receptor family present on pre-osteoclasts and mature osteoclasts. Consistent herewith, an activating mutation of the human RANK gene was found in patients with familial expansile osteolysis, a disease of excess bone resorption [187]. Recent genetic and cell biological studies have begun to elucidate the complex signaling cascade downstream of RANK/ RANKL (reviewed by [181,182,186]). Briefly stated and oversimplified, the binding of RANKL to RANK on the surface of osteoclast precursors recruits the adaptor protein TNF receptor-associated factor (TRAF) 6 to the cytoplasmic domain of RANK, leading to activation of NF-kB and its translocation to the nucleus, and the actions of c-Fos and NFATc1 as outlined above. The actions of RANKL are negatively regulated by a secreted soluble decoy receptor termed osteoprotegerin (OPG) (previously also known as osteoclastogenesis inhibitory factor [OCIF]), also a member of the TNF receptor superfamily. OPG sequesters RANKL molecules and thereby blocks their binding to RANK. As such, OPG protects bone from excessive resorption; this conclusion is supported by the finding that certain homozygous deletions of OPG in humans can cause juvenile Paget’s disease, a disorder characterized by increased bone remodeling, osteopenia, and fractures [188]. The relative concentration of RANKL and OPG in bone is thus a major determinant of bone mass and strength. OPG is widely expressed; not surprisingly, its expression by osteoblasts and stromal cells is positively regulated by bone anabolic or antiresorptive factors, such as estrogen and calcitonin (see [184,186]). The essence of the RANK/RANKL/OPG cascade was clarified by mouse genetic studies. Mice lacking RANKL or RANK and mice with increased circulating OPG by transgenic overexpression were severely osteopetrotic due to a block in osteoclastogenesis. Conversely, targeted mutagenesis of OPG, overexpression of an sRANKL transgene, and administration of RANKL in mice, all led to increased osteoclast formation, activation and/or survival and resulted in an osteoporotic phenotype (see [184,186]). In summary, RANKL and OPG act in an antagonistic fashion to regulate bone resorption and their respective expression levels are under the control of pro- and antiresorptive factors including several hormones, cytokines and growth factors. Other regulatory molecules have been implicated in the late stages of osteoclast differentiation, the fusion process and activation of resorption. Co-stimulatory molecules acting in concert with M-CSF and RANKL to complete osteoclastogenesis include proteins containing an immunoreceptor tyrosine-based activation motif

(ITAM) domain that is critical for the activation of calcium signaling and found in adapter molecules like DNAX-activating protein (DAP)12 and the Fc receptor g (FcRg) [189]. The resultant increase in intracellular calcium is required for activation of NFATc1. The fusion of mononuclear osteoclast precursor cells into mature multinucleated osteoclasts is regulated by a membrane protein called dendritic cell-specific transmembrane protein (DC-STAMP). DC-STAMP-deficient cells failed to fuse, and these mononuclear osteoclasts had reduced resorptive efficiency in vitro. Consequently, DCSTAMP-deficient mice exhibited increased bone mass [190]. Further studies on the regulatory components of osteoclastogenesis will not only expand our basic understanding of the molecular mechanisms of osteoclast differentiation during bone development and remodeling, but also increasingly offer opportunities to develop therapeutic means of intervention in osteoclast-related diseases [191,192]. OPG, as well as soluble recombinant RANK, suppresses osteoclastogenesis, while antibodies to RANK can stimulate osteoclast formation. From the clinical point of view, the RANKL signaling pathway thus holds great promise as a strategy for suppressing excessive osteoclast formation in variety of bone diseases including osteoporosis, autoimmune arthritis, periodontitis, Paget’s disease, and bone tumors/metastases. One drug that has been developed to target RANKL signaling for the treatment or prevention of bone disease is denosumab, a human monoclonal antibody that binds to RANKL and prevents RANKL interaction with RANK [193]. Bone Resorption Bone resorption involves both dissolution of bone mineral as well as degradation of organic bone matrix. Osteoclasts are highly specialized to perform both of these functions. Upon activation of the mature multinucleated osteoclasts, the cells attach themselves firmly to the bone surface, using specialized actin-rich podosomes (actin ring), through cytoskeleton reorganization and cellular polarization [194]. Within these tightly sealed zones of adhesion to the mineralized matrix, the osteoclasts form convoluted, villous-like membranes called “ruffled borders” that drastically increase the surface area of the cell membrane facing the resorption lacuna. Via these ruffled membranes the osteoclasts secrete abundant hydrochloric acid (involving the vacuolar Hþ-ATPase proton pump) mediating acidification of the compartment between the cell and the bone surface, as well as a myriad of enzymes such as lysosomal cathepsins, the phosphatase TRAP, and proteolytic MMPs. The acidity of the environment leads to dissolution of the mineral phase (crystalline hydroxyapatite), activation of the lytic enzymes, and digestion of

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organic matrix compounds. The sealing mechanism allows the localized dissolving and degrading of the mineralized bone matrix, while simultaneously protecting neighboring cells from harm. During the resorption process, dissolution of hydroxyapatite releases large amounts of soluble calcium, phosphate and bicarbonate [195,196]. These complex processes of osteoclast recruitment, polarization on the bone surface, and export of acid and enzymes are orchestrated by many factors, including RANKL, as well as integrin-mediated signaling from bone matrix itself. Particularly the avb3 integrin appeared important for osteoclast functioning based on the finding that inhibition of signaling through this avb3 integrin inhibited osteoclast-mediated bone resorption in vitro and in animal models of osteoporosis and malignant osteolysis [197]. Among the many other molecules that are important for the functional activity of osteoclasts, such as c-Src, cathepsin K, carbonic anhydrase II, TRAP, and several ion channel proteins, many have been found to cause an osteopetrotic phenotype when deleted in mice or altered in humans. The absence of these genes does not affect the differentiation into morphologically normal osteoclasts; however, the osteoclasts are not functional and fail to resorb bone effectively (reviewed in [78,195,196]. For instance, cathepsin K, the key enzyme in the digestion of bone matrix by its activity in degrading type I collagen, is highly expressed by activated osteoclasts and secreted in the resorption lacuna [198,199]. Its deletion in mice led to osteopetrosis, and mutations in the human cathepsin K gene cause pycnodysostosis. Highly selective and potent cathepsin K inhibitors have been shown useful as antiresorptive agents to treat osteoporosis, as well as being promising therapeutic tools to reduce breast cancerinduced osteolysis and skeletal tumor burden [200,201]. Besides cathepsin K, several proteolytic enzyme groups are involved in the degradation of organic components (collagens and proteoglycans) of bone and cartilage matrices after the mineral is dissolved. One of these is the MMP family, which constitutes over 25 members, including secreted collagenases, stromelysins, gelatinases and membrane-type (MT)-MMPs. Several MMPs including MMP9 and MMP14 (also known as MT1-MMP) are highly expressed in osteoclasts/chondroclasts. MMPs are synthesized as latent pro-enzymes that, upon proteolytic activation, can degrade numerous extracellular matrix components. As such, they are involved in development, growth and repair of tissues, but also in pathological conditions associated with excessive matrix degradation such as rheumatoid arthritis, osteoarthritis and tumor metastasis [202e204]. The Coupling Principle in Bone Remodeling Bone remodeling must be tightly controlled to maintain the normal bone homeostasis, but the molecular

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mechanisms that control its initiation, progression, and cessation at any given site remain poorly understood. However, failure of this normal skeletal functioning is one of the most common early manifestations of aging, for example in osteoporosis and osteoarthritis. These and a variety of other pathologic conditions affecting the skeleton (e.g. rheumatoid arthritis, periodontitis, Paget’s disease, and bone tumors) lead to perturbations in the bone remodeling process, predominantly shifting the balance towards more degradation of bone due to local and/or systemic alterations in the levels of hormones or proinflammatory cytokines that stimulate bone resorption [170,205,206]. In normal conditions of development, growth, and bone health, the osteoclast is thought to die via apoptosis after a limited period of resorptive activity [207] and the resorbed area of cartilage or bone is efficiently replaced by newly formed bone through the action of osteoblasts. Hence, skeletal homeostasis remains intact as long as the activities of both osteoclasts and osteoblasts are balanced (“coupled”) and the net bone mass is maintained. This balance implies the existence of mechanisms tightly coordinating the differentiation of osteoblasts and osteoclasts as well as their migration to locations where they function. One prime aspect of the coupling principle is provided by the direct control of osteoclastogenesis by cells of the osteoblast lineage through their expression of M-CSF and RANKL. Conversely, osteoclasts may reciprocally stimulate osteoblast differentiation and function to initiate the anabolic arm of the remodeling process. Osteoclastic bone resorption may well locally release the myriad of growth factors, like for example TGFb, that are stored in the bone matrix, which can subsequently act to attract and stimulate osteoblasts [208]. Direct signaling by osteoclasts to cells of the osteoblast lineage has recently been demonstrated and may participate in coupled osteoblastogenesis and bone formation. For instance, the ephrin/Eph receptor system allows bidirectional signaling between osteoclasts and osteoblasts; osteoclasts express ephrin B2 and osteoblasts express its EphB4 receptor, both membrane-bound proteins [209]. Signaling through EphB4 into osteoblasts (“forward signaling”) enhances osteogenic differentiation in vitro, whereas signaling through ephrin B2 into osteoclast precursors (“reverse signaling”) suppresses osteoclast differentiation [209]. The overall outcome of such interaction thus is predicted to favor bone formation [210]. It has also been reported that the v-ATPase V0 subunit D2 is not only involved in osteoclast fusion but also regulates the secretion by osteoclasts of still unidentified factors that inhibit the differentiation of osteoblast precursors into mature cells [211]. Other cytokines involved in the coupling, such as interleukin-1 (IL-1), IL-2, IL-6, and oncostatin M, transduce their signals

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through the gp130 protein and play an important role in osteoclast and osteoblast physiology [212].

Osteocytes and Mechanosensing Mechanical loading of the skeleton is essential for the development, growth, and maintenance of strong, weight-bearing bones. Altered growth and subsequent deformity resulting from abnormal mechanical loading is often referred to as mechanical modulation of bone growth. This phenomenon has key implications in the progression of infant and juvenile musculoskeletal deformities, such as adolescent idiopathic scoliosis, hyperkyphosis, genu valgum and tibia bowing. Clinical management of these deformities is often directed at modifying the mechanical environment of affected bones. However, there is limited understanding of how bone growth is regulated in response to mechanical loading with respect to growth plate chondrocytes [213]. Better characterized is the concept that mechanical loading improves bone strength particularly by inducing osteoblastic bone formation in regions of high strain energy. In order to withstand loading in the most efficient way (maximal strength for minimal material), the skeleton constantly adjusts its bone mass and architecture in response to load through bone remodeling. It has indeed long been recognized that mechanical stress induced by weight-bearing exercise increases osteoblast activity and induces bone formation. Bone tissue thus has a mechanosensing apparatus that directs osteogenesis to where it is most needed to increase bone strength. The most likely sensors of mechanical loading are the osteocytes, which sense variations in mechanical forces acting on bone and respond to this by signaling via sclerostin secretion and other mechanisms (see below), to coordinate osteogenesis [214,215]. Osteocytes represent the quantitatively dominant cell type in adult bone. Osteocytes, derived from osteoblasts, are buried within the mineralized bone matrix. They are connected to each other and to cells on the bone surface via numerous dendritic processes that run inside lacunar cannaliculi. These processes are thought to allow osteocytes to communicate with other cells including other osteocytes, and osteoblasts and osteoclasts on the bone surface, thereby playing roles in the regulation of bone formation and resorption [92,93,216]. They also allow the osteocytes to sense and respond to mechanical forces, and to secrete proteins involved in systemic phosphate homeostasis (see Chapter 7) including dentin matrix protein (DMP)-1, matrix extracellular phosphoglycoprotein (MEPE) and FGF-23 [92,217]. The importance of viable osteocytes in the maintenance of bone tissue health was shown by experimental destruction of osteocytes in murine bone: activation of the diphtheria toxin receptor, expressed

under the control of the osteocyte-specific DMP-1 promoter, quickly led to targeted ablation of osteocytes through necrosis, and was associated with a large-scale increase in bone resorption, decreased bone formation, and trabecular bone loss. Concomitantly, these mice were resistant to unloading-induced bone loss, indicating the requirement for osteocytes in the response to mechanical signals [218]. Osteocytes have been demonstrated to stimulate bone resorption in vitro and in vivo [219,220]. This modulation of bone remodeling may be elicited by osteocyte apoptosis, which can be consequential to unloading [221]. Conversely, mechanical stimulation is capable of maintaining osteocyte viability [222]. The recent deletion of b-catenin in osteocytes, using a DMP-1-Cre conditional knockout approach, indicated that b-catenin is necessary for normal osteocyte viability and function, and for the maintenance of normal bone. Indeed, these conditional knockout mice had bones with pronounced porosity and fragility due to increased osteoclast number and activity, most likely owing to reduced expression of OPG and an increase in RANKL, both found to be expressed in osteocytes [106]. The mechanisms by which a mechanical stimulus is transduced into biochemical signals in osteocytes and osteoblasts and the means whereby these cells then modulate bone remodeling have not been clearly identified. Influences that have been implicated are shearing forces produced by fluid movement (e.g. in the canaliculi surrounding the osteocytic dendrites) and a variety of membrane proteins, including integrins, connexins and stretch-sensitive ion channels [92,93]. It also has been proposed that the osteocyte senses load through cilia, single flagellar-like structures found on every cell [223]. When Pkd1, the gene encoding the large transmembrane protein polycystin-1 (PC-1) located at the primary cilia, was conditionally deleted in osteocytes using DMP-1-Cre mice, the bone anabolic response to loading was recently found to be dramatically decreased [224]. Osteocytes respond in vitro and in vivo to increased load by altering their signaling. For example, in response to loading, osteocytes upregulate nitric oxide production, release prostaglandin PGE-2 and IGF-1, and decrease glutamate transporter expression [92,93]. DMP-1 expression is also robustly increased upon mechanical stimulation [225]. DMP-1 inactivation in mice is associated with a hypomineralized phenotype linked with elevated FGF-23 expression and defective osteocyte lacuna/canalicular network formation [226]. Of great interest is the recent observation that mechanical loading suppresses the expression of SOST in osteocytes, resulting in a rapid decrease of sclerostin production [215]. As mentioned (see above), the osteocyte-specific protein sclerostin inhibits Wnt signaling

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ENDOCRINE CONTROL OF POSTNATAL BONE GROWTH

through binding to LRP5/6. Consequently, SOST null mice have a very high bone mass [227] whereas, conversely, transgenic mice overexpressing sclerostin in osteocytes suffer severe bone loss [228]. Osteocytes thus appear to use the Wnt/beta-catenin pathway to transmit signals of mechanical loading to cells on the bone surface [92,106].

ENDOCRINE CONTROL OF POSTNATAL BONE GROWTH Several systemic factors are known to profoundly influence postnatal bone growth as well as bone remodeling. These include the principal regulators of mineral homeostasis, such as PTH and 1,25(OH)2 vitamin D that maintain calcium homeostasis by regulating osteoclastic resorption, the actions of which are discussed in detail in other chapters of this book. Here, we will briefly summarize the influences on bone growth and remodeling by the growth hormone /insulin-like growth factor-1 axis, sex steroids, thyroid hormones, and glucocorticoids.

Growth Hormone and Insulin-like Growth Factor-1 Growth hormone (GH) and insulin-like growth factor-1 (IGF-1) are central to the achievement of normal longitudinal bone growth and the acquisition of bone mass during the prepubertal period, and remain important regulators of bone homeostasis throughout life. Although GH may act directly on skeletal cells, most of its effects are mediated by IGF-1, which is present in the systemic circulation and is synthesized by peripheral tissues. Both systemic and local skeletal IGF-1 play a role in bone formation and the maintenance of bone mass [229,230]. GH is a single-chain peptide of 191 amino acids, synthesized and released from the anterior pituitary gland. Its receptor is highly expressed in the liver, adipose tissue, heart, kidneys, intestine, lung, pancreas, cartilage and skeletal muscle where it induces the synthesis of IGF-1, a member of the insulin family of growth factors [229,230]. Systemic IGF-1 is synthesized primarily in the liver and circulates bound by IGFbinding proteins (IGFBPs) that regulate its availability. The triple inactivation of IGFBP-3, -4 and -5 demonstrated that IGFBPs are necessary to maintain appropriate levels of systemic IGF-1 and adequate postnatal growth [231]. IGF-2 shares biochemical and biological properties with IGF-1; it is synthesized by skeletal cells, independent of GH, and is important in skeletal development, but its function in the adult skeleton is not proven [229,230].

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During embryonic development, IGF-1 and IGF-2 are key determinants of growth. IGF-1 deficiency retards both pre- and postnatal growth, and IGF-1 receptor (IGF-1R) null mice, exhibiting a more severe growth deficiency, die shortly after birth [232e235]. In this context, prenatally, IGF-1 signaling is considered to be GH independent, whereas postnatally, IGF-1 is partly or fully GH dependent with phosphorylation of STAT5b having an intermediary role [236]. Knockout of the IGF-2 gene leads to decreased fetal growth in analogous fashion to knockout of the IGF-1 gene [237]. Because of the additional action of IGF-2 on the insulin receptor, knockout of IGF-2 and IGF-1R results in a more severe growth abnormality than knockout of IGF-1R alone [238]. Since IGF-2 acts predominantly in fetal life and GH acts on growth postnatally, there are no expected interactions between IGF-2 and GH in mice. Postnatally and throughout puberty, GH and IGF-1 play a critical role in determining longitudinal skeletal growth. Mice missing both the GH receptor and IGF-1 are only 17% of normal size [235]. These dramatic quantitative effects show that GH and IGF-1 importantly control bone growth in mammals. The crucial role of GH in controlling bone growth dates from the isolation of GH and the evidence that GH administration corrects the growth defects in patients with genetic or acquired GH deficiency (GHD) who display short stature. Patients and mice with mutations in the GH receptor gene (Laron dwarfism) have a similar defect in bone growth [235,239,240]. Studies of mice missing the GH receptor show that the mice, like people with Laron dwarfism, are of normal size at birth but have defective postnatal growth. In mice, the defect is detected days 10e40 after birth [235]. These mice have growth plates with short proliferative columns with fewer chondrocytes than normal, a slower rate of proliferation, and smaller hypertrophic chondrocytes than normal [235]. Thus, these mice display defects both in the proliferation and in the differentiation of chondrocytes. Clear demonstration of the physiologic importance of IGF-1 in bone growth awaited gene ablation studies [233,234,241]; subsequently, one human with IGF-1 gene mutation and a phenotype analogous to that of the knockout mice was reported [232]. Unlike the fairly normal prenatal phenotype of mice missing the GH receptor, mice missing the IGF-1 gene are only 60% of normal weight at birth and have a high perinatal mortality, which varies depending on the genetic background of the particular mouse strain. These mice, like the GH receptor knockout mice, have small hypertrophic chondrocytes [235,242]. Wang et al. [242] found that proliferative columns are of normal size and that the chondrocytes proliferate at a normal rate, whereas Lupu et al. [235] found that the proliferative columns

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are short, with a low rate of proliferation. Although the studies differ in several respects, probably the most important difference that might explain the varying results is that the mice have distinct genetic backgrounds. As noted earlier, the observation that GH stimulates the production of IGF-1 in many cell types and the partial overlap in actions of these two factors has led to the hypothesis that many of the growth-promoting actions of GH on chondrocytes are mediated by IGF-1. The small size of the IGF-1 knockout mice, despite their high GH levels, is consistent with this idea. Nevertheless, the finding that mice missing both the GH receptor and IGF-1 genes have a greater growth defect than either individual knockout mouse is strong genetic evidence that the actions of GH and IGF-1 on growth are predominantly independent and additive [235]. Moreover, the traditional concept that GH primarily acts through stimulation of the IGF-1 production in the liver and increased circulating IGF-1 levels, was contested by mouse genetic studies. Indeed, when the IGF-1 gene was ablated specifically from liver in vivo using the Cre-Lox approach, circulating levels of IGF-1 decreased to approximately 20% of baseline, but the animals grew normally [243,244]. Although these studies demonstrate that a large fraction of circulating IGF-1 derives from the liver, they also suggest that IGF-1 made by the liver may not be vitally important for bone growth. The actions of IGF-1, whether produced locally or systemically, are via the IGF-1 receptor (IGF-1R) expressed on the cell surface of the chondrocytes of the growth plate. The type-2 IGF-1R is expressed equally throughout all maturational zones of the growth plate, whereas the type-1 receptor is more highly expressed by proliferating chondrocytes [245]. These data are consistent with the concept that IGF-1 has regulatory actions on all chondrocytes of the growth plate. Moreover, recently generated cartilage-specific IGF-1R knockout mice, constitutive as well as tamoxifeninducible models, indicate that the IGF-1R in chondrocytes controls cell growth, survival, and differentiation in embryonic and postnatal growth plates in part by suppression of PTHrP expression [246]. Thus, although the functional relationships between GH and IGF-1 actions on the growth plate remain unsettled, it is clear that the IGF-1 signaling pathway has a central function in modulating endochondral bone growth and regulates a number of key chondrocyte physiological processes such as chondrocyte proliferation, matrix synthesis, differentiation, hypertrophy and survival [229,230,235,246]. In addition to the effects on longitudinal growth, GH and IGF-1 are anabolic hormones and have the potential to regulate bone modeling and remodeling. During adolescence, the anabolic effects of GH and IGF-1 in

bone are important for the acquisition of bone mass. Late adolescence and early adulthood are critical periods for the acquisition of bone mass, and the achievement of peak bone mass. This is a critical determinant of future risk of osteoporosis. Adult GHD causes low bone turnover osteoporosis with high risk of vertebral and non-vertebral fractures, and the low bone mass can be partially reversed by GH replacement [230]. IGF1 mediates most of the effects of GH on skeletal metabolism. IGF-1 may reduce osteoblast apoptosis and promote osteoblastogenesis by stabilizing b-catenin, enhancing Wnt-dependent activity. This effect, associated with modest mitogenic properties, causes an increase in the number of osteoblasts, and an increase in osteoblastic function and bone formation [247,248]. Diseases affecting the GH/IGF-1 axis are frequently associated with significant alterations in bone metabolism that often lead to bone loss [229,230].

Sex Steroids: Estrogen and Androgen The effects of sex hormones on growth appear to be very species specific, so lessons from rodents cannot be easily applied to humans. In humans, in association with the increase in sex hormones in boys and girls at the time of puberty, linear bone growth accelerates, followed by the disappearance of the growth plate and permanent lack of further growth. Remarkably, several patients with defective estrogen receptor a (ERa) [249] or defective aromatase (the enzyme that converts testosterone to estradiol) [250,251] have been identified. These patients continued to grow into adulthood owing to a lack of epiphyseal fusion in the long bones, which resulted in increased adult height. Two girls without aromatase presented with signs of androgen excess but lack of a clear pubertal growth spurt until they were treated with estrogen [252]. While estrogen therapy resulted in rapid growth plate closure in patients with aromatase deficiency, it did not in the man with a mutation in the ERa gene [249e251]. In rodents, the growth plates do not fuse directly after sexual maturation, but high-dose estradiol treatment results in a clear reduction of the growth plate height [253]. A mouse model with cartilage-specific inactivation of ERa, the main ER regulating skeletal growth, was recently generated. During sexual maturation, the skeletal growth of these mice was normal, but they continued to grow after 4 months of age, resulting in increased bone length at the age of 1 year. High-dose estradiol treatment of adult mice reduced the growth plate height as a consequence of attenuated proliferation of growth plate chondrocytes in control mice but not in cartilage-specific ERa knockout mice [254]. Sex steroids also have a pronounced influence on the process of bone modeling, remodeling and homeostasis,

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essential for maintenance of the adult skeleton. The precise time of the attainment of peak bone mass is not certain, and it is skeletal-site dependent. The increase in gonadal steroid synthesis at the time of puberty is an important hormonal regulator of bone accretion. Boys with constitutionally delayed puberty achieve lower peak bone mass than normal boys [255]. Androgens are important not only as a source of estrogen, through the action of aromatase, but also for their direct effect in stimulating bone formation [256]. Testosterone, responsible for the male phenotype characterized by a larger skeleton, has complex effects on bone metabolism. As an anabolic steroid, it stimulates bone formation in both male and females. In addition, testosterone can inhibit bone resorption directly, acting through the androgen receptor as well as through conversion by aromatase to estrogens (see below). Androgens also increase periosteal bone formation, leading to larger and therefore stronger bones. Loss of androgens in males from chemical or surgical castration or an ageassociated decline of androgen levels has the same adverse effect on the skeleton as estrogen loss in women, albeit the loss of testosterone in aging men is not as universal or as abrupt as the loss of estrogen at the time of menopause in women [256,257]. Indeed, particularly postmenopausal women suffer from osteoporosis, characterized by low bone mass and high risk of debilitating fractures of the vertebrae and long bones. This disease afflicts millions of people, and becomes increasingly prevalent with the aging of the general population. Skeletal preservation by estrogen in females may be evolutionarily related to the need of calcium stores for embryonic skeletal development. In mammalian adult males and females, including humans, estrogen has been identified as the major inhibitor of bone resorption by reducing osteoclast number [257,258]. Estrogen deficiency increases both the number of sites at which remodeling is initiated, and the extent of resorption at a given site. Increased bone resorption is accompanied by increased bone formation as a result of coupling. However, the increase in bone formation is not sufficient to maintain bone balance and prevent bone loss. The reason for this “skeletal insufficiency” in osteoporosis is not known; it could be due to lack of estrogen or other hormones, such as androgens, required for fully effective bone formation, and is partly due to the above-mentioned kinetic imbalance. In cancellous bone, the loss results in thinner trabeculae, which become rod-shaped rather than plates, and in trabecular discontinuity, which deprives them of mechanical function. In cortical bone, endosteal bone resorption causes thinning of the cortex and sometimes “trabecularization” of the endosteal surface. Enhanced resorption increases the size of haversian canals and the porosity of the cortical bone. The result of these changes is significant

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weakening of the respective bones and increased fracture risk. Treatment of women with hormone replacement therapy (either estrogen alone or estrogen plus progesterone) has been shown to prevent this bone loss [259]. Recent studies have shed light on the mechanism of estrogen action on osteoclasts: osteoclasts express estrogen receptors (ERa and ERb) and estrogen acts directly on osteoclasts to increase apoptosis [260,261]. The regulation of osteoclast life span by estrogen involves induction of the Fas/FasL system causing apoptosis, a pathway that may be involved both in osteoclast cell-autonomous effects [261], as well as in indirect mechanisms mediated via the osteoblast [262]. Furthermore, estrogen can have non-genomic effects, or rapid signaling effects, inducing the phosphorylation of components of various signaling pathways (e.g. the MAPK pathway) or calcium regulation. Such effects may contribute to the induction of osteoclast apoptosis by estrogen without the need for direct binding of ERa to DNA [262,263]. The DNA-binding function of ERa was recently shown to be dispensable also for the effects of estrogen on osteoblastogenesis and on decreasing the prevalence of mature osteoblast apoptosis [264]. In fact, Manolagas and co-workers propose that the protective effects of estrogens on bone result from their ability to attenuate oxidative stress in bone and bone marrow; estrogens diminish the generation of reactive oxygen species (ROS), stimulate the activity of glutathione reductase, and decrease the phosphorylation of p66shc, an adapter protein that serves as a key component of a signaling cascade that is activated by ROS and influences apoptosis and lifespan in invertebrates and mammals. Hence, loss of estrogens may accelerate the effects of aging on bone by decreasing the defense against oxidative stress [264,265]. A variety of other mechanisms contribute to the beneficial effects of estrogen on bone. Estrogen was reported to decrease the responsiveness of osteoclast progenitor cells to RANKL, thereby reducing osteoclastogenesis [266]. Estrogen also regulates a variety of cytokines, indirectly leading to changes in osteoclast number: estrogen suppresses the production of osteoclastogenic cytokines such as IL-1, IL-6, IL-7 and TNF-a in T cells and osteoblasts [267,268]. As well, part of the effects ascribed to estrogen deficiency were suggested to be in fact mediated by the resultant increase in pituitary gland derived follicle stimulating hormone (FSH), acting on osteoclasts [269].

Thyroid Hormones The hypothalamicepituitaryethyroid axis plays a key role in skeletal development, acquisition of peak bone mass and regulation of adult bone turnover. Euthyroid status is essential for maintenance of optimal bone mineralization and strength. In population studies,

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hypothyroidism and hyperthyroidism have both been associated with an increased risk of fracture. In children, thyrotoxicosis advances bone age and accelerates postnatal skeletal development resulting in early cessation of growth and short stature due to premature fusion of the growth plates. Conversely, children and rodents with hypothyroidism have decreased rates of bone lengthening [270]. Although some of this decrease in growth plate function may be caused by associated decreases in GH levels, GH cannot fully correct abnormalities seen in rats and mice with hypothyroidism. These abnormalities include both shortened proliferative layers of chondrocytes and decreased numbers of hypertrophic chondrocytes [271]. At least some of the effects of thyroid hormone are likely to be direct effects on chondrocytes that express thyroid hormone receptors [272] because both growth plates and isolated chondrocytes respond to T3 in vitro by increasing the conversion of proliferating to hypertrophic chondrocytes [273,274]. Genetically engineered mice missing all transcripts from both the thyroid hormone receptor a and b loci have a delay in development of secondary ossification centers, a decrease in the proliferative layer of chondrocytes, and a particularly dramatic decrease in hypertrophic chondrocytes [275]. This phenotype is not as severe as that in mice with congenital absence of thyroid glands, or as in mice missing the pax8 gene encoding an essential transcription factor required for thyroid follicular cell development [276]. These comparisons, as well as comparisons with mice missing selected transcripts from the complicated thyroid hormone receptor loci, suggest that non-ligand-binding variants of these receptors function in the absence of thyroid hormone to generate a growth plate phenotype more severe than that in mice missing all transcripts from these loci [270,275,277,278]. Both receptor loci contribute to growth plate function. Some children with thyroid hormone resistance due to dominant negative mutations in the thyroid hormone receptor b gene have short stature, presumably partly through blockade of receptor action in the growth plate [279].

Glucocorticoids In contrast to sex steroids, glucocorticoid excess is catabolic to bone, as illustrated by glucocorticoidinduced osteoporosis, a devastating consequence of long-term use of glucocorticoids [280]. The mechanisms for glucocorticoid-induced bone loss are complex, including suppression of renal calcium reabsorption, reduction in intestinal calcium absorption, and hypogonadism, all of which lead to increased bone resorption and bone turnover. Perhaps more importantly, glucocorticoids also have direct effects on the skeleton [280]. Glucocorticoids impair the replication, differentiation

and function of osteoblasts and induce the apoptosis of mature osteoblasts and osteocytes, leading to a dramatic suppression of bone formation [281,282]. As well, glucocorticoids act directly on osteoclasts and prolong their life span; in addition, glucocorticoids may sensitize bone cells to regulators of bone remodeling and favor osteoclastogenesis, thus overall leading to increased bone resorption [283,284]. Glucocorticoids are widely used as anti-inflammatory and immunosuppressive drugs in children. Long-term, high-dose glucocorticoid treatment often leads to growth failure. Similarly, systemic administration of glucocorticoid in mice, rats, and rabbits decreases the rate of longitudinal bone growth, likely by inhibiting growth plate chondrocyte proliferation [285,286] and by stimulating chondrocyte apoptosis [287]. Glucocorticoid inhibits longitudinal bone growth, in part, through a direct effect on growth plate chondrocytes that exress glucocorticoid receptor [286]. In addition to its direct action on the growth plate, glucocorticoid may also suppress longitudinal bone growth through an indirect action, mediated, in part, by changes in the GH/IGF-1 axis [285,288]. Discontinuation of glucocorticoid treatment is followed by catch-up growth. Catch-up growth may occur because the decreased cell proliferation during glucocorticoid treatment conserves the proliferative capacity of the chondrocytes, thus slowing growth plate senescence. Following discontinuation of the glucocorticoid treatment, the growth plates are less senescent than normal, and hence show a greater growth rate and grow for a longer time than expected for age, resulting in catch-up growth [285]. Despite catch-up growth, prolonged glucocorticoid administration in children can result in some residual permanent growth deficit.

CONCLUDING REMARKS As outlined above, postnatal bone growth involves a complex spatiotemporal coordination of the proliferation, differentiation, and activity of multiple cell types building the multicomponent skeletal tissue. A myriad of transcription factors, local signaling molecules, endocrine, mechanical and central signals have been implicated in the regulation of bone growth and remodeling. Precise control is mandatory to build and maintain a skeleton that is fully able to fulfill its major functions, including mechanical support and mineral homeostasis. Moreover, a number of recent breakthroughs have shown that bone cells are not merely involved in the acquisition and maintenance of the bone mass, but also execute a number of critical functions that extend beyond the bone tissue proper. Among these are roles of osteoblasts and osteoclasts in the

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interplay between skeletal functioning and the regulation of immunology and inflammation [289,290], hematopoiesis and stem cell biology [291e294], the central and sympathetic nervous system [295e297] and energy metabolism [296,298e300]. Further advances in the molecular and genetic understanding of skeletal biology, including these novel integrative communication pathways with other tissues and organ systems, may offer new insights as well as potential therapeutic options to treat various pediatric bone diseases presented further in this book.

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[287] Chrysis D, Ritzen EM, Savendahl L. Growth retardation induced by dexamethasone is associated with increased apoptosis of the growth plate chondrocytes. J Endocrinol 2003;176:331e7. [288] Nilsson O, Marino R, De Luca F, Phillip M, Baron J. Endocrine regulation of the growth plate. Horm Res 2005;64:157e65. [289] Lorenzo J, Horowitz M, Choi Y. Osteoimmunology: interactions of the bone and immune system. Endocr Rev 2008;29:403e40. [290] Takayanagi H. Osteoimmunology: shared mechanisms and crosstalk between the immune and bone systems. Nat Rev Immunol 2007;7:292e304. [291] Calvi LM, Adams GB, Weibrecht KW, et al. Osteoblastic cells regulate the haematopoietic stem cell niche. Nature 2003;425: 841e6. [292] Lo Celso C, Fleming HE, Wu JW, et al. Live-animal tracking of individual haematopoietic stem/progenitor cells in their niche. Nature 2009;457:92e6. [293] Lymperi S, Ferraro F, Scadden DT. The HSC niche concept has turned 31. Has our knowledge matured? Ann NY Acad Sci 2010;1192:12e8.

[294] Wu JY, Scadden DT, Kronenberg HM. Role of the osteoblast lineage in the bone marrow hematopoietic niches. J Bone Miner Res 2009;24:759e64. [295] Franquinho F, Liz MA, Nunes AF, Neto E, Lamghari M, Sousa MM. Neuropeptide Y and osteoblast differentiation e the balance between the neuro-osteogenic network and local control. FEBS J 2010;277:3664e74. [296] Karsenty G, Oury F. The central regulation of bone mass, the first link between bone remodeling and energy metabolism. J Clin Endocrinol Metab 2010;95:4795e801. [297] Takeda S, Karsenty G. Molecular bases of the sympathetic regulation of bone mass. Bone 2008;42:837e40. [298] Ferron M, Wei J, Yoshizawa T, et al. Insulin signaling in osteoblasts integrates bone remodeling and energy metabolism. Cell 2010;142:296e308. [299] Clemens TL, Karsenty G. The osteoblast: An insulin target cell controlling glucose homeostasis. J Bone Miner Res 2011;26: 677e80. [300] Lee NK, Karsenty G. Reciprocal regulation of bone and energy metabolism. Trends Endocrinol Metab 2008;19:161e6.

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Dental Development and Maturation, from the Dental Crypt to the Final Occlusion Jean-Marc Retrouvey 1, Michel Goldberg 2, Ste´phane Schwartz 3 1

Division of Orthodontics, McGill University, Montreal, Quebec, Canada, 2 UMR-S 747-INSERM, Universite´ Paris Descartes, 3 Dental Clinic, Montreal Children’s Hospital, MUHC (McGill University Health Center), and McGill University

The formation and maturation of dental tissues constitute an important process in craniofacial development. Genes coding the expression of growth factors, transcription factors and extracellular matrix molecules regulate this process. The interactions between the mesoderm and the ectoderm are crucial during the initial steps of tooth germ development, morphogenesis and cell differentiation, leading to crown and later to root formation. These processes are associated with the formation of alveolar bone around the dental follicle and tooth eruption in the oral cavity. Disruption of the normal developmental pattern may be due to gene mutations or to defective expression of intracellular or extracellular structural molecules. The development of the dentition is also a great contributor to the development of the lower face height by enlargement of the dento-alveolar processes. As gene mutations, epigenetic alterations or influenced by post-genomic factors, individual defects can be identified. In such cases, the mutations listed in this review are invariably associated with oral development manifestations. As a more complex phenomenon, the close association of multiple defects interacting closely may produce multifaceted syndromes.

the expression of growth factors and transcription pathways regulate these events. Any mutation in these genes is susceptible to induce morphologic and/or functional alterations in any tooth developmental process. We review here the major events that influence early tooth formation that may shed light on major tooth abnormality such as amelogenesis imperfecta and dentinogenesis imperfecta, which may or may not be associated with major craniofacial alterations or defective mineralization syndromes [2,3]. Within the first branchial arch for all the teeth, and the nasofrontal bud for the maxillary incisors, all tooth buds result from the interactions of the oral ectoderm lining the stomodeal cavity, in specific areas spatially limited as oral placodes, and the dental mesenchyme containing cells derived from the neural crest. Initial Phase In the epithelial cells expressing Otlx2/Pitx2, two families of growth factors (bone morphogenetic proteins [BMPs], fibroblast growth factors [FGFs]) are activated. The epithelial placode then becomes thicker and up to day 11 of gestation is under the influence of Left in the mouse. This leads to the formation of a dental lamina. BMP, FGF, sonic hedgehog (Shh) and Wnt are expressed in the epithelial compartment, whereas the expression of Lhx6-7, Barx1, Msx 1-2, and Pax9 is increased in the odontogenic mesenchyme. The proliferating neural crest-derived cells condense in the dental mesenchyme and express Msx1, Pax9, Left and Cbfa1 (Runx2). This implies that any alteration of the genes coding for these molecules leads to severe alteration or even the lack of tooth formation (anodontia).

THE TOOTH BUD Embryology of the Tooth Three successive steps lead to the formation of a tooth: (1) the initiation of the process, (2) the morphogenesis of the tooth, and (3) the terminal differentiation of two layers of epithelial and mesenchymal cells acquiring their terminal phenotype [1]. Several genes activating

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5. DENTAL DEVELOPMENT AND MATURATION, FROM THE DENTAL CRYPT TO THE FINAL OCCLUSION

Morphogenesis of the Tooth Depending on the position of the germ in the branchial arch, a different type of tooth will be produced. This is related either to a possible gradient of “morphogens”, or to the concomitant burst of cell clones predetermined to form incisors, canines, premolars and molars. The epithelial buds located along the dental lamina proliferate, become caps and later reach the bell stage. During early morphogenesis, three series of events are implicated in the shaping of the tooth crown. Initially, a semi-permeable basement membrane (BM) [4] regulates the diffusion of molecules between the enamel organ and the embryonic pulp compartments. The BM triggers the terminal differentiation of preodontoblasts into polarized secretory odontoblasts. The preameloblasts become differentiated secretory ameloblasts. Metalloproteases and other degradative enzymes trapped in the BM network are shaping the future dentino-enamel junction [5]. Following this initial step, neural crest-derived cells migrate and colonize the dental mesenchyme. Later, they slide toward the periphery of the embryonic pulp and undergo a series of cell divisions. One less division occurs in the inner enamel epithelium compared with the odontoblast layer. This induces a substantial length difference between the two compartments, hence to the folding of the cell layers and consequently to cusp formation. Finally, at an early bell stage, a niche of cells named the “enamel knot” is found in the central part of the tooth. They express the gene Msx2. This group of cells is not proliferative, in contrast with the cells located in the lateral parts of the germ. The central part stays in a fixed position whereas the rest of the enamel organ expands and, because of space limitations, contributes also to the formation of folds and consequently, cusps. The enamel knot expresses Shh, Bmp-2, Bmp-4, Bmp-7, Fgf-4 and Fgf-9 [6]. Terminal Differentiation Post-mitotic polarizing ameloblasts elongate. Intercellular junctions start to develop and the polarized young ameloblasts become functional and implicated in the synthesis and secretion of enamel proteins. Pre-odontoblasts originally bear a fibroblastic appearance. They migrate from the central part of the dental pulp to the periphery and become post-mitotic polarizing odontoblasts. Long protracted processes, characteristic of polarized odontoblasts, emerge from cell bodies associated into a palisade-like structure. They are implicated in the synthesis of collagen and non-collagenous proteins, whereas secretion and re-internalization of fragmented peptides or signal peptides occur along the cell processes, in the predentin and in dentin respectively. In the distal

part of the cell bodies, desmosomes and gap junctions form terminal junctional complexes. Root Formation/Eruption During the pre-eruptive phase, once the crown is formed, at the cervical margin of the enamel organ, epithelial cells proliferate and initiate the formation of the Hertwig’s epithelial root sheath. In the inner part of the sheath, epithelial cells promote the recruitment and cytodifferentiation of pulp progenitors. The outer layer is implicated in the recruitment of cementoblasts that cross the disaggregating intercellular Hertwig’s root sheath. The Hertwig’s root sheath is involved in the recruitment of cells of the periodontal ligament and the construction of the bony crypt [7]. The pulp progenitors migrate toward the inner surface of the Hertwig’s sheath. They become odontoblasts, organized as a palisade-like structure, forming the root dentin. The forming part of the tooth germ keeps its original position during these early stages of root formation. Gradually, the root(s) elongate(s) and the tooth erupts in the oral cavity. This is referred to as the eruptive prefunctional stage. Three possible origins have been reported for the cementoblasts: • during the early stages of formation of the root, epithelial cells of the Hertwig’s root sheath interconvert, become mesenchymal cells and ultimately acquire the cementoblast phenotype • afterwards, during root lengthening, the dissociation in the cervical area of the Hertwig’s root sheath allows some fibroblast-like cells issued from the dental follicle to migrate through these fenestrations and become cementoblasts • later, the tooth comes into contact with its antagonistic teeth, and the late steps of eruption/root formation are referred to as the functional stage. Then progenitors located in the periodontal ligament proliferate under the influence of cementum growth factor, and differentiate into cementoblasts. After stimulation of the cementum adhesion protein, cementoblast progenitors adhere to the cementum surface and contribute to the thickening of the forming cementum. Although the mechanisms implicated in the eruption are not fully elucidated, it is clear that a series of growth and transcription factors intervene in the formation of the bony socket. Colony-stimulating factor-1 (CSF-1), epithelial growth factor (EGF), transforming growth factor (TGF)1, interleukin 1a (Il-1a), receptor activator ligand of the nuclear factor-kappa B (RANKL), c-Fos, osteoprotegerin, parathyroid hormone-related protein, Cbfa1, tumor necrosis factor (TNF), vascular endothelial

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growth factor (VEGF) and bone morphogenetic protein2 (BMP-2) have all been implicated. These factors, together with a series of metalloproteases and disintegrins, contribute to the root lengthening, eruption and functional adaptation [7].

Histology of Dental Tissues Enamel, dentin and cementum constitute the three mineralized dental tissues, whereas the dental pulp is a unique non-mineralized tissue. Enamel The composition of enamel is shown in Table 5.1. ENAMEL STRUCTURE

The interacting presecretory ameloblast and odontoblast layers form distinct compartments on either side of the BM. The BM is then enzymatically degraded, and short ameloblast processes protrude toward the forming dentin, leading to the formation of a scalloped dentino-enamel junction (DEJ). Ameloblasts initiate the formation of the prismatic enamel made by rods (or prisms) and interrods (interprismatic enamel). Tightly packed hexagonal ˚ ngstro¨ms (A ˚ ) in enamel crystallites are about 700 300 A ˚ in cross-section, and vary between 2000 and 10 000 A height. Prisms (rods and interrods), Hunter-Schreger bands and Retzius lines constitute the three characteristic anatomic structures found in enamel (Fig. 5.1). Near the DEJ, in humans, between 10 and 20 prisms are seen bending alternatively to the right or to the left contributing to the formation of Hunter-Schreger’s bands. In transverse sections, Retzius lines appear as continuous lines forming concentric circles, and each enamel segment comprised between two Retzius lines, about 25 mm thick, constitutes a mineralization modulus [8]. This organization allows occlusal forces to be dissipated away from the pressure area. This type of architecture reinforces the mechanical properties of enamel.

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diffuse in the outer layer forming predentin/dentin. The first enamel layer is aprismatic. Accumulation of degraded matrix components contributes to the formation of Tomes’ process. The formation of interrod enamel then starts around the processes, developing a continuous honeycomb structure. The thickness of the forming enamel is gradually increased. In humans, the aprismatic outer enamel layer is about 30 mm. Calcium deficiencies lead to the emergence of structural defects mostly in the outer enamel (hypoplasia). Post-secretory ameloblasts are implicated in enamel maturation. Proteases such as kallikrein-4 (KLK-4) contribute to the hydrolysis of amelogenin. Amelogenin proteins represent the predominate subfamily of gene products found in developing mammalian enamel, and are implicated in the formation of the largest hydroxyapatite crystals in the vertebrate body. Maturation (or postsecretory) ameloblasts appear as ruffle-ended cells, bearing some similarities with the osteoclasts, although some segments display a smooth appearance. The organic matrix expelled from maturing enamel is pushed in deep recesses and internalized into lysosomes. Lysosomal acid phosphatase and other catalytic enzymes contribute to the degradation of this temporary organic matrix, especially amelogenin. Post-secretory ameloblasts allow calcium and phosphate ions to diffuse and contribute to the thickening and lengthening of enamel crystallites. As an enzymatic inhibitor, fluoride alters

ENAMEL FORMATION

Enamel formation results from the secretion of an extracellular matrix (ECM) by secretory ameloblasts and its eventual mineralization. Some enamel proteins TABLE 5.1

Global Composition of Enamel In weight (%)

In volume (%)

Mineral phase

96

87

Organic phase

0.6e1

2

Water

3.4e4 1% free water 2.4% bound water

FIGURE 5.1 Human dental enamel, internal zone. Groups of rods 7e11

cut longitudinally (parazones) alternate with group of rods in crosssection (diazones) at right angles from the parazones. Scanning electron microscope (SEM) preparation.

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these processes leading to mild to severe fluorosis. Defective enamel mineralization is also specifically due to the defective activity of maturation ameloblasts ENAMEL COMPOSITION

The mineral phase of enamel results in the formation of hydroxyapatite (HAp). In addition to calcium and phosphate, HAp composition also includes carbonate, magnesium, fluoride, sodium and many trace elements that are either mineral-associated or enzyme co-factors. Amelogenins constitute the bulk of the forming enamel matrix, while only a residual amount persists in the mature enamel. They are secreted as 28e25 kDa molecules, but are rapidly degraded into 21 kDa or less. They are presumably implicated in the nucleation and orientation of the crystallites. During enamel formation, proteases including enamelysine (MMP20) and other non-specific MMPs are implicated in the degradation of the molecule into small fragments that disappear during enamel maturation. KLK-4 also contributes to restriction of the organic content of mature enamel (0.4e0.6% w/v). Isoforms and spliced forms of amelogenins have been identified. Mutations of amelogenins result in different forms of amelogenesis imperfecta (AI) (Fig. 5.2). Enamelin forms about 5% of the enamel matrix, and the molecule seems to be implicated in nucleation and lengthening of crystallites and also in some forms of AI.

Ameloblastin (or amelin or shethlin) is synthesized by odontoblasts and ameloblasts, and it appears to be an adhesion molecule inhibiting cell proliferation and maintaining cell differentiation. A series of minor molecules are also present in enamel: amelotin, proteins issued from the serum, lipids and/or phospholipids, and calcium-binding proteins. Several enzymes such as MMPs, serine protease (KLK-4), acid and alkaline phosphatases are expressed during enamel formation and maturation. When these molecules are defective, due either to genetic mutations or to impaired enamel protein degradation, structural defects are detectable [9,10]. Dentin The composition of dentin is shown in Table 5.2. DIFFERENT TYPES OF DENTIN [11] OUTER PERIPHERAL DENTIN IN THE CROWN AND ROOT In the crown, odontoblasts at an early stage of

differentiation are implicated in the formation of a thin outer layer, the mantle dentin. Located beneath the dentino-enamel junction, dentin tubules are lacking or only some bended minute tubules are present. Its thickness varies between 15 and 30 mm. This layer displays specific elastic properties because it is less mineralized than the subjacent circumpulpal dentin [12] (Fig. 5.3). The proteins present in this thin layer are not phosphorylated, and a close relation has been established between

FIGURE 5.2 Human tooth displaying an amelogenesis imperfecta (AI). Left: enamel surface observed with the SEM. Hypoplastic enamel is seen, the outer part being missing. Center: longitudinal section of an AI affected tooth. Part of the outer enamel is missing. Right: the entire unsectioned tooth.

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TABLE 5.2 Global Composition of Dentin In weight (%)

In volume (%)

Mineral phase

70

50

Organic phase

20

30

Water

10e11

20

the degree of phosphorylation of the extracellular matrix molecules and their contribution to dentin or bone mineralization [14]. The same process occurs in the root where two distinct superficial layers have been identified for a total thickness of about 30 mm. The Hopewell-Smith hyaline layer and the granular Tomes’ layer are the first outer structures formed by root odontoblasts after their differentiation from the Hertwig’s root sheath. Cell interconnections and the general orientation of dentinal canaliculi along the long axis of the root suggest that these structures are implicated in the shaping and in the gradual reduction in the diameter of the root. As is

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the case in the crown, these peripheral layers are hypomineralized and there is a general increase of mineralization some distance away from the cemento-dentinal junction. CIRCUMPULPAL DENTIN Dentin is produced continuously in adults (4 mm/day) formed by regularly spaced Von Ebner lines appearing as incremental lines, and every 20e24 mm an Owen line is more prominent, implicating the dentin that includes four to six Von Ebner lines. This appearance is related to circadian and other rhythms. The primary dentin is formed until the moment the tooth becomes functional. Then the formation of secondary dentin starts and will continue for the lifetime of the tooth. Apart from the decreasing number of dentinal tubules, there is no structural or chemical difference between primary and secondary dentins. The number of tubules is about 20 000 tubules/mm2. Circumpulpal dentin includes the intertubular and peritubular dentins (Fig. 5.4). Intertubular dentin results from the transformation of predentin into dentin. It forms the bulk of dentin, whereas peritubular dentin is more variable, according to species, and location examined.

FIGURE 5.3 Upper part of the panel: normal dentin. Upper left: SEM of a tooth collected from a patient displaying an X-linked hypophosphatemia. Lower part, left: under the dentino-enamel junction, the outer mantle dentin is unaffected by the disease, whereas in the circumpulpal dentin, empty interglobular spaces are clearly seen between calcospherites. The dentino-enamel junction and the mantle dentin (md) appear to be normal, whereas the circumpulpal dentin interglobular spaces are filled with extracellular matrix partially degraded remnants accumulating between unmerged globular structures. Apparently the mantle dentin is not influenced by phosphorus homeostasis, as seen in X-linked hypophosphatemia [13].

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FIGURE 5.4

The peritubular dentin surrounds the lumen of the tubules. Intertubular dentin forms a continuous collagen-rich network. Left: horse’s dentin observed with the SEM. Right: human dentin observed with the transmission electron microscope (TEM).

Intertubular dentin is a type I collagen-rich structure. Collagen fibrils display a 100e120 nm diameter. Noncollagenous extracellular matrix proteins will be detailed in the paragraph related to the dentin composition. Some of them are phosphorylated and associated with the mineralization process either as promoters or inhibitors. Dentin crystallites appear as needle-like structures 3e4 nm thick and 60 nm long. They are located either along the surface of the collagen fibrils, in association with the so-called “holes” due to the quarter-staggered fibril structure, or filling the empty intercollagen spaces. In contrast, peritubular dentin is formed within the lumen of the tubules independently from the predentin transformation. It does not contain collagen fibrils, but is formed by a network of amorphous non-collagenous proteins, similar to those found in the intertubular dentin. In humans, the surface occupied by peritubular dentin is no more than 10% of the whole dentin volume. This distribution is related to the resistance to abrasion, intertubular dentin being elastic and easily abraded, whereas peritubular dentin is more resistant to attrition. Peritubular dentin has a higher carbonate and magnesium content than the intertubular dentin, and consequently is more soluble in acidic or in chelating solutions. The pathologic dentin defects related to specific gene mutations will be reviewed later in this chapter. They include formation of globular and interglobular

structures, defective dentin formation and altered root length. CELL AND TISSUE ORGANIZATION CELLULAR COMPARTMENT The odontoblasts are implicated in the synthesis of ECM molecules. These cells compose an individual compartment distinct from the pulp. The cells actively secrete predentin which is further transformed into mineralized dentin. THE PREDENTIN COMPARTMENT Another compartment comprises odontoblastic processes and predentin. Secretory vesicles are transferred along the processes and the larger part is secreted in the proximal predentin. Collagen fibrils start to aggregate at the site where they are secreted. The native fibrils elongate by end-to-end self-aggregation of the telopeptides and the diameter is increased by lateral aggregation of subunits. Proteoglycans such as fibromodulin and biglycan are also secreted in the proximal predentin and control collagen fibrillation. THE DENTIN COMPARTMENT The dentin compartment is a mineralized continuous structure starting at the metadentin border at the dentin edge and extending up to the dentino-enamel junction. Odontoblast processes are located within tubules (about 20 000/mm2).

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THE TOOTH BUD

A second group of non-collagenous proteins (NCPs) are secreted along the process, diffusing throughout the porous intertubular dentin and also contributing to the formation of peritubular dentin. DENTIN EXTRACELLULAR MATRIX COMPOSITION

Normal dentin extracellular matrix composition is shown in Table 5.3. Alterations of some small integrin-binding ligand N-linked glycoproteins (SIBLINGs) are related to some dentin pathologies such as dentinogenesis imperfecta or dentin dysplasia. DMP-1, PHEX and FGF-23 mutations are implicated in hyper- or hypophosphatemia. Cement/Cementogenesis [15,16] Cementum global composition in weight of mineral 65%, organic 23%, water 12%. TABLE 5.3

STRUCTURAL DIFFERENCES BETWEEN THE DIFFERENT TYPES OF CEMENT

The formation of cement, the layer that covers the root of the teeth, starts with an acellular afibrillar thin band located at the cervical junction between dentin and enamel (coronal cement). It is regulated by osteopontin [17]. Once the acellular band is completed, the formation of acellular cement begins and includes extrinsic fibrils found on the mid-root cement. The lower part of the root (apical cement) is covered by cellular and acellular cement, which includes intrinsic and/or extrinsic fibrils (Sharpey’s fibers from the periodontal ligament). Cementoblasts are either trapped in cementum, to form cellular cementum, or they slide away from the cementum surface, giving rise to acellular cementum. In the apical part, mixed cement appears as a multilayered structure that includes cellular and acellular successive layers.

Normal Dentin Extracellular Matrix Composition

Collagens 90% Non-collagenous proteins 10%

Phosphorylated proteins

Nonphosphorylated proteins

Type I collagen (89%) þ type I trimer (11%)

þ 1e3% Type III and V collagens

SIBLINGs Genes coding: in human chromosome 4 in rodent chromosome 5 Locus 4q21 Implicated in Dentinogenesis imperfecta and dentin dysplasia

DSPP (between 155 and 95 kDa) cleaved into: >DSP (N-terminal- proteoglycan forming dimers): 100e280 kDa >DPP (C-terminal) 94 kDa DMP-1: 61 kDa (proteoglycan) nucleator BSP: 95 kDa (proteoglycan) nucleation þ crystallite growth OPN: 44 kDa, glycoprotein, mineralization inhibitor MEPE: 66 kDa glycoprotein

Amelogenin

Spliced forms: Aþ 4: 8.1 kDa A-4: 6.9 kDa

Others enamel molecules

Ameloblastine

Osteocalcine DPG : dentin gla-protein (acidic g carboxyglutamic) Matrix Gla protein (MGP)

>5.7 kDa >mineralization inhibitor >14 kDa, not inhibitor of mineralization

Osteonectin- SPARC

43 kDa

Serum proteins

Albumin a2-HS glycoprotein

Small leucine-rich proteoglycans (SLRPS)

CS/DS PGs: decorine, biglycan 42 kDa KS PGs: lumican, fibromoduline, osteoadherine 50 kDa

Growth factors

FGF2, TGFb1, BMPs, ILGF I & II, PDGF

Enzymes

Alkaline and acidic phosphatase, serine proteases, MMPs: collagenases: MMP-1, 8, 13 gelatinases A (MMP-2), and B (MMP9) Stromelysine 1: MMP-3 MT1-MMP, enamelysine: MMP-20 ADAMs et ADAMTS

Proteolipids

ECM Phospholipids

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COMPOSITION OF THE ORGANIC PHASE OF CEMENTUM

(A)

The bulk of the organic fraction is formed by type I collagen (90% of the ECM) and type III collagen (z5%). The rest is formed by proteoglycans (namely decorine, biglycan, and lumican). Phosphorylated proteins such as BSP and osteopontin are implicated in cementum mineralization. Osteonectin, a nonphosphorylated glycoprotein, may also be present. Growth factors such as FGF-2, the cementum-derived growth factor (CDGF) and the cementum attachment protein (CAP) [18] play specific roles in cementum formation. CDGF is thought to be implicated in the recruitment and differentiation of cementoblasts, whereas CAP promotes the adhesion of cementoblasts on the dentin/cementum surface. Dlx-2 and Runx2/ Cbfa 1 regulate the initial cement formation and, consequently, any defective expression of these molecules induces cement alteration. Finally, alkaline phosphatase activity impairment interferes with cement formation.

(B)

MECHANISM OF SINGLE TOOTH ERUPTION Introduction The eruptive path of the dentition has been extensively studied and theories on tooth eruption have been proposed since as early as the 18th century. Teeth are unique specialized structures that develop from discrete invaginations of the oral ectoderm. These structures then invade the mesenchyme of the jaws [19]. The dental follicle, which includes the tooth bud, has to travel through alveolar bone to erupt into the oral cavity. Once the crown of the tooth is close to the gingival tissue, the oral tissues merge with the epithelium of the follicle to form the gingival and periodontal ligament. Specific steps are necessary for a specific tooth to reach and maintain its position on the occlusal plane: following intrabony development, the tooth bud migrates, and erupts in the oral cavity. Once in the oral cavity, the tooth continues its migration until contact is established with the opposing teeth and an occlusal table is established (Fig. 5.5).

Mechanism of Tooth Eruption The formation of the eruption pathway is independent of root formation. This was demonstrated by placing transmandibular wires over premolars in dogs prior to the onset of eruption. An eruption pathway caused by bone resorption formed in the alveolar bone despite the fact that the premolars were maintained stationary. As soon as the wires were removed, the teeth quickly erupted through the formed eruption pathway [7].

FIGURE 5.5 Active eruption of an upper right permanent canine. (A) The tooth is erupting through the gingival tissue slightly buccally. (B) The tooth is now in contact with the mandibular canine. All teeth in the upper quadrant have now at least one point of contact with their mandibular opposing tooth.

Role of the Dental Follicle in Tooth Eruption At the beginning of active tooth eruption (16 weeks postnatal), the dental follicle consists of a well vascularized, discrete connective tissue layer surrounding the tooth [20]. Its importance in tooth eruption was demonstrated by Cahill and Marks in 1980 [21]. They showed that tooth eruption could be interrupted if the dental follicle was removed prior to the onset of eruption. The role of the forming tooth in the formation of the eruptive pathway was also negated when they replaced the developing tooth in the dental follicle by inert (resin or metal) material. This material erupted in the same way as the tooth would have, showing that the presence of a tooth was not necessary for tooth eruption [22]. The dental follicle therefore must possess all, or the majority, of the molecules needed in the normal eruption pattern [7]. Colony stimulating factor 1 (CSF-1) is an essential factor for normal osteoclast differentiation and bone remodeling in mammals. Toothless rats show a severely reduced number of osteoclasts as may be seen in osteopetrosis [23]. Marks et al. have shown that the administration of CSF-1 to the toothless rat has a positive effect

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on tooth eruption, demonstrating the importance of CSF-1 and subsequently proper osteoclast activity on normal tooth eruption pattern [24]. Speed of Intraosseous Eruption The speed of intraosseous eruption depends on several factors. Each group of teeth is programmed to erupt at a different and specific time. However, the speed of intraosseous eruption is slow in relation to intraoral eruption and requires several years to complete. Tooth buds of the majority of permanent teeth are present in the infant, but will go through the gingival tissue only in the preadolescent years. Active eruption will begin after the crown of the tooth is fully completed [25] and will continue until the root is fully formed and the tooth is in contact with the opposing tooth on the occlusal plane Mechanisms of Intraosseous Tooth Eruption Tooth eruption is a complex event that involves many types of tissues. It is the only instance where a developing organ (the tooth) must exit the confines of its bony crypt [7]. To that effect, bone remodeling via osteoblasts and osteoclasts strategically positioned around the dental follicle must take place in order to ensure a normal eruption pattern. The eruption pathway must also continuously compensate for the increase in volume of the dento-alveolar process taking place simultaneously [7]. The dental crypt must resorb bone over the dental follicle by concentrating osteoclasts on its superior border at a specific time in the process of active eruption. Osteoblasts are more concentrated and abundant at the base of the dental follicle to produce alveolar bone. The combination of bone resorption and bone apposition, if properly balanced, will result in active intraosseous eruption [26e28]. The gubernacular canal, first mentioned by Hunter in 1778 [29] and in 1909 by James, is described as an eruption pathway used by the permanent tooth to replace the primary tooth. The gubernacular canals are too small to accommodate the crowns of permanent teeth, but are considerably enlarged by osteoclastic activity during the period of intraosseous tooth eruption [30]. The primary dentition mechanism of eruption via bone resorption is slightly different (Fig. 5.6). Osteoclastic activity is high, even if the tooth is stationary [20] and takes place on the superior aspect of the dental crypt. The gubernacular canal is enlarged and the tooth uses it as a pathway of eruption [31]. Primary teeth are very close to the surface [32] and a true intraosseous eruption pattern does not take place because the tooth buds are never totally covered with dense alveolar bone [33]. The crucial role of active bone resorption by osteoclasts in tooth eruption is demonstrated in osteopetrosis [34]. The teeth of osteopetrotic patients develop to their normal length, but fail to erupt into the oral cavity due to lack of osteoclastic activity, resulting in severe dental impactions.

FIGURE 5.6 Premolar tooth buds developing under primary molars. Note that the permanent molar does not replace a primary tooth but erupts distal to the primary dentition.

Genetic Control of Tooth Eruption According to the Tooth and Craniofacial Development Group of the University of Helsinki (www.bite-it.fi), a large number of genes is involved in the mechanism of tooth development and tooth eruption. Over 300 genes have been sequenced that involve several types of tissue [35]. Pelsmaekers [36] confirmed the genetic control of dental maturation in a study involving monozygotic twins. He showed that twins developed their dentition at the same time and at the same rate, regardless of the environment. It is of no surprise that a large number of heritable syndromes have a large dento-alveolar component and result in significant facial deformities by alteration of the pattern of tooth formation and tooth eruption [37,38]. Hypodontia, a condition in which some teeth fail to develop, is often associated with several syndromes, but it can also be found in the absence of other skeletal or ectodermal anomalies, suggesting the role of specific genes in the formation of the exact number of teeth and their proper positioning on the arch [39]. Mechanisms of Intraoral Tooth Eruption Multiple factors contribute to the onset of tooth eruption. Growth factors, such as insulin-like growth factor-1 (IGF-1), VEGF, tumor necrosis factor a (TNFa), are implicated in the transformation of monocytes into osteoclasts. Colony stimulating factor 1 (CSF-1) and monocyte chemotactic protein-1 (MCF-1) act in synergy with osteoprotegerin (acting as an inhibitor) and the overexpressed RANKL to influence osteoclastogenesis [40]. Four other factors, PTH [41], retarded protein interleukin-1alpha, Cbfa1/Runx2 (transcription factor) and the presence of osteoblasts also play a role. Interactions between growth factors,

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transcription factors and hormones are responsible for the differentiation of mesenchymal cells into osteoblasts at the base of the crypt and osteoclasts at the superior aspect of the dental crypt. These changes are observed during the active part of tooth eruption, and result in the tooth bud erupting through the oral mucosa. Important Factors Contributing to Lack of Tooth Eruption The best model to study proper eruption sequence is a condition that has been termed “primary failure of eruption” [42]. It is a multifactorial event where teeth develop normally but fail to erupt in the oral cavity and remain impacted. Dento-alveolar development continues around these teeth and a normal dento-alveolar height is achieved by periosteal growth of the bone formed around the adjacent teeth. This condition can affect all teeth but is more prevalent in the posterior segment. This phenomenon is also observed in osteogenesis imperfecta types III and IV patients who develop severe lateral open bites during growth (Fig. 5.7 ). The dento-alveolar process fails to develop normally in the area of the non-erupting tooth (Fig. 5.8). Severe occlusal anomalies are observed, such as posterior open bites while adjacent teeth tip into the space of the non-erupting tooth contributing to the creation of a severe malocclusion. The tooth is usually ankylosed and will not respond to orthodontic traction suggesting that the eruptive processes are absent and that the periodontal ligament’s response to mechanical traction does not result in the typical bone remodeling

FIGURE 5.8 Multiple impacted teeth of non-syndromic origin. A 16-year-old patient with only eight permanent teeth erupted, residual primary molars and 21 permanent teeth impacted. Note the lack of development of the maxillary anterior dento-alveolar process.

process observed in normal tooth eruption or orthodontic traction. Ankylosis occurs when cementum from the root of the tooth fuses with the dento-alveolar process. It may either be complete or partial ankylosis as when only some parts of the root fuse with the alveolar process. Bone remodeling stops in the affected area of ankylosis and the tooth fails to erupt. Lack or severely reduced dento-alveolar development is associated with this condition. Orthodontic traction is useless and the tooth either stays impacted or does not totally reach the occlusal plane [42]. True impaction may also occur when the tooth deviates from its normal path of eruption, totally forms in the bone and fails to erupt. Maxillary canines have the highest incidence of impaction, probably due to their complex path of eruption (Fig. 5.9). Once uncovered and provided that adequate space is created on the arch, these impacted teeth can be brought into proper position with orthodontic traction [43].

DEVELOPMENT OF THE DENTAL OCCLUSION Dental occlusion may be static or dynamic. The static phase refers to the stage when the mandibular teeth are brought into light contact with the maxillary teeth by the closing motion of the mandible. Dynamic occlusion occurs when teeth come in contact during chewing, speaking or swallowing.

Timing of Eruption FIGURE 5.7

Dentition of 12-year-old type III osteogenesis imperfecta patient showing bilateral posterior open bites. Permanent molars have erupted several years ago but have failed to come in contact, probably due to lack of growth of the dento-alveolar process. The dento-alveolar process fails to develop normally in the area of the non-erupting tooth.

Modern mammals have a diphyodont teeth arrangement, that is a deciduous and a permanent dentition in succession over their lifespan [44]. In humans, the primary dentition is made up of 20 teeth including incisors, canines and primary molars. Active intraosseous eruption of primary teeth occurs in the first and second

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FIGURE 5.10 Panoramic radiograph demonstrating a normal sequence of eruption in a 9-year-old patient. First permanent molars and upper and lower incisors are present in the mouth. Canines and premolars are resorbing the primary canines and molars.

FIGURE 5.9

Impacted upper right maxillary canine. The tooth has deviated from its normal path of eruption and is now resorbing the maxillary lateral incisor. The lower left second premolar is also impacted and poorly angulated as the lower left second primary molar has failed to resorb.

year of life and by 29 to 30 months, all 20 primary teeth are in place [45]. From the 30th month of life until the late teens, the permanent dentition develops, first with a path of intraosseous eruption followed by intraoral eruption. These movements are of large amplitude in relation to tooth size, occur in the three planes of space and must adjust to the continuous development of the dento-alveolar processes during growth [46]. Once teeth reach the occlusal plane, they continue to adjust with minute movements to stay in proper contact mesiodistally as well as occlusally. As the dental crypt moves toward the oral cavity, bone resorption takes place on the occlusal aspect of the crypt and bone apposition occurs at the base. At the same time, bone formation also increases the size of the dento-alveolar process. Complete remodeling of the alveolar bone occurs when deciduous teeth are replaced by succedaneous teeth. The alveolar bone associated with the primary tooth is completely resorbed together with the roots of the tooth while new alveolar bone is formed to support the newly erupted tooth [47]. The eruption sequence normally follows a path from the anterior to the posterior aspect of the mouth and is always constant during a normal eruption pattern (Fig. 5.10). Factors such as osteoprotegerin, RANKL, CSF-1 and VEGF control the path and sequence of eruption but their interactivity is still a subject of debate [24,48].

Intraosseous eruption of the lower incisors results in the eruption of the lower central incisors at 6 years of age, the first succedaneous teeth to erupt. Continuous eruption of succedaneous teeth occurs in the anterior part of the oral cavity until the age of 12e13 when the upper canines erupt [49]. The dental follicles of the permanent incisors, canines and premolars actively resorb bone and the primary tooth root structure, in order to appear in the oral cavity [50]. Odontoclasts, cells which are very close to osteoclasts, are actively involved in this process [51] and are linked to an increased expression of RANKL and to a decrease in osteoprotegerin (OPG) [52e54]. The absence of activity of odontoclasts results in non-resorption of primary teeth [55]. The succedaneous dentition continues to develop but the lack of resorption of the primary dentition creates an obstacle to eruption, resulting in an alteration of the path of eruption of the tooth buds, and severe impaction [56]. An example of this situation is the dentition of osteogenesis imperfecta types III and IV patients who show multiple impacted teeth caused by an alteration in osteoclastic activity leading to an absence of resorption of the primary dentition (Fig. 5.11) [57e59]. During the replacement of the primary dentition, intraosseous eruption takes place in the distal aspect of the primary dentition where the permanent molars

FIGURE 5.11 Impaction and malformation of premolars in an osteogenesis imperfecta type III patient. The crowns and the roots are no longer aligned as the primary tooth lack of resorption does not allow normal development of the premolar.

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erupt. A large amount of growth in the three planes of space associated with remodeling of the alveolar process and of the anterior aspect of the ramus must take place as the dental arches have to increase their size significantly to allow the eruption of these large teeth. The first molars will normally erupt between the ages of 6 and 8 years while the second ones appear around the age of 12 years. There is considerable variation in the timing of eruption between individuals. However, the sequence has to be consistent to avoid impaction and ectopic eruption of some teeth. The third molars or wisdom teeth are normally programmed to erupt around the age of 18 years but will fail to erupt and remain impacted in about 75% of the population. This is probably due to the lack of sufficient growth of the tuberosity and ramus in modern humans, resulting in a space deficiency to allow a normal eruption pattern for these teeth. The pattern of tooth eruption is an organized bilateral event where pairs of teeth from the right and left part of the jaws will erupt at about the same time [59]. Any significant change in this sequence will compromise the eruption pattern or sequence. Teeth may become impacted if primary teeth are lost prematurely and space becomes inadequate.

Pattern of Tooth Eruption Teeth are genetically designed [60] to erupt in sequence, and position themselves in an orderly manner in the plane of occlusion. The movement of eruption takes place in the three planes of space with vertical (pure eruption), labiolingual (transverse) and mesiodistal (drift) movements [31]. Tooth buds can also rotate during the intraosseous eruption phase. Each tooth shows an individualized pattern of eruption until it reaches its final position on the occlusal plane. The best example is the maxillary canine which tends to erupt in a mesial direction until it reaches the lateral incisor roots. It then turns to follow a more vertical path and may sometimes adopt a more mesial path of eruption [61]. Once in the oral cavity but still away from the occlusal plane, eruption is rapid until the tooth reaches the occlusal plane [62]. Final occlusal positioning is genetically programmed but is very dependent on the functional envelope of muscles and tissues surrounding the alveolar process. Moss [63,64] has described how the dento-alveolar process responds to alterations in the functional matrix, the development of which is guided by the patients’ vital functions such as breathing and feeding. During sleep apnea episodes, patients are unable to breathe through their nose and keep their mouth constantly open. This altered mandibular position results in a potential opening of the gonial angle (the angle formed by the junction of the ramus and the body of

the mandibular bone) and a vertical descent of the upper molars (Fig. 5.12). Constricted maxillary arches, severe crowding and anterior open bite may result from the disturbance in the proper breathing mechanism. Harvold has shown that forcing mouth breathing in monkeys resulted in the development of severe malocclusions [65]. The changes are severe enough to alter the facial appearance and modify mandibular growth [66].

Speed of Eruption While the intraosseous rate of eruption is fairly slow (1e10 mm/day) [62], once in the oral cavity, the speed of eruption is faster until teeth reach the occlusal plane. Proffitt reported a mean daily eruption rate of 25 to 75 microns for premolars out of occlusion [62]. The entire amount of eruption took place at night. Once in contact with the opposing dentition, active eruption stops but drift continues to take place via remodeling of the alveolar bone by the periodontal ligament (PDL) cells to ensure constant contacts in occlusion [67]. The speed of tooth eruption presents several characteristics. First, eruption itself has to proceed at a certain rate to ensure that the eruption sequence is respected and that the dentition develops in an orderly manner. Prolonged delays may prevent eruption and result in ankylosis of tooth to bone. Second, the eruption speed is not uniform. Erupting teeth move at different speeds at different times. Initially, eruption speed is slow within bone; it increases thereafter and becomes very slow as

FIGURE 5.12 Cephalometric radiograph of a 12-year-old male patient diagnosed with moderate to severe sleep apnea. The gonial angle of the mandible is opened and the anterior teeth are not in contact resulting in an anterior open bite.

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the tooth approaches the occlusal plane. These remarkable shifts in speed are also seen in root formation which is fast at first, slows as the apical foramen closes and is very slow thereafter. Third, this range of velocities implies that bone resorption and formation, like root growth, must occur at variable speeds depending upon the stage of eruption. Fourth, during intraosseous eruption, the rate of bone (and root) resorption determines the rate of eruption [25,68]. Continued Tooth Eruption Throughout Life Once the tooth is in occlusion, minute movements in the three dimensions of space will take place to compensate for secondary growth, and abrasion of occlusal and interproximal dental surfaces. Tooth movement takes place more as a displacement caused by dento-alveolar remodeling than tooth eruption. This stage cannot be described as active tooth eruption but as an occlusal compensation to changing conditions in the oral cavity. If an opposing tooth is lost or not present, active tooth eruption may resume with the erupting tooth elevating itself above the plane of occlusion. The dento-alveolar process will follow the erupting tooth and elongate. Tooth movement is present throughout lifetime. Changes in functional factors, such as sleep apnea, periodontal disease or tooth loss will all result in alteration of the occlusion through tooth movement and dentoalveolar remodeling [68].

MATURATION OF THE PERIODONTAL LIGAMENT The Periodontal Ligament: Role and Development of the Dento-Alveolar Complex The development of the periodontal ligament is derived from the inner layer of the dental follicle shortly after initiation of root development [69] and begins with root formation prior to tooth eruption [70]. The developing ligament contains undifferentiated cells capable of differentiating into osteoblasts, osteoclasts and fibroblasts. This combination of cells is responsible for a continuous remodeling of the alveolar bone. Dento-alveolar development from the PDL cells will compensate for the constant change in position of the tooth [68,70]. The periodontal ligament development is a highly organized process [71]. The PDL is a highly vascular and cellular connective tissue situated between the tooth and the alveolar bone. It provides supportive attachment and sensory functions [68]. Cells, vascular elements and an extracellular compartment of matrix proteins and glycosaminoglycans provide unique biophysical functions that enable mammalian teeth to adjust their position while

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remaining attached to the bony socket. Fibroblasts are the predominant cells of the periodontal ligament and have important roles in the development, structure and function of the tooth [72].

The Periodontium The periodontium, which is composed of the root cementum, the dento-alveolar process and the periodontal ligament is a unique structure with a variety of functions [73]. The main one is to provide mastication control and feedback and the second is to ensure optimum positioning of the teeth on the occlusal table. The ligament presents a high cellular turnover rate and Constant remodeling takes place as teeth being subject to masticatory forces and migrate in the dento-alveolar process. During the formation of the orofacial complex, the immature periodontal ligament around the dental follicle participates in the dental eruption and dentoalveolar process development. The maturing fibrils of the periodontal ligament shrink which may contribute to tooth eruption [74,75]. At the end of the intraosseous eruption process, the nascent periodontal ligament fuses with the oral epithelium and forms the periodontium. The ligament is responsible for the intraoral eruption stage of development and guides the tooth into its proper position on the dental arch. Once in position, the PDL will maintain the dental alignment and interdigitation position in relation to the other teeth for the life of the patient by constant remodeling of the dentoalveolar processes [68].

DENTO-ALVEOLAR GROWTH AND DISPLACEMENT OF BONE STRUCTURES Alveolar bone undergoes continuous remodeling due to tooth movement, masticatory forces and dental migration [47]. The periodontal ligament will also constantly remodel the alveolar bone, first rapidly during the period of active eruption and then more slowly once the teeth are in occlusal contact.

DENTAL DEFECTS AND SYNDROMES 1. Dental agenesis and hyperdontia 2. Dental defects • Shape defects • Enamel defects • Dentin defects

Dental Agenesis Tooth formation and morphogenesis are under strict genetic control [76]. Crucial molecules involved in tooth

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formation include RUNX2. In humans, heterozygous loss of function of MSK1 or PAX9 causes oligodontia [76,77]. Agenesis of a single tooth or teeth is a frequent anomaly in human dental development [60]. Oligodontia, hypodontia and anodontia are interchangeable names for this anomaly (Fig. 5.13). Familial tooth agenesis (FTA) is a well known condition in which the most commonly missing permanent teeth are the third molars (20% of the population), followed by the maxillary lateral incisors, the mandibular second bicuspids and the mandibular central incisors respectively [78,79]. Diseases such as scarlet fever or nutritional problems during pregnancy can result in dental agenesis. However, agenesis of several teeth is usually associated with heritable syndromes such as ectodermal dysplasias which constitute a wide and complex group of diseases. Ectodermal Dysplasia The most representative example of this group of conditions is anhydric ectodermal dysplasia with an incidence of between 1/10 000 and 1/100 000 live births. The defective gene is ED1 in humans, and Ta in the mouse [4]. It is X-linked and hemizygotes lack most of their dentition. Dental agenesis is strongly correlated to the absence or severe lack of dento-alveolar bone development as tooth eruption is largely responsible for the increase in volume of the dento-alveolar process. Thus, children affected by ectodermal dysplasia present with no or only a few teeth of conical shape and are deprived of the corresponding alveolar bone development (Fig. 5.14). Vertical face height development is stunted. The upper jaw also exhibits reduced length with a small palate and a correspondingly reduced cranial base width [80]. These patients look older than

FIGURE 5.13

Hypodontia. Very few permanent teeth are present but the ones present are normally shaped. This patient has only the upper central incisor in the maxilla and no lower incisors. Several permanent posterior teeth are also congenitally absent. The dentoalveolar process in the areas of anodontia is not severely reduced in size.

FIGURE 5.14 Ectodermal dysplasia. Hypodontia, abnormally shaped teeth such as conical crowns and short roots, and lack of development of the dento-alveolar processes are common findings in this syndrome.

their age due to sparse scalp hair, pursed lips and depressed nose base. Sofaer [81] found that 1 in 500 females with hypodontia in the permanent dentition and 1 in 50 with hypodontia in the primary dentition may be carriers for anhydric ectodermal dysplasia, suggesting a gene dose effect in ED1 mutations. Incontinentia Pigmenti-Williams syndrome Incontinentia pigmenti (BlocheSulzberger syndrome) is a syndrome similar to ectodermal dysplasia. It is also X-linked and often lethal in males. Its incidence reaches 1/40 000 in females. It affects both the primary and the permanent dentition, and 90% of patients show oral changes such as missing or conical teeth. There is a typical alopecia at the crown of the head, and the eyes are affected [82]. Ocular changes may be serious, such as severe myopia, retinal detachment or optic atrophy. Problems in mental development are found in 15% of the cases. Axelsson [83] examined 62 individuals with Williams syndrome and found that 40.5% of them lacked one or more permanent teeth while 11.6% of them lacked six or more permanent teeth. This appears to be linked to a contiguous gene deletion at chromosome 7 (7q11.23). Cherubism Cherubism is a rare neoplastic disease affecting particularly the jaw bones (Fig. 5.15). Multilocular fibrous dysplasia of both the maxilla and the mandible result in painless enlargement of these craniofacial bones. The dentition is affected by hypodontia and severe malocclusion [84]. The name cherubism comes from the fact that the face becomes deformed and almost circular due to fibrous proliferation. This condition affects mainly growing children and has been shown to regress in adulthood. Long bones are unaffected by the condition [85].

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FIGURE 5.15 Cherubism. Multilocular and radiolucent aspect of the maxillary and mandibular bones. Hypodontia, malformed teeth and severe malocclusion can be observed in this panoramic radiograph.

Cleft and Lip Palate Children born with unilateral or bilateral lip/palate clefts or palatal cleft alone present a high incidence of hypodontia (Figs 5.16 and 5.17) [86]. Lack of fusion between the maxillary and the medial nasal process explains the interruption or fragmentation of the dental lamina causing the agenesis of the lateral incisor on the cleft side. In a 2003 study, 49.8% of the cleft lip/palate children were missing this particular lateral tooth [87]. What is more intriguing is the dental agenesis of the teeth outside the cleft area, where 10.9% of patients lack the contralateral tooth [60]. The second most commonly missing permanent teeth are the maxillary second bicuspids, followed by the mandibular second bicuspids, and finally the first bicuspids [60]. Children with isolated cleft palate also have a high incidence of hypodontia (31.5%). Even in the absence of an alveolar cleft the teeth most frequently absent are the maxillary laterals

FIGURE 5.16 Occlusal view of a unilateral cleft palate. Collapse of the arch on the affected side, maxillary midline deviation and asymetry. The upper left lateral incisor is misshapen. The upper left canine is impacted.

FIGURE 5.17 Bilateral cleft lip and palate. Premaxilla is formed and includes the two central incisors. The clefts extend from the palate, the dento-alveolar process to the lip. Partial anodontia (absence of the permanent canines) and poorly shaped teeth in the area of the cleft are present.

followed by the second bicuspids. The more severe the palatal cleft, the more severe the hypodontia [88]. Down Syndrome The prevalence of tooth agenesis in Down syndrome is high. It is about 10% higher than in the general population and affects predominantly the mandibular permanent incisors [89].

Hyperdontia Hyperdontia (supernumerary teeth) is rarer than oligodontia. In the primary dentition, the maxillary lateral incisor is the only supernumerary tooth (0.10%) [90]. One to 3% of the general population will exhibit hyperdontia in the permanent dentition [91] and the supernumerary tooth is generally the maxillary lateral incisor. A Hong Kong study indicated a 2.7% rate for males and a 6.5% rate for females [104]. There is a higher incidence in Black (6%) than in Caucasian (0.64%) populations. Black children have more supernumerary teeth in the molar and bicuspid areas [92]. Some supernumerary teeth are distorted, conical and small. The most common is the mesiodens, a small conical tooth that is found between the permanent maxillary central incisors (Figs 5.18 and 5.19). If the abnormal tooth appears in the molar area, it is called a paramolar, and if located distal to the third molar, it becomes a supplemental tooth. The last category of non-syndromic hyperdontia is the odontoma. It is considered a hamartomatous

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FIGURE 5.18 The mesiodens is a small tooth usually located between the two upper central incisors and inverted (crown facing the nasal floor).

malformation. The odontoma can be complex (made of dental tissue totally disorganized) or compound (when it bears resemblance with a normal tooth). A supernumerary tooth is the primary cause of the lack of eruption of the maxillary permanent incisor [92]. The presence of multiple supernumerary teeth is often indicative of a syndrome, such as cleidocranial dysplasia [93]. Cleidocranial Dysplasia Cleidocranial dysplasia (CCD) is the best example of a syndrome associated with multiple supernumerary unerupted teeth (Fig. 5.20). Its incidence is about 1/ 1 000 000 with no preference for gender or race [94]. It is an autosomal dominant condition, characterized by delayed closure of the fontanels, brachycephalic skull, hypoplastic or aplastic clavicles and numerous dental anomalies. Multiple supernumerary teeth are part of the syndrome, as are retained primary teeth, and unerupted permanent teeth. Mutations in CBFA1 (RUNX2), mapped to chromosome 6p21, have been identified as

FIGURE 5.19 A rare view of a mesiodens that has erupted in the mouth on the labial aspect of the central incisors.

FIGURE 5.20 Cleidocranial dysplasia. A defect in the dental lamina results in a large number of supernumary teeth. These teeth usually stay impacted and are often misshapen. Primary teeth also fail to resorb especially in the posterior segments.

responsible for CCD [76,93]. CBFA1 is a transcription factor acting as a major controller of osteogenesis. Variable loss of function of CBFA1 may give rise to clinical variability, including classic CCD, mild CCD, and isolated primary dental anomalies. Ooshima et al. [95] reported genotypeephenotype associations such as “the more supernumerary teeth, the shorter the individual”. Mundlos [96] reported that lack of remodeling leads to persistence of the dental lamina, resulting in the formation of multiple supernumerary teeth at the same time that the crowns of the permanent teeth are formed, creating a third dentition [97]. Actually, it is the same type of bone dysplasia that links the supernumerary teeth to the persistence of the primary teeth in the mouth and to the delay, or non-eruption, of the permanent teeth.

Dental Defects Size and Shape Teeth can be abnormally small (microdontia), abnormally large (macrodontia), or misshaped. A study of tooth size in 43 parental pairs and their 100 offspring supported the hypothesis that tooth size is a sex-linked trait [98]. Microdontia is the most common developmental abnormality after hypodontia (Fig. 5.21). There is a significant association with hypodontia, and the more severe the hypodontia, the greater the possibility of microdontia [99]. A strong association was also found between agenesis of the second premolar and the pegged maxillary incisor [79]. Environmental conditions also contribute to variation in human tooth size [100]. In pregnant women affected with hypothyroidism, diabetes or hypertension, odontometric analyses in the offspring showed that maternal hypothyroidism and diabetes resulted in an increase in tooth size, while hypertension was associated with decreased tooth size.

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FIGURE 5.21 Non-syndromic 18-year-old female patient presenting with hypodontia, several severely misshapen and smaller than normal teeth not associated with a syndrome. Lateral and anterior open bite, anterior crossbite are also part of the malocclusion.

Chemotherapy and radiotherapy in infants, babies and young children contribute to changes in both primary and permanent dentition. The defects usually appear as hypodontia, microdontia, enamel hypoplasia and stunted roots. All these defects were present in the dentition of 52 surviving children afflicted by malignant diseases, even though none of them had received radiotherapy to their facial structures. Microdontia was present in 20% of this group, significantly above the 2.5% average for the normal population [101]. Severe and generalized microdontia is associated with several syndromes. Rieger syndrome, an autosomal dominant genetic defect, whose main features are ocular (microphthalmia), craniofacial (hypoplastic alae nasi) and hypospadia, presents with, in addition to hypodontia, an anterior mandibular tooth of smaller size and tapered in shape [102]. Identical dental findings are found in Williams syndrome, where teeth are tapered or incisors are screwdriver-shaped [83]. In Turner syndrome, there is a reduction of the dental crown’s height [103]. Several root anomalies were also reported. Trisomy 21 individuals also have a permanent dentition of reduced size [104]. The opposite was true for the deciduous dentition [104]. The authors hypothesized that, while the decreased mitotic activity of cells could logically explain the general reduction in the permanent teeth size, the phenomenon did not take place at the time of deciduous teeth mineralization (8e10 weeks of gestation) when the fetus experiences a burst of cellular activity. The result of this increased activity would be larger than normal deciduous teeth. Macrodontia is a defect rarely found in the normally developing child. However, there is a positive correlation with body stature and tooth size [100]. In subjects with a 47 XYY phenotype, the extra Y chromosome appears linked to larger teeth, probably because it

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provided a stronger and longer mitotic cell activity during the individual’s prenatal and early postnatal periods [105]. The KBG syndrome is an autosomal dominant trait characterized by short stature, facial anomalies (telecanthus, wide eyebrows, brachycephaly), mental retardation, skeletal anomalies (abnormal vertebrae, short metacarpals, short femoral necks) and macrodontia (mostly large upper incisors) [106]. Dental root anomalies are not frequent, and mostly limited to a reduction in size (short root anomaly). Normal dental roots are approximately twice the length of their corresponding crowns. A ratio of one to one or less defines a short root anomaly. Anomalies often associated with short roots are hypodontia, peg-shaped teeth, dens invaginatus and taurodontism. Short root anomalies may be limited to a few teeth (usually maxillary permanent laterals) or may affect the whole dentition. Chemotherapy, radiotherapy and total body irradiation can be devastating for the permanent dental root development in the pediatric community. A study mentioned earlier found abnormal permanent root development in long-term survivors of malignant diseases, with the root disturbance corresponding to the duration of treatment and dose of irradiation [101]. The addition of chemotherapy treatment further disturbs tooth formation. The extent of the damage seems to depend upon the stage of histodifferentiation of the tooth and the amount of rads administered. Any dose higher than 2000 rads produced serious dental defects, regardless of the child’s age. The authors described a case of a boy who had received 4050 rads to the right middle ear (rhabdomyosarcoma) and to the right cervical lymph nodes when he was 2 years old; the rads were supplemented with vincristine, actinomycin D and cyclophosphamide. At age 13, his panoramic radiograph revealed that his permanent teeth were present in the oral cavity but no root formation was noted on the first molars and several teeth such as the second premolars and molars were not developing normally (Fig. 5.22). Hultta¨ et al. reviewed the radiographs of 52 children who had received stem cell transplantation before the age of 10. The children were treated with anticancer therapy for acute lymphomatic leukemia, acute monocytic leukemia and non-Hodgkin lymphomas. A postoperative assessment took place at an average age of 7.2 years, and 945 teeth were examined. All patients presented with severe disturbances in the root development of the permanent teeth. The most severely affected teeth were found in patients who were 3.1 to 5.0 years at the time of the transplantation. Taurodontism is a dental root anomaly found in 2.5% of the Caucasian population. It is caused by the apical enlargement of the body of the tooth (or coronal part)

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FIGURE 5.22 Rhabdomyosarcoma patient having received radiotherapy at a young age. Note the total absence of development of the roots of the first molars, Conical root shape and malformed second molars and premolars. Lower left second molar is not developing as well as the third molars.

at the expense of the root length. Therefore, the roots appear much shorter. Simultaneously, the classic cemento-enamel constriction lessens or disappears. Taurodontism is often found in individuals with aneuploidy of the X chromosome [107]. Enamel Defects Enamel is a unique, highly mineralized tissue. Its protein constituents, amelogenin (80e90%), ameloblastin (5e10%), enamelin (3e5%) and enamelysin (1%) are produced by ameloblasts. They are responsible for enamel maturation. Deleterious effects of environmental and systemic factors during enamel formation are often traceable when the tooth emerges into the oral cavity. Defects in any of the above proteins can trigger enamel defects without affecting other parts of the structure [108]. Enamel defects are also part of syndromes that are hereafter mentioned. Each group of teeth has a unique anatomy and is easily recognizable. Each crown of a particular tooth may show specific alterations that may or may not endanger its integrity. For example, “dens in dente” or “dens invaginatus” is a developmental anomaly where the enamel of the maxillary incisors enfolds itself on the lingual pits and penetrates into the pulpal space. The “dens in dente” is a structure with its own inner enamel, dentin and blood supply (Fig. 5.23) [109]. As its name implies, it looks like a tooth within a tooth. If the opening is not protected and sealed in time, the invagination may communicate with the pulpal space which may become contaminated by oral bacterial flora. A pulpal infection ensues, requiring endodontic therapy. The incidence of pulpal necrosis is between 0.25 and 9.66%. “Dens evaginatus” or “tubercle shaped cusp” is the opposite of dens in dente. The enamel folds itself outwardly and resembles an extra cusp near the central

FIGURE 5.23 Dens in dente. The dental papilla is forming a second tooth inside the normally developed tooth.

groove of the tooth [110]. This event takes place in the early stage of odontogenesis. Posterior permanent teeth, notably bicuspids, most frequently present with this type of enamel defect. During active intraoral tooth eruption, the opposite tooth will eventually occlude into the central groove and fracture the little tubercle. The dentin and the minuscule pulpal tissue are then exposed, and the tooth becomes infected. This anomaly is frequent in subjects with Asian ethnic background but is also found in other races [111]. The “talon cusp”, which resembles the “dens evaginatus”, limits itself to maxillary incisors (Fig. 5.24). It favors deciduous teeth, but may be found on permanent teeth as well. It can interfere with the patient’s occlusion and presents both an esthetic and a treatment challenge [112]. Enamel defects are also associated with nutritional deficiencies. They appear as linear or irregular spots on the teeth, usually on the edge of the permanent incisors and on the superior third surfaces of the first permanent molars. These surfaces undergo their odontogenesis during the neonatal period and during the first year of life. Vitamin D, calcium and phosphorus deficiencies as well as any severe nutritional deficiency will trigger such defects. Other disturbances may also cause enamel defects. A low calcium concentration during enamel formation is a specific determinant of enamel hypoplasia [113].

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FIGURE 5.25 Hypoplastic phase of enamelogenesis imperfecta FIGURE 5.24 Talon cusp that does not allow for proper tooth positioning as it interferes with arch alignment.

Enamel mineralization defects or amelogenesis imperfecta (AI) [108] may take place at different stages of odontogenesis. When the damage occurs during the secretory time (hypoplastic defect), the enamel is of good quality, but of insufficient quantity, sometimes practically nonexistent. When the damage occurs during the mineralization time (hypomineralized defect), the enamel is of normal thickness but poor quality. When it takes place during maturation time (protein processing and crystallite maturation defect), the enamel retains its normal thickness but quality is even poorer. Most enamel defects have a dominant inheritance. Enamelin (ENAM 4q21) is the main gene involved. A classification of AI was proposed by Witkop in 1988 [108]. He stated that AI, a developmental intrinsic defect in enamel formation, can be classified into four different types: 1. Hypoplastic defect, where the enamel does not develop to its normal thickness (Fig. 5.25) 2. Hypomaturation defects, where the enamel is of normal thickness but of mottled appearance 3. Hypocalcification defects, where the enamel is of normal thickness but friable and yellowish-brown in color 4. Hypomaturationehypoplastic defects associated with taurodontism (hypoplastic and hypocalcified enamel in teeth with atypically enlarged pulp chambers). The complexity and the high intrafamilial variability, together with the difficulties in diagnosing the precise type of AI (such as hypocalcification versus hypomaturation), make it difficult to identify the precise gene defect. Teeth with AI are difficult to maintain, resulting in a high proportion of edentulism among family members. Syndromes associated with AI are rare. In

(see also Fig. 5.2). The enamel is normal but in very small quantity. Notice that the enamel is already fracturing at the incisal edges and is very translucent.

mucopolysaccharidoses (MPS) enamel can be specifically targeted. MPS is a family of metabolic disorders (lysosomal enzyme deficiencies) that share an autosomal recessive inheritance (except for MPS II which is sexlinked). They are all affected with “dysostosis multiplex”. Yet only one MPS, the Morquio syndrome (MPS IVB), is associated with enamel defects, even though the affected individuals show milder phenotypes than their counterparts with MPS IVA. The enamel is yellow, brittle and weak [114]. The tricho-dento-osseous syndrome (TDO), an autosomal dominant trait, is caused by a mutation in DLX3, mapped to 17q21. It presents with a combination of amelogenesis imperfecta (pitted thin and discolored enamel) and taurodontism. The taurodontic tooth is characterized by an abnormally large pulpal space and absence of the constriction between the crown and the root (cemento-enamel junction). TDO individuals have kinky hair (curly at birth), thick dense bones and brittle nails. The dental phenotype is present in all cases [115]. Focal dermal hypoplasia (FDH) or GoltzeGorlin syndrome presents with severe dental defects: the enamel is irregular and misshaped and covers small misaligned teeth. Hypodontia is present as well. It is transmitted as a sex-linked trait on the short arm of the X chromosome [116]. Dentine Defects In dentine, collagen (mostly collagen type I) serves as a network for the forming carbonate apatite. Dentine matrix protein (DMP1), dentin sialoprotein (DSP) and the dentin phosphoprotein (DPP) are the noncollagenous components that regulate dentinogenesis. The most important dentinal defect is dentinogenesis imperfecta (DGI), an autosomal dominant trait (Figs 5.26 and 5.27). In 1973, Shields et al. divided DGI into

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FIGURE 5.26 Dentinogenesis imperfecta. Teeth are brownish in color, fracture easily and are very difficult to restore. Enamel fractures easily as the dentineeenamel bond is weak.

three subgroups [117]. All of them are linked to chromosome 4q21, and result from mutations in DSP [108]. The first group, or DGI type I, is linked to osteogenesis imperfecta (OI); the second group, DGI type II, presents with almost the same clinical picture but with no other systemic involvement; the third type, DGI type III, is a rare isolated disorder known as the “Brandywine isolate”. Two more dentinal defects, dentin dysplasia I [101], an exceedingly rare condition and dentin dysplasia II (DDII) which bares close resemblance with DGI II, complete the list [117]. Dentinogenesis imperfecta type II is one of the most common dominantly inherited disorders. Its incidence is around 1/8000 and its penetrance is high [118].

FIGURE 5.27 Dentinogenesis imperfecta in the mixed dentition. Severe wear and attrition are visible. Permanent molars have a blueish coloration.

In 2003, Sreenath et al. [119] studied the role of the Dspp gene in tooth mineralization in Dspp-null mice and concluded that it regulated proteoglycan synthesis during dentinogenesis. Beattie et al. performed linkage and mutation analysis in studying phenotypic variation in dentinogenesis imperfecta/dentin dysplasia (DDII) linked to 4q21 in a large pedigree spread over four generations. The related individuals all displayed dentin defects associated with DGI and DDII. The pattern of inheritance was autosomal dominant with complete penetrance. They concluded that type II dentin dysplasia and DGI type II as well as DGI type III were non-syndromic heritable dentin defects and should be put in one single category and classified by their respective severities [120]. DSPP is expressed in bone as well as in dentin. However, bone is not involved in DGI type II [3]. It is thus possible that other molecules in bone exert redundancy. DGI is easily recognizable. The teeth are opalescent, with a color varying from gray to yellow. They are fragile and the enamel chips off easily because its poor adhesion to dentin and the dentin itself is defective. Dental fractures can be severe because the circumpulpal dentin (deeper dentin) is weak and does not resist stress. On radiographs, the crowns appear bulbous, the roots are slender and short and the pulp spaces are obliterated by irregular dentin. Dentinogenesis imperfecta type I is associated with the osteogenesis imperfecta syndrome (OI) [58,121] (see Chapter 18). In this heritable disorder, bone brittleness is associated with a decreased bone mass. The syndrome is associated with other anomalies, such as blue sclerae, hearing loss and joint hypermobility [57]. Most patients harbor a mutation in one of the two genes encoding type I collagen [122]. Sillence has proposed a classification into four types which is still frequently used [123]. Type I is the mildest form, type II is lethal, type III is the most severe one compatible with life (Fig. 5.28) and type IV, a heterogeneous group with variable severity. Glorieux and his group were able to describe three more types (V, VI and VII) [122]. When OI is caused by a mutation in a protein other than collagen type I, DI is not present. In the past, OI linked to collagen I defect was further classified as having DI or not. However careful examination of apparently normal dentitions may show subtle signs of DI [124]. Deciduous teeth are more affected than the permanent ones. Other problems are failure of eruption, tooth impaction and severe malocclusion. The most common type of malocclusion in the OI patient is the class III malocclusion. A lack of anteroposterior development associated with a larger mandible results in anterior crossbites. Lack of dentodevelopment, especially in the posterior region, leads to the development of lateral open bites (Figs 5.29 and 5.30) [58]. The latter may occur

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FIGURE 5.28 Osteogenesis imperfecta type III patient presenting

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FIGURE 5.29 Lateral cephalometric radiograph of an osteogenesis imperfecta type III patient presenting the typical lack of anterior projection of the maxilla and lateral posterior open bites.

with dentinogenesis imperfecta. Anterior crossbite is observed.

even in forms of OI not linked to type I collagen abnormalities [3,124]. X-linked hypophosphatemia (XLH), is a heritable disease caused by impaired renal phosphate transport (see Chapter 26) (Figs 5.31 and 5.32). It is linked to mutations in the PHEX gene. It results in rickets, osteomalacia, growth retardation and dentin defects [125, 126]. The latter trigger spontaneous dental abscesses on teeth that appear intact and devoid of carious lesions. Schwartz et al. examined 14 patients with XLH and found a high percentage of these particular abscesses. The teeth showed impaired dentin mineralization that allowed elongated pulp horns to reach the cementoenamel junction and, through minor enamel abrasions, be exposed to the oral bacteria [127]. Shields et al. measured the pulp profile area of the hypophosphatemic teeth on radiographs as a measure of secondary dentin development [125]. The large pulp size, the lack of secondary dentin and the globoid dentin explained

FIGURE 5.30 Intraoral pictures of an osteogenesis imperfecta type III malocclusion in the mixed dentition characterized by an anterior crossbite and lateral open bites. Dentinogenesis imperfecta is also associated with osteogenesis imperfecta in this patient.

FIGURE 5.31

Bite-wing type dental radiograph of a hypophosphatemia patient. The pulp chambers are enlarged. No protective dentin is present leading to spontaneous pulpal necrosis and dental abscesses.

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FIGURE 5.32 Periapical radiograph of a hypophosphatemia patient. Periapical radiolucency is detected on a routine radiograph. No evidence of dental decay or periodontal involvement is present.

the occurrence of dental abscesses. Chaussain-Miller et al. stated that early treatment of XLH with the association of the 1-hydroxylated form of vitamin D and phosphates was beneficial with definite improvement of the dentin tissue [121].

FIGURE 5.34 Periapical radiograph of the anterior aspect of the maxilla of an osteopetrotic patient showing lack of dental eruption, poorly formed teeth and increased density of the alveolar bone.

Osteopetrosis, also known as Albers-Scho¨nberg disease, is characterized by a defect in the differentiation and/or function of osteoclasts [128,129]. The most severe forms tend to have an autosomal recessive inheritance (1 in 250 000 births) while the milder forms have an autosomal dominant inheritance (1 in 20 000 births). Marks [130] demonstrated that tooth eruption was dependent on bone resorption in osteopetrotic rats. When teeth cannot erupt, they change their shape and become dilacerated. In the most severe form of osteopetrosis, called malignant osteopetrosis, the dental findings include delayed tooth eruption, impaction and severe root and crown malformation (Figs 5.33e5.35) [129].

CONCLUSION

FIGURE 5.33 Osteopetrosis in a 6-year-old patient. Extremely dense bone, multiple unerupted and malformed teeth are present. No osteoclastic activity is present to counterbalance the osteoblastic proliferation. Dental eruption patterns are completely disturbed.

Tooth eruption through the dental follicle is a complex phenomenon. It involves osteoblastic and osteoclastic activity that contributes to form dento-alveolar bone. This bone formation and the development of dental occlusion, first for the deciduous, then for the succedaneous dentition, will have a profound impact on the

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REFERENCES

FIGURE 5.35 Intraoral occlusal picture of an osteopetrotic patient. No bone remodeling is present. Dental eruption is incomplete. Proliferation of bone around the dental follicle without bone resorption and remodeling does not allow for normal tooth eruption.

development of the lower part of the face. If teeth are missing or fail to erupt in the proper position, dentoalveolar development will be stunted and the alveolar ridge will either not form or be severely reduced in size. In severe cases of hypodontia, very little dentoalveolar volume is present, resulting in a collapsed lower face height. The mode of tooth eruption, guided by precise genetic control, is also largely influenced by its neuromuscular environment and the response of the periodontal ligaments to the environment. Tooth positioning may be altered by functional imbalance, as the periodontal ligament will respond to forces and rapidly remodel the dento-alveolar complex in order to maintain the necessary physiological distance between the alveolar wall and the dental root. Dentofacial development is intimately linked to proper dento-alveolar development that is itself dependent on a properly developing sequence of dental eruption and PDL influence. The study of heritable conditions has helped to explain why and how dental defects take place. It has also allowed delineation of specific syndromes. As it is the case in other disciplines, a thorough oral examination together with a comprehensive medical history will continue to contribute to further our knowledge of the complex mechanisms underlying dental maturation.

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[47] Sodek J, McKee M. Molecular and cellular biology of alveolar bone. Periodontology. 200;24:99-126. [48] Dobbins DE, Sood R, Hshiramoto A, et al. Mutation of macrophage colony stimulating factor (Csf1) causes osteopetrosis in the tl rat. Biochem Biophys Res Commun 2002;294:1114e20. [49] Smith B, Garn S. Polymorphisms in eruption sequence of permanent teeth in American children. Am J Phys Anthropol 1987;74:289e303. [50] Harokopakis-Hajishengallis E. Physiologic root resorption in primary teeth: molecular and histological events. J Oral Sci. 2007;49:1e12. [51] Sahara N, Okafuji N, Toyoki A, et al. Odontoclastic resorption of the superficial nonmineralized layer of predentine in the shedding of human deciduous teeth. Cell Tissue Res. 1994;277: 19e26. [52] Fukushima H, Kajiya H, Takada K, Okamoto F. Expression and role of RANKL in periodontal ligament cells during physiological root resorption in human deciduous teeth. Eur J Oral Sci. 2003;111:346e52. [53] Bille M, Kvetny M, Kjaer I. A possible association between early apical resorption of primary teeth and ectodermal characteristics of the permanent dentition. Eur J Orthodont 2008;30:346. [54] Oshiro T, Shibasaki Y, Yoda S, et al. Immunolocalization of vacuolar type Hþ ATPase, cathepsin K, matrix metalloproteinase 9, and receptor activator of NFkB ligand in odontoclasts during physiological root resorption of human deciduous teeth. Anat Rec 2001;264:305e11. [55] Yoda S, Suda N, Kitahara Y, Komori T, Onyama K. Delayed tooth eruption and suppressed osteoclast number in the eruption pathway of heterozygous Runx2/Cbfa1 knockout mice. Arch Oral Biol 2004;49:435e42. [56] Iizuka T, Cielinski M, Aukerman SL, Marks Jr SC. The effects of colony-stimulating factor-1 on tooth eruption in the toothless (osteopetrotic) rat in relation to the critical periods for bone resorption during tooth eruption. Arch Oral Biol. 1992;37:629e36. [57] O’Connell A, Marini J. Evaluation of oral problems in an osteogenesis imperfecta population. Oral Surg Oral Med Oral Path Oral Radiol Endodontol 1999;87:189e96. [58] Schwartz S, Tsipouras P. Oral findings in osteogenesis imperfecta* 1. Oral Surg Oral Med Oral Pathol 1984;57:161e7. [59] Mossey P. The heritability of malocclusion: part 2. The influence of genetics in malocclusion. J Orthodont 1999;26:195. [60] Vastardis H. The genetics of human tooth agenesis: new discoveries for understanding dental anomalies. Am J Orthodont Dentofacial Orthoped 2000;117:650e6. [61] Ericson S, Kurol J. Resorption of maxillary lateral incisors caused by ectopic eruption of the canines: a clinical and radiographic analysis of predisposing factors. Am J Orthodont Dentofacial Orthoped 1988;94:503e13. [62] Proffit WR, Prewitt JR, Baik HS, Lee CF. Video microscope observations of human premolar eruption. J Dent Res. 1991; 70:15. [63] Moss M, Salentijn L. The primary role of functional matrices in facial growth* 1. Am J Orthodont 1969;55:566e77. [64] Moss M, Young R. A functional approach to craniology. Am J Physl Anthropol 1960;18:281e92. [65] Harvold EP, Tomer BS, Vargervik K, Chierici G. Primate experiments on oral respiration* 1. Am J Orthodont 1981;79: 359e72. [66] Vargervik K, Miller AJ, Chierici G, Harvold E, Tomer BS. Morphologic response to changes in neuromuscular patterns experimentally induced by altered modes of respiration* 1. Am J Orthodont 1984;85:115e24.

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[91] Davis P. Hypodontia and hyperdontia of permanent teeth in Hong Kong schoolchildren. Commun Dent Oral Epidemiol 1987;15:218e20. [92] Harris E, Clark L. An epidemiological study of hyperdontia in American blacks and whites. J Inform 2008;78:3. [93] Garvey MT. Supernumerary teeth e an overview of classification, diagnostic and management. J Can Dent Assoc. 1999;65:612e6. [94] Baumert U, Golan I, Redlich M, Aknin JJ, Muessig D. Cleidocranial dysplasia: molecular genetic analysis and phenotypic based description of a Middle European patient group. Am J Med Genet A 2005;139:78e85. [95] Ooshima T, Mishima IR. Sobue s K. The prevalence of developmental anomalies of teeth and their association with tooth size in the primary and permanent dentitions of 1650 Japanese children. Int J Paediatr Dent 1996;6:87e94. [96] Mundlos S. Cleidocranial dysplasia: clinical and molecular genetics. J Med Genet. 1999;36:177. [97] Jensen B, Kreiborg S. Development of the dentition in cleidocranial dysplasia. J Oral Pathol Med 1990;19:89e93. [98] Lewis D, Grainger R. Sex-linked inheritance of tooth size: a family study. Arch Oral Biol. 1967;12:539e44. [99] Brook AH. A unifying aetiological explanation for anomalies of human tooth number and size. Arch Oral Biol. 1984;29:373e8. [100] Garn Sm, Osborne RH, Alvesalo L, Horowitz SL. Maternal and gestational influences on deciduous and permanent tooth size. J Dent Res. 1980;59:142e3. [101] Maguire A, Evans RG, Amineddine H, et al. The long-term effects of treatment on the dental condition of children surviving malignant disease. Cancer 1987;60:2570e5. [102] Jorgenson RJ, Levin LS, Cross HE, et al. The Rieger syndrome. Am J Med Genet. 1978;2:307e18. [103] Midtbø M, Halse A. Root length, crown height, and root morphology in Turner syndrome. Acta Odontol 1994;52:303e14. [104] Peretz B, Shapira J, Farbstein H, Arieli E, Smith P. Modification of tooth size and shape in Down’s syndrome. J Anat 1996; 188:167. [105] Alvesalo L, Osborne R, Kari M. The 47, XYY male, Y chromosome, and tooth size. Am J Hum Genet. 1975;27:53. [106] Herrmann J, Pallister PD, Tiddy W, Opitz JM. The KBG syndrome e a syndrome of short stature, characteristic facies, mental retardation, macrodontia and skeletal anomalies. Birth Defect Orig Art Ser 1975;11:7. [107] Jaspers M, Witkop Jr C. Taurodontism, an isolated trait associated with syndromes and X-chromosomal aneuploidy. Am J Hum Genet. 1980;32:396. [108] Witkop Jr C. Amelogenesis imperfecta, dentinogenesis imperfecta and dentin dysplasia revisited: problems in classification. J Oral Pathol Med 1988;17:547e53. [109] Vajrabhaya L. Nonsurgical endodontic treatment of a tooth with double dens in dente. J Endodont 1989;15:323e5. [110] Stewart R, Dixon G, Graber R. Dens evaginatus (tuberculated cusps): genetic and treatment considerations. Oral Surg Oral Med Oral Pathol 1978;46:831e6. [111] Mellor J, Ripa L. Talon cusp: a clinically significant anomaly. Oral Surg Oral Med Oral Pathol 1970;29:225e8. [112] Bailleul-Forestier I, et al. The genetic basis of inherited anomalies of the teeth. Part 2: Syndromes with significant dental involvement. Eur J Med Genet. 2008;51:383e408. [113] Nikiforuk G, Fraser D. The etiology of enamel hypoplasia: A unifying concept. J Pediatr 1981;98:888e93. [114] Gorlin RG, Hennekam RCM. Syndromes of the Head and Neck. Oxford University Press; 2001.

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[115] Price JA, Wright JT, Walker SJ, et al. Tricho-dento-osseous syndrome and amelogenesis imperfecta with taurodontism are genetically distinct conditions. Clin Genet. 1999;56:35e40. [116] Al Ghamdi K, Crawford P. Focal dermal hypoplasia e oral and dental findings. Int J Paediatr Dent 2003;13:121e6. [117] Shields E, Bixler D, El-Kafrawy A. A proposed classification for heritable human dentine defects with a description of a new entity. Arch Oral Biol. 1973;18:543e53. [118] Malmgren B, Lindskog S, Elgadi A, Norgen S. Clinical, histopathologic, and genetic investigation in two large families with dentinogenesis imperfecta type II. Hum Genet. 2004;114:491e8. [119] Sreenath T, Thyagarajan T, Hall B, et al. Dentin sialophosphoprotein knockout mouse teeth display widened predentin zone and develop defective dentin mineralization similar to human dentinogenesis imperfecta type III. J Biol Chem. 2003;278:24874. [120] Beattie ML, Kim JW, Gong SG, et al. Phenotypic variation in dentinogenesis imperfecta/dentin dysplasia linked to 4q21. J Dent Res. 2006;85:329. [121] Chaussain-Miller C, Sinding C, Septier D, et al. Dentin structure in familial hypophosphatemic rickets: benefits of vitamin D and phosphate treatment. Oral Dis 2007;13:482e9. [122] Rauch F, Glorieux F. Osteogenesis imperfecta. Lancet 2004;363:1377e85.

[123] Sillence D, Senn A, Danks D. Genetic heterogeneity in osteogenesis imperfecta. Br Med J 1979;16:101. [124] Lygidakis N, Smith R, Oulis C. Scanning electron microscopy of teeth in osteogenesis imperfecta type I. Oral Surg Oral Med Oral Pathol Oral Radiol Endodontol 1996;81:567e72. [125] Shields Ed, Scriver CR, Reade T, et al. X-linked hypophosphatemia: the mutant gene is expressed in teeth as well as in kidney. Am J Hum Genet. 1990;46:434. [126] Murayama T, Iwatsubo R, Akiyama S, Amano A, Morisaki I. Familial hypophosphatemic vitamin D-resistant rickets: dental findings and histologic study of teeth. Oral Surg Oral Med Oral Pathol Oral Radiol Endodont 2000;90:310e6. [127] Schwartz S, Scriver CR, Reade TM, Shields ED. Oral findings in patients with autosomal dominant hypophosphatemic bone disease and X-linked hypophosphatemia: further evidence that they are different diseases. Oral Surg Oral Med Oral Pathol 1988;66:310. [128] Stark Z. Osteopetrosis. Orphanet J Rare Dis 2009;4:1e10. [129] Moghadam MI, Nemati S, Johan M. Dental radiographic findings of malignant osteopetrosis: report of four cases. Iran J Radiol 2009;6:141e5. [130] Marks Jr S. Tooth eruption depends in bone resorption: experimental evidence from osteoetpotrotic (ia) rats. Metab Bone Dis Relat Res. 1981;3:107e15.

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Parathyroid Hormone and Calcium Homeostasis John T. Potts, Thomas J. Gardella Endocrine Unit, Department of Medicine, The Massachusetts General Hospital and Harvard Medical School Boston, MA, USA

INTRODUCTION Parathyroid hormone (PTH) and PTH-related peptide (PTHrP), along with other calciotropic hormones, play critical roles in calcium homeostasis and bone biology. In contrast to PTH, which is produced by discrete endocrine glands, PTHrP is produced as a paracrine/ autocrine factor in many different adult and fetal tissues and has, unlike PTH, multiple functions (Fig. 6.1). First discovered as a calcium-regulating hormone in the 1920s [1e3], PTH is secreted by the parathyroid glands and is the critical regulator of blood calcium concentration in all terrestrial vertebrate species. PTHrP,

FIGURE 6.1 Biology of the PTH/PTHrP receptor. The PTH/ PTHrP receptor interacts with two ligands, PTH and PTHrP, and can activate several second-messenger signaling pathways, including the cAMP/protein kinase A (PKA) and Ca2þ/inositol 1,4,5-triphosphate/ PKC pathways. The receptor is abundantly expressed in bone and kidney, where it mediates the endocrine actions of PTH, and in the metaphyseal growth plate and numerous other tissues, where it mediates the autocrine/paracrine actions of PTHrP.


a slightly larger molecule than PTH, was discovered more recently through efforts to identify the factor that causes, when produced in excess by certain tumors, the humoral hypercalcemia of malignancy syndrome [4e6]. PTH and PTHrP most likely evolved from a common ancestral precursor, as discussed further below. The polypeptides share only limited overall amino acid sequence identity, yet at least their N-terminal regions are sufficiently homologous to enable them to bind to and activate a common G protein-coupled receptor, the PTH/PTHrP receptor (also referred to as PTH1R) [7e9]. This receptor mediates the most important biologic actions of both peptides: PTH-dependent regulation of calcium homeostasis and PTHrP-dependent regulation of endochondral bone formation [10e14]. This chapter reviews (1) the comparative chemistry of PTH and PTHrP, their genes, and their interactions with PTH1R; (2) the current molecular models of interactions between the two ligands and their common receptor; and (3) the different biologic roles of both peptides on target tissues, such as the role of PTH in calcium homeostasis and bone turnover and the role of PTHrP in bone and cartilage development, as well as the functional characteristics of two novel, closely related receptors and the pharmacologic and physiochemical evidence for several additional, still incompletely characterized receptors for PTH and PTHrP.

PHYSIOLOGICAL ROLE OF PTH To ensure a multitude of essential cellular functions, the extracellular concentration of calcium (Ca2þo) is maintained within narrow limits [15,16]. In terrestrial

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vertebrates, calcium is necessary for adequate mineralization of the skeleton, which provides mechanical support and protection for internal organs and acts as levers for the various muscle groups involved in locomotion. Because of its high calcium content, 99% of the body’s supply, the skeleton also serves as the most important reservoir from which calcium can be rapidly mobilized. Because food intake and thus the nutritional supply of calcium are usually discontinuous, intestinal calcium absorption occurs only intermittently. Maintenance of a constant blood calcium concentration thus constitutes a major homeostatic challenge which, during evolution, led to the development of highly efficient mechanisms to increase intestinal calcium absorption, reduce urinary calcium losses, and facilitate, if necessary, rapid mobilization of calcium from the skeletal reservoir [16]. In contrast to these environmental challenges of most terrestrial vertebrates, marine animals, which are usually exposed to the high environmental calcium concentration of seawater (10 mM) had to adopt mechanisms by which extracellular calcium could be reduced [17,18]. Unlike the diet of terrestrial animals, seawater provides only a very limited supply of phosphate, and this environmental deficiency resulted in the development of mechanisms to conserve phosphate. It thus, appears plausible that the efficient intestinal absorption of phosphate and the impressive capacity of the mammalian kidney to retain phosphate [15,19] are remnants of earlier evolutionary adaptations to life in the low phosphate environment of the oceans (see Chapter 7 for discussion of phosphate homeostasis). PTH and the active form of vitamin D, 1,25-dihydroxyvitamin D3 (1,25(OH)2D3), are the principal physiologic regulators of calcium homeostasis in humans and all terrestrial vertebrates [11,20,21]. Synthesis and secretion of PTH are stimulated by any decrease in blood calcium, and conversely, secretion of the hormone is inhibited by an increase in blood calcium [22,23]. This rapid negative feedback regulation of PTH production, along with the biologic actions of the hormone on different target tissues, represents the most important homeostatic mechanism for minute-to-minute control of calcium concentration in the extracellular fluid (ECF) [24e26]. In contrast to the rapid actions of PTH, 1,25(OH)2D3 is of critical importance for long-term, day-to-day, and week-to-week calcium balance. The actions of both hormones are coordinated, and each influences the synthesis and secretion of the other. At least three distinct, but coordinated, actions of PTH increase the flow of calcium into the ECF and thus increase the concentration of blood calcium [27e29]. Through its rapid actions on the kidney and bone, which are all mediated through the PTH/PTHrP receptor and subsequent secondary messages in specific

and highly specialized cells, PTH increases the release of calcium from bone, reduces the renal clearance of calcium, and stimulates the production of 1,25(OH)2D3 by activating the gene encoding 25-hydroxyvitamin D-1a-hydroxylase (1a-hydroxylase) in the kidney. The relative importance of the first two actions of PTH on the rapid, minute-to-minute regulation of calcium is not definitively resolved, but most physiologists have stressed the importance of the effects of PTH on bone in maintaining hour-to-hour calcium homeostasis in the ECF. Several lines of evidence, such as that provided by calcium kinetic analysis, indicate a transfer between ECF and bone of as much as 500 mg calcium daily, which is equivalent to one-fourth to one-half the total ECF calcium content [15]. Besides regulating this transfer of calcium from bone through direct breakdown of bone tissue (mineral and matrix), PTH influences the rates of exchange of calcium adsorbed to the surface of bone; this exchangeable calcium pool can be stimulated to provide a rapid and substantial rate of entry of calcium into blood. In addition to these actions of PTH on bone, actions of PTH on the kidney may also be extremely important in the precise hourly regulation of ECF calcium. The third action of PTH on calcium homeostasis e namely, enhancement of intestinal calcium absorption e is indirect and involves the synthesis of 1,25(OH)2D3 from the biologically inactive precursor 25(OH)D3. It is difficult, however, to analyze quantitatively or to contrast proportionately the relative physiologic importance of the direct and indirect actions of PTH on the three principal target tissues: kidney, bone, and intestine. The complexity of bone as a tissue and the many detectable rates of exchange of calcium between the skeleton and the ECF have made the action of PTH on the skeleton difficult to analyze. The state of calcium in blood is complex; much of the calcium is present as chelates or is bound to plasma proteins. Because actual filtered loads depend on the ratio of free and bound forms of calcium, it is difficult to calculate renal calcium clearance accurately. The different PTH-dependent actions to promote calcium entry into the ECF are most clearly defined in conditions of deficiency or excess of PTH, such as during experiments in animals or during controlled observations in patients with disorders of parathyroid gland function. The experimental data in these extremes abundantly affirm the crucial calcium homeostatic role of PTH. However, because of continuous and rapid adjustments in mineral ion concentration, it can be difficult to observe the consequences of hormone action under normal physiologic conditions. For example, the rate of PTH secretion changes continually and rapidly so that the controlled variable, calcium, remains constant, and it may, therefore, be difficult experimentally to detect small corrective changes.

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Teleologically, the action of PTH on the regulation of blood phosphate concentration in terrestrial species is best understood as a secondary, rather than a homeostatic, action. Phosphate is abundant in the food chain in terrestrial existence. Phosphate deficiency, unlike calcium deficiency, in the absence of specific organ dysfunction is, therefore, an unlikely environmental challenge (see Chapter 7 for detailed review of the regulation of phosphate homeostasis). To correct a deficiency in calcium, calcium phosphate stores in bone can be rapidly dissolved; such activity results, however, in the simultaneous liberation of ionic calcium and phosphate. Because a high blood phosphate level tends to lower the calcium concentration through multiple mechanisms, the rise in blood calcium that occurs after bone dissolution (desirable homeostatically) is, therefore, beneficial only if the concomitant increase in blood phosphate concentration (undesirable) can be rapidly corrected. To maximize the control of calcium homeostasis, PTH thus has divergent actions on renal tubular handling of the two mineral ions: it increases the retention of calcium and, at the same time, diminishes reabsorption of phosphate. Through these mechanisms, namely, increased renal phosphate clearance to prevent hyperphosphatemia and increased tubular calcium reabsorption, PTH guarantees that an elevation in blood calcium results from the increased release of calcium from bone. The renal action of PTH on phosphate homeostasis is biologically predominant over the increased phosphate flux from bone. Consequently, parathyroidectomy (experimentally in animals) or renal resistance to PTH, as in patients with pseudohypoparathyroidism or renal failure, leads not only to hypocalcemia but also to an increase in blood phosphate and a marked reduction in urinary phosphate excretion. This finding demonstrates the importance of the PTHdependent action on phosphate homeostasis in the kidney, which becomes particularly important in disease states when high bone turnover is the result of dietary calcium deficiency or lack of biologically active vitamin D [30,31].

Chemistry The first extracts from bovine parathyroid glands were described in 1925, and the content of biologically active PTH was assessed by their hypercalcemic and phosphaturic properties [1,2]. However, it was not until 1959, when Aurbach [32] and Rasmussen and Craig [33] developed improved extraction procedures, that it became possible to isolate and purify sufficient quantities to determine the primary structure of bovine, porcine, and human PTH through the protein sequence determination methods [34e39]. Two groups independently determined the sequences of human and bovine

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hormones [34e39]. Shown in Figure 6.2 are the sequences of the bovine, porcine, and human hormones determined by one group [35,36,38,39]. Discordant sequences for the human PTH polypeptide, and in one position for the bovine hormone, published by another group [34,37] are not shown in Figure 6.2, since nucleotide sequence analysis of genomic and complementary DNA confirmed the amino acid sequences of the first group (the only exception was residue 76 in human PTH, which was determined to be glutamine instead of glutamic acid) [35,36,39]. Based on these amino acid sequences, the PTH(1e34) fragments of the different species were synthesized, and their biologic activities were compared in vitro and in vivo with those of highly purified intact PTH from the same species. Molecular cloning techniques then led to the deduction of the amino acid sequences of rat, chicken and dog PTHs [40e43], followed more recently by the identification of PTH molecules in other mammals and fish, discussed below. The synthetic peptides used in parathyroid hormone research today are based largely on the (1e34) regions of the mammalian hormone sequences shown in Figure 6.2 [44,45]. Extensive sequence homology is present in the mammalian PTH species; these molecules consist of a single-chain polypeptide with 84 amino acids and a molecular weight of approximately 9400 Daltons (that of human PTH[1e84] is 9425 Daltons). The N-terminal region of PTH, which is necessary and sufficient for the regulation of mineral ion homeostasis, shows high sequence conservation among all the vertebrate species (see Fig. 6.2). The middle portions of the different molecules exhibit the most structural variation, which could suggest that this region of PTH is only of limited functional importance. The non-mammalian PTH homologs of chicken [41,42] and fish species Danio rerio (zebrafish) [46] and Takifugu ruberipes (puffer fish) [47,48] diverge considerably from the mammalian hormones C-terminal of amino acid residue His32. Interestingly, both fish species have two distinct genes encoding two separate PTH molecules, called PTH1 and PTH2 [46,47]. The zebrafish peptides are considerably shorter than mammalian PTH (67 and 68 residues), while fugu PTH1 is predicted to be 81 residues in length and fugu PTH2 is predicted to be 63 residues [47]. After the original work establishing that the first 34 amino acids of mammalian PTH were sufficient to produce a fully active synthetic peptide [44,45], much work has centered on defining the minimum pharmacophore essential for biologic activity. We describe below how sites of ligand interaction with the PTH/PTHrP receptor were defined by performing assays with products of various combinations of shortened and modified PTH ligands and mutagenized receptors [49]. As also discussed in the section below on hormone/receptor

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FIGURE 6.2 Sequence relationships of PTH family ligands. Comparisons of amino acid sequences of PTH and PTHrP peptides from different

species are shown in panels A and B, respectively. Note numbering indicates alignment position number, and not protein amino-acid sequence number. Comparison of TIP39 and a PTH-like peptide (PTH-L) from the pufferfish (Takifugu rubripes) with human PTH and PTHrP (the [1e84] region only) is shown in panel C. Amino acid sequences were aligned using the ClustalW software program (version 2.012; gap penalties: opening, 15; extending, 5) and further processed using the Boxshade program. Amino acid identities of 50% or more are shown in white type on black field, and similarities are shown in black type on gray field. Sequence identification or accession numbers are shown in the legend to Figure 6.4.

interactions, it has been determined that substitutions of non-naturally occurring amino acids (e.g. alpha-aminoisobutyric acid at positions 1 and 3 in the primary ligand structure) favoring formation of an alpha-helix, even in short peptides, such as PTH(1e14), produce peptides that are highly potent when tested in vitro using cellbased assays and have highly stabilized helical structure in solution (Fig. 6.3) [50e54]. The in vitro activity of the native PTH(1e14), which is quite weak, is improved about 100 000-fold by the modifications indicated in Figure 6.3. Such short-length PTH peptides have also been shown to be active in vivo, since some cause hypercalcemia and are anabolic on bone, although their potency is much less than that of PTH(1e34) due to a more rapid clearance [55]. Replacement of valine-2 in these peptides with bulky amino acids, such as tryptophan or parabenzoyl-L-phenylalanine

(Bpa), results in competitive antagonist peptides defective for AC/cAMP and PLC/IP3/Ca2þ signaling, thus confirming the critical role that this conserved valine plays in receptor activation [56]. Other longer-length PTH or PTHrP analogs having residue-1 (serine or alanine) replaced by glycine [57], Bpa [58] or tryptophan [59] exhibit signal selective properties in that they efficiently stimulate the cAMP cellular pathway but not the inositol triphosphate/intracellular Ca2þ pathway. Because of their generally demonstrated similar potencies at the PTH/PTHrP receptor, it seemed likely that both PTH(1e34) and PTHrP(1e34) ligands would adopt very similar conformations when part of the active hormone-receptor complex. However, recent data, discussed below, suggest that each ligand selectively binds to or induces a distinct receptor confirmation. Ideally, each hormone should be co-crystallized

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FIGURE 6.3 Minimized N-terminal PTH analogs. Shown is the native (rat) PTH(1e14) sequence and the locations of activity-enhancing

substitutions. The six substitutions shown above the sequence, when combined, enhance activity by as much as 100 000-fold; the Bpa2 substitution shown below confers antagonist properties to the peptide. The (1e9) region (non-shaded circles) comprises the minimum-length agonist pharmacophore. Non-encoded amino acids include a-amino-isobutyric acid (Aib), homoarginine (Har), and para-benzoyl-L-phenylalanine (Bpa).

with the PTH/PTHrP receptor to permit analysis by x-ray diffraction of those intermolecular interactions that are characteristic of the biologically active hormonereceptor complexes. G protein-coupled receptors, such as the PTH/PTHrP receptor, have multiple membraneembedded domains and are likely to have complex three-dimensional structures. Interaction with either PTH or PTHrP appears to involve several distinct receptor domains (see later discussion) that may undergo significant conformational changes after ligand binding has occurred, which makes it even more challenging to conduct x-ray or multidimensional NMR analyses. Recent advances, however, have made it possible to co-crystallize the extracellular portion of the PTH/PTHrP receptor with either the carboxyl terminal half of PTH(1e34) or that of PTHrP(1e34) [60,61].

rubripes and Tetraodon fluviatilis, reveal the duplication of the PTH gene in each case [47,65]. Both the PTH1 and PTH2 peptides derived from the zebrafish activate the PTH/PTHrP receptors from different species [48,65] and, indeed, a fugu PTH(1e34) peptide has been shown to induce bone anabolic effects in osteopenic ovariectomized rats [66]. In addition to PTH, the teleost fish also express PTHrP, again encoded by duplicate genes [65,67e70]. Furthermore, PTHrP immunoreactivity has been detected in the cartilaginous sharks and rays [71], and in a more primitive agnathan, the lamprey [72]. The

Evolution To maintain extracellular calcium and phosphate concentrations within narrow limits, the intricate regulatory system outlined above, in which PTH plays the most important role, developed in the terrestrial animals. In mammals, PTH is produced almost exclusively by the parathyroid glands (only small amounts of its messenger RNA [mRNA] have been detected elsewhere [62,63]). During evolution, these glands first appear as discrete organs in amphibians e that is, with the migration of vertebrates from an aquatic to a terrestrial existence e and their appearance most likely represents an evolutionary adaptation to an environment that is, by comparison to seawater, low in calcium [17,18,64]. Parathyroid glands have not been identified in fish or invertebrate species. However, with the rapid advances in characterization of complete genomes of multiple species we have definitive proof of the earlier evolutionary origin of both PTH and PTHrP (Fig. 6.4). Gene analyses of several teleost fish species, including the zebrafish, Danio rerio, and the puffer fishes Takifugu

FIGURE 6.4 Phylogenetic relationships of PTH family ligands. The diagram shows the separate groupings of PTH and PTHrP ligands, with duplications of each ligand in pufferfish (Takifugu ruberipes) and zebra fish (Danio rerio), as well as the relationship to an apparent ancestral, PTH-like ligand (PTH-L) in fugu. Protein data base accession numbers are as follows: human PTH, P01270; chicken PTH, A9YX65; danio PTH-1, Q6WQ25; danio PTH-2, Q6WQ24; fugu PTH-1, Q2PCS7; fugu PTH-2, Q6W9J4; human PTHrP, P12272; chicken PTHrP, Q5TLZ2; danio PTHrP-A, Q1L5E7; fugu PTHrP-A, Q9I8E9; danio PTHrP-B, Q4VVA3; fugu PTHrP-B, Q2PCS8; fugu PTH-L, Q2PCS5.

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teleost PTHrPs contain some amino acid residues characteristic of mammalian PTH; for example, fish PTHrP contains Met at position 8, Trp at position 23, and Leu at position 28, which are amino acid residues found in mammalian PTH. However, there is only one amino acid residue, Gln(Q)25, in fish PTH that is found in some mammalian PTHrP species, but not in mammalian PTH (see Fig. 6.4). This pattern suggests that the fish proteins may be phylogenetically closer to a common PTH/PTHrP precursor than are the mammalian proteins (see below). Indeed, in addition to duplicate copies of PTH and PTHrP genes, the puffer fish genome contains a fifth gene that encodes a protein containing the amino acid residues characteristic of both PTH and PTHrP. This gene, called PTH-L, is phylogenetically an intermediary to PTH and PTHrP, and may thus represent first definitive evidence for an ancestral gene from which the two divergent ligand forms evolved (see Fig. 6.4) [47,65].

The PTH Gene The human PTH gene consists of three exons located on chromosome 11p15 [73e76]. The first exon is 85 nucleotides in length and is non-coding (Fig. 6.5). Exon 2 (90 bp) encodes most amino acids of the prepropeptide sequence, whereas the third exon (612 bp) encodes the remainder of the propeptide sequence and all amino acids of the mature peptide, and it constitutes the 30 non-coding region [77]. Several frequent intragenic polymorphisms (TaqI and PstI [78], BstBI [79], DraIII [80], XmnI [81], and a tetranucleotide repeat ([AAAT]n [82]) have been identified in the human PTH gene, and some were shown to be informative in genetic linkage studies [83e85]. Two mRNAs that are 822 and 793 bp in length are derived in the human gene from the two transcriptional start sites which follow two different functional TATA boxes that are separated by 29 bp [77]. Two closely spaced TATA boxes and two distinct transcripts are also derived from the bovine PTH gene,

FIGURE 6.5 Schematic of the PTH gene. Shown is the gene along several thousand base pairs (approximate length shown by the scale marker for 500 bp). The three exons in the mRNA are represented as numbered rectangles. Control elements are identified in the 50 noncoding region (50 NC). A region responsive to vitamin D is within a few hundred base pairs of exon 1; far upstream are silencers involved in calcium regulation.

while rat and chicken PTH genes give rise to only one transcript; as a consequence of a long 30 non-coding region, the transcript from the chicken PTH gene is unusually long, and comprises 2.3 kilobases [24,86]. The genes encoding zebrafish PTH1 and PTH2 have a similar overall organization as the mammalian PTH genes [46].

Cellular Biosynthesis and Hormonal Processing During the synthesis of the preproPTH molecule, the signal sequence, which comprises the 25-amino acid containing “pre”-sequence, is cleaved off after entry of the nascent peptide chain into the intracisternal space bounded by the endoplasmic reticulum. A heterozygous mutation in this leader sequence, which changes a cysteine to an arginine at position 8 and thus impairs processing of preproPTH to proPTH, has been identified as the most plausible molecular cause of an autosomal dominant familial form of hypoparathyroidism [87,88]. The mutant hormone was found to be trapped intracellularly, predominantly in the endoplasmic reticulum (ER), leading to a marked upregulation of ER stressresponsive proteins (BiP and PERK) and the proapoptotic transcription factor CHOP, indicating that apoptosis-mediated parathyroid cell death is the likely cause of the observed hypoparathyroidism [89]. Subsequent to the removal of the pre-sequence, the pro-peptide is transported to trans-Golgi network where the pro-sequence (amino acid residues 6 through 1) is removed [90]. This latter process may involve furin (paired basic amino acid cleaving enzyme) and/or proprotein convertase-7 (PC-7), which are both expressed in parathyroid tissue; their expression levels do not appear to be regulated by either calcium or 1,25(OH)2D3 [91,92]. After removal of the basic pro-sequence, the mature polypeptide, PTH(1e84), is packaged into secretory granules. Two proteases, cathepsins B and H, are subsequently involved in the intraglandular generation of carboxyl-terminal PTH fragments from the intact hormone; no amino-terminal PTH fragments appear to be released from the gland [93e95]. Since small or intermediate size carboxyl-terminal fragments of PTH are unlikely to be involved in the regulation of calcium homeostasis, the intraglandular degradation of intact PTH is thought to represent an inactivating pathway, at least with regard to the regulation of mineral ion homeostasis. Consistent with this conclusion, hypercalcemia results in a substantial decrease in PTH secretion and, furthermore, favors the secretion of carboxylterminal PTH fragments, including a previously undetected large molecular species that is truncated at the amino-terminus (see section below) [95e98]. However, recent studies have shown that some amino-terminally truncated PTH fragments, such as PTH(7e84), have

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hypocalcemic properties in vivo and can furthermore reduce the formation of osteoclasts in vitro [99]. The pool of stored, intracellular PTH is small and the parathyroid cell must therefore have mechanisms to increase hormone synthesis and release in response to sustained hypocalcemia. One such adaptive mechanism is to reduce the intracellular degradation of the hormone, thereby increasing the net amount of intact, biologically active PTH that is available for secretion. During hypocalcemia, the bulk of the hormone that is released from the parathyroid cell is intact PTH(1e84) [93e95,97,98]. As the level of Ca2þo increases, a greater fraction of intracellular PTH is degraded, and with overt hypercalcemia, most of the secreted immunoreactive PTH consists of biologically inactive C-terminal fragments [10,24,25].

stimulation of intestinal calcium absorption and bone resorption [105e107] and may thus be attractive candidates for treating the hyperparathyroidism of chronic renal insufficiency. Adjustment of the rate of parathyroid cellular proliferation is the third adaptive mechanism contributing to changes in the overall secretory activity of the parathyroid gland. Under normal conditions, parathyroid cells have little or no proliferative activity. The parathyroid glands, however, can enlarge greatly during states of chronic hypocalcemia, particularly in the setting of renal failure, probably because of a combination of hypocalcemia, hyperphosphatemia, and low levels of 1,25(OH)2D3 in the latter condition.

Regulation of PTH Gene Expression

A large number of factors modulate PTH secretion in vitro [11,25,108] but most of these factors are not thought to control hormonal secretion in vivo in a biologically relevant manner. Therefore, we focus in this section principally on factors that are the most physiologically meaningful regulators of PTH secretion e that is, the extracellular ionized calcium concentration itself (Ca2þo), 1,25(OH)2D3, and the level of extracellular phosphate ions. Of these three, Ca2þo is most important in the minute-to-minute control of PTH secretion. Indeed, the actions of 1,25(OH)2D3 and phosphate ions on the secretion of PTH probably result, at least in part, from their effects on hormonal biosynthesis rather than secretion per se [11,25,108]. Ca2þo also modulates several other aspects of parathyroid function that indirectly affect PTH secretion, including PTH gene expression, the hormone’s intracellular degradation, and parathyroid cellular proliferation, as described previously. Recent data have shown that novel factors playing key roles in phosphate homeostasis, especially fibroblast growth factor 23 (FGF-23) and aeKlotho (a co-receptor for FGF receptors), also modulate parathyroid function, inhibiting [109,110] and enhancing [111] parathyroid function, respectively. Our rapidly improving understanding of how these factors participate in phosphate homeostasis is described in detail in Chapter 7. The relationship between PTH and Ca2þo is represented by a steep inverse sigmoidal curve that can be quantitatively described [112e114]. Parathyroid cells can readily detect reductions in Ca2þo of a few percentage points [113] and the percent coefficient of variation in Ca2þo in humans is less than 2% [115]. The set point of the parathyroid gland is the key determinant of the level at which Ca2þo is “set” in vivo [116]. Thus, the parathyroid cell is normally more than half-maximally suppressed at normal levels of Ca2þo and has a large secretory reserve for responding to hypocalcemic stress. Nevertheless, PTH levels in vivo fall dramatically

Another adaptive mechanism of the parathyroid cell to sustained reductions in Ca2þo is to increase cellular levels of PTH mRNA, a response that takes several hours. A reduction in Ca2þo increases, whereas an elevation in Ca2þo reduces the cellular levels of PTH mRNA by affecting both its stability and the transcriptional rate of its gene [11,25,100,101]. Available data suggest that phosphate ions also regulate, directly or indirectly, PTH gene expression. Hypophosphatemia and hyperphosphatemia in the rat, respectively, lower and raise the levels of mRNA for PTH through a mechanism that is independent of changes in Ca2þo or 1,25(OH)2D3. An elevated extracellular phosphate concentration could, thus, contribute importantly to the secondary hyperparathyroidism frequently encountered in patients with end-stage renal failure, who often have chronically elevated serum phosphate concentrations. Metabolites of vitamin D, principally 1,25(OH)2D3, also play an important role in the long-term regulation of parathyroid function and may act at several levels: by affecting the secretion of PTH and regulation of its gene, by regulating transcriptional activity of the genes encoding the calcium-sensing receptor (CaSR) (see below) and the vitamin D receptor (VDR), as well as by regulating parathyroid cellular proliferation [11,25,100,102]. 1,25(OH)2D3 is by far the most important vitamin D metabolite that modulates parathyroid function. It acts through a nuclear receptor, the VDR, often in concert with other such receptors (i.e. those for retinoic acid or glucocorticoids), on DNA sequences upstream from the PTH gene [103,104]. 1,25(OH)2D3induced upregulation of VDR and CaSR expression in the parathyroid could potentiate its inhibitory action(s) on PTH synthesis and secretion [11,25,100]. Non-calcemic or less calcemic analogues of 1,25(OH)2D3 inhibit PTH secretion while producing relatively little

Regulation of Secretion

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(e.g. by 80%) when Ca2þo rises to frankly hypercalcemic levels [112,113], which is thought to contribute importantly to the mineral ion homeostatic system’s defense against hypercalcemia [116]. Furthermore, elevating Ca2þo also decreases the proportion of secreted, intact PTH because of increased intraglandular degradation to inactive fragments (see the earlier section entitled PTH Biosynthesis and Intraglandular Processing and the later section entitled Metabolism of PTH) [96,117]. Even with severe hypercalcemia, however, some residual release of intact PTH(1e84) still occurs in vivo [28,98,118]. This non-suppressible basal component of PTH release may contribute to the hypercalcemia caused by hyperparathyroidism when the mass of abnormal parathyroid tissue is very great (e.g. in patients with renal failure) [114,119e121]. The parathyroid cell has a temporal hierarchy of responses to low Ca2þo that permits it to secrete progressively larger amounts of hormone during prolonged hypocalcemia [11,25,108]. To meet acute hypocalcemic challenges, PTH is released within seconds from preformed secretory vesicles by exocytosis as dictated by the sigmoidal curve. Sufficient PTH is stored in the parathyroid chief cell to sustain maximal, low Ca2þo-stimulated PTH release for about 60e90 minutes [116]. Another rapid response of the parathyroid cell to hypocalcemia that enhances its net synthetic rate of PTH is reduced intracellular hormonal degradation e the opposite of what occurs at high levels of Ca2þo e which occurs within minutes to an hour [96,117]. Hypocalcemia persisting for hours to days elicits increased PTH gene expression, whereas that lasting for days to weeks or longer stimulates parathyroid cellular proliferation [11,25,108,122]. A greater secretory capacity for PTH on a per-cell basis (e.g. as a result of enhanced PTH gene expression) increases maximal hormonal secretion in vivo, as does an increase in cell number as a result of parathyroid cellular proliferation. In severe secondary hyperparathyroidism, very large increases in parathyroid cellular mass can elevate circulating PTH levels by 100-fold or more. The molecular mechanism underlying Ca2þo-regulated PTH secretion involves a G protein-coupled, cell surface Ca2þo-sensing receptor (CaSR) [123]. The CaSR was first isolated from bovine parathyroid glands [124] and subsequently from human parathyroid and several other tissues and species [125]. The receptor exhibits the characteristic “serpentine” motif (seven membranespanning domains) of the superfamily of G proteincoupled receptors. Its long, N-terminal extracellular domain contains the major, but not all, determinants of Ca2þo binding [126e128]. Changes in Ca2þo modulate a number of second messenger systems via coupling of CaSR through its intracellular domains to the relevant G proteins regulating these signaling pathways [123].

These functions include activation of phospholipases C, A2, and D [129], stimulation of several mitogen-activated protein kinases [130] and inhibition of adenylyl cyclase [131]. Despite numerous studies conducted over the past 25 years, a full understanding of the major second messenger pathways through which changes in Ca2þo, acting via the CaSR, regulate various aspects of the function of parathyroid and other CaSR-expressing cells remains elusive. Recent evidence, however, indicates key roles for the G proteins, Gq and G11, but their downstream transduction pathways participating in the control of parathyroid function are not established [132]. In the parathyroid, the CaSR mediates the inhibitory actions of Ca2þo on PTH secretion and gene expression as well as parathyroid cellular proliferation [133,134]. The CaSR is also expressed in several additional tissues involved in systemic mineral ion homeostasis, including the calcitonin-secreting C-cells of the thyroid [135], diverse cells within the kidney [136], bone cells and/or their precursors [100], and intestinal epithelial cells. In the kidney, the CaSR in the cortical thick ascending limb of the nephron mediates direct, high Ca2þo-induced inhibition of the tubular reabsorption of Ca2þ and Mg2þ [136,137]. Therefore, raising Ca2þo both directly inhibits renal tubular reabsorption of Ca2þ via actions on the CaSR expressed in nephron segments involved in hormonal regulation of Ca2þ reabsorption (e.g. by PTH) and indirectly inhibits it by reducing PTH secretion (see Renal Calcium Reabsorption below). The identification of hypercalcemic (e.g. familial hypocalciuric hypercalcemia and neonatal severe hyperparathyroidism) [138] and hypocalcemic (i.e. autosomaldominant hypocalcemia) [139] disorders caused by inactivating and activating mutations of the CaSR, respectively, illustrates the receptor’s central, nonredundant role in setting the serum calcium concentration [140,141]. Targeted disruption of the CaSR gene has also enabled the generation of mouse models of familial hypocalciuric hypercalcemia and neonatal severe hyperparathyroidism via inactivation of one or both alleles of the CaSR [142], further supporting its importance in Ca2þo homeostasis. In addition to directly inhibiting PTH gene expression, 1,25(OH)2D3 also reduces PTH secretion [143,144] (for review [11,25,108]). It is not known whether this latter action is solely secondary to the effect of 1,25(OH)2D3 on biosynthesis of the hormone and/or represents a direct action on the secretory process per se. Increasing the ambient level of phosphate in vitro, independent of concomitant changes in Ca2þo, enhances parathyroid cellular proliferation, PTH gene expression and, ultimately, hormonal secretion [145e147]. Phosphate-induced changes in PTH secretion, however, take several hours and must result secondarily from

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changes in hormonal biosynthesis rather than secretion per se [147]. Finally, Mg2þo clearly functions as a CaSR agonist in vitro when tested in cells containing an endogenous CaSR [148] (e.g. parathyroid cells) or expressing the cloned CaSR [124], although it is twofold to threefold less potent than Ca2þo on a molar basis. Because levels of serum ionized Mg2þo are lower than those of Ca2þo, it is presently unclear whether Mg2þo acts as a physiologically relevant CaSR agonist at the parathyroid gland in vivo under normal circumstances. Patients with inactivating or activating CaSR mutations, however, can exhibit mild hypermagnesemia or hypomagnesemia [140], respectively, thus suggesting that the CaSR does contribute to setting Mg2þo in vivo, as previously suggested [149]. It may do so, at least in part, in the kidney, where Mg2þo in the tubular fluid of the thick ascending limb exceeds that in blood and may be sufficient to activate the CaSR that regulates tubular reabsorption of Ca2þo and Mg2þo in this nephron segment [123,137,150,151]. In addition to the inhibitory effect of elevated Mg2þo on PTH secretion, low concentrations of Mg2þo e as in patients with overt magnesium deficiency e also reduce PTH secretion [152]. The mechanism(s) underlying this effect of hypomagnesemia has recently been suggested to involve increased activity of G proteins to which the CaSR normally couples, probably Gi and Gq/11, thereby leading to increased intracellular signalling and inhibition of PTH secretion [153].

Metabolism of PTH Studies performed over more than three decades by several laboratories have focused on the heterogeneity of circulating forms of PTH, which was first identified by Berson and Yalow in 1968 [154]. From these investigations, it is now apparent that, in addition to the fulllength polypeptide PTH(1e84), which is the biologically active hormone, much of the circulating hormone lacks an intact N-terminus and most of these fragments are, thus, devoid of biologic activity, at least with regard to the PTH/PTHrP receptor-mediated regulation of mineral ion homeostasis [24]. C-terminal PTH fragments are produced in and released from the parathyroid gland, but they are also derived from circulating intact hormone by efficient, high-capacity degradative systems in the liver and kidney and most likely at other peripheral sites (Fig. 6.6). Some PTH fragments, such as PTH(7e84), appear to be generated within the gland and to have some biological activity, perhaps as inhibitors [155e157]. Direct measurement of arterial and venous differences in parathyroid effluent blood (with vigorous conditions to prevent any ex vivo cleavage of hormone after sample collection) were performed in cattle and confirmed that smaller C-terminal fragments and intact

117

FIGURE 6.6 Scheme of peripheral PTH cleavage and the interaction of PTH with the PTH/PTH-related peptide receptor (PTH1R) and with other putative receptors on target cells. Within parathyroid cells, secretory vesicles (circles) and in peripheral organs various patterns of cleavages of PTH(1e84) occur, resulting in multiple circulating fragments, mostly C-terminal ones (solid bars); but not N-terminal ones (suggesting rapid degradation). The carboxyl end of intact PTH and possibly some C fragments interact with a putative C-terminal receptor and, as well, inhibit actions of PTH on the PTH1R. Cyclic adenosine monophosphate, cAMP.

hormone, but not N-terminal fragments, are secreted into the circulation. The relative concentration of these C-terminal fragments released from the gland increases under conditions of systemic hypercalcemia, when overall secretion rates of intact hormone are lower [95,158]. The C-terminal PTH fragments are similar to those generated by peripheral metabolism (see below) but were not chemically characterized. The peripheral metabolism of PTH has been analyzed by injecting intact hormone into the circulation of test animals. Such experiments have not been performed in human subjects, but it is assumed that the similar metabolism of PTH in rats, dogs, and cows is reflective of its metabolism in humans [159]. Clearance of intact PTH from plasma was found to be very rapid (halflife, 2e4 minutes) [160,161]; the major sites of clearance being the liver and kidney. Clearance by the liver predominates over clearance by the kidney; the two organs together account for virtually all clearance of intact hormone. Hepatic clearance of intact hormone has been estimated to be 40e75% and renal clearance, 20e30% [24]. In summary, current evidence indicates that intact PTH and multiple C-terminal fragments (which are derived from glandular and peripheral cleavage and may not be identical) are the principal circulating hormonal forms. In addition, recent data suggest the presence of a previously unrecognized, large,

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N-terminally truncated form of the hormone. Biologically active, N-terminal fragments of PTH, if found in the circulation at all, are likely to circulate only at extremely low concentrations (80% of the Pi filtered load is reabsorbed [101,102]. The proximal tubule is the major site of Pi reabsorption, with approximately 70% of the filtered load reclaimed in the proximal convoluted tubule and approximately 10% in the proximal straight tubule (Fig. 7.1). In addition, a small but variable portion (

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Median

FIGURE 9.17 Influence of protein intake on the impact of increased physical activity on BMC, aBMD and AREA or width of femoral neck (FN) in prepubertal boys. Data, as expressed in Z-scores, are from 237 healthy boys, with mean age 7.4 years. They are distributed in four subgroups according to the median of both physical activity and protein intake. Increased physical activity from 168 40 to 303 54 (SD) kcal/ day was not associated with a significant increment in Z-score of FN variables, when protein intake was below the median (38.0 6.9 g/day). In contrast, increased physical activity of a similar magnitude, i.e. from 167 33 to 324 80 kcal/day was associated with a marked increment in FN variables, when protein intake was above the median (56.2 9.5 g/day). In this situation of relatively high protein intake, the greater FN Zscore associated with more sustained physical activity was þ0.66 and þ0.59 for BMC and AREA. The morphometric difference reflected a larger width of the femoral neck, since for all DXA scans the height of the region of interest parallel to the femoral axis was constant. (Adapted data from Chevalley et al. [262]. See text for further details.)

In “well-nourished” children and adolescents, the question arises whether or not variations in the protein intake within the “normal” range can influence skeletal growth and thereby modulate the influence of genetic determinants on peak bone mass attainment [6]. In the relationship between protein intake and bone mass gain, it is not surprising to find a positive correlation between these two variables [242,249,250]. Similar to calcium intake, the association appears to be particularly significant in prepubertal children [242]. These observations suggest that relatively high protein intakes could favor bone mass accrual during childhood. Positive relationships were reported between increased calcium intake from dairy foods and bone mineral acquisition during childhood and adolescence [234,251e254]. The associated increase in protein intake from dairy foods may have substantially contributed to such positive relationships. Likewise, in the often-quoted study carried out in two Yugoslav populations, the difference in young adult bone mass has usually been ascribed to calcium intake [255]. However, both protein intake and physical activity were also associated with peak bone mass [255]. The reasons why protein intake was not considered early on as an essential nutrient for calcium economy and bone health appears to stem from longterm prejudice or “belief sytems in the conduct of nutritional science” as thoughtfully analyzed by Heaney [256]. Claims against dietary proteins, particularly against those from animal food sources were mainly

based on a questionable dietary acid load hypothesis which has been refuted by several analyses or metaanalyses of experimental data [257e261]. In order to determine the quantitative relationship between protein intake and bone mass acquisition during childhood and adolescence, interventional studies testing different levels of protein intakes in otherwise isocaloric diets remain to be conducted. Furthermore, calcium requirement for optimal bone mass accrual could vary according to the level of protein intake. Thus, the possible positive interaction between protein and calcium intake deserves to be investigated in the perspective of increasing peak bone mass by modifying bone trophic nutrients. Interaction of Protein Intake and Physical Activity As already underscored, growing bones are usually more responsive to mechanical loading than adult bones. Increased physical activity was shown to increase bone mineral mass accumulation in children and adolescents. The positive impact of increased physical activity on bone acquisition might be greater before than during or after the period of pubertal maturation [6], although this pubertal maturation modulation may depend upon the skeletal site (axial vs appendicular) and/or structural (cortical vs trabecular) components examined [166]. Adequate nutritional supply can be expected to sustain the anabolic effect of mechanical loading on bone tissue as it does on skeletal muscle development.

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CONCLUSIONS

Among nutrients, high calcium intake was shown to enhance the response to physical activity in healthy children aged 3e5 years [221]. In healthy children and adolescents aged 6e18 years, long-term protein consumption exerts a stronger impact than calcium intake on bone mass and strength acquisition [249]. That high protein intake may enhance the bone response to increased physical activity has been recently reported in 8-year-old prepubertal boys [262]. At the femoral neck level, the increased aBMD and BMC was associated with a wider external perimeter (Fig. 9.17) [262]. Such an effect on the macroarchitecture of the femoral neck would be compatible with a greater resistance to mechanical load [166,263]. Potential of Increased Protein Intake in the Management of a Genetic Bone Disorder An interesting observation has been reported regarding the effect of a specific nutritional intervention to correct a defect in skeletal development. The CoffineLowry syndrome is a rare X-linked disorder in which males show severe mental retardation associated with several skeletal defects including short stature, kyphosis and/or scoliosis [264]. The skeletal manifestations worsen over time. The genetic defect is caused by loss-of-function mutations in a gene (RPS6KA3) coding for a growth factor-regulated protein kinase hRSK2 [265]. One mechanism whereby RSK2 favors skeletal development and bone formation appears to be by phosphorylating ATF4. This transcriptional factor itself regulates osteoblast differentiation during development and favors bone formation postnatally [266]. ATF4 exerts a stimulatory effect on amino acid import in eukaryotic cells [267]. Likewise, in osteoblasts, this transcription factor stimulates the amino acid import and regulates the synthesis of type I collagen, the main constituent of the bone matrix [266]. ATF4 deficiency in mice results in delayed bone formation during embryonic development and low bone mass throughout postnatal life [266]. Interestingly, a high protein diet in ATF4 deficient mice normalizes osteoblast differentiation and collagen synthesis in bone [268]. Furthermore, both bone formation and bone mass are enhanced. These observations suggest that the severe expression of genetic defect could be alleviated by simple dietary manipulation [269]. Dietary Protein Recommendations According to several national and international agencies, the daily recommended protein allowance for children and adolescents declines slightly from age 5 (1.05 g/kg b.w.) to 17 years (0.80 g/kg b.w.) [270e273]. In sharp contrast to a recommended amount of approximately 0.9e1.0 kg/b.w. per day for children aged 7e12 years, surveys made from several countries

in Europe, as well as in North America and Australia have recorded much higher daily protein intake of about 1.9e2.0 g/kg b.w. [24,212,249,250,262,273,274]. It has been underscored that the literature is essentially void of studies specifically directed at quantifying protein needs of healthy children between 7 and 12 years of age [275]. Current recommendations for protein intake in children remain speculative, since they correspond to estimates derived from interpolation of very shortterm nitrogen balance studies carried out in infants and young adults [272,275,276]. As described above in prepubertal boys aged 7.4 years (see Fig. 9.17), increased physical activity from about 170 to 315 Kcal/day in the presence of a spontaneous protein intake of z2.0 instead of 1.5 g/kg b.w. per day was associated with higher mean BMC at six skeletal sites of þ 0.64 vs þ 0.15 Z-score. This difference of z0.5 Z-score could well be expressed by a substantial shift of the BMC trajectory, resulting in a higher PBM and possibly in a reduction of fragility fractures in adulthood [15]. Likewise, such an upward shift in bone mass acquisition could also reduce the incidence of fractures during growth [46,147].

CONCLUSIONS Bone mass and strength achieved at the end of the growth period, simply designated as “peak bone mass (PBM)”, play an essential role in the risk of osteoporotic fractures occurring in adulthood. It is considered that an increase of PBM by 1 standard deviation would reduce the fracture risk by 50%. As estimated from twin studies, genetics is the major determinant of PBM, accounting for about 60e80% of its variance. Numerous polymorphisms of “candidate” genes have been found to be associated with the areal bone mineral density (aBMD), so far the most convenient measurable surrogate of bone mass and strength. The studied genes code for molecules implicated in bone function and structure such as circulating endocrine factors, hormone receptors, local regulators of bone modeling and remodeling or matrix molecules. None of these genes appears to account for more than a few percent of PBM variance. Before puberty, there is no substantial gender difference in aBMD when adjusted for age, nutritional factors and physical activity. During pubertal maturation, the size of the bone increases whereas the volumetric bone mineral density remains constant in both genders. At the end of puberty, the sex difference is essentially due to a greater bone size in male than female subjects. This is achieved by larger periosteal deposition in boys, thus conferring at PBM a better resistance to mechanical forces in men than in women. Sex hormones and the IGF-1 system are implicated in the bone sexual dimorphism occurring

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9. PEAK BONE MASS AND ITS REGULATION

during pubertal maturation. The genetically determined trajectory of bone mass development can be modulated to a certain extent by modifiable environmental factors. Interventions aimed at increasing the physical activity of children have been shown to exert a positive impact on cortical bones of the weight-bearing appendicular skeleton. Likewise, interventional studies using either calcium salt or milk supplementation have documented an increased bone mass gain, particularly at weightbearing cortical bones. Some studies suggest a positive interaction between calcium supplementation and increased physical activity on bone development. In healthy prepubertal boys, the impact of increased physical activity affecting bone acquisition is enhanced by protein intake within limits above the usual recommended allowance. Prepuberty appears to be an opportune time to modify environmental factors that impinge on bone mineral mass acquisition and eventually peak bone mass and strength.

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[12] Duan Y, Wang XF, Evans A, Seeman E. Structural and biomechanical basis of racial and sex differences in vertebral fragility in Chinese and Caucasians. Bone 2005;36:987e98. [13] Cummings SR, Black DM, Nevitt MC, et al. Bone density at various sites for prediction of hip fractures. The Study of Osteoporotic Fractures Research Group. Lancet 1993;341:72e5. [14] Melton 3rd LJ, Atkinson EJ, O’Fallon WM, Wahner HW, Riggs BL. Long-term fracture prediction by bone mineral assessed at different skeletal sites. J Bone Miner Res 1993;8:1227e33. [15] Marshall D, Johnell O, Wedel H. Meta-analysis of how well measures of bone mineral density predict occurrence of osteoporotic fractures. Br Med J 1996;312:1254e9. [16] Adams JE, Shaw N. A Practical Guide to Bone Densitometry in Children. National Osteoporosis Society, Camerton, Bath, UK; 2004. [17] Baim S, Leonard MB, Bianchi ML, et al. Official Positions of the International Society for Clinical Densitometry and executive summary of the 2007 ISCD Pediatric Position Development Conference. J Clin Densitom 2008;11:6e21. [18] Boutroy S, Bouxsein ML, Munoz F, Delmas PD. In vivo assessment of trabecular bone microarchitecture by high-resolution peripheral quantitative computed tomography. J Clin Endocrinol Metab 2005;90:6508e15. [19] Sornay-Rendu E, Boutroy S, Munoz F, Delmas PD. Alterations of cortical and trabecular architecture are associated with fractures in postmenopausal women, partially independent of decreased BMD measured by DXA: the OFELY study. J Bone Miner Res 2007;22:425e33. [20] Boutroy S, Van Rietbergen B, Sornay-Rendu E, Munoz F, Bouxsein ML, Delmas PD. Finite element analysis based on in vivo HR-pQCT images of the distal radius is associated with wrist fracture in postmenopausal women. J Bone Miner Res 2008;23:392e9. [21] Chevalley T, Bonjour JP, Ferrari S, Rizzoli R. Influence of age at menarche on forearm bone microstructure in healthy young women. J Clin Endocrinol Metab 2008;93:2594e601. [22] Chevalley T, Bonjour JP, Ferrari S, Rizzoli R. Deleterious effect of late menarche on distal tibia microstructure in healthy 20-yearold and premenopausal middle-aged women. J Bone Miner Res 2009;24:144e52. [23] Kirmani S, Christen D, van Lenthe GH, et al. Bone structure at the distal radius during adolescent growth. J Bone Miner Res 2009;24:1033e42. [24] Bacchetta J, Boutroy S, Vilayphiou N, et al. Early impairment of trabecular microarchitecture assessed with HR-pQCT in patients with stage IIeIV chronic kidney disease. J Bone Miner Res 2010;25:849e57. [25] Burghardt AJ, Issever AS, Schwartz AV, et al. High-resolution peripheral quantitative computed tomographic imaging of cortical and trabecular bone microarchitecture in patients with type 2 diabetes mellitus. J Clin Endocrinol Metab 2010;95: 5045e55. [26] Briot K, Roux C. What is the role of DXA, QUS and bone markers in fracture prediction, treatment allocation and monitoring? Best Pract Res Clin Rheumatol 2005;19:951e64. [27] Specker BL, Schoenau E. Quantitative bone analysis in children: current methods and recommendations. J Pediatr 2005;146: 726e31. [28] Binkley TL, Berry R, Specker BL. Methods for measurement of pediatric bone. Rev Endocr Metab Disord 2008;9:95e106. [29] Bonjour JP, Theintz G, Buchs B, Slosman D, Rizzoli R. Critical years and stages of puberty for spinal and femoral bone mass accumulation during adolescence. J Clin Endocrinol Metab 1991;73:555e63.

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Pregnancy and Lactation Ann Prentice Medical Research Council Human Nutrition Research, Elsie Widdowson Laboratory, Cambridge, UK

INTRODUCTION The fluxes of the primary bone-forming minerals, calcium, phosphorus, magnesium, and zinc, that occur between mother and offspring during pregnancy and lactation place considerable demands on maternal mineral economy. There are several possible biological strategies for meeting these extra requirements, including increased food consumption, elevated gastrointestinal absorption efficiencies, decreased mineral excretion, and mobilization of tissue stores. This chapter presents a review of the evidence on the extent to which these strategies apply in the human situation, the mechanisms by which they occur, the limitations imposed by maternal diet, and the possible consequences for the growth of the infant and bone health of the mother. It also discusses the importance of maternal vitamin D status in the mineral metabolism of the mother and infant. The evidence suggests that pregnancy and lactation are associated with physiological adaptive changes that are independent of maternal mineral supply, within the range of normal dietary intakes. These processes appear to provide the minerals necessary for fetal growth and breast milk production without requiring an increase in maternal dietary intake or compromising maternal bone health in the long term. More research is needed to define the limitations of these processes in women with marginal mineral intakes and poor vitamin D status.

MINERAL FLUXES FROM MOTHER TO OFFSPRING At birth, the skeleton contains approximately 20e30 g calcium [1e3]. This represents 98e99% of the total body content of this mineral. The proportion of calcium in fetal ash increases during early gestation and reaches a plateau of approximately 27% (g/g) by 4 months [2]. Substantial skeletal growth occurs from mid-gestation


and maximal fetal calcium accretion occurs during the third trimester. Quantitatively, fetal calcium accretion increases from approximately 50 mg/day at 20 weeks of gestation to 330 mg/day at 35 weeks [4]. For the third trimester of pregnancy, 200 mg/day is considered a typical calcium accretion rate. After delivery, skeletal growth and calcium accretion continue at a slower pace. A typical child accretes approximately 140 mg/ day of calcium during the first year of life, with the rate being highest in the first months and slowing progressively with age [3,5]. The flux of calcium from mother to child across the placenta and via breast milk needs to be sufficient to match this accretion rate and, in the child, to meet any additional requirements imposed by gastrointestinal absorption and obligatory losses in urine, feces, and sweat. Breast milk calcium secretion averages approximately 200 mg/day at peak lactation but can be as high as 400 mg/day in some individuals [6]. Similar estimates can be made for the fluxes of the other primary bone-forming minerals e phosphorus, magnesium, and zinc [3]. A newborn baby contains approximately 16 g of phosphorus, of which approximately 80% is contained in the skeleton. A typical whole-body phosphorus accretion rate in the first year of life is 70 mg/day. The magnesium content at birth is approximately 750 mg, of which 60% is in the skeleton, and a typical whole-body accretion rate in infancy is 3 mg/day. For zinc, typically the whole-body content is around 50 mg at birth, of which 30% is in the skeleton, and the accretion rate in infancy is around 0.4 mg/day [7,8]. Breast milk secretion during full lactation averages approximately 120, 25, and 1.6 mg/day for phosphorus, magnesium, and zinc, respectively, but the ranges of values between individuals are wide. These mineral fluxes from mother to child represent a significant proportion of the mineral intakes of the mother, especially for calcium [9]. Intakes of these minerals vary in different areas of the world and range

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widely between individuals, but average intakes for women are of the order of 300e1500 mg/day calcium, 1000 mg/day phosphorus, 250 mg/day magnesium, and 10 mg/day zinc [10e13]. There are several possible biological strategies for meeting the extra demands that pregnancy and lactation make on the mineral economy of the mother, including increases in food consumption, elevated gastrointestinal absorption efficiencies, decreased mineral excretion, and mobilization of tissue stores. The extent to which these strategies apply in the human situation, the mechanisms by which they occur, the limitations imposed by maternal diet, and the possible consequences for the growth of the baby and bone health of the mother are discussed next.

PREGNANCY Mineral Metabolism and Bone Biochemistry Human pregnancy is associated with major changes in mineral and bone metabolism (Table 10.1). These changes occur from early in gestation, in advance of the mineral requirement for skeletal development of the fetus. Calcium During pregnancy, intestinal calcium absorption efficiency and urinary calcium excretion approximately double compared to the non-pregnant state (Fig. 10.1) [14e20]. The elevation in calcium absorption is associated with increased intestinal expression of calbindinD9K, a vitamin D-dependent calcium-binding protein [21,22]. The increased expression of calbindin-D9K is a result of increased transcription and reduced posttranslational degradation mediated by 1,25-dihydroxyvitamin D, which is elevated in pregnancy, and by other factors [21]. 1,25-Dihydroxyvitamin D also acts at the enterocyte brush border to open voltage-gated calcium channels (primarily TRPV6). However, there is evidence TABLE 10.1

from animal studies to suggest that this mechanism may not be fully responsible for the upregulation of intestinal calcium absorption in pregnancy, which has been shown to occur in the absence of 1,25-dihydroxyvitamin D and its receptor [23]. The increase in urinary calcium excretion is largely due to the combined effects of the increases in glomerular filtration rate and calcium absorption. Fasting calcium excretion, corrected for creatinine clearance, is normal or decreased [16,24e26]. Measured calcium balance in the later stages of pregnancy is generally positive and retention approximates to that required for fetal growth [27]. The proportion of serum calcium circulating in the ionized form is increased in pregnancy. Serum ionized calcium concentration is unchanged or decreases slightly but remains within a narrow physiological range throughout [28,29]. The concentration of total serum calcium declines to a greater extent, with a slight increase toward the end of gestation. The decrease in total serum calcium largely reflects the change in serum albumin concentration associated with the increased intravascular fluid volume of pregnancy and the resulting hemodilution [29,30]. Calcium transport across the placenta is predominantly through an active transcellular pathway involving TRPV6 expressed on the apical membrane of the trophoblast, intracellular calcium-binding proteins (primarily calbindin-D9K) and active transport into the fetal circulation at the basolateral membrane through the calcium pump PMCA3 [31,32]. The fetal calcium concentration is maintained at a higher level than the maternal concentration through the actions of fetal parathyroid hormone (PTH) and parathyroid hormone-related peptide (PTHrP) on the expression of the intracellular calbindins [32,33]. Other regulators of calbindin expression include progesterone and estrogen, but studies in animals do not support an essential role for the vitamin D system in placental calcium transfer, although it may have a role as a negative modulator [31e33].

Calcium and Bone Metabolic Changes in Human Pregnancy, Compared to the Non-Pregnant, Non-Lactating State

Calcium absorption

[

Serum 1,25 (OH)2 vitamin D (free and bound)

[

Urinary calcium excretion, daily

[

Serum parathyroid hormone (intact)

4Y

Fasting urinary calcium, creatinine corrected

4Y

Serum parathyroid hormone-related protein

[

Serum calcium (ionized)

4(Y)

Nephrogenous cyclic AMP

4

Serum calcium (total)

Y

Serum calcitonin

[4

Tubular phosphate reabsorption

4Y

Bone resorption histology and markers*

[

Urinary phosphate excretion

[4

Bone formation markers (except Oc)**

[

Serum phosphate

4Y

Osteocalcin (intact)

Y

* Urinary collagen cross-links, telopeptides, hydroxyproline, serum tartrate-resistant acid phosphatase. ** Serum bone alkaline phosphatase and procollagen peptides

PEDIATRIC BONE

PREGNANCY

↑ 1,25 (OH)2D ↑ PTHrP ↑ Prolactin ↑ Placental lactogen

pregnancy to term. This has been attributed to hemodilution, decreased concentrations of the zinc binding protein, hormonal changes in pregnancy and active transport of zinc to the fetus [8]. There is evidence that zinc absorption and urinary zinc excretion are increased, although individual responses are highly variable [13,41e43].

↑ intestinal Ca++ absorption ↑ Ca-release from bone ↑ Ca-uptake into bone

↑ urinary Ca excretion

225

Bone Turnover

Ca transport to fetus

FIGURE 10.1 Schematic representation of the changes in calcium and bone metabolism that occur during pregnancy.

Phosphorus, Magnesium, and Zinc The patterns of change in the metabolism of the other bone-forming minerals largely parallel that of calcium, with decreases in serum concentrations coupled with increases in intestinal absorption and urinary excretion. There is no evidence for increased renal conservation of any of these minerals. Phosphorus balance becomes increasingly positive (i.e. absorption exceeds excretion) as gestation advances [14], and net absorption is higher in pregnant women compared with non-pregnant women [34]. Some studies have indicated a decrease in serum phosphorus concentration [16] and the renal phosphate threshold in the second and third trimesters of pregnancy, with a corresponding increase in urinary phosphate excretion [16,18]. The data are inconsistent, and other studies have shown no changes in phosphate metabolism [18,24,35]. Little is known about the mechanisms of transporting phosphate to the fetus [32]. As with calcium, fetal phosphate concentrations are maintained at a higher level than in the maternal circulation which suggests that there is active transport across the placenta, although the mechanisms involved have yet to be fully elucidated [32]. The data on magnesium absorption and retention in pregnant women are very limited and inconclusive [10]. Ionized and total serum magnesium concentrations decrease with increasing gestational age [36e39] and are lower in pregnant women than preconception [18] or compared to non-pregnant controls [40]. These effects parallel the changes in serum proteins due to hemodilution and are not considered to represent alterations in maternal magnesium status [10]. Lymphocyte magnesium concentration is unchanged or decreased in pregnancy [38,39]. Urinary magnesium excretion is elevated in late pregnancy [18]. Serum zinc concentration decreases in pregnancy by around 35% from early

Bone resorption increases during pregnancy. This has been indicated both histologically [44] and biochemically by urinary markers, such as collagen cross-links, telopeptides, and hydroxyproline, serum markers such as tartrate-resistant acid phosphatase [17e19,45,46] and stable isotope kinetic studies [47]. After an initial decrease, increases are also noted in biochemical indices of bone formation, such as serum bone alkaline phosphatase and procollagen peptides, to levels higher than those observed prepregnancy [17e19,45,46,48e50]. However, serum osteocalcin concentration, a commonly used marker of bone formation, is decreased relative to preconception levels [17,18], although its concentration in late gestation is higher than in early pregnancy [17e19,51]. Increases in bone turnover indices are observed in early gestation. Their levels increase by 50e200% during pregnancy [17e19,46,48,52]. The increases in markers of bone resorption occur before those of bone formation [19,53], suggestive of bone mineral mobilization. In support of this, a stable isotope study of Brazilian women at 10e12 weeks of pregnancy recorded a net deficit in bone calcium turnover [47]. A study of women with multifetal pregnancies demonstrated that selective fetal reduction reduced circulating concentrations of the cross-linked carboxy (C)-terminal telopeptide of type-1 collagen (ICTP), a marker of bone resorption, without corresponding changes in the C-terminal propeptide of type I procollagen (PICP), an index of bone formation [45]. This suggests that factors derived from the fetoplacental unit are involved in the stimulation of maternal bone turnover, primarily via an effect on bone resorption. Decreases in bone resorption markers have been noted in pregnant women following calcium supplementation [54,55], indicating that the physiological response to an increased calcium load remains intact during pregnancy. There are problems in interpreting changes in biochemical indices during pregnancy because of the effects of hemodilution, alterations in creatinine excretion and renal clearance, and the contribution to and metabolism of these markers by the products of conception [22]. The disparity between the different indices of bone formation, for example, may be due to the degradation or uptake of osteocalcin by the placenta [49,56]. It has been suggested that the reduction in osteocalcin

PEDIATRIC BONE

226


concentration may facilitate bone formation [28]. Measurements of an osteocalcin metabolite e octeocalcin N terminal fragment, Ocf e adjusted for alterations in creatinine clearance, have indicated that despite the low measurable concentrations of the intact protein, osteocalcin production is not decreased in pregnancy [19]. Total alkaline phosphatase is not a useful index of bone formation in pregnancy because of the contribution from the placental isoform [22]. The increase in resorption markers may partly reflect a contribution from the turnover of the fetal skeleton. However, a study of the ratio of a to b isomers of the C-terminal telopeptide of type 1 collagen (CTX), a bone resorption marker, suggested that the fetal contribution to maternal CTX excretion is small, amounting to less than 10% of a-CTX and only 2% of b-CTX [19]. Calciotropic Hormones The concentration of serum 1,25-dihydroxyvitamin D is increased substantially throughout pregnancy [16e18,57], while observational studies suggest there is no change or a modest decline in 25-hydroxyvitamin D concentration [23]. The elevation in 1,25-dihydroxyvitamin D concentration occurs in both the free and protein-bound forms [24,58]. The increase is apparent in the first trimester and continues to rise during gestation by several-fold. Until late in pregnancy, the increase in 1,25-dihydroxyvitamin D parallels an increase in the concentration of vitamin D-binding protein, such that the proportion of the free hormone in the circulation is only elevated in the last trimester [23,59,60]. It is unlikely, therefore, that the early rise in 1,25-dihydroxyvitamin D is fully responsible for the increase in intestinal absorption at that time, and there is animal evidence to suggest that neither this hormone nor its receptor are necessary for the higher calcium absorption efficiency of pregnancy [23]. The mechanisms underlying the increase in 1,25dihydroxyvitamin D are unclear but may involve placental or fetal synthesis of the hormone from maternal 25-hydroxyvitamin D, upregulation of maternal renal 1a-hydoxylase by a variety of pregnancy-associated hormones, or an alteration in the balance between the production of 1,25-dihydroxyvitamin D and 24,25-dihydroxyvitamin D [20,22,61e64]. It is likely that the production of 1,25-dihydroxyvitamin D by the maternal kidneys plays the greater role because low serum concentrations of this hormone have been reported from an anephric individual during pregnancy [22]. Fetal cord blood concentrations of 25-hydroxyvitamin D are similar to or up to 20% lower than maternal concentrations [23], while fetal 1,25-dihydroxyvitamin D concentrations are low. There is evidence of placental transfer of 25-hydroxyvitamin D, but the quantities are thought to be small and considered unlikely to compromise the vitamin D status of pregnant women [10,65]. Furthermore,

although vitamin D and its metabolites are essential for mineral ion homeostasis in adults, there is growing evidence from animal studies that fetal mineral ion homeostasis appears to be mostly independent of this hormonal system [32], or that it plays only a minor modulating role in placental calcium transfer [33]. Although serum 1,25-dihydroxyvitamin D is raised, there is no evidence of an increase in intact PTH concentration in pregnancy [16,20,24,35,52,66], and it may be decreased [17,18,53,67]. There may be a nadir in PTH concentration in early gestation followed by an increase in later pregnancy [28]. Early studies reported high concentrations of PTH during pregnancy, but these have had to be reinterpreted in light of research conducted after the advent of a sensitive two-site immunoassay specific for the intact molecule of PTH [20]. The elevated concentrations reported in the earlier studies are likely explained by the detection of multiple fragments of PTH, many of which are biologically inactive. Although increases in PTH production and turnover cannot be discounted, it appears that human pregnancy is not associated with an increase in PTH bioactivity [20,68]. This is supported by normal nephrogenous cyclic adenosine monophosphate (NcAMP) production, a marker of PTH-like bioactivity [16,35,66]. Consequently, the view of pregnancy as a period of physiological hyperparathyroidism driven by the fetal demand for calcium [69] is no longer regarded as tenable [16]. However, the homeostatic response to a calcium load by an increase in PTH appears to remain intact [70]. Fetal serum concentrations of PTH, which originates from the fetal parathyroids and thymus, are low and decrease further towards the end of gestation, although fetal serum ionized calcium remains stable [20]. Increased concentrations of PTHrP are detected in the maternal circulation during pregnancy [71], probably originating from fetal, placental, or mammary tissues [20]. PTHrP, or more specifically its amino-terminal fragments, has close homology with the N-terminal 1e34 amino acid sequence of PTH. It has the ability to activate the PTH/PTHrP receptor [72] and, consequently, has PTH-like characteristics. It stimulates renal 1a-hydoxylase activity and NcAMP production, thereby promoting 1,25-dihydroxyvitamin D synthesis and calcium reabsorption [21]. In addition, N-terminal PTHrP promotes bone resorption via the classical PTH/PTHrP receptor, although the C-terminal fragment PTHrP (107e139) inhibits osteoclastic bone resorption through a different receptor [72]. The role of PTHrP during pregnancy is unclear, however, but its presence may account, at least in part, for the increase in 1,25-dihydroxyvitamin D, which occurs even though intact PTH concentrations are reduced. Non-pregnant women administered PTHrP (1e36) subcutaneously have elevated 1,25-dihydroxyvitamin D levels, urinary calcium excretion, and

PEDIATRIC BONE

PREGNANCY

NcAMP production with no alteration in serum PTH or calcium concentrations [73] e a response that resembles some of the biochemical changes of pregnancy. The response of calcitonin (CT) to pregnancy appears to be highly variable [74], with some studies reporting elevations [20,40,75] and others reporting no changes [17,28,61,74]. Increases in circulating CT have been observed in thyroidectomized women during pregnancy, probably as a result of CT synthesis by mammary and placental tissues [20]. The physiological function of CT is not fully understood, although a role in protecting the maternal skeleton from resorption during pregnancy has been proposed [76], and it may promote renal calcium excretion [22]. Calcitonin is secreted by the fetal thyroid and concentrations in the fetal circulation are higher than maternal concentrations [32]. Nevertheless, animal studies do not suggest a major role for calcitonin in fetal mineral homeostasis or bone metabolism [32]. Many other hormones, growth factors, and cytokines are elevated in the maternal circulation during pregnancy that could stimulate or drive the observed changes in calcium absorption, bone turnover, and 1,25-dihydroxyvitamin D synthesis. These include prolactin, estrogen, progesterone, placental lactogen, placental growth hormone, tumor necrosis factor-a, insulin-like growth factor-1 (IGF-1), components of the OPG/RANKL/RANK system and the ratio of osteoprotegerin to the other components [19,20,50,77,78]. Their relative contributions to mineral and bone metabolism in human pregnancy, and the interactions between their effects, have yet to be established. Positive correlations with bone turnover markers and kinetically derived measures of bone mineral mobilization have been observed in pregnant women for maternal IGF-1, estrogen and placental lactogen concentrations [47,79e81)] but not for components of the OPG/ RANKL/RANK system [78].

Maternal Skeleton Physiological Changes Biochemical data suggest that human pregnancy is accompanied by alterations in the uptake and release of calcium and other minerals from the maternal skeleton. Whether the balance between storage and mobilization is sufficient to result in an overall increase or decrease in bone mineral content is not clear. Direct assessments of skeletal changes during pregnancy are restricted by the fact that the most sensitive methods for the measurement of bone mineral content are based on the attenuation of ionizing radiation, such as dualand single-energy x-ray absorptiometry, and quantitative computed tomography. Although the radiation doses incurred with modern instruments are generally

227

low and similar to that received from background radiation, these techniques are not useful for measurements of the axial skeleton in pregnant women because of the involvement of fetal tissues. As a consequence, investigations of the axial skeleton and whole body are limited to estimating the integrated skeletal response over the whole of pregnancy by measuring bone mineral status before conception and after delivery. Pregnancy is accompanied by changes in weight and advances in age, both of which can have independent effects on the bone mineral content of the skeleton, and these should be factored into any conclusions about bone mineral changes during pregnancy [82]. In addition, the bone mineral content of the skeleton postpartum is affected by lactation (discussed later) and, consequently, the timing of the postpartum measurement can confound the interpretation of the bone changes during pregnancy in women who breastfeed. To date, of the prospective studies that have been undertaken, most involved relatively small numbers of individuals [17e19,46,52,53,82e89], and only a few have compared the results with a group of non-pregnant women, measured over the same time period, to consider age and weight changes [82,89]. Changes in bone mineral content have been reported in many of these studies, but the observed response differs widely between studies, between individual women and between skeletal sites. The average effect of pregnancy on the axial skeleton reported in these studies ranges from no change in bone mineral content to a decrease of around 4e5%. Nevertheless, a pattern is emerging for pregnancy to be associated with a reduction in bone mineral content at one or more axial sites, reflected in a decrease in whole-body bone mineral content of around 2%. In a longitudinal study of pregnant women in the UK, the decrease in bone mineral content at the spine and whole body was largely independent of concomitant changes in weight and age [82], but the decrease at the femoral neck was similar to that experienced by non-pregnant women studied over the same time period, and therefore could not be ascribed to pregnancy [82]. The calculated release of calcium from the maternal skeleton over the course of pregnancy was estimated to be have been sufficient to cover most of the calcium required for fetal bone mineral accretion [82]. In the case of women entering pregnancy during or after a period of extended lactation, substantial increases in bone mineral content have been reported [83,85], suggesting that maternal adaptative processes are sufficient to supply mineral for both maternal and fetal mineral accretion. Measurements at peripheral skeletal sites, using single- or dual-energy absorptiometry (DXA), peripheral quantitative computed tomography or bone ultrasonography, have also been used to investigate the pattern

PEDIATRIC BONE

228


of skeletal response during pregnancy [18,24,28,48, 52,82,86,90e94]. Many of these studies are difficult to interpret because the initial measurements were made when the women were already pregnant rather than before conception and, as is now well recognized, major changes in bone metabolism occur in the early stages of pregnancy [44]. Decreases in bone mineral content over the course of pregnancy have been noted in ultradistal scans of the forearm, a region rich in trabecular bone [28,52,86,91,92], but generally not at more proximal appendicular sites and not in all studies [18,24,28,82]. Ultrasound studies of the os calcis and phalanges have reported decreases in the measured variables in the later stages of pregnancy [48,90,93,94]. Speed of sound and broadband ultrasound attenuation, measured by bone ultrasonography are regarded as indices of bone mineral density for fracture risk prediction in older people. However, the validity of this assumption for quantifying changes in bone mineral content during pregnancy is not known, particularly in the presence of peripheral edema, and has been called into question by a longitudinal study of lactating women in which ultrasound measurements of the os calcis were not concordant with the skeletal bone mineral changes observed by DXA [95]. The reasons for the variability in maternal skeletal response to pregnancy have yet to be determined. It is possible that the changes in bone mineral content are governed by a variety of influences, such as the mother’s age or parity and her nutritional or endocrinological status prior to or after conception. For example, in studies of women who conceived during or soon after a period of extended lactation, increases in bone mineral content occurred during pregnancy that were similar to those required for the recovery of lactational bone losses [83,85]. However, those women who conceived after recovery of lactational bone loss had taken place showed little further change in bone mineral content by the end of the subsequent pregnancy. Slim pregnant women (body mass index

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